BIOLOGICAL FIELD STATION Cooperstown, New York

46th ANNUAL REPORT 2013

1 mm

Gloeotrichia echinulata collected in Panther Lake, Oswego County, New York

Scanning electron micrograph of a newly discovered species of nematode from Redbreast sunfish in Otsego Lake.

STATE UNIVERSITY OF NEW YORK COLLEGE AT ONEONTA

OCCASIONAL PAPERS PUBLISHED BY THE BIOLOGICAL FIELD STATION

No. 1. The diet and feeding habits of the terrestrial stage of the common newt, Notophthalmus viridescens (Raf.). M.C. MacNamara, April 1976 No. 2. The relationship of age, growth and food habits to the relative success of the whitefish (Coregonus clupeaformis) and the cisco (C. artedi) in Otsego Lake, New York. A.J. Newell, April 1976. No. 3. A basic limnology of Otsego Lake (Summary of research 1968-75). W. N. Harman and L. P. Sohacki, June 1976. No. 4. An ecology of the Unionidae of Otsego Lake with special references to the immature stages. G. P. Weir, November 1977. No. 5. A history and description of the Biological Field Station (1966-1977). W. N. Harman, November 1977. No. 6. The distribution and ecology of the aquatic molluscan fauna of the Black River drainage basin in northern New York. D. E Buckley, April 1977. No. 7. The fishes of Otsego Lake. R. C. MacWatters, May 1980. No. 8. The ecology of the aquatic macrophytes of Rat Cove, Otsego Lake, N.Y. F. A Vertucci, W. N. Harman and J. H. Peverly, December 1981. No. 9. Pictorial keys to the aquatic mollusks of the upper Susquehanna. W. N. Harman, April 1982. No. 10. The dragonflies and damselflies (Odonata: Anisoptera and Zygoptera) of Otsego County, New York with illustrated keys to the genera and species. L.S. House III, September 1982. No. 11. Some aspects of predator recognition and anti-predator behavior in the Black-capped chickadee (Parus atricapillus). A. Kevin Gleason, November 1982. No. 12. Mating, aggression, and cement gland development in the crayfish, Cambarus bartoni. Richard E. Thomas, Jr., February 1983. No. 13. The systematics and ecology of Najadicola ingens (Koenike 1896) (Acarina: Hydrachnida) in Otsego Lake, New York. Thomas Simmons, April 1983. No. 14. Hibernating bat populations in eastern New York State. Donald B. Clark, June 1983. No. 15. The fishes of Otsego Lake (2nd edition). R. C MacWatters, July 1983. No. 16. The effect of the internal seiche on zooplankton distribution in Lake Otsego. J. K. Hill, October 1983. No. 17. The potential use of wood as a supplemental energy source for Otsego County, New York: A preliminary examination. Edward M. Mathieu, February 1984. No. 18. Ecological determinants of distribution for several small mammals: A central New York perspective. Daniel Osenni, November 1984. No. 19. A self-guided tour of Goodyear Swamp Sanctuary. W. N. Harman and B. Higgins, February 1986. No. 20. The Chironomidae of Otsego Lake with keys to the immature stages of the subfamilies Tanypodinae and Diamesinae (Diptera). J. P. Fagnani and W. N. Harman, August 1987. No. 21. The aquatic invertebrates of Goodyear Swamp Sanctuary, Otsego Lake, Otsego County, New York. Robert J. Montione, April 1989. No. 22. The lake book: a guide to reducing water pollution at home. Otsego Lake Watershed Planning Report #1. W. N. Harman, March 1990. No. 23. A model land use plan for the Otsego Lake Watershed. Phase II: The chemical limnology and water quality of Otsego Lake, New York. Otsego Lake Watershed Planning Report Nos. 2a, 2b. T. J. Iannuzzi, January 1991. No. 24. The biology, invasion and control of the Zebra Mussel (Dreissena polymorpha) in North America. Otsego Lake Watershed Planning Report No. 3. Leann Maxwell, February 1992. No. 25. Biological Field Station safety and health manuel. W. N. Harman, May 1997. No. 26. Quantitative analysis of periphyton biomass and identification of periphyton in the tributaries of Otsego Lake, NY in relation to selected environmental parameters. S. H. Komorosky, July 1994. No. 27. A limnological and biological survey of Weaver Lake, Herkimer County, New York. C.A. McArthur, August 1995. No. 28. Nested subsets of songbirds in Upstate New York woodlots. D. Dempsey, March 1996. No. 29. Hydrological and nutrient budgets for Otsego lake, N. Y. and relationships between land form/use and export rates of its sub -basins. M. F. Albright, L. P. Sohacki, W. N. Harman, June 1996. No. 30. The State of Otsego Lake 1936-1996. W. N. Harman, L. P. Sohacki, M. F. Albright, January 1997. No. 31. A Self-guided tour of Goodyear Swamp Sanctuary. W. N. Harman and B. Higgins (Revised by J. Lopez),1998. No. 32. Alewives in Otsego Lake N. Y.: A Comparison of their direct and indirect mechanisms of impact on transparency and Chlorophyll a. D. M. Warner, December 1999. No.33. Moe Pond limnology and fish population biology: An ecosystem approach. C. Mead McCoy, C. P. Madenjian, V. J. Adams, W. N. Harman, D. M. Warner, M. F. Albright and L. P. Sohacki, January 2000. No. 34. Trout movements on Delaware River System tail-waters in New York State. Scott D. Stanton, September 2000. No. 35. Geochemistry of surface and subsurface water flow in the Otsego lake basin, Otsego County New York. Andrew R. Fetterman, June 2001. No. 36 A fisheries survey of Peck Lake, Fulton County, New York. Laurie A. Trotta. June 2002. No. 37 Plans for the programmatic use and management of the State University of New York College at Oneonta Biological Field Station upland natural resources, Willard N. Harman. May 2003. No. 38. Biocontrol of Eurasian water-milfoil in central New York State: Myriophyllum spicatum L., its insect herbivores and associated fish. Paul H. Lord. August 2004. No. 39. The benthic macroinvertebrates of Butternut Creek, Otsego County, New York. Michael F. Stensland. June 2005. No. 40. Re-introduction of walleye to Otsego Lake: re-establishing a fishery and subsequent influences of a top Predator. Mark D. Cornwell. September 2005. No. 41. 1. The role of small lake-outlet streams in the dispersal of zebra mussel (Dreissena polymorpha) veligers in the upper Susquehanna River basin in New York. 2. Eaton Brook Reservoir boaters: Habits, zebra mussel awareness, and adult zebra mussel dispersal via boater. Michael S. Gray. No. 42. The behavior of lake trout, Salvelinus namaycush (Walbaum, 1972) in Otsego Lake: A documentation of the strains, movements and the natural reproduction of lake trout under present conditions. Wesley T. Tibbitts. No. 43. The Upper Susquehanna watershed project: A fusion of science and pedagogy. Todd Paternoster. No. 44. Water chestnut (Trapa natans L.) infestation in the Susquehanna River watershed: Population assessment, control, and effects. Willow Eyres. No. 45. The use of radium isotopes and water chemistry to determine patterns of groundwater recharge to Otsego Lake, Otsego County, New York. Elias J. Maskal.

Annual Reports and Technical Reports published by the Biological Field Station are available from Willard N. Harman, BFS, 5838 St. Hwy. 80, Cooperstown, NY 13326.

46th ANNUAL REPORT 2013

BIOLOGICAL FIELD STATION COOPERSTOWN, NEW YORK bfs.oneonta.edu

STATE UNIVERSITY COLLEGE AT ONEONTA

The information contained herein may not be reproduced without permission of the author(s) or the SUNY Oneonta Biological Field Station

BFS 2013 ANNUAL REPORT CONTENTS

INTRODUCTION: W. N. Harman…………………………………………………………………...….1

ONGOING STUDIES:

OTSEGO LAKE WATERSHED MONITORING: 2013 Otsego Lake water levels. W.N. Harman and M.F. Albright………………………..8 Otsego Lake limnological monitoring, 2013. H.A. Waterfield and M.F. Albright..….…11 A survey of Otsego Lake’s zooplankton community, summer 2013. C. Tanner and M.F. Albright ………….……………………………..…….…..22 Chlorophyll a concentrations in Otsego Lake, summer 2013. T. Bianchine and C. Tanner………………………………….……...... 34 Water quality monitoring of five major tributaries in the Otsego Lake watershed, summer 2013. C. Teter. ..………………………………………….42

SUSQUEHANNA RIVER MONITORING: Monitoring the water quality and fecal coliform bacteria in the upper Susquehanna River, summer 2013. T. Bianchine………………..…….……….59

ARTHROPOD MONITORING: Mosquito data from light traps, Thayer Farm. W.L. Butts…………………………....74

REPORTS:

Aquatic invasive species present in Otsego County, NY water bodies. A. Yoo, K. Herzog and H.A. Waterfield …………………………….……………….75 Zebra mussel (Dreissena polymorpha) monitoring using navigation buoys. A. Yoo, P.H. Lord and W.H. Wong………..…………………………..…….……….95 Zebra mussels and other benthic organisms in Otsego Lake in 2008. J. Vanassche, W.H. Wong, W.N. Harman and M.F. Albright...... …………………103 Trap net monitoring of fish communities within the weedy littoral zone at Rat Cove and rocky littoral zone at Brookwood Point, Otsego Lake. S.G. Stowell.……110 Population assessment of fresh-water mussels (Unionidae) in Otsego Lake since the introduction of zebra mussels (Dreissena polymorpha). D. Caracciolo……….…....115 Gastropods and fish as hosts of digenetic trematodes in Otsego Lake and nearby waters. E. Darpino, R. Russell and F. Reyda……………………………123 Nematodes of fishes of Otsego Lake, New York, including a species that is new to science. A. Borden and F. Reyda……...……… ……………….…….129 An examination of the morphological diversity within a new genus of tapeworm from stingrays (Class: Cestoda). K. Herzog, R. Russell and F. Reyda..……………..135 Cestodes of the fishes of Otsego Lake and nearby waters. A. Sendkewitz, I. Delgado and F. Reyda..…………………………………………...140 Forest bryophyte reproduction and dispersal: An update. A. Lawrence and S. C. Robinson……………………………………...…………….144

Monitoring the effectiveness of the Cooperstown wastewater treatment wetland, 2013. M.F. Albright…………………….……………………….150 Groundwater flow and geochemistry at Greenwoods Conservancy. M. Moore and L. Hasbargen…………………………………………….……...……161 Utilizing environmental DNA to identify aquatic invasive species. L. Newton.……………186 A biosurvey of Allen’s Lake, Richfield Springs, NY. P. H. Lord ……...……..…….………192 Chronological field observations at various BFS sites, 2013. W.L. Butts..…………………194 Dynamics of Galerucella spp. and purple loosestrife (Lythrum salicaria) in Goodyear Swamp Sanctuary, summer 2013 update. H.A. Waterfield……………....197 Walleye (Sander vitreus) movement and depth utilization in response to changes in alewife (Alosa pseudoharengus) abundance in Otsego Lake J.R. Foster and D.J. Drake……………………………………………..………….....204 Hydroacoustic survey of Otsego Lake’s pelagic fish community, spring 2013.H.A. Waterfield and M. Cornwell………………..…………………….211 Natural recruitment of lake trout (Salvelinus namaycush) in Otsego Lake. N.M. Sawick and J.R. Foster………………………………………………………...219 Monitoring the Moe Pond ecosystem and population estimates of largemouth bass (Micropterus salmoides) post unauthorized introduction. S.G. Stowell…………….226 Otsego Lake, NY ice phenology 1843-2014. H.A. Waterfield………………………………237 Presence of mercury and comparison to other metals in lakes, rivers, and streams in central New York. M. Moore and D. Castendyk……….……………241 The effects of zebra mussels on benthic macroinvertebrates in Otsego Lake. J.M. Vanassche, W.H., W.N. Harman, and M.F. Albright…………………….…….253 Control and eradication of water chestnut (Trapa natans, L.) in an Oneonta wetland, 2013 progress report. H.A. Waterfield and M.F. Albright……………………….….265 Aquatic macrophyte management plan facilitation, Lake Moraine, Madison County, NY 2013.W.N. Harman and M.F. Albright………………………267 Otsego Lake fry sampling, 2013. H.A. Waterfield and M.F. Albright………………………278

46th Annual Report of the Biological Field Station

INTRODUCTION

Willard N. Harman

This year a new faculty member, shared with the Biology Department, joined the BFS. Kiyoko Yokota specializes in algal community dynamics which are important in the primary production of fresh-waters. She is particularly interested in blue-green algae (cyanobacteria) that are becoming more of a problem in our inland lakes causing Harmful Algal Blooms. She compliments current BFS faculty members, David Wong, (Invasive species; particularly zebra and quagga mussels and biostatistics), and Bill Harman, (benthic ecology, lake management). All three are lake managers, certified by the North American Lake Management Society as are BFS staff members Matt Albright and Holly Waterfield. Florian Reyda conducts basic research with fish parasites while collaboration with John Foster and Mark Cornwell from SUNY Cobleskill provide expertise in aquaculture and fisheries management.

Our original wood frame laboratory on the Upper Site adjacent to Moe Pond has been completely rebuilt for year around use. We have yet to occupy the structure for permanent use because of problems with control systems associated with local off-grid utility services.

Interns:

Kaylee Herzog, a SUNY Oneonta Biology major, received the SUNY Oneonta Biology Department Internship. She was sponsored in part by the National Science Foundation. Under the direction of Florian Reyda and with Student Research Assistant Rebecca Russell, she evaluated a new genus of tapeworm from stingrays. Kaylee also worked with Annie Yoo, a SUNY Oneonta Biology major who was sponsored by the Otsego Land Trust. They conducted a survey of nuisance aquatic species of Otsego County lakes and streams. Annie also worked under the direction of David Wong to evaluate the use of navigational buoys for zebra mussel monitoring. Jennifer Vanassche, a SUNY Oneonta Biology major, conducted a benthic invertebrate survey of Otsego Lake in an attempt to gain insight into the influence of the establishment of zebra mussels. She was supported by the Village of Cooperstown. Deanna Caracciolo, a SUNY Oneonta Environmental Science major, held the OCCA W.N. Harman Internship. She conducted a freshwater clam survey of Otsego Lake. Christopher Teter, also a SUNY Oneonta Environmental Science major, held the Otsego County Conservation Association-sponsored Rufus J. Thayer Otsego Lake Internship. He conducted water quality monitoring throughout Otsego Lake’s watershed to evaluate the effectiveness of agricultural Best Management Practices. Lisa Newton, a SUNY Oneonta Biology/Psychology major, was supported by a contract with the NYC Department of Environmental Protection. She, under the direction of Jeff Heilveil, worked on using environmental DNA to identify aquatic invasive species. Austin Borden and Erica Darpino, both SUNY Oneonta Biology majors and BFS Internship recipients, worked with Florian Reyda. Austin surveyed nematodes of the fishes of Otsego Lake, while Erica surveyed trematodes of Otsego Lake fishes and snails. Myles Moore, a SUNY Oneonta Water Resources major, worked with Les Hasbargen. They studied groundwater flow and geochemistry at Greenwoods Conservancy and were assisted by Student Research Assistants Joe

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Spaulding and Alayna Fuess, both SUNY Oneonta Earth Science majors. This work was supported by the Peterson Family Conservation Trust. Alexander Lawrence, a SUNY Oneonta Environmental Science major, worked under the direction of Sean Robinson as they continued their research on moss reproduction and dispersal. Steve Stowell, a Fisheries and Aquaculture major from SUNY Cobleskill, held the Robert C. MacWatters Internship. He continued monitoring in Moe Pond, evaluating the impacts of the establishment of largemouth bass in the system. He also monitored the littoral fish community of Otsego Lake. Clara Tanner, a recent graduate from Schenevus Central School and Tyler Bianchine, from Sharron Springs Central School, both received FHV Mecklenburg Fellowships. Clara, supported by the Otsego County Conservation Association, surveyed the zooplankton community in Otsego Lake. Tyler, sponsored by the Village of Cooperstown, monitored fecal coliform bacteria and water quality in the upper Susquehanna River. Together, Clara and Tyler monitored chlorophyll a concentrations in Otsego Lake.

Faculty and staff activities:

Jeffrey Heilveil: In addition to the mentoring and supervision of Lisa Newton (see contribution), which satisfies a requirement for a NYC DEP Invasive Species contract that Bill Harman and I are doing, I led a 16-day course in Field Entomology during the Summer 2013 semester. Students in the course camped near the Upland Interpretive Center and worked in the UIC, Hop House, and Boat House. The students learned about insect identification, life history, and ecology, focusing on the species that live on and near the Biological Field Station properties. The weather was fabulous, allowing students to take full advantage of the habitats available. Taxa lists were kept by habitat to facilitate future entomological research on the properties. The Biology Club held two outings at the Biological Field Station this past year. Both events had students collecting organisms and going on faculty-led nature walks. Each event culminated in a group dinner, an evening of biologically-oriented trivia, and students having a chance to camp out on the grounds and listen to the coyotes and the owls.

Holly Waterfield published a paper with colleagues; Kocovsky PK, LG Rudstam, DL Yule, DM Warner, T Schaner, B Pientka, JW Heller, HA Waterfield, LD Witzel, PJ Sullivan (2013) Sensitivity of fish density estimates to standard analytical procedures applied to Great Lakes hydroacoustic data. Journal of Great Lakes Research 39:655-662.

Kiyoko Yokota utilized the BFS resources throughout Fall 2013 to teach BIOL 385 (Limnology), including two extended labs at BFS for Otsego Lake sampling and subsequent analyses and an all-day field trip at the Upper Site/Moe Pond. A site visit to the Cooperstown Wastewater Treatment Plant and its nutrient removal wetland was also made possible through existing research collaboration between the plant and BFS. In Spring 2014, she is teaching both BIOL 575 (Phytoplankton Ecology and Analysis) and BIOL 691 (Management of Aquatic Biota) at BFS. In addition to mentoring research projects on fish parasites, Florian Reyda brought his SUNY Oneonta parasitology class to Thayer Farm for a field trip last fall. He and approximately 20

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students encountered and examined parasites of fish, amphibians and invertebrates during their intensive science and camping trip.

Paul Lord and Tim Pokorny continued monitoring Eurasian milfoil (Myriophyllm spicatum) conditions in Woodridge Lake, Litchfield County, CT. The Biological Field Station pearly mussel team performed limited surveys sampling previously sampled sites for pearly mussel persistence. These sites included locations in Otsego Lake, the Otego Creek, the Butternut Creek, the Sangerfield River, and the Otselic River. Additionally, we responded to two reports of whey spills to ascertain pearly mussel impacts. Finally, we participated in the NYSDEC review of pearly mussel species of greatest conservation need and recommended the up-listing of one species (Alasmidonta varicosa) and the down-listing of another (Lasmigona subviridis). Unfortunately, our survey of Otsego Lake leads us to believe pearly mussels will be extirpated from that waterbody in the next year or two. The highlight of the season’s effort was the discovery of another set of recently spent shells of Margaritifera margaritifera in the Otselic River headwaters within 10 m (downstream) of the set found in 2008.

David Wong supervised Student Research Assistant Emily Davidson, a student from Oneonta’s Environmental Science program to determine zebra mussel abundance in Otsego Lake. They collected 37 benthic samples along 7 transects. The samples and data are now being analyzed. The project will be completed during the next ice free season. Paul Lord, collaborating with Joseph W. Zarzynski, a professional underwater archaeologist, led the BFS volunteer divers, Lee Ferrara, Chip Dunn, Jim Vogler, Kaylee Herzog and Dale Webster, in locating the sunken wreckage of a 1948 plane crash in Otsego Lake. Zarzynski funded the 2012 side scan sonar survey, conducted by Garry Kozak (GK Consulting). The dive was part of a long-term project to map the lake to gain a greater understanding of the cultural and biological resources in the historic waterway. The aluminum aircraft, distinguished by its unique twin-tail configuration, crashed into Otsego Lake around 3 pm on July 13, 1948. Two World War II veterans, Edward Francis and Harold Caulkins, both aged 24, were killed in the aviation accident. During another dive, Paul, Bjorn Eilertsen and Zarzynski visited the wreck of W.T. Sampson Smith’s Leatherstocking, a 24 ft. long classic wooden Gar Wood that caught fire in 1940 and sunk. The slopes of the regression lines in the below graphic, constructed by Holly Waterfield, reflect the warming of our local winters since 1845. It illustrates the numbers of days annually that Otsego Lake has been covered by ice from 1850 to 2014. The scatter documents annual data, the larger circles 10 year averages. The regression line to the right shows the trend since 1960. Note that the lake had 0 days of total ice cover during the winters of 2001-02 and 2011-12, the only times in recorded history. The annual maximum duration of ice cover, between 125 - 131days, occurred 4 times in the 1870s.

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As the Master of Science in Lake Management degree program enters its 3rd year, it appears it will meet its target number of 15 students soon. The program intensively uses the resources of the Biological Field Station, most courses being taught on site in Cooperstown. Individuals enrolled are developing comprehensive management plans for selected lakes and their watersheds throughout the State. Students are now active are; Carter Bailey, hailing from Mt. Vision, N.Y. He did his undergraduate work at the SUNY College of Environmental Science and Forestry and is now studying Canadarago Lake, south of Richfield Springs, specializing in algal community dynamics. Ben German, a member of the US Air Force Reserves from Oneonta, graduated from SUNY Cobleskill in Wildlife and Fisheries Technology. He is working on Moraine Lake in Madison County, near Hamilton, where the BFS has conducted aquatic plant management research and monitoring for years. Derek Johnson, from Ava, N.Y. graduated from Paul Smith’s College in the Adirondacks after previous work at Virginia Commonwealth University. He is developing a plan for Panther Lake north of Oneida Lake in Oswego County. Dan Kopec was a SUNY Oneonta undergraduate in the Earth Sciences. He is now developing a groundwater based comprehensive management plan for Cazenovia Lake in Madison County near Syracuse. Jason Luce is doing research on two connected lakes in Madison County; Hatch Lake and Bradley Brook Reservoir. He comes from East Freetown, N.Y. and graduated from Cazenovia College. Caitlin Stroosnyder is working on Goodyear Lake on the Susquehanna River near Oneonta. She lives in Worcester, N.Y., having held a position at Delaware Engineering since her graduation from Cornell University. Owen Zaengle is from Cooperstown, having done undergraduate work at Davis College, Broome County Community College, SUNY Oneonta and most recently, the SUNY College of Environmental Science and Forestry. He is working on one of the Indian River Lakes Conservancy waters, Grass Lake, near the St. Lawrence River north of Tug Hill. Eight applicants have been accepted to the program for fall 2014. Maxine Verteramo, from Ware, Massachusetts has been employed by a consulting firm, Water Resources Services, since 2011. She received her undergraduate degree from Hampshire College in Amherst, Mass. We first met her at the North American Lake Management Society meetings last fall in San Diego.

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Four students are coming to us from SUNY Environmental Science and Forestry, all with backgrounds in fisheries ecology. Christian Jenne is from Richford, N.Y., Luke Gervase from Malverne, N.Y, Edward Kwietriewski, from Lake View, N.Y. and Michael Greco from Glenmont, N.Y. Michael has also done undergraduate work at both Hudson Valley and Columbia Green Community Colleges. Jenna Leskovec, from Fort Edward, N. Y., did her undergraduate work at SUNY Geneseo in geology. Kathleen Marean, from Jamaica, N.Y., graduated from Cornell University in 2010. Since then she has been employed by the NYSDEC in their Region 1 Freshwater Fisheries Unit. Alejhandro Reyes from Putnam Valley, N.Y., did his undergraduate work at SUNY Plattsburgh and has been engaged in several management related activities on Lake Champlain, in Colorado and on the West Coast.

Shane Pickering, a graduate student enrolled in the MS in Biology program, is from Star Lake, N.Y., and earned an undergraduate degree from SUNY Potsdam. Shane is working with the effects of environmental variables on the maturation and spawning of zebra mussels in Otsego Lake under the direction of David Wong.

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Otsego Lake boat census data:

Year 1975 1976 1977 1978 1979 1980 1981 1991 Date 28-Jul 22-Jul 22-Jul 13-Aug 31-Jul Sailboats 224 186 129 101 92 95 230 243 Rowboats 145 236 160 94 86 42 87 285 Canoes 59 52 28 75

Outboards 636 515 436 456 378 197 445 470 Inboards 73 38 22 36 60

Inboard-Outboards 213

Personal Watercraft 61 Misc. cruisers/houseboats 65 41 40 33 24 23 Total 1070 978 765 783 679 408 896 1332

Year 1992 1993 1994 1995 1996 1997 1998 1999 Date 5-Aug 5-Aug 27-Jul 14-Jul 23-Jul 18-Jul 7-Aug 29-Jul Sailboats 220 181 208 208 207 183 236 238 Rowboats and 243 266 311 313 325 312 372 309 canoes Outboards 407 405 461 430 378 371 377 412 Inboards 22 27 16 13 36 13 20 15 Inboard-Outboards 219 215 227 267 260 275 261 265 Personal Watercraft 32 28 29 47 51 62 84 66 Misc. 40 57 49 Total 1158 1145 1285 1315 1272 1226 1351 1317

Year 2000 2001 2002 2005 2007 2008 2009 2010 Date 10-Aug 9-Aug 22-Jul 23-Aug 27-Aug 26-Aug 31-Aug Sailboats 187 190 171 198 192 153 178 162 Rowboats and 349 389 384 450 383 422 407 458 canoes Outboards 381 375 319 380 344 340 349 363 Inboards 23 9 36 21 24 25 30 14 Inboard-Outboards 287 285 216 297 277 280 251 272 Personal Watercraft 19 23 18 15 22 16 17 9 Misc. 53 66 43 51 43 38 48 44 Total 1299 1337 1187 1412 1285 1274 1280 1322

Year 2011 2012 2013 Date 9-Sep 15-Aug 22-Aug Sailboats 118 140 113 Rowboats 450 545 520 Canoes Outboards 227 334 329 Inboards 15 16 31 Inboard-Outboards 190 274 247 Personal Watercraft 14 22 17 Misc. 40 40 41 Total 1054 1371 1298

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Public support makes our work possible. Funding for BFS research and educational programs was procured in 2013 from many citizens and organizations. Special thanks go to the Clark and Scriven Foundations who generously support our annual needs. The OCCA, the Peterson Family Charitable Trust, the Village of Cooperstown, the Otsego Lake Association, The Otsego Land Trust, SUNY Oneonta, and the SUNY Graduate Research Initiative have also supported our endeavors. A diversity of Lake Associations, and the New York State Federation of Lake Associations, contribute to the support of students in our Lake Management program.

Willard N. Harman, CLM

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ONGOING STUDIES:

OTSEGO LAKE WATERSHED MONITORING:

2013 Otsego Lake water levels W.N. Harman and M.F. Albright

Graphs represent Otsego Lake elevation readings at Rat Cove, in centimeters, above or below “0”, which equals the level considered optimal (364.1 m, or 1194.5 ft, above mean sea level).

Ice on (24 Jan)

Ice off (13 Apr)

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Otsego Lake limnological monitoring, 2013

Holly A. Waterfield CLM1, Matthew F. Albright CLM2

INTRODUCTION

Otsego Lake is a glacially formed, dimictic lake (max depth 51m) supporting a cold water fishery. The Lake is generally classified as being chemically mesotrophic, although flora and fauna characteristically associated with oligotrophic lakes are present (Iannuzzi, 1991).

This study is the continuation of a year-round monitoring protocol that began in 1991. The data collected in this report run for the calendar year and are comparable with contributions by Homburger and Buttigieg (1992), Groff et al. (1993), Harman (1994; 1995), Austin et al. (1996), Albright (1997; 1998; 1999; 2000; 2001; 2002; 2003; 2004; 2005; 2006; 2007; 2008), Albright and Waterfield (2009), and Waterfield and Albright (2010; 2011; 2012; 2013). Concurrent additional work related to Otsego Lake included estimates of fluvial nutrient inputs (Teter 2014), and descriptions of the zooplankton community (Tanner and Albright 2014), chlorophyll a (Bianchine and Tanner 2014), benthic invertebrate communities (Caracciolo 2014, Vanassche et al. 2014, Yoo et al. 2014) and nekton communities (Stowell 2014, Waterfield and Cornwell 2014).

MATERIALS AND METHODS

Physiochemical data and water samples were collected near the deepest part of the lake (TR4-C) (Figure 1), which is considered representative of whole-lake conditions, as past studies have shown the Lake to be spatially homogenous with respect to the factors under study (Iannuzzi 1991). Data and sample collection occurred approximately bi-weekly during open water conditions, 05 February through 03 December 2013. Physical measurements were recorded at 2-m intervals between 0 and 20 m and 40 m to the bottom; 5-meter intervals were used between 20 and 40 m. Measurements of pH, temperature, dissolved oxygen (mg/l and % saturation), specific conductance, Oxidation-Reduction Potential (ORP), and Chlorophylla concentration were recorded with the use of a YSI® 650 MDS with a 6-Series multiparameter sonde which had been calibrated according to the manufacturer’s instructions prior to use (YSI Inc. 2009). Samples were collected for chemical analyses at 4-m intervals between 0 and 20 m and 40m and 48m; 10-m intervals were used between 20 and 40 m. Methodologies employed for sample preservation and chemical analyses are summarized in Table 1. Nutrient and chlorophylla concentrations were determined for all sampling dates; alkalinity, calcium, and chloride concentrations were determined for one profile date per month.

1 Research Support Specialist: SUNY College at Oneonta Biological Field Station, Cooperstown, NY. 2 Assistant to the Director: SUNY College at Oneonta Biological Field Station, Cooperstown, NY.

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46th Annual Report of the Biological Field Station

TR4-C

Figure 1. Bathymetric map of Otsego Lake showing sampling site (TR4-C).

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Table 1. Summary of laboratory methodologies.

Parameter Preservation Method Reference Persulfate digestion followed by Total Phosphorus H SO to pH < 2 Liao and Marten 2001 2 4 single reagent ascorbic acid Cadmium reduction method following Pritzlaff 2003; Total Nitrogen H SO to pH < 2 2 4 peroxodisulfate digestion Ebina et al. 1983

Nitrate+nitrite-N H2SO4 to pH < 2 Cadmium reduction method Pritzlaff 2003

Ammonia-N H2SO4 to pH < 2 Phenolate method Liao 2001 Calcium Store at 4oC EDTA trimetric method EPA 1983 Chloride Store at 4oC Mercuric nitrate titration APHA 1989 Alkalinity Store at 4oC Titration to pH= 4.6 APHA 1989 Filter Buffered acetone extraction followed Chlorophylla immediately; Welschmyer, 1994 o by flourometric detection store at 0 C

RESULTS AND DISCUSSION

Temperature Figures 2a and 2b depict temperatures measured in profile (0 to 48m) at site TR4-C from 05 February through 16 July and 16 July through 03 December 2013, respectively. Observed surface temperature ranged from a low of 1.10oC on 04 April to 25.97oC on 16 July, at which point the epilimnion extended through 6m depth (Figure 2a). Temperatures at 48m reached the annual minimum of 2.79oC on 26 February, maximum of 4.82oC on 3 December. Complete ice- cover formed on 24 January; the lake was completely ice-free on 13 April. Spring mixing was underway prior to the 17 April sampling event and thermal stratification was evident by 21 May. Surface temperatures began to decrease after the profile collected 16 July and the thermocline occurred at greater depth until fall turnover, which was ongoing as of the 03 December sampling event (Figure 2b).

Dissolved Oxygen Isopleths of oxygen concentration based on the profiles for the calendar year are presented in Figure 3. On 17 April, prior to the onset of thermal stratification (in May), dissolved oxygen ranged from 11.46 mg/l at bottom to 12.41 mg/l at the surface. The minimum observed DO concentration in 2013 was 4.99 mg/l recorded on 15 October at 48m. In most years between 1995 and 2009, the bottom minimum concentration was near or below 1.0 mg/l. The areal hypolimnetic oxygen depletion rate (AHOD), calculated at 0.061 mg/cm2/day (between 21 May and 15 October), remains well below the historical average for the fourth consecutive year (Table 2).

Alkalinity Alkalinity concentrations followed a typical pattern of seasonal variation, with concentrations decreasing in the epilimnion during the growing season. Mean annual

- 13 - 46th Annual Report of the Biological Field Station concentration at TR4-C was 127 mg/l, ranging from 108 mg/l at 8m and 12m on 23 April to 142 mg/l at 48m on 4 March.

Calcium Calcium concentrations followed a typical seasonal pattern of fluctuation similar to that of alkalinity. Mean annual concentration at TR4-C was 50.3 mg/l, ranging from 40.1 mg/l just below the ice to 54.5 mg/l at 44m on 14 March.

Temperature (oC) 0 5 10 15 20 25 2a. 0 2/5/2013 5 2/26/2013 10 3/14/2013 15 4/4/2013

20 4/17/2013

5/2/2013 25 5/21/2013 30 (meters) Depth 6/5/2013 35 6/18/2013 40 7/3/2013

45 7/16/2013 50

Temperature (oC) 0 5 10 15 20 25 2b. 0 7/16/2013 5 7/30/2013

10 8/14/2013

15 8/27/2013

20 9/11/2013 9/24/2013 25 10/15/2013 30 (meters) Depth 11/7/2013 35 11/21/2013 40 12/3/2013

45 50

Figure 2. Otsego Lake temperature profiles (oC) observed at TR4-C 5 February through 16 July (2a) and 16 July through 3 December (2b) 2013.

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Figure 3. Distribution of dissolved oxygen (isopleths in mg/L) as recorded in 2013 at site TR4-C on Otsego Lake. Points along the x-axis approximately indicate profile observation dates.

Chlorides Mean chloride concentrations in Otsego Lake from 1925 to 2013 are shown in Figure 4. Between 1994 and 2005 mean concentration increased steadily at of rate of 0.5 to 1.0 mg/l per year (Figure 4). Since then, mean annual concentrations have been variable and have actually trended slightly downwards, likely reflecting flushing of the system that has occurred during major flooding events (2006, 2011, 2013). The mean lake-wide concentration in 2013 was 14.0 mg/l. Chlorides in Otsego Lake have generally been attributed to road salting practices, with the greatest influx of the ion during spring snowmelt events or early-winter snow storms.

Nutrients Total phosphorus averaged 7 µg/l in 2013, ranging from below detection (< 4 µg/l) on multiple dates to 30 µg/l at 8m on 18 June. Concentrations were nearly homogeneous from surface to bottom on many dates during the growing season while higher, more variable, concentrations were observed occasionally (2 and 21 May, 18 June, and 11 September). No phosphorus release from the sediments was observed prior to fall turnover, as dissolved oxygen was present at concentrations sufficient to maintain iron-phosphorus bonds in sediment materials.

Nitrite+nitrate-N averaged 0.55 mg/l; ammonia-N was not measured, as it is generally below detectable levels (<0.02 mg/L) when dissolved oxygen exists in the bottom of the

- 15 - 46th Annual Report of the Biological Field Station hypolimnion. Total nitrogen analyses, yielding a mean of 0.72 mg/l, indicate an average organic nitrogen concentration of about 0.17 mg/l over the year. The concentrations of nitrate-N and Total Nitrogen were higher than in recent years, while the organic fraction was nearly identical (Waterfield and Albright 2011, 2012, and 2013).

Table 2. Areal hypolimnetic oxygen deficits (AHOD) for Otsego Lake, computed over summer stratification in 1969, 1972 (Sohacki, unpubl.), 1988 (Iannuzzi, 1991), and 1992-2013.

Time Interval AHOD (mg/cm2/day) 05/16/69 - 09/27/69 0.080 05/30/72 - 10/14/72 0.076 05/12/88 - 10/06/88 0.042 05/18/92 - 09/29/92 0.091 05/10/93 - 09/27/93 0.096 05/17/94 - 09/20/94 0.096 05/19/95 - 10/10/95 0.102 05/14/96 - 09/17/96 0.090 05/08/97 - 09/25/97 0.101 05/15/98 - 09/17/98 0.095 05/20/99 - 09/27/99 0.095 05/11/00 - 09/14/00 0.109 05/17/01 - 09/13/01 0.092 05/15/02 - 09/26/02 0.087 05/16/03 - 09/18/03 0.087 05/20/04 - 09/24/04 0.102 05/27/05 - 10/05/05 0.085 05/4/06 - 09/26/06 0.084 05/18/07 - 9/27/07 0.083 05/8/08 - 10/7/08 0.088 05/27/09 - 10/19/09 0.082 05/26/10 - 10/7/10 0.053 05/19/11 – 10/12/11 0.060 05/24/12 – 10/05/12 0.056 05/21/13 – 10/15/23 0.061

- 16 - 46th Annual Report of the Biological Field Station 20 18

16

14 12 10 8

Chloride (mg/l) (mg/l) Chloride 6 4 2

0

1920 1940 1960 1980 2000 2020

Year

Figure 4. Mean chloride concentrations at TR4-C, 1925-2013. Points later than 1990 represent yearly averages (figure modified from Peters 1987).

Chlorophyll a and Secchi Disk Transparency Chlorophylla concentrations were determined for samples collected on 10 dates from May through September 2013. Average 0-20m composite chlorophyll a concentration was 2.0µg/l (range= 0.6 to 5.4 µg/l). Temporal and spatial distribution of chlorophyll a was studied in June and July is discussed by Tanner and Bianchine (2014).

Secchi disk transparency measurements, presented in Figure 5, ranged from a low of 1.7m on 3 July to a growing season-maximum of 10.5m on 21 May. The temporal variation of transparency differed from that observed since 2010; May-September transparency measurements for 2010 through 2013 are presented in Figure 5. Mean summer Secchi transparencies for all years available (1935-2013) are given in Figure 6. Transparency was lower on average in 2013 than in previous years, with a growing season mean of 5.65m.

CONCLUSIONS

Lake conditions have been variable from year to year as interactions between management efforts and invasive species continue to develop. The establishment of zebra mussels (Dreissena polymorpha) (around 2007) coincided with the walleye (Sander vitreus) stocking program that was intended to control the population of alewife (Alosa pseudoharengus) (an invasive forage fish). While either event on its own would have ultimately led to increased water clarity, their simultaneous occurrence resulted in dramatic changes at the time when zebra mussel abundance increased and alewife abundance declined substantially. Algal biomass (inferred from chlorophylla concentrations), seasonal AHOD rate, Secchi disk transparency all declined, cladoceran mean size and abundance and lake trout abundance increased markedly. The lake trout (Salvelinus namayacush) population is currently being evaluated by the NYS DEC Region 4 Fisheries Biologists; the sudden decrease in alewife abundance, while a management success, was unexpected and has left the lake trout without an important component of its winter diet.

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2010 2011

0 0

2 2 4 4 6 6 8 8 10 10

Depth (meters) Depth 12 (meters) Depth 12 14 14 16 16

2012 2013

0 0

2 2 4 4 6 6 8 8 10 10

12 (meters) Depth 12 Depth (meters) Depth 14 14

16 16

Figure 5. May through September Secchi transparencies at TR4C, Otsego Lake, 2010 through 2013.

Year '35 '68 '69 '70 '71 '72 '73 '75 '76 '77 '78 '79 '80 '81 '82 '84 '85 '86 '87 '88 '92 '93 '94 '95 '96 '97 '98 '99 '00 '01 '02 '03 '04 '05 '06 '07 '08 '09 '10 '11 '12 '13 0.0

1.0

2.0

3.0

4.0

5.0

6.0

7.0

8.0

Figure 6. Mean summer (May through September) Secchi disk transparency collected at TR4-C, 1935-2013.

- 18 - 46th Annual Report of the Biological Field Station

REFERENCES

Albright, M.F. 1997. Otsego Lake limnological monitoring, 1996. In 29th Ann. Rept. (1996). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 1998. Otsego Lake limnological monitoring, 1997. In 30th Ann. Rept. (1997). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 1999. Otsego Lake limnological monitoring, 1998. In 31st Ann. Rept. (1998). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2000. Otsego Lake limnological monitoring, 1999. In 32nd Ann. Rept. (1999). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2001. Otsego Lake limnological monitoring, 2000. In 33rd Ann. Rept. (2000). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2002. Otsego Lake limnological monitoring, 2001. In 34th Ann. Rept. (2001). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2003. Otsego Lake limnological monitoring, 2002. In 35th Ann. Rept. (2002). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2004. Otsego Lake limnological monitoring, 2003. In 36th Ann. Rept. (2003). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2005. Otsego Lake limnological monitoring, 2004. In 37th Ann. Rept. (2004). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2006. Otsego Lake limnological monitoring, 2005. In 38th Ann. Rept. (2005). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2007. Otsego Lake limnological monitoring, 2006. In 39th Ann. Rept. (2006). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2008. Otsego Lake limnological monitoring, 2007. In 40th Ann. Rept. (2007). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. and H.A. Waterfield. 2009. Otsego Lake limnological monitoring, 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

APHA, AWWA, WPCF. 1989. Standard methods for the examination of water and wastewater, 17th ed. American Public Health Association. Washington, DC.

Austin, T., M.F. Albright, and W.N. Harman. 1996. Otsego Lake monitoring, 1995. In 28th Ann. Rept. (1995). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Bianchine, T. and C. Tanner. 2014. Chlorophyll a concentrations in Otsego Lake, summer 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Caracciolo, D. 2014. Population assessment of fresh-water mussels (Unionidae) in Otsego Lake since the introduction of zebra mussels (Dreissena polymorpha). In: 46th Ann. Rept. (2013) SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Ebina, J., T. Tsutsi, and T. Shirai. 1983. Simultaneous determination of total nitrogen and total phosphorus in water using peroxodisulfate oxidation. Water Res. 17(12):1721-1726.

EPA. 1983. Methods for the analysis of water and wastes. Environmental Monitoring and Support Lab. Office of Research and Development. Cincinnati, OH.

Groff, A., J.J. Homburger and W.N. Harman. 1993. Otsego Lake limnological monitoring, 1992. In 24th Ann. Rept. (1991). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W.N. 1994. Otsego Lake limnological monitoring, 1993. In 26th Ann. Rept. (1993). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W.N. 1995. Otsego Lake limnological monitoring, 1994. In 27th Ann. Rept. (1994). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Homburger, J.J. and G. Buttigieg. 1992. Otsego Lake limnological monitoring. In 24th Ann. Rept. (1991). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Iannuzzi, T.J. 1991. A model plan for the Otsego Lake watershed. Phase II: The chemical limnology and water quality of Otsego Lake, Occasional Paper #23. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Leach, J. H. 1993. Impacts of the zebra mussel (Dreissena polymorpha) on water quality and fish spawning reefs in western Lake Erie. In: Zebra Mussels: Biology, Impacts, and Control. Lewis Publishers, Boca Raton, FL p 381-397.

Liao, N. 2001. Determination of ammonia by flow injection analysis. QuikChem ® Method 10- 107-06-1-J. Lachat Instruments, Loveland, CO.

Liao, N. and S. Marten. 2001. Determination of total phosphorus by flow injection analysis colorimetry (acid persulfate digestion method). QuikChem ® Method 10-115-01-1-F. Lachat Instruments, Loveland, CO.

Peters, T. 1987. Update on chemical characteristics of Otsego lake water. In 19th Ann. Rept. (1986). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Pritzlaff, D. 2003. Determination of nitrate/nitrite in surface and wastewaters by flow injection analysis. QuikChem ® Method 10-107-04-1-C. Lachat Instruments, Loveland, CO.

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Slater, C. and M.F. Albright. 2013. Chlorophyll a concentrations in Otsego Lake, summer 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Stowell, S.G. 2014. Trap net monitoring of fish communities within the weedy littoral zone at Rat Cove and rocky littoral zone at Brookwood Point, Otsego Lake. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Teter, C. 2014. Water quality monitoring of five major tributaries in the Otsego Lake watershed, summer 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Vanassche, J., W.H. Wong, W.N. Harman, and M.F. Albright. 2014. Zebra mussels and other benthic organisms in Otsego Lake in 2008. In: 46th Ann. Rept. (2013) SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., and M.F. Albright. 2010. Otsego Lake limnological monitoring, 2009. In 42nd Ann. Rept. (2009). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., and M.F. Albright. 2011. Otsego Lake limnological monitoring, 2010. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., and M.F. Albright. 2012. Otsego Lake limnological monitoring, 2011. In 43rd Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., and M.F. Albright. 2013. Otsego Lake limnological monitoring, 2012. In 44th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2013. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Welschmyer, N.A. 1994. Fluorometric analysis of chlorophyll a in the presence of chlorophyll b and pheopigments. Limnol. Oceanogr. 39:1985-1992.

Yoo, A., P. Lord, and W.H. Wong. 2014. Zebra mussels (Dreissena polymorpha) monitoring using navigational buoys. In: 46th Ann. Rept. (2013) SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

YSI Incorporated. 2009. 6-Series multiparameter water quality sonde user manual. Yellow Springs, OH.

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A survey of Otsego Lake’s zooplankton community, summer 2013

C. Tanner1 and M.F. Albright

INTRODUCTION

This is a continuation of a study that has entailed monitoring the zooplankton community that exists in Otsego Lake, in part to evaluate efforts implemented to control alewife (Alosa pseudoharengus) by the addition of walleye (Sander vitreus) as well as influences by the zebra mussel (Dreissena polymorpha). Historically, Otsego Lake has been characterized as oligo-mesotrophic based on various trophic state indicators (Harman et. al 1997). In the 1970s, data collected on Otsego Lake to evaluate algal standing crops were indicative of oligotrophic conditions (water that has low nutrients along with density of algae, but high dissolved oxygen readings) (Godfrey 1977). However, there was evidence of phosphorus loading rates more indicative of a mesotrophic state (where water contains moderate amounts of dissolved nutrients, promoting moderate algal growth and leading to deep-water oxygen declines) (Godfrey 1977). This disjunct had been attributed to high rates of algal grazing by the crustacean zooplankton community in the lake that had been larger-bodied and more abundant compared to other lakes in New York studied at that time (Godfrey 1977). In 1986, alewife was documented in Otsego Lake (Foster 1990); by 1990 it was the dominant forage fish (Warner 1999). Being efficient grazers, they virtually eliminated the larger bodied crustacean plankton (Warner 1999). The zooplankton community changed from crustacean dominance to rotifers gaining dominance (Foster and Wigens 1990). Rotifers sequester fewer nutrients and have substantially lower algal grazing rates than crustaceans (Warner 1999). Through the 1990s and early 2000s, higher algal standing crops lead to lower transparencies in the summer and the increased rates of hypolimnetic oxygen depletion (Harman et al. 2002). Though there were mitigative efforts set forth to reduce the nutrient inputs in the lake (Albright 2005), the efforts seemed to be overshadowed by the indirect influence of the still- dominant alewife. Walleye (Sander vitreus) have been stocked into Otsego lake since 2000 (Cornwell 2005) at a targeted rate of 80,000 per year (though most years the numbers have been lower; Sanford 2012). The expectation was that predation on alewife might allow for the re-establishment of crustacean zooplankton through trophic cascading, returning oligotrophioc conditions to Otsego Lake (Cornwell 2005). Zebra mussels were first documented in Otsego lake in 2007 (Waterfield 2009) and by 2010, adults had become widespread throughout substrate all over the lake (Albright and Zaengle 2012). This study helps give insight on the zebra mussel reproductive timing even though the composite samples are not suggestive of the entire lakes condition and it is not certain of the affects by the zebra mussel in the zooplankton community.

1 F.H.V. Mecklenburg Conservation Fellow, summer 2013. Present affiliation: State University at Albany. Funding provided by the Otsego County Conservation Association.

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This report summarizes surveys of the zooplankton community of Otsego Lake through 2013. Aside from conventional sampling at the lakes deepest spot, on three dates samples were collected at two additional sites to evaluate the spatial heterogeneity of their distribution.

METHODS

From 2 May to 24 September 2013, samples were taken biweekly at TR4-C (Figure 1) to evaluate the temporal distribution of the zooplankton community at the deepest location of Otsego Lake. This site historically has been monitored regularly for physical, chemical and biological parameters. On 3 July, 17 July and 30 July, zooplankton samples were also collected at sites TR3-C and TR5-C (Figure 1). These additional sites were included this year to gain insight into the spatial variations of the plankton community.

At each site, a conical 63 µm plankton net with a 0.2m diameter opening was used for collecting zooplankton. The end of the cup was weighted and the net was lowered to, then hauled up from, 12 m (the approximate depth to the thermocline by late summer). A G.O.™ mechanical flow meter was mounted across the net opening, allowing for calculation of the volume of lake water filtered. The concentrated samples were preserved with ethanol.

Samples were analyzed one ml at a time on a gridded Sedgwick Rafter cell. Zooplankton were identified, measured and enumerated using a research grade compound microscope with digital imaging capabilities. Typically, at least 100 organisms were viewed per sample.

After each slide was assessed as above, cross polarized light was employed and the cell was viewed again to enumerate zebra mussel veligers as described by Johnson (1995).

Mean densities as well as lengths for cladocerans, copepods and rotifers were used to calculate dry weight (Peters and Downing 1984), daily filtering rate (Knoechel and Holtby 1986) and phosphorus regeneration (Esjmon-Karabin 1983) on each date sampled according to the equations provided in Table 1.

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Figure 1. Otsego Lake, New York, showing the three sample sites (TR3-C, TR4-C and TR5-C).

Table 1. Equations used to determine zooplankton dry weight (Peters and Downing 1984), filtering rates (Knoechel and Holtby 1986), and phosphorus regeneration rates (Esjmon-Karabin 1983).

Dry weight: D.W. = 9.86*(length in mm)2.1 Filtering Rate: F.R. = 11.695*(length in mm)2.48 Phosphorus regeneration: Cladocerans: P.R. = .519*(dry weight in ug)-.023*e0.039*(temp. in C) Copepods: P.R. = .229*(dry weight in ug)-.645*e0.039*(temp. in C) Rotifers: P.R. = .0514*(dry weight in ug)-1.27*e0.096*(temp. in C)

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RESULTS AND DISCUSSION

Table 2 summarizes the data collected from TR4-C over the summer of 2013, including mean epilimnetic temperature (which influences phosphorus regeneration rates), zooplankton densities, mean lengths and dry weights, dry weights per liter, phosphorus regeneration and filtration rates.

Figure 2 provides the calculated dry weights of rotifers, copepods and cladocerans on each date sampled at TR4C over the summer of 2013. Figures 3, 4 and 5 provide comparable data from 2012, 2011 and 2010, respectively. It is difficult to discern any seasonal pattern over the recent years, though rotifers continue to comprise only a minor part of the community, in contrast to the period during which alewife were dominant (Warner 1999). An exception was encountered on 17 July 2013 when the relatively large-bodied rotifer Asplanchna priodontus was common.

Over the past several years, the zooplankton community in Otsego Lake has changed significantly. In the 2000s the community of cladocerans was predominantly Bosmina longirostris, a small bodied organism, typically around 0.3mm. When Daphnia were present, their measurements were around 0.6 to 0.7 mm (Harman et al. 2002). In the more recent years, the Daphnia have increased relative to Bosmina, and have increased in mean size of over 1.0 mm, leading to an increase in the mean cladoceran length. Throughout the 2012 season, the cladocerans were comprised of 98 percent of Daphnia sp., averaging 21.5/1 and having a mean length of 1.19 mm (Albright 2012). In 2013, the crustacean community was split equally between Bosmina and Daphnia and averaged 5.5 crustaceans/l and 1.0 mm length. Table 3 summarizes the mean crustacean density, mean cladoceran size and mean dry weight, percent of the epilimnion filtered per day and phosphorus regeneration by crustaceans in 2000 and 2002 – 2013 for Samples collected at TR4-C.

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Avg Avg Mean Dry Phos. Regen. Rate Phos. Regen. Filtering % Temp. #/L length Dry Wt (ugP*mgdrywt-1 Rate Rates Epilimnion (°C) (mm) Wt (µg) (µg/L) *ind*h-1) (µg/l/day) (ml/ind/day) filtered/day 5/2 7.27 Cladocera 1.46 0.952 15.29 22.37 0.368 0.198 10.355 1.51 Copepoda 23.57 0.490 2.87 67.59 0.154 0.250 1.990 4.69 Rotifers 0.65 0.147 0.18 0.12 0.000 0.000 0.100 0.01 Total 90.08 0.448 6.21 5/21 10.14 Cladocera 8.72 1.065 13.45 117.27 0.424 1.193 13.682 11.93 Copepoda 16.63 0.493 3.45 57.41 0.153 0.211 2.028 3.37 Rotifers 0.16 0.153 0.19 0.03 0.188 0.000 0.110 0.00 Total 174.71 1.404 15.31 6/5 13.65 Cladocera 6.70 1.130 14.31 95.93 0.479 1.103 15.847 10.62 Copepoda 14.85 0.517 3.67 54.53 0.655 0.858 2.273 3.37 Rotifers 2.39 0.088 0.06 0.15 1.683 0.006 0.028 0.01 Total 150.60 1.967 14.01 6/18 14.19 Cladocera 2.55 1.241 18.30 46.73 0.463 0.519 19.976 5.10 Copepoda 5.89 0.797 6.97 41.08 0.114 0.112 6.671 3.93 Rotifers 15.91 0.088 0.06 0.97 0.934 0.022 0.028 0.05 Total 88.78 0.653 9.08 7/3 16.74 Cladocera 1.32 1.182 18.45 24.32 0.462 0.269 17.700 2.33 Copepoda 15.82 0.280 1.46 23.15 0.312 0.173 0.497 0.79 Rotifers 1.88 0.146 0.21 0.39 0.196 0.002 0.099 0.02 Total 47.86 0.444 3.14 7/17 18.66 Cladocera 5.82 0.845 15.15 88.13 0.483 1.022 7.705 4.48 Copepoda 37.48 0.298 1.60 59.90 0.294 0.423 0.582 2.18 Rotifers 139.57 0.252 0.77 106.88 0.038 0.096 0.385 5.37 Total 254.91 1.541 12.03

Table 2. Summary of site TR4-C of 2013 for mean epilimnetic temperature, zooplankton densities and mean length per taxa, as well as derived values for mean weight per individual and per liter, phosphorus regeneration per individual and per liter, filtering rates per individual and the percent epilimnion filtered per day.

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Avg Avg Mean Dry Phos. Regen. Rate Phos. Regen. Filtering % Temp. #/L length Dry Wt ugP*mgdrywt-1 Rate Rates Epilimnion (°C) (mm) Wt (µg) (µg/L) *ind*h-1 (µg/l/day) ml/ind/day filtered/day 7/30 18.57 Cladocera 30.35 0.42 3.48 105.69 0.804 2.038 1.333 4.04 Copepoda 32.30 0.29 1.55 49.94 0.357 0.428 0.532 1.72 Rotifers 80.27 0.12 0.14 11.00 0.396 0.105 0.066 0.53 Total 166.63 2.571 6.29 8/14 18.36 Cladocera 0.21 0.716 5.86 1.23 0.707 0.021 5.112 0.11 Copepoda 14.03 0.465 2.87 40.23 0.237 0.229 1.749 2.45 Rotifers 0.84 0.127 0.13 0.11 0.415 0.001 0.070 0.01 Total 41.57 0.251 2.57 8/27 18.12 Cladocera 0.61 1.003 9.92 6.01 0.621 0.090 11.775 0.71 Copepoda 30.90 0.385 2.15 66.43 0.283 0.452 1.098 3.39 Rotifers 27.27 0.109 0.09 2.59 0.621 0.039 0.048 0.13 Total 75.03 0.580 4.24 9/11 17.85 Cladocera 1.02 1.076 11.76 12.05 0.591 0.171 14.037 1.44 Copepoda 34.84 0.317 1.52 53.03 0.350 0.446 0.680 2.37 Rotifers 31.77 0.111 0.10 3.14 0.584 0.044 0.050 0.16 Total 68.22 0.661 3.96 9/24 16.09 Cladocera 1.32 1.378 20.08 26.44 0.522 0.331 25.912 3.41 Copepoda 22.72 0.464 2.71 61.47 0.242 0.357 1.742 3.96 Rotifers 15.14 0.103 0.08 1.29 0.708 0.022 0.041 0.06 Total 89.20 0.710 7.43

Season mean Cladocera 5.461 1.001 13.278 49.651 0.538 0.632 13.039 4.155 Copepoda 22.638 0.436 2.802 52.250 0.287 0.358 1.804 2.929 Rotifers 28.714 0.132 0.183 11.515 0.524 0.031 0.093 0.576 Total 113.42 1.021 7.66 Table 2. Summary of site TR4-C of 2013 for mean epilimnetic temperature, zooplankton densities and mean length per taxa, as well as derived values for mean weight per individual and per liter, phosphorus regeneration per individual and per liter, filtering rates per individual and the percent epilimnion filtered per day.

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400 Rotifera 350 Copepoda

300 Cladocera 250 200 150 100 Dry weight (ug/l) 50 0 5/2 5/21 6/5 6/18 7/3 7/17 7/30 8/14 8/27 9/11 9/24

Figure 2. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake over the summer of 2013 at TR4-C. 400 Rotifera 350 Copepoda

300 Cladocera 250 200 150 100 Dry weight (ug/l) 50 0 5/9 5/24 6/7 6/21 7/4 7/19 8/2 8/16 9/5 9/19

Figure 3. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake over the summer of 2012 (Albright 2013) at TR4-C. 400 Rotifera 350 Copepoda

300 Cladocera 250 200 150 100 Dry weight (ug/l) 50 0 5/19 6/1 6/15 6/28 7/13 7/26 8/8 8/24 9/9 9/27

Figure 4. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake over the summer of 2011. (Albright and Zaengle 2012) at TR4-C.

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400 Rotifera 350 Copepoda

300 Cladocera 250 200 150 100 Dry weight (ug/l) 50 0 5/18 6/4 6/15 7/1 7/15 8/2 8/12 8/26

Figure 5. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake over the summer of 2010 at TR4-C (Albright and Leonardo 2011).

Table 3. Mean crustacean density, mean cladoceran size and mean dry weight, percent of the epilimnion filtered per day and phosphorus regeneration by crustaceans in 2000 and 2002 – 2013. Samples collected at TR4-C.

2000 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 Mean cladoceran size (mm) 0.29 0.30 0.36 0.53 0.55 0.55 0.34 0.54 0.69 0.81 0.76 1.19 1.00 Mean crustacean density (#/l) 208 146 132 163 159 159 154 178 97 56.7 59.4 21.5 28.10 Mean crustacean dry weight (ug/l) 175 145 177 261 206 206 128 321 142 143 155 122 102 Mean % of epilimnion filtered/day 11.9 9.9 12.7 25.1 19.2 19.2 12.2 31.9 9.5 10.8 12.1 11.5 7.70 Mean phosphorus regeneration 4.49 2.60 3.10 4.40 2.70 2.40 3.00 5.80 1.49 1.90 1.80 1.17 1.02 (ug/l/day)

Tables 6, 7 and 8 summarize the dry weight combined by rotifers, copepods and cladocerans in Otsego Lake on 3 July, 13 July and 30 July 2013 at TR3-C, TR4-C and TR5-C. These data suggest that the dry weights contributed by each group on each date were similar, though the small sample size (one vertical haul per site per date) does not allow for a statistical evaluation. One exception to the above was noted on 30 July when the dry weight contributed by cladocerans at TR4-C was markedly higher than at the other two sites.

Figure 9 illustrates the abundance of zebra mussel veligers in the 0-12 m composite samples collected over the summer of 2013 at TR4-C, and on the three dates sampled at TR3-C, and TR5-C. The mean density at TR4-C was 3.3 veligers/l, similar to the mean count in 2012 (4.0/l). The peak density was observed on 27 August, approximately two weeks later than in 2012 (21 June; Albright 2013). In 2010, the mean count was 8.8 veligers/l and the density peaked on 24 August (Albright and Leonardo 2011). (Data are not available for 2011).

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400 3 July 2013 Rotifera Copepoda

300 Cladocera

200

100 Dry weight (ug/l)

0 TR3C TR4C TR5C

Figure 6. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake on 3 July 2013 at TR3-C, TR4-C and TR5-C.

400 13 July 2013 Rotifera Copepoda

300 Cladocera

200

100 Dry weight (ug/l)

0 TR3C TR4C TR5C

Figure 7. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake on 13 July 2013 at TR3-C, TR4-C and TR5-C.

400 30 July 2013 Rotifera Copepoda

300 Cladocera

200

100 Dry weight (ug/l)

0 TR3C TR4C TR5C

Figure 8. Dry weight combined by rotifers, copepods and cladocerans in Otsego Lake on 30 July 2013 at TR3-C, TR4-C and TR5-C.

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30

TR4-C 25 TR3-C 20 TR5-C 15

10

Veliger density (#/liter) 5

0 5/1 5/15 5/29 6/12 6/26 7/10 7/24 8/7 8/21 9/4 9/18

Figure 9. Abundance of zebra mussel veligers in the 0-12 m composite samples collected over the summer of 2013 at TR4-C, and on the three dates sampled at TR3-C, and TR5-C.

CONCLUSION

Through the 1990s, when alewives were dominant, there were very low numbers of larger bodied crustaceans; plankton filtering rates were low, algal standing crops were high, transparencies were low and hypolimnetic oxygen demand was high (Harman et al. 2002). Following the establishment of walleye, alewife were virtually eliminated from the lake (Waterfield and Cornwell 2013; Stowell 2014) and the above trends were reversed. Secchi transparency was high (mean = 5.6m) and oxygen depletion rates were low (AHOD = 0.61 mg/cm2/day) (Waterfield and Albright 2013) and chlorophyll a was low, generally < 2 µg/l (Bianchine and Tanner 2014). While at a lower density than observed in 2012, Daphnia continued to be common and large bodied through the first half of the summer of 2013.

The density of zebra mussel veligers at the mid-lake site has not increased since 2010. The influence the mussels on the zooplankton community is not well understood.

REFERENCES

Albright, M.F. 2005. A report on the evaluation of changes in water quality in a stream following the implementation of agricultural best management practices. In 37th Ann. Rept. (2004). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. 2013. A survey of Otsego Lake’s zooplankton community. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. and O. Zaengle. 2012. A survey of Otsego Lake’s zooplankton community, summer 2011. In 44th Ann. Rept. (2011). SUNY Oneonta. Biol. Fld. Sta., SUNY Oneonta.

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Albright, M.F. and M. Leonardo. 2011. A survey of Otsego Lake’s zooplankton community, summer 2010. In 43rd Ann. Rept. (2010). SUNY Oneonta. Biol. Fld. Sta., SUNY Oneonta.

Cornwell, M.D. 2005. Re-introduction of walleye to Otsego Lake: Re-establishing a fishery and subsequent influences of a top predator. Occas. Pap. No. 40. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Esjmont-Karabin, J. 1984. Phosphorus and nitrogen excretion by lake zooplankton (rotifers and crustaceans) in relation to the individual body weights of the animals, ambient temperature, and presence of food. Ekologia Polska 32:3-42.

Foster, J.R. 1990. Introduction of the alewife (Alosa pseudoharengus) in Otsego Lake. In 22nd Ann. Rept. (1989) SUNY Oneonta Bio Fld. Sta., SUNY Oneonta.

Foster, J.R. and J. Wigen.1990. Zooplankton community as an ecological indicator in cold water fish community of Otsego Lake. In 22nd Ann. Rept. (1989). SUNY Oneonta Bio Fld. Sta., SUNY Oneonta.

Godfrey, P.J. 1977. An alalysis of phytoplankton standing crop and growth: Their historical development and trophic impacts. In 9th Ann. Rept. (1976). SUNY Oneonta Bio Fld. Sta., SUNY Oneonta.

Johnson, L.E. 1995. Enhanced early detection and enumeration of zebra mussel (Dreissena sp.) veligers using cross-polarized light microscopy. Hydrobiologia 312:139-147.

Harman, W.N., M.F. Albright and D.M. Warner. 2002. Trophic changes in Otsego Lake, NY following the introduction of the alewife (Alosa pseudohargenous). Lake and Reserv. Manage. 18(3)215-226.

Iannuzzi, T.J. 1991. A model plan for the Otsego Lake watershed. Phase II: The chemical limnology and water quality of Otsego Lake, Occasional Paper #23. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Knoechel, R. and B. Holtbly.1986 Construction of body length model for the prediction of cladoceran community filtering rates. Limnol. Oceanogr. 31(1):1-16.

Peters, R.H. and Downing, J.A. 1984 Empirical analysis of zooplankton filtering and feeding rates. Limnology and Oceanography, 29 (4). pp. 763-784.

Sanford, S. 2012. Pers. Comm. Sanford Bait Farm, Wolcott, NY.

Stowell, S.G. 2014. Trap net monitoring of fish communities within the weedy littoral zone at Rat Cove and rocky littoral zone at Brookwood Point, Otsego Lake. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Warner, D.M. 1999. Alewives in Otsego Lake, NY: a comparison of their direct and indirect mechanisms of impact on transparency and chlorophyll a. Occas. Pap. No.32. SUNY Oneonta Bio Fld. Sta., SUNY Oneonta.

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Waterfield, H.A. 2009. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake, 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.F. Albright. 2013. Otsego Lake limnological monitoring, 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2013. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Chlorophyll a concentrations in Otsego Lake, summer 2013

T. Bianchine1 and C. Tanner2

INTRODUCTION

Historically, Otsego Lake has been considered meso-oligotrophic based upon its morphology, algal standing crop, transparency and hypolimnetic dissolved oxygen concentrations (Godfrey 1977). This was attributed to high densities of larger crustacean zooplankton which effectively filter algae from the water. Godfrey (1977) suggested that if crustacean plankton were to be reduced in size or number, the lake would show signs of moving towards a more eutrophic state. Alewife (Alosa pseudoharengus) were first documented in the lake in 1986 (Foster 1990) and were abundant by the early 1990s (Harman et al. 2002). Alewife are efficient planktivores which caused substantial reductions in crustacean zooplankton size and abundance throughout the 1990s. The resultant reduction in algal grazing lead to higher algal standing crop (estimated by chlorophyll a concentrations), reduced Secchi transparencies, and greater rates of hypolimnetic oxygen depletion (Harman 2002), all of which are consistent with Godfrey’s (1977) postulation.

Otsego Lake has been stocked with walleye since 2000; the primary intent was to take advantage of the forage base provided by alewife to re-establish this popular sports fish. Concurrent monitoring has attempted to document any changes that might be related to this trophic modification to the lake, including measurements of alewife abundance (Waterfield and Cornwell 2014), descriptions of the zooplankton community (Tanner and Albright 2014) and physical/chemical profiles of the lake (Waterfield and Albright 2014).

In addition to the influence of alewife on Otsego’s algal standing crop, zebra mussels (Dreissena polymorpha ) were first documented in Otsego lake in 2007 (Waterfield 2009). This exotic bivalve has a high filtering rate and, following its establishment, algal standing crops and chlorophyll a typically decline while transparency increases (D’Itri 1997). Indeed, a marked increase in Otsego Lake’s transparency was coincident with the establishment of zebra mussels (Waterfield and Albright 2013).

Chlorophyll a is a photosynthetic pigment common to all algae and cyanobacteria which, on average, constitutes 1-2% of the dry weight of planktonic algae (APHA 2012). Algal biomass data provide insight into water quality and the trophic status of a lake. It is a function of both nutrient availability and grazing by primary consumers. Excessive production results in hypolimnetic oxygen depletion due to the decomposition of the organic matter (Wetzel 1975),

1 F.H.V. Mecklenburg Conservation Fellow, summer 2013. Present affiliation: Sharron Springs Central School. Funding provided by the Village of Cooperstown..

2 F.H.V. Mecklenburg Conservation Fellow, summer 2013. Present affiliation: State University at Albany. Funding provided by the Otsego County Conservation Association.

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jeopardizing cold water fish species such as lake trout (Salvelinus namaycush). This survey is a continuation of work to evaluate chlorophyll a concentrations in profile in Otsego Lake.

This project is intended to evaluate chlorophyll a concentrations in profile at the lake’s deepest point, TR4-C, and to evaluate composite (surface to 20 m depth) at two additional points to gain insight into spatial variation in its concentration. A more complete record of composite chlorophyll a concentrations at TR4-C is provided in Waterfield and Albright (2014).

METHODS

Chlorophyll a samples were collected from TR3-C, TR4-C and TR5-C (see Figure 1) throughout the summer of 2013. On 16 July, 30 July and 14 August, at TR4-C, samples were collected from surface to 20 m at 1 m intervals. On 16 and 30 July, composite surface-to-20 m samples were collected at all three sites.

Figure 1. Otsego Lake, NY, indicating sites where chlorophyll a concentrations were profiled over summer, 2013.

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Discrete samples at TR4-C were taken from the surface of the lake to 20 meters using a Kemmerer sampler. Composite samples were collected using a weighted garden hose lowered to a depth of 20 m and retrieved from a line attached to the hose’s bottom. The samples were then stored in dark, Nalgene bottles and kept in a cooler on ice to prevent degradation of chlorophyll a. Analysis followed the methods of Welschmeyer (1994). An aliquot of each sample (250ml) was vacuum-filtered through a 47mm Whatman® GF/A Glass Micro Fiber filter, which was subsequently folded in half and patted dry between paper towels to absorb excess water. Filters were placed in individual labeled petri dishes, wrapped in foil and placed in the freezer until further processing. On the day of analysis, filters were cut into small pieces and put in a grinding tube with roughly 3 to 4 ml of buffered acetone (90% acetone and 10% saturated MgCO3). Each filter was ground to a homogenous slurry using a snug-fitting Teflon pestle attached to a drill. The slurry was diluted to a volume of 10 ml with buffered acetone and allowed to sit for 30 minutes. Afterwards, the tubes were centrifuged for 10 minutes at 10,000 X G. The sample supernatant was then transferred into a cuvette and measured in a Turner Designs™ TD-700 fluorometer. Samples were processed under low light to prevent the chlorophyll from degrading. Sample chlorophyll a (ug/l) was determined by dividing the measured value by the sample volume (250 ml) and multiplying by the concentrate volume (10 ml).

RESULTS AND DISCUSSION

Figure 2 illustrates chlorophyll a concentrations vs. depth at TR4-C from 2002 to 2013. 2013 chlorophyll a concentrations from the surface to 10m were higher than those reported in recent years (2007, 2010-2012) while concentrations from 15m to 20m depth were the lowest reported since 2002. Extremely wet conditions in early summer are expected to have played a role in this increased algal production, as sediment and nutrient contributions likely increased during the intense storm events (i.e. Teter 2014; Waterfield and Albright 2014. Figure 3 depicts the mean chlorophyll a concentrations throughout the summer of 2013 for the three dates sampled at TR4-C. Figures 4 and 5 display composite chlorophyll a concentrations at the three sample sites, TR3-C, TR4-C and TR5-C, on 17 July and 30 July 2013, respectively.

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Average Chlorophyll a Concentrations Otsego Lake at TR4-C Concentration (ppb) 0 2 4 6 8 10 0

2

4

2002 6 2003 2004 8 2005

2006 2007 10 2010 2011 Depth (m)

12 2012 2013

14

16

18

20

Figure 2. Average chlorophyll a concentrations per depth at sample site TR4-C from 2002 to 2013, excluding 2008 and 2009. Data were retrieved from 2002 (Wayman 2003), 2003 (Schmitt 2004), 2004 (Murray 2005), 2005 (Zurmuhlen 2006), 2006 (Stevens 2007), 2007 (Ottley 2008), 2010 (Bauer 2011), 2011 (Levenstein 2012), 2012 (Slater 2013), 2013 (Bianchine/Tanner 2013).

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2013 Chlorophyll a Concentrations Otsego Lake at TR4-C Concentration (ppb) 0 2 4 6 8 10 0

2

4

6

16-Jul-13 8

30-Jul-13

10

14-Aug-13

12 Depth (m) Depth

14

16

18

20

Figure 3. Average chlorophyll a concentrations throughout the water column at TR4-C (see Figure 1) on 16 July, 30 July and 14 August 2013.

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Chlorophyll a Concentrations 16 July 2013 Sites TR3-C, TR4-C and TR5-C 6

5

4

3

2

1 Concentration (ppb)

0 TR3-C Comp TR4-C Comp. TR5-C Comp. Figure 4. Chlorophyll a concentrations in composite samples at sample sites on 16 July 2013.

Chlorophyll a Concentrations 30 July 2013 Sites TR3-C, TR4-C and TR5-C 6

5

4

3

2

1 Concentration (ppb)

0 TR3-C Comp TR4-C Comp. TR5-C Comp. Figure 5. Chlorophyll a concentrations in composite samples at sample sites on 30 July 2013.

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CONCLUSION

Data collected this year shows that the chlorophyll a levels are within the range typically observed in recent years, though the pattern from surface to 20m showed greater variation. Summer mean values at TR4-C, at less than 2 ug/l, are still indicative of low algal production and oligotrophic conditions (Carlson 1977). Highly variable weather conditions, including intense rain events and periods of extreme heat, could have played a role.

REFERENCES

APHA, AWWA, WPCF. 2012. Standard methods for the examination of water and wastewater, 22nd ed. American Public Health Association. Washington, DC.

Bauer, H. 2011. Chlorophyll a analysis of Otsego Lake, summer 2010. In 43rd Annual Report (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Carlson, R.E. 1977. A trophic state index for lakes. Limnol. Oceanogr. 22(2)362-369.

D’Itri, F.M. 1997. Zebra mussels and aquatic nuisance species. Ann Arbor Press, Inc.

Foster, J.R. 1990. Introduction of the alewife (Alosa pseudoharengus) in Otsego Lake. In 22nd Annual Report (1989). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Godfrey, P.J. 1977. An analysis of phytoplankton standing crop and growth: Their historical development and trophic impacts. In 9th Annual Report (1976). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W.N., M.F. Albright, and D.M. Warner. 2002. Trophic changes in Otsego Lake, NY following the introduction of the alewife (Alosa Pseudoharengus). Lake and Reservoir Management. 18(3):215-226.

Levenstein, A. 2012. Chlorophyll a concentrations in Otsego Lake, summer 2011. In 44th Annual Report (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Murray, K. 2005. Chlorophyll a concentrations in Otsego Lake, summer 2004. In 37th Annual Report (2004). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Ottley, S.G. 2008. Chlorophyll a concentrations in Otsego Lake, summer 2007. In 39th Annual Report (2007). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Schmitt, R. 2004. Chlorophyll a concentrations in Otsego Lake, summer 2003. In 36th Annual Report (2003). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Stevens, K. 2007. Chlorophyll a analysis of Otsego Lake, summer 2006. In 39th Annual Report (2006). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Tanner, C. and M.F. Albright. 2014. A survey of Otsego Lake’s zooplankton, 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2014. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, summer 2012. In 46th Annual Report (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. 2009. Update on zebra mussel (Dreissena Polymorpha) invasion and establishment in Otsego Lake, 2008. In 41st Annual Report. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.F. Albright. 2014. Otsego lake limnological monitoring, summer 2013. In 46th Annual Report (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.F. Albright. 2013. Otsego lake limnological monitoring, summer 2012. In 45th Annual Report (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Wayman, K. 2003. Chlorophyll a concentrations in Otsego Lake, summer 2002. In 35th Annual Report (2002). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Welschmyer, N.A. 1994. Fluorometric analysis of chlorophyll a in the presence of chlorophyll b and pheopigments. Limnol. Oceanogr. 39:1985-1992.

Wetzel, R.G. 1975. Limnology. W.B. Sanders Company, Philadelphia.

Zurmuhlen, S. 2006. Chlorophyll a analysis of Otsego Lake, summer 2005. In 38th Annual Report (2005). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Water quality monitoring of five major tributaries in the Otsego Lake watershed, summer 2013

Christopher Teter1

INTRODUCTION

Pollution that enters a water body from diffuse sources is commonly called non-point source pollution. Overland runoff flows through soils, picking up pollutants and carrying them downward into water bodies. At lower elevations, pollutants may become concentrated in streams, rivers, ponds and lakes. Possible contaminants include excess fertilizers (phosphorus and nitrogen), herbicides, insecticides, sediment, salts, and bacteria (USEPA 2012). Because of heavy agricultural use in the Otsego Lake watershed, monitoring efforts focus on the nutrients phosphorus and nitrogen.

Otsego Lake was previously classified as mesotrophic, with oligotrophic characteristics (Iannuzzi 1991). Through the 1990s, Otsego Lake had been exhibiting increasing characteristics of eutrophy (Albright 2001), namely increasing rates of deep water oxygen depletion during summer stratification. Nutrients added to the lake promote algal growth which, upon death, will decompose, consuming dissolved oxygen.

The above situation has been compounded by the introduction of two exotic species. The introduction of the invasive fish species, alewife (Alosa pseudoharengus), by 1986 (Foster 1989) created eutrophication-like conditions in Otsego Lake because they effectively removed larger bodied crustacean zooplankton, which previously had been efficient algal grazers (Harman et. al. 1997). The additional introduction of zebra mussels (Dreissena polymorpha) in about 2008 increased lake clarity by decreasing phytoplankton populations through high rates of filtration (Waterfield 2009). Monitoring of the nutrients in the lake itself has not been wholly effective in determining the success of watershed nutrient control efforts because of the addition of these exotic fauna. Therefore, the direct monitoring of the available nutrients within Otsego Lakes tributaries is an important component of assessing effectiveness of agricultural best management practices (BMPs).

This study is an annual continuation of work initiated in a 1996 study conducted by the SUNY College at Oneonta’s Biological Field Station to evaluate the effectiveness BMPs that are currently in place within the Otsego Lake watershed (Hewett 1996). BMPs are conservation techniques recommended by the United States Department of Agricultures (USDA). In 1998, the municipalities surrounding the lake agreed upon a management plan, with priority given to reducing phosphorus inputs (Anon. 1998). Because 44 percent of the Otsego Lake watershed is being used for agricultural purposes (Harman et al. 1997), loading by this source was targeted for management. To date, 23 farms within close proximity to stream in the watershed have employed BMPs. The funding for these BMPs was provided for in part by the USDA

1 Rufus J. Thayer Otsego Lake Research Assistant, summer 2013. Current affiliation: SUNY Oneonta. Funded by the Otsego County Conservation Association.

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Environmental Quality Incentive Program (Otsego County Water Quality Coordination Committee 1998). Techniques such as conservation tillage, crop nutrient management, pest management, manure storage management and conservation buffers are used to prevent soil erosion of, and reduce sediment and nutrient loading to streams.

METHODS

This year’s data were collected in accordance with the methods of previous years (i.e., Mehigan 2013). The five tributaries monitored and the numbers of sample sites on each are: White Creek (3), Cripple Creek (5), Hayden Creek (8), Shadow Brook (5), and Mount Wellington (2). A list of sample sites is displayed in Table 1, along with their GPS coordinates and physical descriptions. The locations of the sample sites and their relation to current BMPs in use in the watershed are shown in Figure 1. These sites were chosen because of their accessibility and proximity to farms using BMPs.

Each site was visited on the Tuesday of every week from 30 May to 6 August 2013. Physical parameters were recorded at each sample site using a YSI (6820 V2) multi-parameter water quality sonde. Before each use, the sonde was calibrated using the manufacturer’s specifications. Water quality parameters measured included temperature (°C), pH, specific conductivity (µs/cm), oxidation reduction potential (mv), percent dissolved oxygen, oxygen concentration (mg/l), and turbidity (NTU).

Chemical analyses were conducted using water samples collected at each point in 125ml acid washed plastic bottles. Samples were chilled until brought to the laboratory, where they were preserved to pH<1. Analysis of the samples included nitrite+nitrate, total nitrogen and total phosphorus concentrations. The analysis was conducted using Lachat® QuikChem FIA+ Water Analyzer.

Table 1. GPS coordinates and physical descriptions of sample locations (modified from Putnam 2010).

White Creek 1 (WC1): N 42º 49.612’ W 74º 56.967’ South side of Allen Lake on County Route 26 over a steep bank.

White Creek 2 (WC2): N 42º 48.93’ W 74º 55.29’ Plunge-pool side of stream on County Route 27 (Allen Lake Road) where there is a large dip in the road.

White Creek 3 (WC3): N 42º 48.407’ W 74º 54.178’ West side of large stone culvert under Route 80, just past the turn to Country Route 27.

Cripple Creek 1 (CC1): N 42º 50.878 W 74º 55.584’ Weaver Lake accessed from the north side of Route 20 just past outflow of beaver dam.

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Table 1 (cont.). GPS coordinates and physical descriptions of sample locations (modified from Putnam 2010).

Cripple Creek 2 (CC2): N 42º 50.603’ W 74º 54.933’ Young Lake accessed from the west side of Hoke Road. The water at this site is shallow; some distance from shore is required for sampling.

Cripple Creek 3 (CC3): N 42º 49.418’ W 74º 54.007’ North side of culvert on Bartlett Road.

Cripple Creek 4 (CC4): N 42º 48.837’ W 74º 54.032’ Large culvert on west side of Route 80. The stream widens and slows at this point; this is the inlet to Clarke Pond.

Cripple Creek 5 (CC5): N 42º 48.805’ W 74º 53.768’ Dam just south of Clarke Pond accessed from the Otsego Golf Club road. Samples were collected on the downstream side of the bridge.

Hayden Creek 1 (HC1): N 42º 51.658’ W 74º 51.010’ Summit Lake accessed from the east side of Route 80, north of the Route 20 and Route 80 intersection. Small pull off but researcher must wade in the water to place the probe.

Hayden Creek 2 (HC2): N 42º 51.324’ W 74º 51.294’ Downstream side of culvert on Dominion Road.

Hayden Creek 3 (HC3): N 42º 50.890’ W 74º 51.796’ Culvert on the east side of Route 80 north of the intersection of Route 20 and Route 80.

Hayden Creek 4(HC4): N 42º 50.267’ W 74º 52.175’ North side of large culvert at the intersection of Route 20 and Route 80.

Hayden Creek 5 (HC5): N 42º 49.996’ W 74º 52.501’ Immediately below the Shipman Pond spillway on Route 80.

Hayden Creek 6 (HC6): N 42º 49.685’ W 74º 52.773’ East side of the culvert on Route 80 in the village of Springfield Center.

Hayden Creek 7 (HC7): N 42º 49.279’ W 74º 53.984’ Large culvert on the south side of County Route 53.

Hayden Creek 8 (HC8): N 42º 48.869’ W 74º 53.291’ Otsego Golf Club, above the white bridge adjacent to the clubhouse. The water here is slow moving and murky.

Shadow Brook 1 (SB1): N 42º 51.831’ W 74º 47.731’ Small culvert on the downstream side off of County Route 30 south of Swamp Road.

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Table 1 (cont.). GPS coordinates and physical descriptions of sample locations (modified from Putnam 2010).

Shadow Brook 2(SB2): N 42º 49.891’ W 74º 49.067’ Large culvert on the north side of Route 20, west of County Route 31.

Shadow Brook 3 (SB3): N 42º 48.799’ W 74º 49.839’ Private driveway (Box 2075) off of County Route 31, south of the intersection of Route 20 and Country Route 31 leading to a small wooden bridge on a dairy farm.

Shadow Brook 4 (SB4): N 42º 48.337’ W 74º 50.608’ One lane bridge on Rathbun Road. This site is located on an active dairy farm. The streambed consists of exposed limestone bedrock.

Shadow Brook 5 (SB5): N 42º 47.441’ W 74º 51.506’ North side of large culvert on Mill Road behind Glimmerglass State Park.

Mount Wellington 1 (MW1): N 42º 48.843’ W 74º 52.608’ Stone bridge on Public Landing Road adjacent to an active dairy farm.

Mount Wellington 2 (MW2): N 42º 48.77’ W 74º 53.004’ Small stone bridge is accessible from a private road off Public Landing Road; at the end of the private road near a white house there is a mowed path which leads to the bridge. Water here is stagnant and murky. Sampled on the side opposite the lake before the confluence of Mount Wellington stream and the brook beside the golf course.

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Figure 1. Map showing sampling locations of five tributaries in the northern watershed of Otsego Lake, as well as locations of BMPs (asterisks).

RESULTS AND DISCUSSION

Temperature

Aquatic species, especially fishes, are sensitive to temperature changes. Highs and lows in temperature in a given water body can result in mortality or displacement of aquatic organisms (Mason 2002). In order to prevent these fluctuations, watershed managers may promote the growth of conservation buffers. These buffers allow vegetation to grow unmolested along stream and river banks, which allows streams to run cooler. Traditionally, the removal of woody plant species and tall herbaceous species along stream banks has allowed more sunlight to be absorbed by streambeds. Over the summer of 2013 the lowest value of 13.17 °C was located at site CC3, and the highest value of 31.9 °C was located at HC2. The mean values ranged from 16.20 °C to

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22.38 °C. Last year’s mean values ranged from 15.44°C at MW1 to 22.56°C at HC1 (Mehigan 2013). Figure 2 displays the mean temperatures recorded for the summer of 2013. Mean Temperature

24.0

22.0

20.0

18.0 Temperature (C) Temperature 16.0

14.0 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km) White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington Figure 2. Mean Temperatures of sampling sites along the stream gradients of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph.

pH

Most organisms are sensitive to pH, which is an expression of the concentrations of positive hydrogen ions on a negative logarithmic scale from 0-14. Aquatic organisms usually can only survive in the midranges of this scale (EPA 2012). The pH of water is heavily influenced by atmospheric, geological and biological processes. The pH of water also affects the availability of dissolved oxygen, nutrients, and toxic heavy metals. The lowest recorded value was 7.42 at site CC1, and the highest was 8.66 at HC1. The minimum and maximum mean pH values for sites in the summer of 2013 are 7.78 at CC2 and 8.33 at CC1 respectively. Last year’s mean pH ranged from 7.82 at CC1 to 8.28 at HC4 in 2012 (Mehigan 2013).The northern reaches of the Otsego Lake watershed are fed by carbonate rich groundwater seepage (Fetterman & Burgin 1997), making samples more basic than those of other streams. Figure 3 displays the mean pH recorded for sites during the summer of 2013.

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Mean pH 8.5 8.4 8.3 8.2 8.1 8.0 pH pH 7.9 7.8 7.7 7.6 7.5 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km) White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington

Figure 3. Mean pH of sampling sites along the stream gradients of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph. Conductivity Conductivity is a measure of a solution’s ability to conduct electricity, which increases with increasing ion content (Wetzel 2001). This measurement can be used to identify possible polluted runoff. The lowest value recorded was 0.135 ms/cm at site CC1, and the highest was 0.602 ms/cm at site CC3. The minimum and maximum mean specific conductance values for sites in the summer of 2013 are 0.208 ms/cm at WC1 and 0.602 ms/cm at CC3 respectively. Mean conductivity readings in the summer of 2011 ranged from 0.233 ms/cm at WC1 to 0.451 ms/cm at MW2 (Zaengle 2012). Figure 4 displays the mean specific conductivity of sampling sites in the summer of 2013.

Dissolved Oxygen Measurements of dissolved oxygen (DO) content show whether waters are favorable or unfavorable for fish and other organisms. Fish typically require more than 6mg/l (Murphy &Willis 1996). The amount of oxygen present in the tributaries of Otsego Lake can be affected by nutrient loading and a lack of shade for streams, two aspects that the USDA BMP’s attempt to improve. Cooler waters can hold more DO (Wetzel 2001). The lowest value recorded was 3.64 mg/l at site CC1, and the highest was 14.55 mg/l at site HC1. The minimum and maximum mean values for sites in the summer of 2013 were 6.47 mg/l at CC1 and 10.20 mg/l at HC1 respectively. Mean site DO concentrations in 2012 ranged from 5.04 mg/L at CC1 to 11.16 mg/L at SB4 (Mehigan 2013). Figure 5 shows the mean DO values for all sites sampled in 2013.

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Mean Conductivity 0.6

0.5

0.4

0.3

0.2 Conductivity (ms/cm) Conductivity

0.1 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km) White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington

Figure 4. Mean conductivity of sampling sites along the stream gradients of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph.

Mean Dissolved Oxygen 12.0 11.0 10.0 9.0 8.0 7.0 6.0

Disolved Oxygen (mg/l) Oxygen Disolved 5.0 4.0 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km) White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington

Figure 5. Mean dissolved oxygen of sampling sites along the stream gradients of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph.

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Turbidity

Turbidity is a lack of clarity of water, usually due to the absorption and scattering of light by suspended inorganic particles (Wetzel 2001). These particles can contribute to siltation of substrates, disrupting lake and stream benthic communities. High concentration of suspended particles can affect fish species ability to absorb dissolved oxygen, find food, and reproduce (Jorgensen 2013). In the study of the Otsego Lake watershed, this has only been the second year of turbidity data collection. This data collection was facilitated by a turbidity sensor which has been added to the BFS’ YSI probe to collect in situ turbidity data. The minimum and maximum mean values for sites in the summer of 2013 were 1.47 NTU’s at WC1 and 49.1 NTU’s at SB5 respectively. The lowest values recorded were typically bellow detection levels for the probe used. The highest turbidity value recorded was 225.7 NTU’s at site SB5. In 2012, turbidity ranged from 3.68 NTU at CC3 to 26.03 NTU at MW2 (Mehigan 2013). Figure 6 shows the mean turbidity of all sites sampled in 2013. Mean Turbidity 80 70

60 50 40 30 Turbidity (NTU) Turbidity 20 10 0 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km)

White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington Figure 6. Mean turbidity of sampling sites along the stream gradients of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph.

Nitrogen

Nitrogen is an essential nutrient for the growth of plants. Unnaturally high levels of nitrogen can be found in water bodies that receive runoff from agricultural lands. The addition of excess nitrogen to a system can increase its productivity (Wetzel 2001). Algae and plant growth will increase due to the increased availability of nitrogen as a fertilizer. Upon reaching the end of their growing season, decomposing plants and algae will contribute to the eutrophication of the lowest regions of the water column. The minimum and maximum mean values for the total nitrogen content of sites sampled in 2013 were 0.336 mg/l at WC2 and 2.33 mg/l at SB2 respectively. The lowest value of 0.21 mg/l was recorded at WC1, and the highest value of 3.64 mg/l was recorded at SB2. In 2012, mean total nitrogen values ranged from 0.37 mg/L at WC1 to

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2.3 mg/L at HC8 (Mehigan 2013). Figure 7 shows the mean total nitrogen of all sites sampled in 2013. Figure 8 shows the mean nitrite and nitrate concentrations for all sites sampled in 2013. Figure 9 shows the mean nitrite and nitrate concentrations at stream outlets since 1998, including data from 1991. Table 1 shows a comparison of mean nitrate concentrations since 1998, including data from 1991. Mean Total Nitrogen 3.5

3.0 2.5 2.0 1.5 1.0

Total Nitrogen (mg/L) Nitrogen Total 0.5 0.0 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km) White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington

Figure 7. Mean total nitrogen of sampling sites along the stream gradient of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph.

Mean Nitrite + Nitrate 2.5

2.0

1.5 (mg/L) 1.0

0.5 Nitrate + Nitrite Concentrations Concentrations Nitrite + Nitrate 0.0 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Distance from Otsego Lake (km) White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington

Figure 8. Mean nitrite + nitrate of sampling sites along the stream gradient of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph.

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Mean Nitrite + Nitrate at Stream Outlet 3

2.5

2

1.5

1

Nitrate (mg/L) Nitrate 0.5

0 White Creek Cripple Creek Hayden Creek Shadow Brook Mount Stream Outlets Wellington 1991 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

Figure 9. Mean nitrate+nitrite at stream outlets of 5 tributaries in the northern watershed of Otsego Lake 1991, 1998-2013.

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Comparison of Mean Nitrate Concentrations (mg/L) 1998-2013 1991 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 WC1 0.10 0.05 0.11 0.25 0.31 0.29 0.15 0.27 0.01 0.22 0.21 0.06 0.25 0.09 0.10 0.10 WC2 0.31 0.30 0.12 0.16 0.25 0.24 0.15 0.09 0.04 0.11 0.09 0.12 0.10 0.06 0.08 0.08 WC3 1.09 0.37 0.41 0.19 0.22 0.33 0.24 0.35 0.31 0.12 0.35 0.24 0.16 0.24 0.13 0.13 0.13 CC1 0.07 0.07 0.06 0.08 0.12 0.18 0.13 0.04 0.00 0.02 0.01 0.01 0.03 0.01 0.23 0.23 CC2 0.04 0.02 0.24 0.04 0.16 0.34 0.22 0.20 0.01 0.00 0.00 0.01 0.97 0.01 0.28 0.28 CC3 1.54 1.19 0.89 1.63 1.20 1.12 1.06 0.60 0.86 0.88 0.97 1.16 1.17 0.97 1.32 1.32 CC4 1.42 0.97 0.92 1.77 1.07 1.37 1.05 0.56 0.88 0.97 0.77 1.15 1.19 1.01 1.28 1.28 CC5 0.69 0.99 0.37 0.68 1.41 0.77 0.80 0.77 0.27 0.83 0.39 0.38 0.99 0.81 0.69 1.18 1.18 HC1 0.82 0.29 0.82 0.68 0.64 0.52 0.26 0.02 0.72 0.07 0.01 0.47 0.33 0.30 0.90 0.90 HC2 0.72 0.24 0.71 0.66 0.76 0.52 0.24 0.03 0.84 0.06 0.01 0.59 0.34 0.36 1.04 1.04 HC3 1.35 0.64 0.96 1.62 1.44 1.43 1.11 0.60 1.11 0.51 0.44 0.62 0.86 0.70 1.45 1.45 HC4 1.34 0.95 1.17 1.73 1.41 1.27 1.11 0.66 1.10 0.55 0.46 0.68 0.88 0.73 1.39 1.39 HC5 1.36 0.85 1.19 1.87 1.18 1.34 1.39 0.98 1.64 0.59 0.36 0.94 0.89 0.72 1.41 1.41 HC6 1.45 0.90 1.29 1.87 1.51 1.27 1.51 1.38 1.58 0.69 0.45 1.02 0.95 0.77 1.51 1.51 HC7 1.45 0.95 1.33 2.00 1.50 1.46 1.31 1.05 2.52 1.22 0.57 1.93 1.00 0.94 1.57 1.57 HC8 1.11 1.63 1.21 1.48 1.56 2.09 1.62 1.62 1.31 1.69 0.89 0.70 1.62 1.22 1.17 1.77 1.77 SB1 0.21 0.31 0.66 0.53 0.33 0.34 0.32 0.21 0.25 0.09 0.14 0.16 0.21 0.17 - - 0.96 SB2 1.86 1.21 1.45 1.40 1.80 1.33 1.39 1.55 0.61 0.98 0.95 1.43 1.34 1.57 1.79 1.79 SB3 1.56 0.77 1.57 1.37 1.38 1.36 1.19 0.73 0.94 0.57 0.44 1.34 0.65 0.89 1.69 1.69 SB4 1.39 0.87 1.56 1.55 1.43 1.47 1.02 0.73 0.88 0.63 0.57 1.31 0.77 0.94 1.72 1.72 SB5 0.90 1.20 0.58 1.27 1.27 1.11 1.05 1.04 0.47 0.87 0.35 0.39 1.22 0.51 0.78 1.48 1.48 MW1 0.91 1.11 0.78 1.14 2.31 2.46 1.17 0.67 0.70 0.55 0.23 0.79 0.32 0.50 1.24 1.24 MW2 1.47 0.68 1.10 1.06 1.66 2.70 1.58 1.60 1.18 0.83 0.35 0.89 1.07 1.15 1.15 1.15 * - - stream flow was too low for sample collection; no nutrient data exists for Site SB1 in 2012

Table 1. Comparison of mean nitrate concentrations (mg/L) 1998- 2013. 1991 is also included, but only the values for stream outlets were taken, where available.

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Phosphorus

Phosphorus is a component of many agricultural fertilizers. Phosphorus is an important component of all life, and is an essential mineral for the growth of flora and fauna. In Otsego Lake the addition of excess phosphorus can create increases in productivity, due to it being the limiting nutrient (Harman et al. 1997). In 2013 the minimum and maximum mean values for sites sampled in 2013 were 18.5 µg/L and 233.6 µg/L. The lowest value of 5 µg/L was recorded at HC2 and the highest 1090 µg/L was recorded at MW2. In 2012, mean total phosphorus values ranged from 20 µg/L at HC4 to 71µg/L at MW2 (Mehigan 2013). Figure 10 shows the mean total phosphorus content of samples taken at each site in 2013. Figure 11 shows mean total phosphorus at stream outlets from 1996 to 2013. Table 2 is a comparison of phosphorus concentrations since 2000.

Mean Total Phosphorus 300.0

250.0

200.0

150.0

100.0 Total Phosphorus (ug/L)

50.0

0.0 0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 Distance from Otsego Lake (km)

White Creek Cripple Creek Hayden Creek Shadow Brook Mount Wellington

Figure 10. Mean total phosphorus of sampling sites along the stream gradients of five major tributaries in summer 2013. Distance from the lake increases moving from left to right on the graph. Note – High phosphorus readings at site MW2 required Y axis scale to be increased from 90 µg/L to 300 µg/L.

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Mean Total Phosphorus at Stream Outlets 300

250 g/L) 200

150

100

Total Phosphorus (µ Total 50

0 White Creek Cripple Creek Hayden Creek Shadow Brook Mount Stream Outlets Wellington

1996 1997 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

Figure 11. Mean total phosphorus concentration (µg/L) at the most downstream sites (outlets) of 5 tributaries in the northern watershed of Otsego Lake 1996-2013.

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Comparison of phosphorus concentrations (µg/L), 2000-2012 Site 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 WC1 31 34 72 25 33 51 17 66 46 33 33 25 22 18 WC2 28 33 23 26 39 61 33 37 34 24 25 25 33 29 WC3 19 24 12 23 26 36 40 38 19 22 17 21 21 27 CC1 45 36 112 30 49 49 33 86 89 38 63 49 62 26 CC2 48 23 46 124 144 172 37 36 25 24 25 28 22 22 CC3 25 24 10 25 39 37 62 40 22 26 41 30 21 49 CC4 28 35 19 22 46 55 40 39 34 27 45 30 37 126 CC5 42 45 51 28 46 70 37 58 59 34 41 40 51 45 HC1 26 25 60 21 43 33 33 48 43 35 28 53 22 25 HC2 20 17 14 13 23 34 57 30 27 18 24 20 52 23 HC3 25 28 47 26 34 39 50 35 54 24 31 24 21 27 HC4 20 23 17 26 29 41 22 38 27 24 31 24 20 24 HC5 28 27 27 22 33 43 46 41 37 22 31 27 41 32 HC6 24 24 21 33 28 40 40 49 32 26 26 27 25 31 HC7 34 26 19 30 44 54 73 40 42 27 32 28 30 40 HC8 32 37 54 31 51 120 89 43 71 30 37 32 62 42 SB1 52 39 57 21 27 103 54 28 19 36 30 33 - - 23 SB2 56 43 24 31 45 63 50 17 32 34 29 21 27 37 SB3 28 36 46 24 37 40 30 35 30 25 35 24 32 36 SB4 48 37 27 27 62 62 22 26 39 38 26 22 42 39 SB5 39 54 40 34 63 85 38 45 44 37 38 31 45 92 MW1 38 45 36 50 83 51 23 54 33 29 45 25 26 40 MW2 142 192 99 136 88 214 69 65 38 57 68 46 71 234 * - - stream flow was too low for sample collection; no nutrient data exists for Site SB1 in 2012

Table 2. Comparison of total phosphorus concentrations (ug/L) 2000- 2013

CONCLUSION From the middle of June, 2013, to July there were several weeks of record setting rainfall for Otsego and Herkimer Counties. This influenced this year’s data by increasing the variability of the data set. Phosphorus and nitrogen values were higher than they have been for several tributaries compared to recent years. A cause of this could be found in the washing out of several dirt roads that cross Mount Wellington Creek, or unknown human disturbance.

Physical parameters were not significantly different this year than those of the last. Temperature averages did seem to drop in the Hayden Creek and Shadow brook compared to last year. The pH values for all sites seemed to be less basic than previously reported. Dissolved oxygen seems to be slightly higher overall at most sites. Turbidity values are considerably higher than they have been in past years as well. Overall, these differences are typical of streams that receive increased rainfall. Increased strength and speed of a stream can: lower temperatures, lower pH values, introduce more ions into a stream, create more turbulence for oxygen to

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dissolve into the water, and suspend more sediment in the water for longer (Ward & Elliot 1995). All of which are observed in this years data.

It may also be worth noting that rainfall events seemed to mirror the sampling regime of our researcher. Sampling was done every Tuesday morning for the length of the study. Unfortunately, rain events seemed to happen often on Monday evenings. This rhythmic deviation from normal weather patterns may have skewed trends in our data set. This is an example as to why continued monitoring of the Otsego Lake tributaries is beneficial and necessary. Continued data collection will define more accurate trends in the water quality of a tributary. By recording data for several decades, researchers can paint a better picture of the effects of BMP placement in their watershed.

REFERENCES

Albright, M.F. 2001. Otsego Lake limnological monitoring, 2001. In 34th Ann. Rept. (2001). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Anonymous. 2007. A plan for the management of the Otsego Lake watershed. Otsego County Water Quality Coordinating Committee. Otsego County. New York.

Fetterman, A. R., Burgin, B. 1997. Preliminary geochemical analysis of surface and groundwater in Cripple Creek, a tributary to Otsego Lake, Otsego Co., New York. . SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Foster, J.R. 1989. Introduction of alewife (Alosa pseudoharengus) in Otsego Lake. In 22nd Ann. Rept. (1988). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W.N., L.P. Sohacki, M.R Albright and D.L. Rosen. 1997. The State of Otsego Lake, 1936-1996. Occasional Paper #30. SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Hewett, B. L. 1996.Water quality monitoring and the benthic community in the Otsego Lake watershed. In 29th Annual Report (1997). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Iannuzzi, T.J. 1991. A model plan for the Otsego Lake watershed. Phase II: The Chemical limnology and water quality of Otsego Lake. Occasional Paper #23. SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Jorgensen, S., Tundisis, J. G., Matsumura-Tundisi, T. 2013. Handbook of inland aquatic ecosystems management.

Mason, Christopher Frank. Biology of freshwater pollution. Pearson Education, 2002.

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Mehigan, K. 2013. Water quality monitoring of five major tributaries in the Otsego Lake watershed, summer 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Murphy, B.R., Willis, D. W. Fisheries techniques. 1996. American Fisheries Society. Bethesda, MD.

United States Environmental Protection Agency. 2012. What is Non-point Source Pollution? Web. http://water.epa.gov/polwaste/nps/whatis.cfm

United States Environmental Protection Agency. 2012. CADDIS Volume 2: Sources, Stressors & Responses (pH). Web. http://www.epa.gov/caddis/ssr_ph_int.html

Ward, A. D., Elliot, W. J. Environmental Hydrology. 1995. CRC Press, Boca Raton, FL.

Waterfield, H.A. 2009. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake, 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Wetzel, R.G. 2001. Limnology, Lake and River Ecosystems, 3rd edition. Academic Press. San Diego, California. Zaengle, O. 2012. Water quality monitoring of five major tributaries in the Otsego Lake watershed, summer 2011. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Upper Susquehanna River Water Quality Monitoring:

Monitoring water quality and fecal coliform bacteria in the upper Susquehanna River, summer 2013

T. Bianchine1

INTRODUCTION

The Susquehanna River starts at Otsego Lake near Cooperstown, New York. The River travels southward through Pennsylvania until it reaches the Chesapeake Bay at Havre De Grace, Maryland and then into the Atlantic Ocean. The water of the Susquehanna River travels a distance of 444 miles. The Susquehanna is one of the largest rivers on the eastern coast of the United States. More than three-quarters of the entire river basin (20,960 square miles) is located in Pennsylvania. The Susquehanna River contributes about 446 million gallons of water per day at peak use (SRBC 2013).

Each summer during the summer months, the upper Susquehanna River is sampled at nine sites and tested for temperature, dissolved oxygen, specific conductivity, nitrate+nitrite, total nitrogen, total phosphorous and fecal coliform levels. Testing for these parameters is an important part in determining the health of the Susquehanna River and ensures that the Village of Cooperstown, which indirectly discharges its secondarily treated sewage into the river, is not jeopardizing the river. Monitoring could also help recognize any unauthorized discharges.

METHODS

The nine sites in Table 1 were monitored weekly, from 27 June to 13 August, between 8:00a.m. and 12:00p.m. The nine sites were located between Otsego Lake and Oaks Creek (Table 1; Figure 1). A YSI® 6820 V2-2 multiprobe was used to measure temperature, pH, conductivity, dissolved oxygen, and turbidity at each site. Water samples for nutrient analyses were collected in 125ml Nalgene® bottles from each site and processed by a Lachat® QuickChem FIA + Water Analyzer to determine nitrate+nitrite (Pritzlaff 2003), total nitrogen (Pritzlaff 2003) and total phosphorous (Liao and Marten 2001).

1 FHV Mecklenburg Conservation Intern, summer 2013. Present affiliation: Sharron Springs Central School. Funding provided by the Village of Cooperstown.

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Table 1. Locations and descriptions of the nine Upper Susquehanna sites.

Site Distance from Description source 3 144m Under the Main Street Bridge; accessed via slope beside the bridge. 6a 1012m Below the dam at Bassett Hospital; accessed from the northern corner of the lower parking lot of Bassett Hospital. 7 1533m Below the dam at Bassett Hospital; accessed from the southern corner of the lower parking lot of Bassett Hospital. 8 1724m Under the Susquehanna Ave. bridge west of the Clark Sports Center; accessed via the slope beside the bridge. 12 4119m Just above the sewage discharge of the Cooperstown Wastewater Treatment Plant, near Cooperstown High School. Accessed by an opening in the fence. 16 5460m Small bridge perpendicular to the road on Clark Property. Accessed by crossing a gated bovine grazing area (cow field). 16a 5939m Distinct bend in river alongside road on Clark Property, in field directly across from large house with hay rolls in front. Accessed by long path found on the right side of the field. Be cautious of barbed wire. 17 8143m Abandoned bridge on Phoenix Mill Road. 18 9867m Railroad trestle about 200m north of the railroad crossing on Rt. 11 going out of Hyde Park, accessed by walking on the railroad tracks.

Water samples for fecal coliform bacteria analysis were also collected at each site. The samples were kept in a cooler until processing. The membrane filter technique was used for each sample for coliform bacteria (APHA 1992). A total of six subsamples ranging from 10 ml to 50 ml were used from each sample and run through a pre-sterilized filter with a low-pressure vacuum. Multiple volumes were used to increase the likelihood that the optimum range of colonies (20-80) would be encountered at one of the volumes (APHA 1992). The Milipore® filter pads were then placed in culture dishes of the same brand, saturated with nutrient-laden broth, and placed in a 44.5 degrees Celsius water bath for a period of 24 hours. After the incubation period, the number of colonies on each culture dish were counted and recorded as colonies per 100ml. All lab equipment was sterilized in 70% ethanol and washed in hot water and dilution water between sample sites.

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Figure 1. Upper Susquehanna sites, summer 2013.

RESULTS AND DISCUSION Temperature

Temperature exerts a major influence on biological activity and growth. Mean temperature at each site (+/- standard error) over 2013 is given in Figure 2. Figure 3 compares mean temperature over 2013 with the summers 2004 through 2012.

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26

25

24

23

22

21

20 Temperature C) Temperature (Degrees 19

18 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 2. Average temperature profile for the upper Susquehanna, summer 2013.

2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 26

25

24

23

22

21

Temperature (Celsius) Temperature 20

19

18 0 2000 4000 6000 8000 10000 Distance from site (meters)

Figure 3. Average temperature profile for the upper Susquehanna, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

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pH pH is a measure of the acidic or basic nature of a solution. The concentration of the hydrogen ion activity in a solution determines the pH. Pollution can change a waterbody's pH, which in turn can harm animals and plants living in the water (Wetzel 2001). Mean pH at each site (+/- standard error) over 2013 is given in Figure 4. Figure 5 compares mean pH over 2013 with the summers 2004 through 2012.

Conductivity Conductivity measures the water’s ability to transmit electricity based upon the amount of dissolved ions within it (Wetzel 2001). Discharges to streams can change the conductivity depending on their make-up. A failing sewage system would raise the conductivity because of the presence of chloride, phosphate, and nitrate. Mean conductivity at each site (+/- standard error) over 2013 is given in Figure 6. Figure 7 compares mean conductivity over 2013 with the summers 2004 through 2012.

8.7

8.5

8.3

pH 8.1

7.9

7.7

7.5 0 2000 4000 6000 8000 10000 Distance from source (meters)

Figure 4. Average pH profile for the upper Susquehanna River, summer 2013.

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2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 8.5

8.3

8.1

pH

7.9

7.7

7.5 0 2000 4000 6000 8000 10000 12000 Distance from site (meters)

Figure 5. Average pH profile for the upper Susquehanna, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

0.4

0.35

0.3

0.25 Conductivity (mmho/cm)

0.2 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 6. Mean conductivity levels of the upper Susquehanna River, summer 2013.

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2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 0.4

0.35

0.3

0.25 Conductivity (umho/cm)

0.2 0 2000 4000 6000 8000 10000 Distance from source (meters)

Figure 7. Average conductivity levels of the upper Susquehanna, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

Dissolved Oxygen Dissolved oxygen is essential for the survival of all aquatic organisms (not only fish but also invertebrates). The main factor contributing to changes in dissolved oxygen levels is the decomposition of organic material (Wetzel 2001). Mean dissolved oxygen at each site (+/- standard error) over 2013 is given in Figure 8. Figure 9 compares mean dissolved oxygen over 2013 with the summers 2004 through 2012.

10

9

8

7

Dissolved Dissolved Oxygen (mg/l) 6

5 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 8. Average dissolved oxygen content of the upper Susquehanna, summer 2013.

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2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 10

9

8

7

(mg/l) oxygem Dissolved 6

5 0 2000 4000 6000 8000 10000 Distance from source (meters)

Figure 9. Dissolved oxygen levels of the upper Susquehanna, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

Turbidity

Turbidity is a measure of water clarity, influenced by how much the material suspended in water decreases the passage of light through the water. Higher turbidity increases water temperatures because suspended particles absorb more heat. Higher turbidity reduces the concentration of dissolved oxygen because warm water holds less dissolved oxygen than cold. Higher turbidity also reduces the amount of light penetrating the water, which reduces photosynthesis. This year is the first year for which turbidity has been tested. Mean turbidity at each site (+/- standard error) over 2013 is given in Figure 10.

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16

14

12

10

8

Turbidity 6

4

2

0 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 10. Average Turbidity content of the upper Susquehanna, summer 2013.

Total Phosphorus

Phosphorus comes from both point and nonpoint sources (Albright et al. 1996). Point sources include municipal waste treatment plants, industrial discharge, large confined livestock operations, and urban storm water. These sources are regulated by federal and state laws. Nonpoint sources of phosphorus include soil erosion and water runoff from cropland, lawns and gardens, home waste treatment systems, livestock pastures, rangeland, and even forests. Urban areas may produce significant nonpoint source phosphorus runoff due to over-application of fertilizer to lawns and gardens, pet waste, and other materials that collect on impervious surfaces (e.g. parking lots, rooftops, etc.) and are washed off during precipitation events. Mean total phosphorus at each site (+/- standard error) over 2013 is given in Figure 11. Figure 12 compares mean total phosphorus over 2013 with the summers 2004 through 2012.

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250

200

150

100

Total Phosphorus (ug/l) 50

0 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 11. Average phosphorus concentrations along the upper Susquehanna River, summer 2013.

2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 250

200

150

100 Total Phosphorus (ug/l) 50

0 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 12. Average phosphorus concentrations along the upper Susquehanna River, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

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Nitrogen

Nitrogen is a very common element; it is present in our water, our air, in plants and animals (Wetzel 2001). Small amounts of nitrogen are essential to the life, growth, and reproduction of all organisms. Inorganic nitrogen may exist in the free state as a gas N2, or as nitrate NO3-, nitrite NO2-, or ammonia NH3+. Organic nitrogen is found in proteins and is continually recycled by plants and animals. Mean nitrite+nitrate at each site (+/- standard error) over 2013 is given in Figure 13. Figure 14 compares mean nitrite+nitrate over 2013 with the summers 2004 through 2012. Mean total nitrogen at each site (+/- standard error) over 2013 is given in Figure 15. Figure 16 compares mean total nitrogen over 2013 with the summers 2004 through 2012.

1

0.8

0.6

0.4 Nitrite+Nitrate (mg/l) 0.2

0 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 13. Nitrate and nitrite concentrations of the upper Susquehanna River, summer 2013.

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2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 1

0.8

0.6

0.4 Nitrite+Nitrate (mg/l)

0.2

0 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 14. Average nitrite and nitrate concentrations of the upper Susquehanna River, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

2

1.5

1

Total Nitrogen (mg/l) 0.5

0 0 2000 4000 6000 8000 10000 Distance from source (meters)

Figure 15. Total nitrogen levels of the upper Susquehanna River, summer 2013.

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2005 2006 2007 2008 2009 2010 2011 2012 2013 2.00

1.50

1.00 Total Nitrogen (mg/l) 0.50

0.00 0 2000 4000 6000 8000 10000

Distance from source (M)

Figure 16. Average total nitrogen levels of the upper Susquehanna River, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

Fecal Coliform

Fecal coliform bacteria are the most common microbiological contaminants of natural waters. Fecal coliform live in the digestive tracks of warm-blooded animals. A fecal coliform test is used to determine whether water has been contaminated with fecal matter, though fecal coliforms bacteria are not pathogens themselves. The presence of fecal coliform indicates the possible presence of organisms that can cause illness. Mean fecal coliform at each site (+/- standard error) over 2013 is given in Figure 17. Figure 18 compares mean fecal coliform over 2013 with the summers 2004 through 2012.

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2000

1500

1000

500 Fecal coliform (colonies/100ml)

0 0 2000 4000 6000 8000 10000 Distance from source (meters)

Figure 17. Fecal coliform levels of the upper Susquehanna River, summer 2013.

2004 2005 2006 2007 2009 2010 2011 2012 2013 2000

1500

1000

500 Fecal Coliform (col./100 ml)

0 0 2000 4000 6000 8000 10000

Distance from source (meters)

Figure 18. Average fecal coliform levels of the upper Susquehanna River, summers 2004 (Hill 2005), 2005 (Bauer 2006), 2006 (Zurmuhlen 2007), 2007 (Coyle 2008), 2008 (Matus 2009), 2009 (Heiland 2010), 2010 (Bauer 2011), 2011 (Scott 2012), 2012 (Katz 2013), and 2013.

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REFERENCES

Albright, M.F. 2012 Monitoring the effectiveness of the Cooperstown wastewater treatment wetland. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F., L.P. Sohacki and W.N. Harman. 1996. Hydrological and nutrient budgets for Otsego Lake, NY and relationships between land form/use and export rates of its sub- basins. Occas. Pap. No. 29. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Bauer, E. 2006. Monitoring the water quality and fecal coliform in the upper Susquehanna River, summer 2005. In 38th Ann. Rept. (2005). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Bauer, H. 2011 Monitoring the water quality and fecal coliform in the upper Susquehanna River, summer 2010. In 43th Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Coyle, O.L. 2008. Monitoring water quality and fecal coliform bacteria in the Upper Susquehanna River, summer 2007. In 40th Annual Report (2007), SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Ebina, J.T. Tsutsui, and T. Shirai. 1983. Simultaneous determination of total nitrogen and total phosphorus in water using peroxodisulfate oxidation. Water Res. 17(12):1712-1726.

Eureka Environmental Engineering. 2004. Manta water quality probe startup guide. Austin, TX.

Hill, J.2005. Monitoring the water quality in the upper Susquehanna River, summer 2004. In 37th Ann. Rept. (2004). SUNY Oneonta Bio. Fld Sta., SUNY Oneonta.

Heiland, L. 2010. Monitoring water quality in the upper Susquehanna River, summer 2009. In 42st Ann. Rept. (2009). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta

Liao, N. 2001. Determination of ammonia by flow injection analysis. QwikChem® Method 10- 115-01-0-F. Lachat Instruments. Loveland, Colorado.

Liao, N. and S. Marten. 2001. Determination of total phosphorus by flow injection analysis chloriometry (acid persulfate digestion method). QwikChem® Method 10-115-01-1-F. Lachat Instruments. Loveland, Colorado.

Matus, J.E. 2009. Monitoring water quality and fecal coliform bacteria in the upper Susquehanna River, summer 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta

Pritzlaff, D. 2003. Determination of nitrate+nitrite in surface and wastewaters by flow injection analysis. QwikChem® Method 10-115-01-1-F. Lachat Instruments. Loveland, Colorado.

Susquehanna River Basin Commision. 2009. http://www.srbc.net/about/index.htm.

Wetzel, R.G. 2001. Limnology: Lake and reservoir systems. Academic Press, San Diego.

Wetzel, R.G. and G.E. Lichens. 1991. Limnological Analysis, 2nd ed. Springer-Verlag New York.

Zurmuhlen, S. J. 2007. Monitoring water quality in the upper Susquehanna River, summer 2006. In 39th Annual Report (2006), SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

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ARTHROPOD MONITORING

Mosquito data from light traps, Thayer Farm

William L. Butts

Light traps were deployed overnight on several dates from June to July 2013 at the Thayer Farm in Springfield, NY. Table 1 summarizes the results; Figure 1 illustrates approximate trap locations.

Table. 1. Summary of mosquito trapping efforts, 2013.

Date Site Results VI.25.2013 1 Low area beyond pond drainage No catch VII.2.2013 2 By path to boat dock No catch VII.9.2013 3 Above step ponds No catch VII.16.2013 4 Edge upper step pond No catch VII.30.2013 5 Step pond Anopheles sp. ♂

Figure 1. Approximate trap locations (1-5), Thayer Farm, 2013.

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Aquatic invasive species present in Otsego County, NY water bodies Annie Yoo1, Kaylee Herzog2, and Holly Waterfield CLM3 INTRODUCTION Otsego County, located in central New York, experiences increased populations in the summer months largely related to tourism and outdoor recreation. Abundant water resources in the county, combined with this increase in summer traffic contribute to the risk of invasive species transport between water bodies and major watersheds. Knowledge of the current distribution of invasive species will allow natural resource managers to prioritize transport prevention and population control measures.

Exotic invasive species are those not native to an area that outcompete native species. Such exotic, non-native species typically possess traits which, in combination with lack of natural disease and/or predators, allow them to out-compete native species (Santos et al. 2011) and in-so-doing have negative ecological and economic impacts. Some of these impacts include productivity losses in agriculture, forestry, and other segments of the U.S. economy (Pimentel 2005). Aquatic invasive species (AIS) pose major threats to biodiversity in ecosystems. They often overtake important native aquatic plants and animals, cause habitat degradation and loss, and interfere with water-based recreational activities (Zhang and Boyle 2010). Pimentel (2005) suggests it is difficult to estimate the full extent of the environmental damages caused by invasive species and the number of species extinctions they have caused because little is known about each of the ~750,000 species present in the United States. In most cases AIS are initially introduced to watersheds through recreational boating activities or unintentional “hitchhiking ” (Horvath 2008), which can occur through international trade, with invaders stowed in ships, planes, trucks, or packing materials (McNeely 2001).

Aquatic invasive species and their impacts have been the subject of BFS research since its inception in 1968. In terms of surveys to document AIS distribution, surveys were conducted in 2011 to assess AIS in the Catskills region (Harman 2012), but no studies have involved a county-wide assessment in Otsego County. This study was conducted to evaluate the presence/absence of aquatic invasive algae, vascular plants, zooplankton, and invertebrate benthos present in the water bodies of Otsego County, New York.

MATERIALS AND METHODS Eighteen lentic systems and 9 lotic systems (Table 1, Figure 1) were assessed for the presence of 24 exotic species (Table 2). Survey sites were chosen based on accessibility and also to achieve relatively even coverage of water bodies across the county, ensuring that the major water ways and water bodies were assessed. Sites chosen for the survey included NYS DEC Boat Launches and Public Fishing Access Points, as human traffic (and associated activities and

1 BFS Intern, summer 2013. Current affiliation: SUNY Oneonta. Funding provided by Otsego Land Trust. 2 SUNY Oneonta Biology Department Intern, summer 2013. Current affiliation: SUNY Oneonta. Funded, in part, by NSF. 3 Research Support Specialist, Biological Field Station (CLM = Certified Lake Manager).

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equipment) is the main vector for transport of invasive species from one locale to another. Additional survey sites for lotic systems were included at all accessible road-crossings. A complete listing of all site locations is provided in Appendix 1. Species not listed in Table 2 were not specifically searched for, though would have been noted if found.

Bottom composition and substrate conditions were characterized for each site, and the presence of a Department of Environmental Conservation angling access or boat launch site was noted. GPS coordinates (datum: NAD 1987) for each survey site were recorded using a Garmin GPSmap 76CSx.

Table 1.Otesgo County water bodies surveyed in 2013 for the presence/absence of 20 aquatic invasive species; Lentic (left) and lotic (right) systems

Number of Lentic Systems (Lakes & Ponds) Lotic Systems (Rivers & Streams) Sites Sampled Allen Lake (Richfield Springs, NY) Butternut Creek 13 Arnold Lake (Hartwick, NY) Cherry Valley Creek 8 Basswood Pond (Burlington, NY) Oaks Creek 8 Belvedere Lake (Cherry Valley, NY) Otego Creek 10 Canadarago Lake (Richfield Springs, NY) Schenevus Creek 8 Crumhorn Lake (Milford, NY) Susquehanna River 9 Gilbert Lake (New Lisbon, NY) Unadilla River 13 Goey Pond (Hartwick, NY) Unnamed creek, Lull Hill Rd. (Laurens, NY) 1 Goodyear Lake (Portlandville, NY) Wharton Creek (tributary of Otego Creek) 6 Hunt Union Pond (Oneonta, NY) Larchwood Lake (New Lisbon, NY) Neahwa Pond (Oneonta, NY) Otsego Lake (Otsego, NY) Silver Lake (New Berlin, NY) Summit Lake (Springfield, NY) Susquehanna State Park Pond (Milford, NY) Wetland, unnamed (Oneonta, NY) Wilber Lake (Oneonta, NY)

Lentic Systems Sixteen of the 18 lentic systems in this survey were examined via canoe. Permission was sought for access from private sites or onto water bodies with restricted public access (i.e., reservoirs, private residential communities). Observations were also made on shore. Presence of invasive emergent plants was assessed along shorelines, and benthic species were assessed by scanning the available hard substrates. Submerged vegetation was collected by hand-picking shallow rake tosses (two to four sites per system, dependent upon water body size and variability of substrate). Plankton samples were gathered using tow nets (80µm mesh). Otsego and Canadarago Lakes were not directly sampled during this survey due to the size and complexity

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of the systems; presence of invasive species was determined based on recent surveys by BFS personnel (i.e., Canadarago: Albright and Waterfield 2012; Otsego: McShane and Mehigan 2013, Tanner and Albright 2014, Vanassche and Wong in prep).

Figure 1. Map depicting all sites surveyed in the 2013 AIS survey. White dots represent lentic, or still-water systems, black dots indicate lotic, or flowing water, systems.

Lotic Systems Several locations were sampled along each of the nine lotic systems in this survey, resulting in a total of 76 sampling sites. Riparian areas were assessed at each site for the presence of invasive emergent plants. Seining, dip-netting, hand-sieving and hand-picking were also employed at each site to assess the presence of invasive invertebrates.

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Equipment Sterilization and Sample Preservation

At the conclusion of sampling in each location, all equipment was removed from the water and rinsed on-shore with a diluted bleach or rock salt solution. Any organisms which were inconclusively identified on site were saved, marked with collection data, and transported on ice to the laboratory for definitive identification. Plankton samples were transported on ice to the laboratory, preserved with 70% ethanol, and analyzed under a dissecting microscope.

Mapping and Data Portrayal Waypoints stored at each site were uploaded electronically using Global Mapper™ software. Information regarding the presence or absence of invasive species at each site was attached to each waypoint and graphically represented using ESRI ArcMap software. Data will also be uploaded to iMapInvasives, an online, GIS-based data management system used by citizen scientists, researchers, and natural resource managers to document, assess, and coordinate the management of invasive species.

Table 2. Invasive species for which presence or absence was assessed at each survey site. Algae Didymosphenia germinate Didymo Nitellopsis obtusa starry stonewort Vascular Plants Egeria densa Brazillian elodea Hydrilla verticillata Hydrilla Myriophyllum spicatum Eurasian watermilfoil Myriophyllum aquaticum parrot’s feather Myriophyllum heterophyllum variable-leaved watermilfoil Fallopia japonica Japanese knotweed Lythrum salicaria purple loosestrife Phragmites australis common reed Trapa natans water chestnut Hydrocharis morsus-ranae European frog bit Potamogeton crispus curly leaf pondweed Zooplankton Bythotrephes cederstroemi spiny water flea Cercopagis pengoi fish hook water flea Invertebrate Benthos Cordylophora caspia freshwater hydroid Corbicula fluminea Asiatic clam Dreissena polymorpha zebra mussel Dreissena bugensis quagga mussel Bithynia tentaculata faucet snail Cipangopaludina chinensis Chinese mystery snail Potamopyrgus antipodarium New Zealand mud snail Orconectes rusticus rusty crayfish Eriocheir sinensis Chinese mitten crab

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Figure 2. Survey sheet used to document observations of AIS presence/absence at each site surveyed in the 2013 AIS Survey of Otsego County.

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RESULTS AND DISCUSSION

Observations are summarized graphically in Figures 3-7 and in tabular form in Tables 3 and 4. Eight of the 24 surveyed species were observed in at least one location; this survey did not document the presence of any species previously undocumented in the county, though water chestnut, a high-priority species, was found in a new location. The species found were Orconectes rusticus, Trapa natans, Fallopia japonica, Myriophyllum spicatum, Lythrum salicaria, Potamogeton crispus, Nitellopsis obtusa, and Dreissena polymorpha.

Of the sites surveyed, invasive species were found at 48 stream sites, representing 8 of the 9 lotic systems, and in nine of the 18 surveyed lentic waterbodies (Figure 3, Tables 3 and 4). Sites containing the greatest number of AIS were lentic systems, the top three of which were the three largest lakes included in the survey, Canadarago Lake (7 spp.), Goodyear Lake (5 spp.), and Otsego Lake (7 spp.). A brief overview of each species observed is provided in the following paragraphs.

Orconectes rusticus (rusty crayfish) is the most prolific invasive crayfish in the US. Native to the Ohio River drainage, rusty crayfish have been spread to 15 states outside of their native range, primarily though bait-bucket introductions and also by releases from science classrooms (Olden et al. 2006). It inhabits lakes, streams, and rivers, preferring habitats where debris for cover is present (Gunderson 2008). Once introduced to a water body, rusty crayfish quickly displace native species of crayfish and result in cascading trophic impacts due in large part to their aggressive behavior and higher metabolism and food consumption (Gunderson 2008). Rusty crayfish were first documented in the upper Susquehanna River drainage in 1991 (Kuhlmann and Hazelton 2007).

Trapa natans (water chestnut) is a Eurasian macrophytic plant that occurs in dense beds of floating rosettes. It was originally introduced into North America as an ornamental plant, but is now present in lentic water bodies from Virginia north to Vermont in the US and in the province of Quebec in Canada (U.S. Dept. of Agriculture 2013). The dense beds impede recreational activities, displace submerged native plant species, alter littoral bentic inverteberate and epiphytic community composition, and greatly alter the environmental conditions the waters underneath, including increasing summer temperature, creation of anoxic conditions, and decreased light availability (Hummel and Kiviat 2004, Kornijow et al. 2010).

Native to Asia, Fallopia japonica (Japanese knowtweed) was first introduced into Europe in the mid-19th century and had rapidly invaded areas in the USA by 1877. It is easily identifiable by its hollow, bamboo-like stems that grow in excess of 3 m high. It is most commonly found in riparian zones where water is readily available, but is also common along upland roadsides. Its invasion occurs through the process of colonizing rhizomes, allowing the plant to quickly displace native plant species and reduce area for other animals to survive (Forman and Kesseli 2003).

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Figure 3. 2013 Otsego County AIS Survey results. Pie charts mark the location of each lotic survey site or lentic waterbody. The total number of AIS observed at each site is indicated by number of pie “slices” in the chart; each species is represented by a different pattern. In order to avoid overlap of symbols, pie charts indicate the approximate location of sample sites that are in close proximity to another. See insets (Figures 4, 5, 6 and 7) for an enlarged view.

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Figure 4. 2013 AIS Survey results in Northwestern Otsego County, NY. Pie charts indicate AIS observed at each survey site. The total number of AIS observed at a particular site is indicated by number of pie “slices” in the chart; each species is represented by a different pattern. In order to avoid overlap of symbols, pie charts indicate the approximate location of sample sites that are in close proximity to another.

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Figure 5. 2013 AIS Survey results in Northeastern Otsego County, NY. Pie charts indicate AIS observed at each survey site. The total number of AIS observed at a particular site is indicated by number of pie “slices” in the chart; each species is represented by a different pattern. In order to avoid overlap of symbols, pie charts indicate the approximate location of sample sites that are in close proximity to another.

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Figure 6. 2013 AIS Survey results in Southwestern Otsego County, NY. Pie charts indicate AIS observed at each survey site. The total number of AIS observed at a particular site is indicated by number of pie “slices” in the chart; each species is represented by a different pattern. In order to avoid overlap of symbols, pie charts indicate the approximate location of sample sites that are in close proximity to another.

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Figure 7. 2013 AIS Survey results in Southeastern Otsego County, NY. Pie charts indicate AIS observed at each survey site. The total number of AIS observed at a particular site is indicated by number of pie “slices” in the chart; each species is represented by a different pattern. In order to avoid overlap of symbols, pie charts indicate the approximate location of sample sites that are in close proximity to another.

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Table 3. 2013 Otsego County AIS Survey results for lentic (still-water) systems. Multiple sites were assessed within each waterbody. Species present are indicated by an “x” while those absent are blank. Systems in which no AIS were observed during the survey are listed below the species table. Eurasian Japanese purple water curly leaf zebra rusty starry Lentic Systems (Lakes & Ponds) milfoil knotweed loosestrife chestnut pondweed mussel crayfish stonewort Basswood Pond (Burlington, NY) x Canadarago Lake (Richfield Springs, NY) x x x x x x x Crumhorn Lake (Milford, NY) x Goodyear Lake (Portlandville, NY) x x x x x Larchwood Lake (New Lisbon, NY) x Neahwa Pond (Oneonta, NY) x Otsego Lake (Otsego, NY) x x x x x x x Silver Lake (New Berlin, NY) x x Wetland, unnamed (Oneonta, NY) x x x

No AIS Observed: Allen Lake (Richfield Springs, NY) Arnold Lake (Hartwick, NY) Belvedere Lake (Cherry Valley, NY) Gilbert Lake (New Lisbon, NY) Goey Pond (Hartwick, NY) Hunt Union Pond (Oneonta, NY) Summit Lake (Springfield, NY) Susquehanna State Park Pond (Milford, NY) Wilber Lake (Oneonta, NY)

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Table 4. 2013 Otsego County AIS Survey results for lotic systems. Species present are indicated by an “x” while those absent are blank. Sites where no AIS were observed are listed. Site Eurasian Japanese curly leaf zebra rusty Lotic Systems (Rivers & Streams) Number milfoil knotweed pondweed mussel crayfish Buttnernut Creek, 13 sites sampled Butternut Creek 6 x Butternut Creek 7 x Butternut Creek 10 x Butternut Creek 11 x Butternut Creek 12 x Butternut Creek 13 x No AIS found: Butternut Creek 1-5, 8, 9 Cherry Valley Creek, 8 sites sampled Cherry Valley Creek 1 x Cherry Valley Creek 2 x Cherry Valley Creek 4 x No AIS found: Cherry Valley Creek 3, 5-8 Oaks Creek, 8 sites sampled Oaks Creek 1 x x x Oaks Creek 3 x x x Oaks Creek 4 x Oaks Creek 5 x Oaks Creek 6 x Oaks Creek 7 x x Oaks Creek 8 x No AIS found: Oaks Creek 2 Otego Creek, 10 sites sampled Otego Creek 3 x Otego Creek 5 x Otego Creek 8 x Otego Creek 9 x Otego Creek 10 x No AIS found: Otego Creek 1-2, 4, 6-7 x Schenevus Creek, 8 sites sampled Schenevus Creek 1 x x Schenevus Creek 2 x x Schenevus Creek 3 x Schenevus Creek 5 x Schenevus Creek 6 x x Schenevus Creek 7 x x Schenevus Creek 8 x No AIS found: Schenevus Creek 4

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Table 4 cont’d. 2013 Otsego County AIS Survey results for lotic systems. Species present are indicated by an “x” while those absent are blank. Sites where no AIS were observed are listed. Site Eurasian Japanese curly leaf zebra rusty Lotic Systems (Rivers & Streams) Number milfoil knotweed pondweed mussel crayfish Susquehanna River, 9 sites sampled Susquehanna River 1 x x Susquehanna River 2 x x x Susquehanna River 3 x x x Susquehanna River 4 x x x Susquehanna River 8 x Susquehanna River 9 x No AIS found: Susquehanna River 5, 6, 7 Unadilla River, 13 sites sampled Unadilla River 1 x Unadilla River 2 x Unadilla River 3 x x Unadilla River 4 x Unadilla River 5 x Unadilla River 6 x x Unadilla River 7 x Unadilla River 8 x Unadilla River 9 x Unadilla River 10 x Unadilla River 11 x x Unadilla River 12 x Unadilla River 13 x x Wharton Creek, 6 sites sampled Wharton Creek 1 x No AIS found: Wharton Creek 2-6

Myriophyllum spicatum (Eurasian watermifoil) and Potamogeton crispus (crispy pondweed) were introduced to North America from Europe (Santos et al. 2011). These are submerged plant species that have the ability to rapidly dominate aquatic systems through the formation of dense beds that displace native vegetation, reduce phytoplankton densities and open water nutrient concentrations, and impede recreational activities. Past studies have proposed that their competitive advantage and ability to rapidly colonize an area lies in their use of auto fragmentation and lower light and CO2 requirements (Gross et al. 1996).

Lythrum salicaria (purple loosestrife) was initially introduced to North America in the early 1800s and has since spread throughout the continent. It is now present in all lower 48 states with the exception of Florida. It has been declared as a noxious weed because of its negative impacts to wetlands, bog turtles, and mammals (Blossey et al. 2001). Many actions have been taken to stop spread of this plant or control established populations, though physical controls such as flooding, burning, and mowing are expensive and have had little or no long term

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effect. Biological control has been successful in reducing the density of L. salicaria and controlling populations over decades of monitoring (Albright et al. 2004). Populations documented in this survey may be targeted for establishment of Galerucella beetles in order to combat spread and recover native wetland plant communities.

Nitellopsis obtusa (starry stonewort) is an exotic charoid macroalga that has been present in Lake St. Clair since 1986. It can be identified by its starry rhizoids that are present in all parts of the plant at all times of the year in Michigan Lakes. It is light green colored when actively growing and its stem-like thallus is comprised of a single cell. Like previously described invasive submerged plants, it forms dense beds up to 2 m in thickness, out-competing native plants and drastically altering the littoral zone habitat (Pullman and Crawford 2010).

Dreissena polymorpha (zebra mussel) was observed in a number of locations throughout the county and is widespread across the continental US. It is one of the more dramatic invasive species due to its rapid range expansion and immediate economic impacts. Zebra mussels have directly affected native unionid clams; the species displaces native clams by attaching to their hard outer shell, essentially causing them to suffocate and/or starve. Unfortunately, it is easily transported to new water bodies as a free-floating larva or in the adult form attached to equipment, plants, etc. For example, any activity that can move water within or between water bodies can greatly accelerate the spread of D. polymorpha (Johnson and Carlton 1996).

This study along with other research on the transport, establishment, and growth of AIS may help natural resource, specifically lake, managers understand the biology of invasive species and prioritize prevention and control measures within Otsego County. Furthermore, it may help raise public awareness of impacts on biotic resources, water quality, cultural and recreational activities.

REFERENCES

Albright MF, WN Harman, H Meehan, S Fickbohm, S Groff and T Austin. 2004. Recovery of native flora and behavioral responses by Galerucella spp. following biocontrol of purple loosestrife. Am. Midl. Nat. 152:248-254.

Albright MF and HA Waterfield. The State of Canadarago Lake, 2011. 30th Tech. Rept. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Blossey B, L.C. Skinner and J. Taylor. 2001. Impact and management of purple loosestrife (Lythrum salicaria) in North America. Biodiversity and Conservation. 10(10):1787-1807.

Forman J. and R.V. Kesseli. 2003. Sexual reproduction in the invasive species (Fallopia japonica) Polygonaceae. American Journal of Botany. 90(4):586-592.

Gunderson, J. 2008. Rusty crayfish: a nasty invader; biology, identification, and impacts. Publication Number X34 of Sea Grant Minnesota. www.seagrant.umn.edu

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Gross E.M., H. Meyer and G. Schilling. 1996. Release and ecological impact algicidal hydrolysable polyphenols in Myriophyllum spicatum. Phytochemistry. 44(1):133-138.

Harman WN. 2012. 2011 Catskill region aquatic nuisance species survey for the Catskill Center for Conservation and Development. In: 44th Annual Report (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Horvath T. 2008. Economically viable strategy for prevention of invasive species introduction: Case study of Otsego Lake, New York. Aquatic Invasions. 3(1):3-9.

Hummel, M and E Kiviat. 2004. Review of world literature on water chestnut with implications for management in North America. J. Aquat. Plant Manage. 42:17-28

Johnson L.E. and J.T. Carlton. 1996. Post-establishment spread in large-scale invasions: dispersal mechanisms of the zebra mussel, Dreissena polymorpha. Ecology. 77(6):1686- 1690.

Kornijow R., D.L. Strayer and N.F. Caraco . 2010. Macroinvertebrate communities of hypoxic habitats created by an invasive plant (Trapa natans) in the freshwater tidal Hudson River. Fundamental and Applied Limnology/Archivfur Hydrobiologie. 176(3):199-207.

Kuhlmann, M.L. and P.D. Hazelton. 2007. Invasion of the upper Susquehanna River watershed by rusty crayfish (Orconectes rusticus). Northeastern Naturalist. 14(4):507-518.

McNeely J. 2001. Invasive species: a costly catastrophe for native biodiversity. Land Use and Water Resources Research. 1(2):1-10.

McShane, D. and K. Mehigan. 2013. 2012 aquatic macrophyte survey of Otsego Lake. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Olden J.D., J.M. McCarthy, J.T. Maxted , W.W. Fetzer and M.J. Vander Zanden. 2006. The rapid spread of rusty crayfish (Orconectes rusticus) with observations on native crayfish declines in Wisconsin (U.S.A.) over the past 130 years. Biological Invasions. 8:1621- 1628.

Pimentel D., R. Zuniga and D. Morrison. 2005. Update on the environmental and economic costs associated with alien-invasive species in the United States. Ecological Economics 52(2005):273-288.

Pullman, G.D. and G. Crawford. 2010. A decade of starry stonewort in Michigan. Lakeline. 36- 42.

Santos M., L. Anderson and S. Ustin. 2011. Effects of invasive species on plant communities: an example using submersed aquatic plants at the regional scale. Biological Invasions. 13(2):443-457.

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Tanner, C. and M.F. Albright. 2014. A survey of Otsego Lake’s zooplankton community, summer 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

U.S. Dept. of Agriculture. 2013. Plants profile for Trapa natans. PLANTS Database, US Department of Agriculture. http://plants.usda.gov . Accessed 04 Feb 2014.

Zhang C. and K. Boyle. 2010. The effect of an aquatic invasive species (Eurasian watermilfoil) on lakefront property values. Ecological Economics. 70(2):394-404.

Appendix A: Site locations of surveyed Otsego County lotic and lentic systems Location oN oW Butternut Creek, Co. Rt. 16, 1 mile N of Rt. 18 & 16 intersection 42o44.125 75o7.409 Butternut Creek, Burlington 42o43.397 75o07.346 Butternut Creek, Patent Rd. 42o42.583 75o07.202 Butternut Creek, Miller Rd. / El Jen Kay Rd. 42o40.650 75o08.316 Butternut Creek, BC fishing access site #2 42o40.232 75o08.648

Butternut Creek, Backus Rd./ Coles Bridge Rd. 42o38.878 75o09.815 Butternut Creek, Ben Hill Rd. 42o37.704 75o10.891 Butternut Creek, 1/4 mile E of Rt. 51 on Rt. 12 42o35.357 75o11.594 Butternut Creek, Rt. 49 42o33.113 75o13.410 Butternut Creek, Rt. 23 in Morris Village 42o32.691 75o14.292 Butternut Creek, Peet Rd. 42o31.454 75o15.338 Butternut Creek, Lovers Ln. 42o28.644 75o19.153 Butternut Creek, Rt. 3 bridge 42o26.163 75o20.810 Oneida St. wetland in Oneonta 42o26.624 75o06.275 Susquehanna River, Under the Main St. bridge 42o41.993 74o55.215 Susquehanna River, Below the dam at Bassett Hospital 42o41.579 74o55.283 Susquehanna River, Below the dam at Bassett Hospital 42o41.617 74o55.321 Susquehanna River, West of Clark Sports Center 42o41.554 74o55.638 Susquehanna River, Small bridge perpendicular to road on Clark property 42o40.728 74o56.266 Susquehanna River, Distinct bend in river alongside Clark property 42o40.543 74o56.319 Susquehanna River, Aban. Bridge on Pheonix Mill Rd 42o40.020 74o56.707 Susquehanna River, Railroad trestle 42o39.761 74o56.408 Susquehanna River, Just above sewage discharge 42o41.158 74o55.937

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Susquehanna River State Park, Crumhorn Fishing Access 42o33.386 74o56.564 Gilbert Lake, New Lisbon, NY 42o35.017 75o07.759 Gilbert Lake, New Lisbon, NY 42o34.666 75o07.603 Goey Pond, Hartwick, NY 42o37.630 74o58.578 Goey Pond, Hartwick, NY 42o37.517 74o58.553 Goey Pond, Hartwick, NY 42o37.308 74o58.442 Arnold Lake, Hartwick, NY 42o36.858 75o00.434 Summit Lake, Springfield, NY 42o51.743 74o50.963 Summit Lake, Springfield, NY 42o51.585 74o50.867 Summit Lake, Springfield, NY 42o51.676 74o50.092 Belvedere Lake 42o44.697 74o45.687 Belvedere Lake 42o44.853 74o45.462 Crumhorn Lake 42o33.632 74o55.139 Crumhorn Lake 42o33.531 74o55.428 Allen Lake 42o49.634 74o56.976 Allen Lake 42o49.773 74o56.896 Allen Lake 42o49.875 74o56.953 Allen Lake 42o49.899 74o57.137 Larchwood Lake 42o 33.157 74o 10.416 Larchwood Lake 42o 33.286 75o10.486 Larchwood Lake 42o 33.334 75o10.516 Larchwood Lake 42o 33.292 75o10.272 Larchwood Lake 42o 33.048 75o10.442 Silver Lake 42o 35.871 75o19.768 Silver Lake 42o 36.090 75o19.842 Silver Lake 42o 35.955 75o19.919 Neahwa Park Pond 42o27.001 75o03.531 Hunt Union Pond 42o28.294 75o03.676 Basswood Pond 42o45.059 75o07.381 Basswood Pond 42o45.248 75o07.345 Basswood Pond 42o45.183 75o07.296 Basswood Pond 42o45.115 75o07.319 Goodyear Lake, RT 35 Boat Launch Access 42o30.299 75o58.546 Goodyear Lake, RT 35 Boat Launch Access 42o30.304 75o58.796 Goodyear Lake, Portlandville Fishing Access 42o31.693 75o58.268 Goodyear Lake, Portlandville Fishing Access 42o31.477 74o58.360

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Wilber Lake 42o30.738 75o03.254 Wilber Lake 42o30.756 75o03.170 Unnamed creek, Lull Hill Rd. 42o31.786 75o06.106 Wharton Creek, Laurens, NY. Off of New Rd 42o31.478 75o07.137 Wharton Creek, Laurens, NY. Off of New Rd 42o31.683 75o07.415 Wharton Creek, Laurens, NY. Off of New Rd 42o31.131 75o06.949 Wharton Creek, Laurens, NY. Off of New Rd 42o30.853 75o06.759 Wharton Creek, Laurens, NY. Off of New Rd 42o30.675 75o06.449 Wharton Creek, Laurens, NY. Off of New Rd 42o32.269 75o08.657 Wharton Creek, Laurens, NY. Off of New Rd 42o32.359 75o08.746 Otego Creek, Thayer Rd 42o44.172 75o03.370 Otego Creek, Bristol Rd 42o42.992 75o02.672 Otego Creek, Wiley Town Rd 42o41.243 75o03.027 Otego Creek, County Highway 11 42o39.570 75o03.207 Otego Creek, Jones Crossing Rd 42o37.699 75o03.283 Otego Creek, County Highway 11D 42o36.996 75o03.454 Otego Creek, Angel Rd 42o35.198 75o03.694 Otego Creek, County Highway 11A 42o31.916 75o05.069

Otego Creek, County Highway 23 DEC Fishing Access Site 42o28.259 75o06.506 Otego Creek, Rt 9 Public Park Access Site 42o26.850 75o06.298 Schenevus Creek, South Hill Rd off of County Highway 39 42o35.309 74o45.023 Schenevus Creek,, Tannery Rd, town of Maryland 42o32.778 74o49.880 Schenevus Creek Tributary, DEC Fishing Access Site off of Rt 7 42o32.668 74o50.518 Schenevus Creek DEC Fishing Access Site off of Rt 7 42o32.647 74o50.524 Schenevus Creek, Steven Rd DEC Fishing Access Site 42o32.454 74o51.171 Schenevus Creek, Loft Rd off of Rt 7 42o32.113 74o53.097 Schenevus creek, Leonard Rd DEC Fishing Access Site 42o31.119 74o54.530 Schenevus Creek, Highway 28 in Milford (prior to Junc. w/ 88) 42o29.166 74o58.120 Oaks Creek, off of Highway 28 under the bridge before Rt 11 42o39.967 74o57.550 Oaks Creek, Lower Toddsville Rd under abandoned bridge 42o40.949 74o57.469 Oaks Creek, Fork Shop Rd under bridge 42o42.117 74o58.530 Oaks Creek, Allison Rd under bridge 42o42.615 74o59.319 Oaks Creek, off of Highway 28 in the town of Oaksville 42o43.465 75o00.138 Oaks Creek, Keating Rd under bridge 42o43.831 75o00.131 Oaks Creek, Keating Rd before Panther Mt. Rd fork in the road 42o45.299 75o00.804 Oaks Creek, Rt 22 under bridge 42o46.845 75o01.051

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Cherry Valley Creek, off of Highway 166; Village of Cherry Valley near Fish and Game Rd 42o47.572 74o45.443 Cherry Valley Creek, Highway 165 under bridge in Roseboom 42o44.423 74o46.381 Cherry Valley Creek, Dubbens Cross Rd off of Highway 166 42o42.245 74o48.810 Cherry Valley Creek, County Highway 35 off of Moore Rd 42o41.390 74o50.539 Cherry Valley Creek, Rt 43 42o37.954 74o52.955 Cherry Valley Creek, Rt 35B 42o35.582 74o55.627 Unadilla River, Chesapeake Dr. off of Rt 7 42o19.452 75º24.588 Unadilla River, County Highway 35 in Chenango County leading to Highway 18 in Otsego County 42º20.459 75º23.923 Unadilla River, Chenango County Highway 40 42º19.453 75 º24.587 Unadilla River, DEC Fishing Access Site off of rt 8 on edge of town of Unadilla towards Butternuts 42º23.615 75º23.873 Unadilla River, Old Rte 8 North off of Rt 8 42º27.656 75º23.891 Unadilla River, St highway 23, South New Berlin, DEC Fishing Access site 42º31.716 75º22.960 Unadilla river, DEC Fishing Access/boat launch off of Rt 8 in Morris 42º33.177 75º21.955 Unadilla river, route 80 in Pittsfield 42º33.176 75º21.955

Unadilla River, County Highway 20 next to Chobani factory 42º41.076 75 º19.156 Unadilla River, off of Rt 8 in Edmeson DEC Fishing Access Site 42º44.707 75º17.699 Unadilla River, Center St off of Rt 8 in Madison/Otsego County 42º48.541 75º14.931 Unadilla River, County Highway 18B, Unadilla Forks 42º50.535 75º14.578 Unadilla River, St Highway 20 in Plainsfield 42º50.536 75º14.577 Canadarago Lake N/A N/A Otsego Lake N/A N/A

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BFS 2013 ANNUAL REPORT CONTENTS

INTRODUCTION: W. N. Harman…………………………………………………………………...….1

ONGOING STUDIES:

OTSEGO LAKE WATERSHED MONITORING: 2013 Otsego Lake water levels. W.N. Harman and M.F. Albright………………………..8 Otsego Lake limnological monitoring, 2013. H.A. Waterfield and M.F. Albright..….…11 A survey of Otsego Lake’s zooplankton community, summer 2013. C. Tanner and M.F. Albright ………….……………………………..…….…..22 Chlorophyll a concentrations in Otsego Lake, summer 2013. T. Bianchine and C. Tanner………………………………….……...... 34 Water quality monitoring of five major tributaries in the Otsego Lake watershed, summer 2013. C. Teter. ..………………………………………….42

SUSQUEHANNA RIVER MONITORING: Monitoring the water quality and fecal coliform bacteria in the upper Susquehanna River, summer 2013. T. Bianchine………………..…….……….59

ARTHROPOD MONITORING: Mosquito data from light traps, Thayer Farm. W.L. Butts…………………………....74

REPORTS:

Aquatic invasive species present in Otsego County, NY water bodies. A. Yoo, K. Herzog and H.A. Waterfield …………………………….……………….75 Zebra mussel (Dreissena polymorpha) monitoring using navigation buoys. A. Yoo, P.H. Lord and W.H. Wong………..…………………………..…….……….95 Zebra mussels and other benthic organisms in Otsego Lake in 2008. J. Vanassche, W.H. Wong, W.N. Harman and M.F. Albright...... …………………103 Trap net monitoring of fish communities within the weedy littoral zone at Rat Cove and rocky littoral zone at Brookwood Point, Otsego Lake. S.G. Stowell.……110 Population assessment of fresh-water mussels (Unionidae) in Otsego Lake since the introduction of zebra mussels (Dreissena polymorpha). D. Caracciolo……….…....115 Gastropods and fish as hosts of digenetic trematodes in Otsego Lake and nearby waters. E. Darpino, R. Russell and F. Reyda……………………………123 Nematodes of fishes of Otsego Lake, New York, including a species that is new to science. A. Borden and F. Reyda……...……… ……………….…….129 An examination of the morphological diversity within a new genus of tapeworm from stingrays (Class: Cestoda). K. Herzog, R. Russell and F. Reyda..……………..135 Cestodes of the fishes of Otsego Lake and nearby waters. A. Sendkewitz, I. Delgado and F. Reyda..…………………………………………...140 Forest bryophyte reproduction and dispersal: An update. A. Lawrence and S. C. Robinson……………………………………...…………….144

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Monitoring the effectiveness of the Cooperstown wastewater treatment wetland, 2013. M.F. Albright…………………….……………………….150 Groundwater flow and geochemistry at Greenwoods Conservancy. M. Moore and L. Hasbargen…………………………………………….……...……161 Utilizing environmental DNA to identify aquatic invasive species. L. Newton.……………186 A biosurvey of Allen’s Lake, Richfield Springs, NY. P. H. Lord ……...……..…….………192 Chronological field observations at various BFS sites, 2013. W.L. Butts..…………………194 Dynamics of Galerucella spp. and purple loosestrife (Lythrum salicaria) in Goodyear Swamp Sanctuary, summer 2013 update. H.A. Waterfield……………....197 Walleye (Sander vitreus) movement and depth utilization in response to changes in alewife (Alosa pseudoharengus) abundance in Otsego Lake J.R. Foster and D.J. Drake……………………………………………..………….....204 Hydroacoustic survey of Otsego Lake’s pelagic fish community, spring 2013.H.A. Waterfield and M. Cornwell………………..…………………….211 Natural recruitment of lake trout (Salvelinus namaycush) in Otsego Lake. N.M. Sawick and J.R. Foster………………………………………………………...219 Monitoring the Moe Pond ecosystem and population estimates of largemouth bass (Micropterus salmoides) post unauthorized introduction. S.G. Stowell…………….226 Otsego Lake, NY ice phenology 1843-2014. H.A. Waterfield………………………………237 Presence of mercury and comparison to other metals in lakes, rivers, and streams in central New York. M. Moore and D. Castendyk……….……………241 The effects of zebra mussels on benthic macroinvertebrates in Otsego Lake. J.M. Vanassche, W.H., W.N. Harman, and M.F. Albright…………………….…….253 Control and eradication of water chestnut (Trapa natans, L.) in an Oneonta wetland, 2013 progress report. H.A. Waterfield and M.F. Albright……………………….….265 Aquatic macrophyte management plan facilitation, Lake Moraine, Madison County, NY 2013.W.N. Harman and M.F. Albright………………………267 Otsego Lake fry sampling, 2013. H.A. Waterfield and M.F. Albright………………………278

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Zebra mussel (Dreissena polymorpha) monitoring using navigation buoys

Annie Yoo1, Paul Lord2, and Wai Hing Wong3

ABSTRACT

In the present study, navigational buoys were used to monitor Dreissena polymorpha (zebra mussel), an aquatic invasive species in Otsego Lake, New York. Zebra mussels from the two navigational buoys were collected at sites in the north and south ends of the lake. Buoys were taken out of the water on 29 December and 31 December 2012 after the growing season. All sampled mussels were frozen until further analyzed; an electronic caliper and compound microscope were used to measure shell lengths. We observed growth, settlement, and density of each colonized buoy and determined that bottom sides of the buoy had the most colonization whereas the top sides had the least. Shell length of mussels from the south site was significantly larger than those from the north site. The mussels from the metal ring were the largest while no difference was found among the rest. Colonization of zebra mussels can be due to specific substrate types and amount of nutrition available in the habitat.

INTRODUCTION

Over the past few decades, at least 36 mollusk species were introduced to Atlantic, Pacific, and Gulf coasts of North America (Johnson and Carlton 1996). Some of these species caused abiotic and biotic changes in inland waterways (MacIssac 1996). One species greatly impacting its surrounding environment is Dreissena polymorpha (zebra mussel) (Pallas, 1771). Although not native to the Eastern United States, zebra mussels were discovered in the Hudson River in 1993 and later detected in Massachusetts in 2008 (Wong et al. 2012). They now clog water pipelines, attach to boats, colonize dam gates, and foul other substrates (Wong et al. 2012). Zebra mussels are found living on rock surfaces, macrophytes, native molluscs, canal and dock walls, and watercraft and motor outdrives; they quickly outcompete the native population and decrease recreational water activities (MacIssac 1996).

Dreissena bugensis (quagga mussel) (Andrusov 1897) also over-filters water in areas such as Lake Erie where phytoplankton populations decreased. Quagga mussel invasion led to alteration in food webs due to changes in fish and zooplankton populations in Lake Huron (Mueting et al. 2010). Zebra and quagga mussels have become serious nonindigenous pests in North America causing environmental harm and associated damage costs (Pimental et al. 2005). Both species are native to Eastern Europe and were first introduced in 1986 into the Great Lakes in North America. They entered the Great Lakes from ballast water dumping by large ocean-going ships from Europe (Hebert et al. 1989; May and Marsden 1992; Mills et al. 1993; Carlton 2008).

Both dreissenid mussels have invaded many lakes and rivers in North America. The spread of dreissenid mussels will presumably continue for many years until their entire potential range is filled (Strayer 2009). Zebra mussels were found in Otsego Lake in 2007 (Horvath 2008; Waterfield 2008). No systematic monitoring program was in place evaluating colonization and growth of these invasive pests in Otsego Lake. Basic biological information is needed for this species such as

1 BFS Intern, summer 2013. Current affiliation: SUNY Oneonta. Funding provided by the Otsego County Land Trust. 2 BFS Researcher & Adjunct Instructor of Environmental Science, SUNY Oneonta. 3 BFS Researcher & Assistant Professor of Biology, SUNY Oneonta.

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growth and settlement rate (Wong et al. 2012). Settlement and growth rates of these invasive zebra mussels in Otsego Lake need investigation.

In New York State, buoys are permitted to be deployed to delineate no-wake zones (New York State Navigation Law 2013). The four municipalities bordering the Otsego Lake passed identical laws extending the NY State 100' no-wake zone out to 200' for Otsego Lake (Otsego Lake Association 2013). Navigation buoys have been used to monitor invasive mussels and other benthic invertebrates in St. Lawrence River, Lake Ontario, and the Welland Canal (Conn and Conn 2007); we therefore hypothesize that these buoys can be a good tool for evaluating zebra mussel settlement and growth in Otsego Lake, New York.

METHODS

Monitoring buoys were made with polyethylene (Taylor Made Products 2013). The two navigation buoys we monitored were deployed on 14 April, 2012 with one at the Lake Front (N 42º42.223 W 74º55.237) at the south end of Otsego Lake and the other one at Springfield Landing (N 42º48.451 W 74º53.022) at the north end of Otsego Lake. The buoys were 2.3 m off the lake bottom. The Springfield Landing and Lake Front buoy were taken out of the water on 29 December and 31 December 2012, respectively. Colonized mussels on these two buoys were used for size analysis and density calculations. All mussels collected from these two buoys are young- of-the-year since the two buoys were mussel free before being deployed. Mussels were taken from seven locations for each navigation buoy (Table 1).

Mussels collected from a specific region of a navigation buoy (Figure 1) were stored in a freezer until analyzed. An electronic caliper (Mitutoyo Absolute digital caliper, 965 Corporate Boulevard Aurora, Illinois 60502) was used to measure mussels with shell lengths larger than 4 mm. Mussels equal or smaller than 4 mm were measured using a compound microscope (Zeiss compound microscope, Carl-Zeiss-Strasse 2273447 Oberkochen, Germany). Each mussel cohort was estimated using the modal progression of Fish Stock Assessment Tool II. FiSAT is the official program used by United Nations’ Fisheries and Aquaculture Department to estimate population dynamics of finfish and shellfish. FiSAT II applies the maximum likelihood concept to separate the normally distributed components of size-frequency samples, allowing accurate demarcation of the component cohorts from the composite polymodal population size of finfish or shellfish (Gayanilo et al. 2005). Densities of mussels (mussels/m2) on different parts of the buoy were also calculated (Wong et al. 2012). All the statistics were performed using SAS (Version 9.2, SAS Institute Inc. Cary, NC).

RESULTS

The no-wake zone buoys were colonized by zebra mussels. The settlement rate of zebra mussels is shown in Table 1, and length summary is shown in Table 2. The buoys and the seven locations used for the analyses are shown in Figure 1. The bottom sides of the buoys had the most colonization whereas the top sides had the least colonization. The computed shell length means of the two identified cohorts were 2.5 mm and 5.3 mm for the Lake Front buoy subpopulation (Figure 2). The means were 2.5 mm and 5.2 mm for the mussels on the Springfield Landing buoy (Figure 2).

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Table 1. Density of zebra mussels in the two Otsego Lake buoys. Location Buoy location Mussels (#/m2) Lake Front Large Side 756 Large Bottom 64,029 Small Bottom 12,590 Metal Ring 5,289 Small Inside Bottom 113,046 Small Side 14,920 Large Top 412

Springfield Large Side 139 Large Bottom 93,676 Small Bottom 5,820 Metal Ring 688 Small Inside Bottom 106,812 Small Side 5,296 Large Top 0

Table 2. Length summary of zebra mussels on the two monitoring buoys in Otsego Lake, from Lake Front and Springfield Landing. Lengths in mm. Standard Buoy Position Mean Min Max deviation n Lake Front Large Side 2.6 1.2 6.0 1.0 114 Large Bottom 2.8 1.0 7.4 1.1 622 Small Bottom 2.6 1.0 6.3 1.0 106 Metal Ring 3.8 1.2 9.8 1.5 292 Small Inside Bottom 3.1 0.2 7.9 1.2 780 Small Side 2.5 0.6 6.9 1.1 431 Large Top 3.0 1.4 5.2 1.2 16

Springfield Large Side 2.8 1.6 3.8 0.7 21 Large Bottom 2.7 1.0 8.2 1.0 910 Small Bottom 2.5 1.4 3.8 0.5 49 Metal Ring 2.1 1.2 5.5 0.8 38 Small Inside Bottom 2.6 0.8 7.4 1.0 737 Small Side 2.6 1.2 7.8 1.0 153 Large Top 0.0 0.0 0.0 0.0 0.0

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Figure 1. Two buoys from which zebra mussels were collected (top panel) and locations from which samples were collected (bottom panel): 1 Large Side; 2 Large Bottom; 3 Small Bottom; 4. Metal Ring; 5. Small Inside Bottom; 6. Small Side; 7 Large Top.

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Figure 2. Frequency of zebra mussels on two monitoring buoys in Otsego Lake (Lakefront= above, Springfield= below).

Shell length of mussels from Lake Front was significantly larger than those in Springfield Landing (Two-way ANOVA; DF = 1; F = 82.11; p < 0.0001; α = 0.05). The mussels from the metal ring were the largest whereas no difference was found among the rest (Two-way ANOVA; DF = 6; F = 39.45; p < 0.0001; α = 0.05).

Overall density means for Lake Front and Springfield Landing buoys were 30,148 and 30,347 mussels/m2, respectively; however, there was no significant difference in mussel density between Lake Front and Springfield Landing (T-test; DF = 12; t = -0.01; p = 0.99; α = 0.05).

DISCUSSION

Our study demonstrates that no-wake zone buoys can be used to monitor colonization and growth of invasive zebra mussels. Karatayev et al. (2006) observed zebra mussel growth increases in water columns on upper surfaces than on the bottom such as buoys, cages, or floating objects. Substantial differences in growth were displayed between lakes and reservoirs – mussels grow faster in reservoirs than lakes (Table 3; Karatayev et al. 2006). Larger zebra mussels were found on the Lake Front buoy than the Springfield Landing buoy. One possible reason for this is that it is siltier in Springfield Landing than Lake Front, and zebra mussels do not grow well in an environment with higher sediment content (Schneider et al. 1998). After one growing season, the average size of mussels attached to the Lake Front and Springfield Landing buoy were 2.9 mm and 2.6 mm with maximum shell length of 9.8 mm and 8.2 mm, respectively.

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Settled mussels on the bottom of both buoys were more abundant than those settling on top. This is in agreement with other studies demonstrating that mussels do not like to be exposed directly to light (Marsden and Lansky 2000). Mueting et al. (2010) found substrate preference is based more on the depth of the surface in the water column than the texture or composition of the substrate.

This study and other works documenting zebra mussel settlement and growth rates can help lake managers understand the biology of this invasive species in the Eastern United States (Wong et al. 2012). Growth rate of dreissenid mussels depend on water temperature, season of the year, location in the water column, trophic conditions, and water velocity (Karatayev et al. 2006). Further studies should be implemented to research different substrates where dreissenid colonies occur. Diet and nutritional values may also be another factor in settlement and growth rate; mussels are herbivores with phytoplankton as their primary diet (Wong et al. 2012). Wong et al. (2012) hypothesized that natural populations in deeper areas grow slowly due to lower productivity of phytoplankton. In Otsego Lake, future research may help raise awareness on zebra mussel impacts on biotic resources (e.g., fisheries, benthos, and planktonic community), infrastructure (e.g., water quality and water-delivery facilities) and recreational values (unfavorable odors from decaying mussels and boat/propeller contamination).

ACKNOWLEDGEMENTS

The authors gratefully acknowledge and thank the Biological Field Station, Volunteer Diver Team, and their tenders in Cooperstown, New York.

REFERENCES

Carlton, J.T. 2008. The zebra mussel Dreissena polymorpha found in North America in 1986 and 1987. Journal of Great Lakes Research 34: 770-773.

Conn, D.B. and D.A. Conn. 2007. Navigational buoy survey of invasive and native benthic invertebrates of the St. Lawrence River, Lake Ontario, and the Welland Canal. Abstracts of the 15th International Conference on Aquatic Invasive Species, Nijmegen, the Netherlands.

Gayanilo, F.C., P. Sparre and D. Pauly. 2005. FAO-ICLARM stock assessment tools II: user’s guide.

Hebert, P.D.N., B.W. Muncaster and G.L. Mackie. 1989. Ecological and genetic studies on Dreissena polymorpha (Pallas) - a new mollusk in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 46: 1587-1591. Horvath, T. 2008. Economically viable strategy for prevention of invasive species introduction: Case study of Otsego Lake, New York. Aquatic Invasions 3(1): 3-9.

Johnson, L.E. and J.T. Carlton. 1996. Post-establishment spread in large scale invasions:

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dispersal mechanisms of the zebra mussel Dreissena polymorpha. Ecology 77(6): 1686- 1690.

Karatayev, A.Y., L.E. Burlakova and D.K. Padilla. 2006. Growth rate and longevity of Dreissena polymorpha (Pallas): a review and recommendations for future study. Journal of Shellfish Research 25(1): 23-32.

MacIssac, H.J. 1996. Potential abiotic and biotic impacts of zebra mussels on the inland waters of North America. American Zoology 36: 287-299.

Marsden, J.E. and D.M. Lansky. 2000. Substrate selection by settling zebra mussels, Dreissena polymorpha, relative to material, texture, orientation, and sunlight. Can J Zool 78(5): 787- 793.

May, B. and J.E. Marsden. 1992. Genetic identification and implications of another invasive species of dreissenid mussel in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 49: 1501-1506.

Mills, E.L., R.M. Dermott, E.F. Roseman, D. Dustin, E. Mellina, D.B. Conn and A.P. Spidle. 1993. Colonization, ecology, and population structure of the quagga mussel (Bivalvia, Dreissenidae) in the lower Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 50: 2305-2314.

Mueting, S.A., S.L. Gerstenberger and W.H. Wong. 2010. An evaluation of artificial substrates for monitoring the quagga mussel (Dreissena bugensis) in Lake Mead, Nevada-Arizona. Lake and Reservoir Management 26: 283-292.

New York State Navigation Law (2013) Section 45. http://www.nyss.com/NYS.html. Date viewed: 31 July 2013.

Otsego Lake Association (2013) Otsego Lake Regulations. http://otsegolakeassociation.org/toc.htm. Date viewed: 14 Aug 2013.

Pimental, D., R. Zuniga and D. Morrison. 2005. Update on the environmental and economic costs associated with alien-invasive species in the United States. Ecological Economics 52: 273- 288.

Schneider, D.W., S.P. Madon, J.A. Stoeckel and R.E. Sparks.1998. Seston quality controls zebra mussel (Dreissena polymorpha & hair sp.) energetics in turbid rivers. Oecologia 117: 331- 341.

Strayer, D.L. 2009. Twenty years of zebra mussels: lessons from the mollusk that made headlines. Frontiers in Ecology and the Environment 7: 135-141.

Taylor Made Products. 2013. 2013 Product Catalog: Sur-Mark™ Marker Buoys. www.taylormadeproducts.com/catalog/. Date viewed: 15 Aug 2013.

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Waterfield, H. 2008. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake 2008. In 41st Annual Repor. SUNY at Oneonta Bio. Fld. Sta., SUNY Oneonta.

Wong, W.H., S. Gerstenberger, W. Baldwin and B. Moore. 2012. Settlement and growth of quagga mussels (Dreissena rostriformis bugensis Andrusov, 1897) in Lake Mead, Nevada- Arizona, USA. Aquatic Invasions 7(1): 7-19.

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Zebra mussels and other benthic organisms in Otsego Lake in 2008

Jennifer Vanassche1, Wai Hing Wong2, Willard N. Harman2, and Matthew F. Albright2

ABSTRACT

The zebra mussel, Dreissena polymorpha (Pallas 1771), was first documented in North America in the 1980s. Zebra mussels were documented in Otsego Lake in 2007 and were considered abundant by 2010. In 2008, benthic samples were collected at 50 sites on Otsego Lake and species composition was described in order to provide baseline data prior to the community being influenced by zebra mussel establishment. Sampling efforts undertaken in 2013 will evaluate the extent to which zebra mussel colonization has subsequently influenced the benthic community.

INTRODUCTION

Dreissenid mussels, including the zebra mussel (Dreissena polymorpha Pallas, 1771), originating from the Ponto-Caspian area (Black, Asov, and Caspian Sea), and the quagga mussel (Dreissena rostriformis bugensis Andrusov, 1897), originating from the mouths of the Rivers Southern Bug and Dnieper, are both native to Eastern Europe (Van der Velde, Rajagopal, bij de Vaate 2010). Both zebra and quagga mussels were introduced into the Laurentian Great Lakes in North America in the 1980s, most through ballast water exchanges (Ludyanskiy et al. 1993; Carlton 2008; Van der Velde, Rajagopal, bij de Vaate 2010). Zebra mussels were first reported in North America in 1988 in Lake St. Clair (Hebert et al. 1989). However, a recent paper (Carlton 2008) provides convincing evidence that they were first established as early as 1986 in Lake Erie. The first occurrence of the quagga mussel in North America was documented in 1989 in Lake Erie (Mills et al. 1993), but it was first identified as a separate species and given the common name “quagga” in a paper by May and Marsden (1992).This species was later identified from morphological and genetic material as “Dreissena bugensis” (Spidle et al. 1994, Rosenburg and Ludyanskiy 1994). However, a recent genetic comparison between D. bugensis and D. rostriformis indicated no distinct differences between the two taxa (Therriault et al. 2004).

Otsego Lake in Otsego County, New York is an oligo-mesotrophic dimictic lake formed by glacial over-deepening of the Susquehanna River Valley (Harman 1997). Zebra mussels were first documented in Otsego Lake in 2007 (Waterfield 2009) and were considered abundant by 2010 (McShane and Mehigan 2012). Zebra mussel colonization may favor some macroinvertebrate species while negatively impacting others; i.e., they affect the environment directly by altering the substrate they colonize and indirectly by increasing water clarity (Ward and Ricciardi 2010). Colonies of zebra mussels increase the complexity of substrates and provide a more diverse habitat and more refugia, and they can increase food availability by releasing feces and pseudofeces (Ozersky et al. 2011).

1 BFS Intern, summer 2013. Present affiliation: SUNY Oneonta. Funding provided by the Village of Cooperstown. 2 SUNY Oneonta Biological Field Station.

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The ability of zebra mussels to easily colonize hard substrates is especially detrimental to native mussels such as those in the family Unionidae (freshwater pearly mussels). Unionids, which have been in decline since the mid-1800s, compete with zebra mussels for food and space (Cope et al. 2003). Zebra mussels often colonize the shells of unionid mussels, preventing the native mussels from eliminating waste and feeding; this contributes to the further decline of native mussels (Cope et al. 2003). Several other species are negatively impacted by zebra mussels, including animals in the family Sphaeriidae (fingernail clams) and amphipods in the genus Diporeia (Nalepa et al. 2009; Ward and Ricciardi 2010). Declines in populations of some macroinvertebrates may contribute to declines in populations of fish that have inflexible diets (Owens and Dittman 2003). Because zebra mussels are a low-energy alternative to the animals they displace, fish species that become dependent on zebra mussels for food are smaller and weigh less than fish of the same species that eat other macroinvertebrates (Owens and Dittman 2003). Dreissenid mussels can have a positive effect on other macroinvertebrates, especially when their distribution is still patchy, by increasing the complexity of the substrate, providing more cover for macroinvertebrates (Ozersky et al. 2011; Nalepa et al. 2009; Ward and Ricciardi 2010). This more complex substrate is beneficial to invertebrates in that it offers more opportunities for protection from predators (Ward and Ricciardi 2010). Feces and pseudofeces released by zebra mussels may also benefit macroinvertebrates that can use them as a food source (Hecky et al. 2004). Therefore, the introduction of the zebra mussel into Otsego Lake will likely result in changes of the benthic community. Benthic samples were taken in 2008 to provide baseline data for future surveys to document how zebra mussels affected species richness in benthic macroinvertebrates.

METHODS

Benthic samples were collected from Otsego Lake via the research vessel Anadontiodes between 10 July 2008 and 21 July 2008. An Ekman dredge was used to collect samples from the benthic zone along three transects (Figure 1) at depths of 0m to 50m; a total of 50 samples were collected. Samples were preserved in jars in 70% ethanol. In June and July 2013, rose bengal was added to each jar and macroinvertebrates were separated from the substrate and sorted. If less than 50% of a single organism was present, the organism was discarded. Macroinvertebrates were identified to subclass, order, or family. Morphological differences within these broader taxonomic groups were used to estimate more specific taxonomic richness, which was then calculated for each site (Harman 1997).

Nematodes were not identified further than phylum. Annelids and Platyhelminthes were identified to subclass excluding one unknown annelid. All other taxa were identified to family excluding two unknown arthropods.

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Figure 1. Transects and sites that were sampled in Otsego Lake, New York during July 2008 (from Iannuzzi 1990). Transects TR-2, TR-4, and TR-6 were sampled.

RESULTS

Table 1 provides a summary of taxonomic richness across each of the three transects of Otsego Lake and Table 2 lists the taxa collected. Macroinvertebrates were found in the phyla Annelida, Arthropoda, Mollusca, Nematoda, Platyhelminthes, and Porifera. Overall, 16 families of insects, 6 families of mollusks, 2 families of amphipods, and one family each of isopods, sponges, and springtails were found. A total of 34 taxa were identified. (Table 2).

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Of the 50 samples surveyed, a total of eight zebra mussels were collected among five sites at depths of 2m, 4m, and 5m. At site TR2E, 2 mussels were found at a depth of 2m and 3 mussels were found at 4m. At TR2W, 1 mussel was found at 5m. At sites TR4E and TR6W, one mussel was found at each site at a depth of 2m.

Table 1. Taxonomic richness at each site sampled in Otsego Lake. The total richness is the total taxonomic richness for all transects combined.

Transect ID Depth (m) Richness TR2 0.5-20 25 TR4 0-40 18 TR6 0.5-15 14

Unlabeled 1

DISCUSSION

The 2008 macrobenthic community data provide baseline information that will be used to assess the impacts of Dreissena polymorpha on the benthic community in Otsego Lake. At the time of sampling in 2008, lake-wide zebra mussel abundance was low and their influence on the benthic community would have been minimal. Changes in benthic macroinvertebrates in Otsego Lake have been seen as a result of other invasive species, such as Eurasian milfoil, Myriophyllum spicatum L. (Harman 1997). With the introduction of zebra mussels, the benthic community may have drastically changed since 2008. Changes in the environment of the lake may benefit species that feed from the feces and pseudofeces deposited by the mussels (Atalah et al. 2010; Hecky et al. 2004). Abundance and family richness of macroinvertebrates have been found to increase with the presence of zebra mussels; increases in surface area, complexity, and heterogeneity of the substrate provide more refuges for macroinvertebrates to hide from predators and more habitat to live in (Horvath et al.1999; Ozersky et al. 2011). Zebra mussels, while beneficial to some macroinvertebrates, are detrimental to other populations of macroinvertebrates. Declines in unionid mussel populations may have occurred more rapidly from the colonization of zebra mussels. Future surveys will help determine how D. polymorpha has influenced the benthic community of Otsego Lake.

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Table 2. Otsego Lake macrobenthos identified in the 2008 samples.

Phylum Class Subclass Order Suborder Family Genus Porifera Demospongiae Haplosclerida Spongillidae Platyhelminthes Turbellaria Nematoda Mollusca Bivalva Veneroida Dreisseni dae Sphaeriidae Gastropoda Basommatophora Physidae Planorbidae Heterostropha Valvatidae Neotaenioglossa Hydrobiidae Annelida Clitellata Hirudinia Oligochaeta Unknown Unknown Arthropoda Arachnida Trombidiform es Branchiopoda Diplostraca Cladocera Entognatha Collembola Isotomidae Insecta Coleoptera Unknown Diptera Ceratopogonidae Chironomidae Tipulidae Unknown Ephemeroptera Caenidae Ephemeridae Heptageniidae Hexagenia sp. MacCaffertium sp. Leptophlebiidae Stenacron sp. Megaloptera Sialidae Leptophlebia sp. Odonata Coenagrionidae Sialis sp. Corduliidae Trichoptera Leptoceridae Polycentropodidae Malacostraca Amphipoda Gammaridae Talitridae Isopoda Asellidae Maxillopoda Copepoda

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REFERENCES

Atalah J., M Kelly-Quinn, K. Irvine and T.P. Crowe. 2010. Impacts of invasion by Dreissena polymorpha (Pallas, 1771) on the performance of macroinvertebrate assessment tools for eutrophication pressure in lakes. Hydrobiologia 654: 237-251.

Carlton, J. 2008. The zebra mussel (Dreissena polymorpha) found in North America in 1986 and 1987. Journal of Great Lakes Research 34: 770-773.

Cope, W.G., T.J. Newton and C.M. Gatenby. 2003. Review of techniques to prevent introduction of zebra mussels (Dreissena polymorpha) during native mussel (Unionoidea) conservation activities. Journal of Shellfish Research 22(1): 177-184.

Harman, W.N. 1997. Changes in the Otsego Lake macrobenthos communities between 1935 and 1994. Lake and Reservoir Management 13: 160-169.

Hebert, P.D.N., B.W. Muncaster and G.L. Mackie. 1989. Ecological and genetic studies on Dreissena polymorpha (Pallas) - a new mollusk in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 46: 1587-1591.

Hecky, R.E., R.E.H. Smith, D.R. Barton, S.J. Guildford, W.D. Taylor, M.N. Charlton and T. Howell. 2004. The nearshore phosphorus shunt: a consequence of ecosystem engineering by dreissenids in the Laurentian Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 61: 1285–1293.

Horvath, T.G, K.M. Martin and G.A. Lamberti. 1999. Effect of zebra mussels, Dreissena polymorpha, on macroinvertebrates in a lake-outlet stream. The American Midland Naturalist Journal 142: 340-347.

Iannuzzi, T.J. 1991. A model plan for the Otsego Lake watershed. Phase II: The chemical limnology and water quality of Otsego Lake. Occasional Paper #23, Rept. No. 2a. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Ludyanskiy, M.L., D. McDonald and D. Macneill. 1993. Impact of the zebra mussel, a bivalve invader – Dreissena polymorpha is rapidly colonizing hard surfaces throughout waterways of the United-States and Canada. Bioscience 43: 533-544.

May, B. and J.E. Marsden. 1992. Genetic identification and implications of another invasive species of dreissenid mussel in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 49: 1501-1506.

McShane, D. and K. Mehigan. 2013. 2012 aquatic macrophyte survey of Otsego Lake. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Mills, E.L., R.M. Dermott, E.F. Roseman, D. Dustin, E. Mellina, D.B. Conn and A.P. Spidle. 1993. Colonization, ecology, and population structure of the quagga mussel (Bivalvia, Dreissenidae) in the lower Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 50: 2305-2314.

Nalepa, T.F., S.A. Pothoven and D.L. Fanslow. 2009. Recent changes in benthic macroinvertebrate populations in Lake Huron and impact on the diet of lake whitefish (Coregonus clupeaformis). Aquatic Ecosystem Health & Management 12(1): 2-10.

Owens, R.W. and D.E. Dittman. 2003. Shifts in the diets of slimy sculpin (Cottus cognatus) and lake whitefish (Coregonus clupeaformis) in Lake Ontario following the collapse of the burrowing amphipod Diporeia. Aquatic Ecosystem Health & Management 6(3): 311-323.

Ozersky, T., D.R. Barton and D.O. Evans. 2011. Fourteen years of dreissenid presence in the rocky littoral zone of a large lake: effects on macroinvertebrate abundance and diversity. Journal of the North American Benthological Society 30(4): 913-922.

Rosenberg G. and M.L. Ludyanskiy. 1994. A nomenclatural review of Dreissna (Bivalve, Dreissenidae), with identification of the quagga mussel as Dreissena bugensis. Canadian Journal of Fisheries and Aquatic Sciences 51: 1474-1484.

Spidle, A.P., J.E. Marsden and B. May. 1994. Identification of the Great Lakes quagga mussel as Dreissena bugensis from the Dnieper River, Ukraine, on the basis of allozyme variation. Canadian Journal of Fisheries and Aquatic Sciences 51: 1485-1489.

Therriault, T.W., M.F. Docker, M.I. Orlova,D.D. Heath and H.J., MacIsaaca. 2004. Molecular resolution of the family Dreissenidae (Mollusca : Bivalvia) with emphasis on Ponto- Caspian species, including first report of Mytilopsis leucophaeata in the Black Sea basin. Molecular Phylogenetics and Evolution 30: 479-489.

Van der Velde, G., S. Rajagopal and A. Bij de Vaate. 2010. The zebra mussel in Europe. Backhuys Publishers, Leiden, The Netherlands, 490 pp.

Ward, J.M. and A. Ricciardi. 2010. Community-level effects of co-occurring native and exotic ecosystem engineers. Freshwater Biology 55: 1803-1817.

Waterfield, H.A. 2009. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Trap net monitoring of fish communities within the weedy littoral zone at Rat Cove and rocky littoral zone at Brookwood Point, Otsego Lake

Stephen G. Stowell1

INTRODUCTION

This study was a continuation of yearly monitoring of the littoral fishes of Otsego Lake. The long term goal of the study is to assess the littoral fish community and determine population dynamics of species utilizing littoral habitats. Rat Cove has been studied since 1979 (MacWatters 1980) and Brookwood Point since 2002 (Wayman 2003). Littoral habitats of sizeable lakes such as Otsego Lake are essential, providing spawning and nursery habitats for many species of fish. The illegal introduction of alewife (Alosa pseudoharengus) in 1986 (Foster 1990) altered the trophic balance and physical/chemical characteristics of Otsego Lake, due to the species’ opportunistic behavior and over-effective grazing of the lake’s zooplanktonic community (Harman 2002). Alewives are efficient, opportunistic, epilimnetic planktivores that feed on microcrustaceans, insects, ichthyoplankton, zooplankton and their own eggs (Cornwell 2005). Long term monitoring of littoral fishes helps to assess the effect alewives have on the lake’s fish communities, as well as alewife abundance and spawning activity. Additionally this study provides useful long term data on non-alewife species. In order to mitigate the detrimental effects alewife have imposed on the lake’s ecosystem, predatory walleye (Sander vitreus) have been re-established through stocking, which began in 2000. During summer stratification, alewife utilize only the top layer of water (the epilimnion), resulting in a spatial separation between them and the cold water predators of Otsego Lake. Walleye, however, have been known to forage in the epilimnion, so during summer stratification alewife would be ideal prey. This study continues to document littoral fish communities that could provide insight into changes occurring in Otsego Lake.

1 Robert C. MacWatters Internship in the Aquatic Sciences, summer 2013. Present affiliation: Department of Fisheries and Wildlife Technology, SUNY Agriculture and Technical College, Cobleskill, NY.

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METHODS & MATERIALS Winged Indiana trap nets with a single throat were set out Monday through Friday and checked daily, at both Rat Cove and Brookwood Point (Figure 1) from 4 June to 25 July. At Rat Cove the trap was deployed perpendicular to the north shore and at Brookwood Point the trap was deployed due east from the middle of the point. The catch was transferred from the nets into totes, all metrics were taken on site and the fish were promptly returned to the water. Each fish was identified, weighed in grams, and measured (mm).

Figure 1. Bathymetric contour map of Otsego Lake, NY. Trap nets were set perpendicular to the shore at Brookwood Point and Rat Cove.

RESULTS & DISCUSSION The overall mean catch per week at both Rat Cove and Brookwood Point increased between 2005 and 2011, dropping off somewhat in 2013 (data are not available for 2012) (Tables 1 and 2). There are several factors that could be the cause of this abrupt change. The first factor that likely influenced the 2013 number was the utilization of new nets for the 2011 season (German 2011). Although the nets were the same general design used in previous years (Indiana style traps), the new nets have wings which appear to improve catch rate. Additionally, the older nets used earlier were in regular need of repair. Even with extensive repairs to the old nets, it is evident that new functional equipment improved catch rates.

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Table 1. Mean weekly catch at Rat Cove and catch contributed by each species, 2000-2013 (modified from German 2011).

Species 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2013 Alewife 120 68 8 45 2 <1 0 3 1 <1 <1 <1 0 Golden Shiner <1 <1 <1 <1 <1 <1 0 0 <1 <1 <1 <1 1.25 Pumpkinseed 10 21 15 33 13 5 2 2 4 5 5 16 8.6 Blue Gill 2 3 4 2 2 1 <1 3 6 7 5 7 1.25 Redbreast Sunfish <1 <1 <1 <1 <1 <1 0 0 <1 <1 0 <1 0 Rock Bass 2 2 4 <1 2 <1 <1 <1 <1 <1 1 2 <1 Largemouth Bass <1 <1 <1 <1 <1 <1 0 <1 <1 <1 <1 <1 <1 Chain Pickerel <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 Atlantic Salmon 0 <1 0 <1 0 0 0 0 0 0 0 0 0 Yellow Perch 3 <1 1 <1 1 <1 <1 <1 <1 0 <1 4 3.38 White Sucker 1 <1 1 <1 2 <1 <1 0 0 0 <1 <1 <1 Common Carp <1 <1 <1 <1 <1 <1 <1 0 0 0 0 <1 <1 Brown Bullhead 2 <1 6 3 2 <1 0 0 0 <1 <1 <1 <1 Spottail Shiner 0 0 <1 0 0 0 0 <1 0 0 0 0 0 Smallmouth Bass 0 0 <1 0 0 0 0 0 0 0 0 0 0 Emerald Shiner 0 0 0 0 <1 0 0 0 0 0 0 <1 <1 European Rudd <1 0 <1 <1 <1 0 <1 0 <1 <1 1 <1 0 Common Shiner 0 0 0 0 0 0 0 0 0 0 0 <1 0 Walleye 0 0 0 0 0 0 0 0 0 0 0 <1 0 Total 141 96 41 87 25 9 5 11 14 15 14 35 18

Table 2. Mean weekly catch at Brookwood Point and catch contributed by each species, 2000- 2013 (modified from German 2011)

Species 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2013 Alewife 224 137 77 95 13 6 1 5 <1 <1 1 0 0 Golden Shiner <1 <1 1 2 2 <1 <1 0 0 0 <1 <1 <1 Pumpkinseed 3 7 12 13 12 1 <1 <1 2 2 1 10 5.13 Blue Gill 7 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 15 <1 Redbreast Sunfish <1 0 <1 <1 <1 <1 <1 <1 0 <1 <1 4 0 Rock Bass 8 4 4 4 3 1 <1 <1 <1 2 2 14 3 Largemouth Bass <1 <1 <1 <1 0 <1 0 0 <1 <1 0 <1 0 Chain Pickerel <1 0 <1 <1 <1 <1 0 <1 <1 0 0 <1 <1 Atlantic Salmon 0 <1 0 0 0 <1 0 0 0 0 <1 0 0 Yellow Perch 2 <1 <1 0 <1 <1 <1 0 <1 <1 0 1 1.63 Walleye 0 0 0 <1 0 0 0 0 0 <1 0 <1 <1 White Sucker 5 0 2 <1 <1 <1 <1 0 0 0 <1 <1 <1 Common Carp 2 <1 <1 <1 <1 0 <1 0 0 0 0 0 <1 Bluntnose Minnow <1 0 0 0 0 <1 0 0 0 0 0 0 0 Brown Bullhead 7 0 <1 4 4 0 <1 0 0 0 0 <1 <1 Spottail Shiner 0 <1 0 0 0 0 0 <1 0 0 <1 3 0 Smallmouth Bass 0 0 0 <1 <1 0 0 0 <1 0 <1 <1 0 European Rudd 0 <1 0 <1 <1 0 <1 0 <1 0 0 0 0 Common Shiner 0 0 0 0 0 <1 0 0 0 0 0 0 0 Emerald Shiner 0 0 0 0 0 0 0 0 0 0 0 <1 0 Lake Trout 0 0 0 0 0 0 0 0 0 0 0 <1 0 Total 259 152 101 121 37 10 4 8 4 5 6 50 12

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A total of 30 fish were caught per week at Rat Cove and Brookwood Point, which was much lower then the 2011 study. Not one Alewife was caught in the 2013 study (Figure 2). The decline of alewife is shown in Figure 2.

250 Rat Cove Brookwood Point 200

150

100

Alewives per set 50

0 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2013

Year Figure 2. Mean weekly alewife catch in Rat Cove and Brookwood Point trap nets 2000-2013. (Modified from German 2011).

DISCUSSION

A total of 249 fish were caught between Rat Cove and Brookwood Point over the 2013 sampling season. A total of 161 fish were caught in a weedy littoral habitat represented by Rat Cove, which was significantly lower than 2011. Brookwood Point is a rocky shoal that caught a total of 88 fish with the dominate fishes being pumpkinseed and yellow perch. Again, this was a decrease from the 2011 season when over 800 fish were captured in total.

CONCLUSION Otsego Lake has seen an increase in clarity, potentially due to two separate factors, first being the introduction and establishment of zebra mussels (Dreissena polymorpha) first documented in 2007 (Harman 2008). Zebra mussels have been documented to cause ecological changes, including increased water clarity, following a successful introduction to a water body (D’Itri 1996). In 2009 (Gillespie 2010) and 2010 (Albright and Leonardo 2011), cladoceran zooplankton mean size and Daphnia sp. abundance had increased, correlating with increased water clarity. (Transparencies through 2011 were even greater (Waterfield and Albright 2012)). This change in the plankton community is likely due to reduced grazing by alewife. Reduced competition would allow for increased length of the (fewer) remaining alewife (German 2011). Alewife declined steadily following the initiation of the walleye stocking effort and they appear to have been virtually eliminated by the mid-2000s (Figure 2). This is corroborated by hydroacoustics surveys by Waterfield and Cornwell (2013).

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REFERENCES Albright, M.F. and M. Leonardo. 2011. A survey of Otsego Lake’s zooplankton, summer 2010. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F. and H.A. Waterfield. 2012. Otsego Lake water quality monitoring, 2011. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Bowers, B. 2011. Summer 2010 trap net monitoring of littoral fish communities at Rat Cove & Brookwood Point, Otsego Lake. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Cornwell, M.D. 2005. Re-introduction of walleye to Otsego Lake: Re-establishing a fishery and subsequent influences of a top predator. Occas. pap. #40, SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

D'Itri, Frank. 1996. Zebra Mussels and Aquatic Nuisances Species. Chelsea, NY: Ann Arbor Press. 161-163. Print.

Foster, J.R. 1990. Introduction of alewife (Alosa pseudoharengus) in Otsego Lake. In 22nd Ann. Rept. (1989). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Gillespie, S. 2010. A survey of Otsego Lake’s zooplankton community, summer 2009. In 42nd Ann. Rept. (2009). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

German, B. 2011. Summer 2011 trap net monitoring of fish communities utilizing the weedy littoral zone at Rat Cove and rocky littoral zone Brookwood Point, Otsego Lake. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W.N, M.F. Albright and D.M. Warner. 2007. Trophic changes in Otsego Lake, NY, following the introduction of alewife (Alosa psuedoharengus). Lake and Reserv. Manage. 18(3):215-226.

Harman, W.N. 2008. Introduction. In 40th Ann. Rept. (2007). SUNY Oneonta Biol. Fld., SUNY Oneonta.

MacWaters, R. C. 1980. The fishes of Otsego Lake. Occas. Paper #7. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2013. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Wayman, K. 2003. Rat Cove and Brookwood Point littoral fish survey, 2002. In 35th Ann. Rept. (2002). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Population assessment of fresh-water mussels (Unionidae) in Otsego Lake since the introduction of zebra mussels (Dreissena polymorpha)

Deanna Caracciolo1

INTRODUCTION

Fresh-water mussels are native to Lake Otsego, which is located north of Cooperstown, New York. These mussels are suspension feeders, removing particulates from the water column. They also act as bio-indicators due to sensitivity to pollution, invasive species and siltation (Harman 2014). They move very little throughout their lifespan and are therefore extremely susceptible to being buried under sediment deposition. The six species previously known to be present in the lake are Alasmidonta undulata, Anodontoides ferussacianus, Elliptio complanata, Lampsilis radiata, Pyganodon cataracta, and Strophitus undulatus. Surveys were conducted in 1969 by Harman (1970) as well as in 2000 by Ferrara (2001). Since Ferrara’s survey, the invasive zebra mussel (Dreissena polymorpha) was introduced into Lake Otsego and is thought to have had an established population as early as 2006 (Waterfield 2009). Dreissena polymorpha is known to colonize the posterior end of native mussels and starve it of nutrients. It was hypothesized that since the introduction of zebra mussels, the pearly mussel community in Otsego Lake has dramatically decreased. Since total unionid biomass and diversity was also in decline between 1969 and 2000, it was believed that siltation was the cause (Ferrara 2001). In 2013, historical survey sites were revisited to evaluate populations of freshwater mussel in the lake.

METHODS

Past studies involving Otsego Lake unionids used both free diving as well as SCUBA diving to complete surveys. High levels of rainfall during the summer of 2013 resulted in poor visibility. This led to the use of SCUBA as the main method of surveying, increasing focus and time beneath the water. Harman was the first to survey 28 sites around the lake, finding all 6 aforementioned species (Harman 1970). Ferrara then re-surveyed the same sites in 2000 and found 20 sites that yielded live unionids, but comprising only 3 of the original 6 species (Ferarra 2001). The 2013 study surveyed 31 sites, including the 20 historical sites that had live unionids in 2000. Additional sites were selected based upon bottom composition and proximity to historical sites surveyed to gain more extensive data.

A group of divers (1-3) began the survey in June 2013 using time interval transects to sample. This consists of divers descending to approximately one to four meters and using a compass to orient a straight transect underwater. Divers then followed this direction for exactly ten minutes, surveying substrate for any trace of unionids, live or dead. Time was extended if the site was promising for live valves. Empty valves were collected and brought to the surface for identification and kept for further analysis. Live specimens were identified underwater if possible. Any unidentifiable live valves were brought to the surface, identified and immediately

1 BFS W.N. Harman Internship, summer 2013. Current affiliation: SUNY Oneonta.

- 117 - 46th Annual Report of the Biological Field Station returned and placed back in the sediment, posterior end uncovered. Catch Per Unit Effort (CPUE) was also recorded (the number of clams collected per hour). Depth, GPS coordinates and substrate information was collected at each site (displayed in Table 1 and plotted in Figures 1 and 2).

Empty valves brought back to the lab were processed as follows. Zebra mussels from each unionid were removed and counted. Only zebra mussels that were attached to a live or dead unionid were considered. The level of sediment accumulation on the shell was also noted due to the possibility that increased siltation can cause the death of a mussel (Table 2). The declining population of native bivalves in the lake is a cause for concern since they are the lakes natural bio-indicators and provide an indication of water quality and overall ecosystem health.

Table 1. Coordinates of 2013 Otsego Lake collection sites.

Site Location Coordinates 4 East Woody Cove N42 44.088 W74 53.918 5 North Sunken Island N42 47.844 W74 53.588 6 Central East Sunken Island N42 47.804 W74 53.489 7/8 S/W Sunken Island N42 47.589 W74 53.602 9 South 5 Mile Point N42 45.834 W74 53.993 10 North 5 Mile Point N42 45.877 W74 53.962 11 S/E Sunken Island N42 47.647 W74 53.467 12 Steamboat Ln Cove N42 48.047 W74 53.973 13 White Creek N42 48.427 W74 53.728 14 Cripple Creek N42 48.733 W74 53.620 15 Hayden Creek N42 48.746 W74 53.312 16 Clarke Point N42 47.445 W74 52.770 17 Hyde Bay Cove N42 47.491 W74 52.356 18 Clarke Point N42 47.461 W74 52.801 19 Clarke Point N42 47.461 W74 52.822 20 Brookwood Point N42 43.564 W74 54.996 21 Rat Cove N42 43.174 W74 55.462 22 Susquehanna Outlet N42 46.696 W74 52.774 23 Peggs Point N42 42.093 W74 55.107 24 Harmans House N42 47.320 W74 53.836 25 6 Mile Point N42 47.037 W74 53.832 26 5 Mile Point North N42 46.074 W74 53.994 27 South of 3 Mile Point N42 44.366 W74 54.611 28 Hayden Creek West N42 48.751 W74 53.448 29 Eel Island Point, East Bank N42 47.931 W74 53.024 30 South of Eel Island N42 47.799 W74 52.935 31 Shadowbrook Outlet N42 47.100 W74 52.161 32 3 Mile Point, East Bank N42 44.762 W74 53.604 33 Point Judith North N42 43.831 W74 54.171 34 South of Ratcove N42 42.774 W74 55.400

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Figure 1. Enlarged view of Sunken Island collection sites, 2013.

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015

014 029 013 012

005 019 006 016 007 018 008 017 011

024

031 025

022 026

010

009

004

027 032

020 033

021

034

023

Figure 2. Unionid collection sites, Otsego Lake, 2013.

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Table 2. Siltation level descriptions used in Table 3 (from Box and Mossa 1999).

Average Siltation Description

Shell color and texture are easily seen, virtually no visible silt, sand or Low mud, Valve is not filled with silt Majority of shell is clean of sediment, most of shell markings, color and Medium texture are viisable, some silt accumulation inside empty valve Could have effect on health of unionid. Color and texture of shell are comlpeatly covered, Majority of valve High buried when found, Empty valve filled with sediment Could cause severe health effects/ likely kill unionid

RESULTS

Overall, the CPUE of the 2000 survey was 1.28 unionids/hour (Ferarra 2001). In the 2013 survey, the CPUE was 0.34 unionids/hour. This dramatic decrease in CPUE indicates that unionids populations are dwindling and soon may not be present in the lake. This hypothesis is also supported by the 2013 survey findings. No living or empty Elliptio complata, A. undulata, A. jerussiacianus, and S. undulates were found. Also L. radiata was the most commonly found empty valve followed by P. cataracta. Live and empty (dead) unionids are listed in Table 2 along with average zebra mussel coverage per site.

A total of 518 unionids were found in the survey, with only two found alive. Both live specimens were L. radiata, the first found on North Sunken Island, the second on the west slope of Eel Island. The first live specimen found had 123 zebra mussels attached to its posterior end, and looked to be in poor condition. The second live mussel found appeared better, only having three zebra mussels attached, one of which was dead. The healthier unionid was burrowing into the sediment, possibly to attempt to remove the zebra mussels. This is also believed to be a new behavior first noticed in Oaks Creek by Lord in 2012 (Lord 2013).

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Table 2. Collected data from 2013 Unionid survey. Only zebra mussels attached to live/dead unionids were considered. (Avg) Average; (#ZM) zebra mussels found on unionid shell. See Table 1 for explanation of “Avg siltation on shells”.

Location Living Unionids Dead Unionids Site Date Depth Bottom Composition # Found #ZM # Found Avg ZM Avg siltation on shells 5 6/12/13 1.5m Gravel and Silt 1 L. radiata 123 59 L. radiata 27 High 1 P. cataracta 6 6/13/13 2m Silt/Gravel/Rock 0 21 L. radiata 65 High 7,8 6/13/13 1m Gravel and Cobble 0 2 L. radiata 0 Low 9 6/13/13 Organic Matter, gravel 0 15 L. radiata 10 6/13/13 2m Cobble 0 4 L. radiata 3 Medium 11 7/1/13 1.5m Silt and Rock 0 12 L. radiata 2 Low 12 7/11/13 1.5m Silt 0 0 0 13 7/11/13 >1m Solid Rock 0 0 0 14 7/11/13 >1m Silt 0 0 0 15 7/17/13 >1m Silt 0 0 0 16 7/17/13 3.5m Gravel,Sand,Cobble 0 35 L. radiata 11 low 17 7/17/13 3m Silt 0 0 0 18 7/17/13 3m Gravel,Sand,Cobble 0 27 L.radiata 13 Medium 1 P. cataracta 19 7/17/13 4.5m Gravel, Sand, Cobble 0 38 L. radiata 7 Low 20 7/12/13 >1m Silt 0 0 21 7/12/13 >1m Silt 0 0 22 7/21/13 2m Sand, Gravel 0 37 L. radiata 11 Medium 1 P. cataracta 23 7/21/13 3.5m Gravel, Cobble 0 26 L. radiata 11 Low 1 P. cataracta 24 7/29/13 3.5m Sand, Gravel 0 54 L. radiata 2 Medium 1 P. cataracta 25 7/19/13 3.5m Gravel 0 11 L. radiata 3.5 Medium 1 P. cataracta 26 7/29/13 3m Silt 0 18 L. radiata 16 High 1 P. cataracta 27 7/29/13 2.5m Gravel, Sand 0 47 L. radiata 4 Medium 28 7/30/13 1m Silt 0 0 29 7/30/13 2.5m Cobble, Gravel 1 L. radiata 3 55 L radiata 4 Low 1 P. cataracta 30 7/30/13 3m Gravel 0 38 L. radiata 7 Low 31 7/30/13 1m Silt 0 0 32 7/30/13 3-7m Steep slope, Cobble 0 16 L. radiata 11 Medium 33 7/30/13 2.5m Cobble, Sand 0 23 L. radiata 3 Medium 34 7/30/13 2.5m Silt, Sand 0 11 L. radiata 40 Medium

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DISCUSSION

Data collected during the 2013 survey clearly show a continued decline of the unionid community in Otsego Lake. The 1969 survey indicated that dense populations of all 6 species were present in the lake (Harman 1969). Thirty one years later, the community was re-surveyed and 4 of the 6 species were no longer present (Ferrara 2001); forty-four years after the initial survey, only two individual representing one species of the original 6 were found alive in the lake. Possible reasons for this decline in total unionid mass are introduction of zebra mussels into the lake in 2007, as well as the increase in sediment runoff accumulating in the lake during heavy rain occurrences.

The 2000 survey found live specimens bordering macrophyte beds (Ferarra 2001); this was not seen in 2013. Many of the macrophyte beds noted in 2000 were visibly covered with a thick layer of silt and most unionids surrounding these beds were empty. Water with high silt suspension is known to interfere with unionid feeding and can also smother them (Ferrara 2001). Under a controlled environment it was found that some species are more tolerant to siltation than others (Downing et al. 2000). Lampsilis radiata, E. complanata, and P. cataracta were found to have a higher survival rate when placed randomly in a tank containing patches of silt and sand. This is consistent with 2013 findings; only L. radiata was found alive and P. cataracta was the only other valve found.

The summer of 2000 average Secchi disk reading was 2.7 m. Between 2009 and 2013, a period following the establishment of zebra mussels, annual mean Secchi transparencies have been between 5 and 7 m (Waterfield and Albright 2014). Zebra mussels, like pearly mussels, are suspension feeders. As the population of zebra mussels increased, the amount of suspended particles decreases. Much of the dive surveying in 2000 needed the use of flashlights to aid in searching below 3m. Divers in 2013 needed no flashlight for visual aid but did encounter poor visibility after storm events.

Current living unionids seem to be moving from sand substrate to gravel substrate. One live L. radiata found at site 29 seemed to be burrowing into the sediment. Lord (2012) observed similar behavior in Oaks Creek in 2012 in some L. radiata found just downstream of Canadarago Lake. Zebra mussels were first documented in Canadarago in 2002 (Horvath and Lord 2003), six years before being documented in Otsego lake (Waterfield 2009).

Overall, a majority of the historical survey sites surveyed by Harman have since been silted over. With the constant threat of sediment loading into the lake from local roadside ditches and construction projects, as well as the new threat of zebra mussels, the future of native pearly mussels seems bleak. Without immediate mediation of these problems it is possible that unionids will be extirpated from the lake within the next decade.

REFERENCES

Box, J.B. and J. Mossa. 1999. Sediment, land use and mussels. J. of North Amer. Benth. Soc. Vol. 18 No. 1.

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Ferrara, L. 2001. Population survey of the fresh-water mussels (Unionidae) of Otsego Lake. In 33rd Ann. Rept. (2000) SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Harman, W.N. 2014. Personal communication. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W. N. 1970. Biological studies -Otsego Lake. In 3rd Ann. Rept. SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Harman, W. N. 1971. Mollusks of Otsego Lake. Nautilus. Vol 85. pp.71.

Harman, W. N., L. P. Sohaki, M. F Albright, and D. L.Rosen, 1997. The state of Otsego Lake 1936-1996. pp.226-229.

Horvath, T.G. and P.H. Lord. 2003. First report of zebra mussels (Dreissena polymorpha) in Canadarago Lake. In 35th Ann. Rept. (2002). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Lord, P.H. 2012. Personal communication. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. Strayer, D. L. and K. L. Jirka, 1997. The pearly mussels of New York State. New York State Education Department. Fort Orange Press Incorporated. pp.I-I13; plates 1-27.

Waterfield, H. 2009. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake 2008. In 41st Ann. Rept. (2008) SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Weir, G. P. 1977. An ecology of the Unionidae in Otsego Lake with special references to immature stages. Occas. Pap. No.4. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Gastropods and fish as hosts of digenetic trematodes in Otsego Lake and nearby waters

Erica Darpino¹, Rebecca Russell1, and Florian Reyda2

INTRODUCTION This study of digenetic trematodes is part of a survey of the intestinal parasites of fishes of Otsego Lake and its tributaries (Cooperstown, New York) from 2008 to 2014. The survey included a total of 27 fish species, consisting of six centrarchid species, one ictalurid species, eleven cyprinid species, three percid species, three salmonid species, one catostomid species, one clupeid species, and one esocid species. Among the fish examined, seven fish species were infected with adult digenetic trematode species in the alimentary canal, whereas 18 fish species were infected with metacercaria in other organs. Another component of this survey included molluscs as an intermediate host of the digenetic trematodes. The survey included six species of snails, three of which were infected with larval trematodes. Many digenetic trematodes have a similar life cycle in that their first intermediate host is a mollusc—usually a snail--, the second intermediate host is often a fish and, in some trematodes, the final host is a bird. An example of this is Clinostomum marginatum whose definitive host is a great blue heron and intermediate hosts are a species of snail (Helisoma spp.) and a multitude of fish species including largemouth bass, Micropterus salmoides (Hazen and Esch 1977). Some digenes will fully develop as adults in the fish digestive system and will not move on to a bird host, like Azygia angusticauda which utilizes a snail, Amnicola limosa as an intermediate host and fish such as yellow perch, Perca flavescens, as a definitive host (Sillman 1962). The goals of this study were to identify the adult digenetic trematodes in the area and, using their known life cycles, determine what trematodes are present as larva within the gastropod hosts in the area. METHODS In total, 400+ individual fish were collected by hook and line, seine, gill net, or ElectroFisher from Otsego Lake and surrounding water bodies and subsequently examined for intestinal parasites, and in many cases, for parasites in other fish organs. Fish were collected and placed in aquaria at the Biological Field Station until they were anesthetized and examined. Dorsal incisions exposed the internal organs, which were then inspected for parasites. In total, 367 snails were collected. The snails were most prevalent in shallow waters on submerged vegetation, logs or rocks. Snails were collected by hand after prolonged visual observation and placed in a bucket with water to be taken to the lab. Upwards of 100 snails could be collected within ten minutes if sampling in a good area. When back in the lab, snails were identified using a key from Harman and Berg (1971). The different species were separated

¹ SUNY Oneonta undergraduate student, Biology Department, SUNY Oneonta, SUNY Oneonta Research Foundation 2Assistant Professor of Invertebrate Zoology and Researcher, Biology Department and Biological Field Station, SUNY Oneonta

- 125 - 46th Annual Report of the Biological Field Station into dishes of lake water. Each snail was dissected, using forceps and probe to separate the visceral mass from the shell, and checked for parasites. Adult and larval stages of trematodes were processed the same way. Some were preserved in 4% neutral-buffered Formalin and later switched to 70% ethanol to be made into slides to be viewed with light microscopy. Others were preserved the same way initially but were later processed with conventional methods (e.g. Reyda & Caira 2006) in preparation for scanning electron microscopy. In addition one of each of the specimens that we thought to be a different species was saved in 100% ethanol and sent to colleagues in Canada for sequencing of the mitochondrial oxidase I gene.

RESULTS

Adult trematodes can be more easily identified than the other stages because they possess more diagnostic structures, especially reproductive structures. Azygia angusticauda was found in four host fish species: Perca flavescens, Ambloplites rupestris, Lepomis macrochirus, and Lepomis gibbosus. Azygia longa was found in Esox niger. Cryptogonimus chili was found in two fish species: A. rupestris and Micropterus dolomieu. Crepidostomum cornutum was found in six fish species: Micropterus salmoides, Lepomis gibbous, L. auritus, L. macrochirus, P. flavescens, and Notemigonus crysoleucas. Crepidostomum metoecus was found in Salvelinus fontinalis. Bunoderina sacculata was found in P. flavescens. Allocreadium lobatum was found in two fish species: Semotilus atromaculatus and Pimephales promelas. Metacercarial stages are harder to identify than are adult stages because they are sexually immature but given the DNA sequencing work of colleagues in Canada we now have some identifications. Posthodiplostomum sp. were found in seven fish species: Ambloplites rupestris, M. salmoides, M. dolomieu, L. macrochirus, S. atromaculatus, Pimephales notatus and Rhinichthus cataractae. Ornithodiplostomum sp. were found in three fish species: Luxilus cornutus, Catostomus commersonii and Notropis stramineus. Apharyngostrigea cornu were found in C. commersonii. Apatemon sp. were found in Cottus bairdii. Two different larval stages have been observed in the snails: redia and sporocysts. The infected snails contained at least one of these stages. The sporocyst, redia and cercarial stages in the snails are even harder to identify than metacercaria. We are still awaiting molecular identifications and are currently working on other techniques to help us identify the ones we have collected. We have collected six different species of snails and all of them had different rates of infection (see Table 1). Table 1. Snail species and number of snail specimens examined and infected with the sporocyst or redia trematode larval stages. Infections included either or both stages.

Physella Lymnaea Gyraulus Amnicola Physella Helisoma heterostropha humilis parvus limosa integra trivolvis Number of 147 159 7 2 39 13 snails Number 26 4 0 0 23 3 infected

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DISCUSSION Life cycle information, including specific information on many of the larval stages of these trematode species are known and published (Schell 1985). Using these data we can infer what sporocysts or redia are likely to be present in the different snail species that we have been examining. Clinostomum marginatum uses Helisoma spp. as an intermediate host so it is possible that we have some of these larval stages present. Ornithodiplostomum sp. uses physid snails as an intermediate host, so it is likely that some of the sporocysts we acquired from these snails represent species of that genus in our collection. Posthodiplostomum sp. and Apatemon sp. use physid or lymnaeid snails as an intermediate host so it is possible that we have some of these as well. Species of Bunoderina and Allocreadium, and Crepidostomum cornutum all use some sort of freshwater clam as their first intermediate host. We can therefore assume that these larval stages are present in the lake, but they are likely not present in our collection since we did not sample bivalves in this study. We were able to identify all of the seven trematode species that reach sexual maturity in the digestive system of fishes. Because life cycles are known for each of these seven species (see Schell 1985), we were able to infer what hosts, in addition to fish, are present in these water bodies. For example, the life cycle of Bunoderina sacculata (Figure 1) includes two hosts, Pisidium noveboracense, a type of fingernail clam for the intermediate host and Perca flavescens, as the definitive host. Allocreadium lobatum (Figure 2) and Crepidostomum cornutum (Figure 3) also use fingernail clams. Crepidostomum metoecus (Figure 4) uses Lymnaea spp., as intermediate hosts and based on their previously documented life cycles, we should therefore find sporocyst and redia stages in the molluscs. Azygia angusticauda (Figure 5) and Azygia longa (Figure 6) utilize prosobranch snails as their intermediate host and use several fish species as their definitive hosts. The seventh adult digene, Cryptogonimus chili (Figure 7), utilizes Lymnaea humilis as the intermediate host and one of two possible fish species as the definitive hosts. We currently have not encountered fingernail clams, but due to the presence of the adult trematodes, we can assume that these molluscs are present in the Otsego watershed. Other studies (see Caracciolo, 2014) have addressed the decline of bivalves in Otsego Lake. Our work provides parasitological evidence that these bivalves are present in the Otsego watershed, at least in numbers adequate to sustain the continued life cycles of these trematodes. Clinostomum marginatum (Figure 8) uses Helisoma spp. as an intermediate host so it is possible that we have some of the larval stages from our Helisoma trivolvis samples. Clinostomum marginatum is widely occurring in North America and is and is known among fisherman for reducing the appeal of fish for consumption. Ornithodiplostomum sp. uses physid snails as an intermediate host so we may have some of these redia and sporocysts in our collection. We may also have Posthodiplostomum sp. and Apatemon sp. in our collection. Posthodiplostomum sp. and Apatemon sp. use physid or lymnaeid snails as a fish intermediate host, and a fish and frog species, respectively, as second intermediate host, and herons, such as great blue herons and black-capped night herons, as their definitive hosts. There are some limitations when working with the larval stages. The technique for identification requires live cercarial or larval stages and all of the specimens already collected are dead and preserved, therefore additional sampling is needed. Lignin pink dye can be used to stain the live specimens and highlight the flame cells, which can be used to identify the cercaria at least to family. Given this, we must collect more snails and observe the larval stages while they are alive and use this new technique to help identify them. This work will be continued the

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summer of 2014, and hopefully will help us in our ultimate goal to identify all the larval stages of these seven trematode species from their mollusc hosts.

ACKNOWLEDGMENTS We would like to thank the students of Dr. Reyda’s Lab, the numerous summer interns from SUNY Oneonta, Steve Stowell, (SUNY Cobleskill) and Tim Pokorny (NYS DEC), for their help in fish and snail collections. We thank Stephen Curran (University of Mississippi) for species identification and lignin pink dye protocols. Anindo Choudhury (St. Norbert College) assisted with species identifications. Sean Locke (International Barcode of Life Project) obtained DNA sequences for some metacercaria and adult trematodes. Snail identifications were confirmed by W. Harman. Funding for this research was provided in party by a Student Research grant to E. D. from the SUNY Oneonta Research Foundation, and in part by an NSF FSML grant to W. Harman (DBI 1034744). Also, special thanks to Dr. Bill Harman, Matt Albright, and Holly Waterfield of the Biological Field Station, and Dr. Janine Caira of the University of Connecticut. SUNY Biological Field Station provided the necessary equipment and work space to carry out this project. This work was done as an independent study and BFS internship from Spring 2013 through Spring 2014.

REFERENCES Caracciolo, D. 2014. Population assessment of fresh-water mussels (Unionidae) in Otsego Lake since the introduction of zebra mussels (Dreissena polymorpha). In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W. and Berg, C. 1971. The Freshwater Snails of Central New York. Vol.1 No. 4., Ithaca, NY: Cornell University.

Hazen, T. and Esch, G. 1978. Observation on the ecology of Clinostomum marginatum in largemouth bass (Micropterus salmoides). J. Fish Biol. 12, 411-420.

Reyda, F., and Caira, J. N. 2006. Five new species of Acanthobothrium (Cestoda: Tetraphyllidae) from Himantura uarnacoides (Myliobatiformes: Dasyatidae) in Malaysian Borneo. Comparative Parasitology 73: 49-71.

Schell, S. C. 1985. Trematodes of North America. Moscow, ID: University Press of Idaho.

Sillman, E. I. 1962. The life history of Ayzgia longa (Leidy 1851) (Trematoda: Digenea), and notes on A. acuminata Goldberger 1911. Transactions of the American Microscopical Society Vol. 81, No. 1 (Jan., 1962), pp. 43-65.

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Nematodes of fishes of Otsego Lake, New York, including a species that is new to science

Austin Borden1 and Florian Reyda2

INTRODUCTION

This study of freshwater nematodes is part of a survey of the intestinal parasites of fishes of Otsego Lake and its tributaries (Cooperstown, New York) from 2008 to 2014. The survey included a total of 27 fish species, consisting of six centrarchid species, one ictalurid species, eleven cyprinid species, three percid species, three salmonid species, one catostomid species, one clupeid species, and one esocid species. Five species of nematodes were encountered in the alimentary canal of fishes examined. These include Spinitectus gracilis, Spinitectus carolini, Spinitectus micracanthus, Spinitectus n. sp., and Dichelyne cotylophora. Additionally, Eustrongylides tubifex was found outside of the alimentary canal.

The goal of the study was to identify all nematode species encountered in fishes in Otsego Lake. This study emphasized specimens that were previously collected with a variety of methods, but also included the acquisition of additional specimens. This work is part of a larger survey that was conducted on the entire parasite population of Otsego Lake (Reyda 2010; Szmygiel and Reyda 2011).

METHODS

Collection and dissection. Over 400 fish samples were collected from various locations on Otsego Lake and its tributaries. Fish were collected through gill netting, angling, and electrofishing, with the latter used as the main method used in tributaries of Otsego Lake. Fish were anesthetized using Tricaine-S. All of the fish were measured and photographed prior to dissection. A mid-ventral cut the length of the fish was made to expose the body cavity. The stomach and intestines were examined under a dissecting microscope at a magnification of 20 times their actual size for all of the specimens. Parasites encountered were fixed in either hot 10% neutral-buffered formalin, followed by storage in 70% ethanol, or in 100% ethanol for molecular analysis. Specimens initially stored in 70% ethanol were subsequently transferred to a mixture of 5% Glycerin in 70% ethanol solution. Nematodes were cleared in glycerin (see Choudhury and Pérez-Ponce de León 2001). Some samples of nematodes that were stored in 70% ethanol were subsequently prepared for scanning electron microscopy (SEM) following conventional methods (e.g., Reyda and Caira 2006).

Identification of Nematodes: Two major references, Parasites of North American Freshwater Fishes (Hoffman 1999), and Nematodes of Freshwater Fishes of the Neotropical Region (Moravec 1998), were used to identify the nematodes to genus. After the genera were identified, an exhaustive literature search was done for species information. Several original description papers of North American nematodes of the genera previously mentioned were used in identification of the species.

¹ SUNY Oneonta undergraduate student, Biology Department, SUNY Oneonta Research Foundation 2Assistant Professor of Invertebrate Zoology and Researcher, Biology Department and Biological Field Station, SUNY Oneonta

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RESULTS

Four intestinal Spinitectus species were identified. Of these, three had previously been encountered during the survey: Spinitectus carolini Holl, 1928, Spinitectus gracilis Ward and Magath, 1916, and Spinitectus micracanthus Christian, 1972. Spinitectus carolini (Figure 1) samples were found in Micropterus dolomieu (Smallmouth bass), and Ambloplites rupestris (Rock bass). Spinitectus gracilis (Figure 2) was found in three different fish hosts, A. rupestris, M. dolomieu, and Lepomis auritus (Redbreast sunfish). Spinitectus micracanthus (Figure 3) samples were obtained from Lepomis macrochirus (Bluegill), Lepomis gibbosus (Pumpkinseed), L. auritus, A. rupestris, M. dolomieu, and Perca flavescens (Yellow perch). A fourth species of Spinitectus (Figure 4) that is new to science was observed in L. auritus, and L. gibbosus. One intestinal Dichelyne species, Dichelyne cotylophora (Figure 5), was found in P. flavescens and in L. macrochirus. Eustrongylides tubifex Fastzkie and Crites, 1977 (Figure 6), was found in the body cavity of the following hosts: Esox niger (chain pickerel), P. flavescens, L. auritus, L. gibbosus, and L. macrochirus. Scanning electron microscope or light microscopes images of these nematodes (See figures 1–6) show key morphological features of each species. These unique features were previously described (Ward and Magath 1916, Holl 1928, Keppner 1975, Moravec et al. 2011), though not consistently illustrated with SEM prior to this study. Comparing data collected in this study to the previous description of the Spinitectus species, the data collected from this study suggest that the nematodes from Otsego Lake have a notably smaller total body length and a greater variation in this measurement.

DISCUSSION

Although five nematode species have been found in Otsego Lake prior to this study (Reyda 2010; Szmygiel and Reyda 2011), they were not previously identified to species. Among the nematodes encountered, we found the intestinal Spinitectus species to be the most intriguing, and we will continue to study this genus in future work. Spinitectus species have a unique morphology characterized by their spiny cuticle (Ward and Magath 1916). Spinitectus is a diverse genus, however, in Otsego Lake only 3 species were encountered during previous survey work (Reyda 2010). This study has confirmed there are actually four species, one of which is new to science.

The three previously known species of Spinitectus found in Otsego Lake were differentiated by comparing data from our specimens with data from past descriptions and SEM images provided in other studies. For instance, S. gracilis is known to have a spiny cuticle that can contain 30-50 spines per row, and S. carolini has 20-30 spines per row (Jilek and Crites 1982). Spinitectus gracilis has a shorter stoma than S. carolini, which has a stoma that extends posterior to the first circlet of spines (Mueller and Van Cleave 1932). Another defining characteristic is the orientation of the vagina in females and the size of the spicule in males. Spinitectus micracanthus is easily distinguishable due to the smaller anterior region of the worm, as well as the abundant cuticular spines, which range 50-100 spines per row (Keppner 1975). In this study, most of the measurements taken for each species matched those provided in the original descriptions, however, in a few cases there were measurements that were missing, or

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did not fit within the ranges provided in previous studies. Out of the three previously known species of Spinitectus, S. micracanthus is in need of a re-description because of the lack of information in the original description, such as number of sectors, any number of spines per sector, both of which are distinct characteristics used in recent species Spinitectus descriptions.

Scanning electron microscope images of the fourth Spinitectus species (Figure 4) demonstrated an external morphology similar to that of Spinitectus macrospinosus, a species that also possesses relatively large cuticular spines, but that was only reported from catfish, in southern Manitoba, Canada and the Midwestern USA (Choudhury and Perryman 2003). This fourth species can easily be distinguished from S. macrospinosus on the basis of its internal morphology. The fourth species of Spinitectus is new to science and a description of this enigmatic species has been started.

Another nematode species found in fishes in the lake was Eustrongylides tubifex, which are not found in the intestine, but in a cyst-like capsule in the body cavity. This nematode is commonly known as “red worm” and is known among fisherman for reducing the appeal of fish for consumption. The definitive hosts of this nematode are piscivorous birds such as kingfishers (Measures 1988).

A species of a third genus, Dichelyne, had been encountered in the lake in a previous study (Reyda 2010); however, it was not identified as Dichelyne cotylophora until this study. This worm was initially identified (Reyda 2010) as a species of the genus Cucullanus due to their similar morphology. This nematode was encountered throughout the study in P. flavescens and L. macrochirus, and with the help of SEM images and numerous slide specimens, it was identified to species on the basis of distinctive characteristic such as number of postanal papillae (Moravec et al. 2011).

In addition to E. tubifex, other nematodes found outside of the intestine, i.e., as juveniles, await identification to species. This will be challenging, because juvenile nematodes are immature and have fewer structures that can be used to distinguish them, and because many of them are too large to mount on slides–the usual approach for the study of fish parasites. Further research, including DNA sequencing, is needed to identify these nematodes. In addition to identification, an updated study of the morphology through re-description of selected species may be beneficial to the further understanding of these nematodes.

ACKNOWLEDGMENTS

I would like to thank Dr. Janine Caira from the University of Connecticut for the use of her lab as well the use of the University of Connecticut electron microscopy facility. Also, thanks to Dr. Anindo Choudhury for his collaboration and help with identifications. Tim Pokorny (NYS DEC) provided many fish for this study. Thanks to Dr. Bill Harman, Matt Albright, and Holly Waterfield for the use of the Biological Field Station and logistical help. Also, I would like to thank Erica Darpino, Illari Delgado, Elsie Dendrick, Nathan Heller, Kaylee Herzog, Annie Murphy, Rebecca Russell, Amanda Sendkewitz and Joseph Westenberger for their help and support during this study. Funds for this project were provided in part by a student

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research grant to A. B. from the SUNY Research Foundation, and in part by an NSF FSML grant to W. Harman (DBI 1034744).

REFERENCES

Christian FA. 1972. Spinitectus micracanthus sp. n. (Nematoda: Rhabdochonidae) from the Bluegill, Lepomis macrochirus Rafinesque, in Ohio. Helminthol Soc Wash, Proc. 39:51- 54.

Choudhury A, Pérez-Ponce de león G. 2001. Spinitectus osorioi n. sp. (Nematode: Cystidicolidae) From Chirostoma spp. (Osteichthyes: Atherinidae) In Lake Pátzcuaro, Michoacán, México. J Parasitol. 87:648-655.

Choudhury A, Perryman BJ. 2003. Spinitectus macrospinosus n. sp. (Nematoda: Cystidicolidae) From The Channel Catfish Ictalurus punctatus in Southern Manitoba and distribution in other Icalurus Spp. J Parasitol. 89:782- 791.

Fastzkie J, Crites JL. 1977. A Redescription of Eustrongylides tubifex (Nitzsch 1819) Jägerskiöld 1909 (Neamtoda: Dictophymatidae) from Mallards (Anas platyrhynchos). J Parasitol. 63:707-712.

Hoffman, GL. 1999. Parasites of North American Freshwater Fishes. Cornell University. Second Edition. 322 pp.

Holl FJ. 1928. Two New Nematode parasites. J Elisha Mitchell Sci Soc. 43:184-187.

Jilek R, Crites JL. 1982. Comparative morphology of the North American Species of Spinitectus (Nematoda: Spriurida) Analyzed by Scanning Electron Microscopy. Trans Am Microsc Soc. 101:126-134.

Keppner EJ. 1975. Life Cycle of Spinitectus micracanthus Christian 1972 (Nematoda: Rhabdochonidae) from the Bluegill, Lepomis macrochirus Rafinesque, in Missouri with a Note on Spinitectus gracilis Ward and Magath, 1917. Am Midl Nat. 93:411-423.

Measures LN. 1988. The Development of Eustrongylides tubifex (Nematoda: Dictophymatoidea) in Oligochaetes. J Parasitol. 74:294-304.

Moravec F. 1998. Nematodes of freshwater fishes of the Neotropical Region. Academia. 464 pp.

Moravec F, Levron, C, Buron I. 2011. Morphology and taxonomic Status of Two Little-Kown Nematode Species Parasitizing North American Fishes. J Parasitol. 97:297-304.

Mueller JF, Van Cleave HJ. 1932. Parasites of Oneida Lake fishes. II. Descriptions of new species and some general considerations, especially concerning the trematode family Heterophyidae. Roosevelt Wildlife Bull 3:79-137.

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Reyda FB. 2010. Parasitic worms of fishes of Otsego Lake and nearby water bodies, 2009. 42nd Annual Report of the SUNY Oneonta Biological Field Station. 42:276–281.

Reyda FB, Caira JN. 2006. Five new species of Acanthobothrium (Cestoda: Tetraphyllidea) from Himantura uarnacoides (Myliobatiformes: Dasyatidae) in Malaysian Borneo. Comp Parasitol. 73:49-71.

Szmygiel C, Reyda FB. 2011. A survey of the parasites of Smallmouth bass (Micropterus dolomieu). 43rd Annual Report of the SUNY Oneonta Biological Field Station. 43:235- 240.

Ward HB, Magath TB. 1916. Notes on Some Nematodes from Fresh-Water Fishes. J Parasitol. 3:57-64.

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An examination of the morphological diversity within a new genus of tapeworm from stingrays (Class: Cestoda)

Kaylee Herzog1, Rebecca Russell2 & Florian Reyda3

INTRODUCTION

Despite their unfortunate reputation, parasites are vital members of any biological community. Nearly every free-living species on the planet is known to play host to one, if not several, parasite species (Price 1980). In part because of this ubiquity, parasites can be used to facilitate investigations ranging from elucidating community food web patterns to acting as indicators of ecosystem health (Marcogliese 2005). In a recent international effort to expand our understanding of marine biodiversity, researchers have been working cooperatively with the goal of identifying new species of elasmobranchs (i.e., sharks, skates and stingrays) and their parasites. The majority of these elasmobranch parasites have been discovered to be tapeworms (Eyring et al. 2012).

Tapeworms, or cestodes, make up one of the more common parasite subgroups, with more than 5,000 described species, over 1,400 of which are exclusively marine (Caira & Reyda 2005). As new elasmobranchs are identified, it is becoming evident that their associated tapeworm populations are highly diverse and mostly consist of undescribed species (Reyda & Caira 2006). These tapeworm specimens are so radically different from previously described species that entirely new taxonomic classifications are needed in order to describe them (e.g., Healy et al. 2009). These new tapeworm specimens must be correctly identified and formally described before trends such as host specificity can be examined (Caira & Jensen 2001). Formal species descriptions are thus a stepping-stone towards the ultimate goal of understanding ecological interactions within this parasite-host community.

This study highlights a portion of the larger, ongoing international effort and assesses a newly-established group of tapeworm species, i.e., a genus, temporarily referred to as Rhinebothriinae New Genus 3 (as referred to by Healy et al. 2009). Our goal is to produce a comprehensive organization of the morphological diversity that exists within this new genus using both traditional light microscopy and scanning electron microscopy (SEM). Assessment of diversity will include examination of the bothridial and proglottid characteristics of multiple individuals within the genus, as combinations of these features have in the past aided in recognizing several potential morpho-types or species. In this report, we provide observations of the scolex, a prominent feature of the tapeworm anatomy. The scolex (see Figure 1) is the structure a tapeworm uses to attach to the intestinal wall of its vertebrate host. In the tapeworms studied here, the scolex consists of four cup-like organs called bothridia (Figure 1) that are structurally modified for attachment to the intestinal walls of stingrays.

1 SUNY Oneonta Biology Department Intern, summer 2013. Present affiliation: SUNY Oneonta. 2 Student Research Assistant (funded by NSF). Present affiliation: SUNY Oneonta. 3 Assistant Professor of Invertebrate Zoology, Biology Department and Biological Field Station, SUNY Oneonta.

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Figure 1. An image of scolex features in Rhinebothriinae New Genus 3 (line drawing by Danielle Willsey).

MATERIALS AND METHODS

Elasmobranch specimens were collected by collaborators from the University of Connecticut and elsewhere, and a subset of the collected tapeworm specimens were sent to the Reyda lab at the main lab of the Biological Field Station for morphological analyses. Specimens were preserved in 70% ethanol (following fixation in the field with 4% neutral-buffering formalin) and organized according to host stingray species. Examined specimens also included permanently-mounted slides on loan from the Royal Museum of Ontario (voucher numbers available upon request).

Specimens to be studied with light microscopy were permanently mounted on glass slides in Canada balsam using conventional methods (see Reyda & Olson 2003). Slides were examined using a Leica DM2500 compound light microscope (Leica Microsystems, Wetzlar, Germany) at 100x, 200x, 400x, 630x and 1,000x magnifications. Associated Leica microscope software was used for obtaining and recording specific measurements. In addition, a representative bothridium of the scolex (Figure 1) of each potential species was drawn with the aid of a drawing tube on the Leica microscope.

Scolex features (Figure 1) that were examined included length and width of the bothridia;

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whether the bothridia was constantly constricted; whether the anterior loculus did or did not overhang the remainder of the bothridium; number of complete transverse septa in the anterior region; presence or absence of incomplete transverse septa (septa that do not connect completely from one edge of the bothridium to the other) in the anterior region; presence or absence of complete longitudinal septa in the posterior region; and presence or absence of lateral posterior subloculi (Figure 1). Proglottid features that were examined included the presence or absence of an external seminal vesicle; a thick- or thin-walled cirrus sac (the male tapeworm organ); overall cirrus sac size in relation to the proglottid (segment of the tapeworm body); presence or absence of a seminal receptacle; and a vagina that did or did not recurve anteriorly to the cirrus sac.

A subset of the preserved specimens was also prepared for SEM examination using conventional methods (see Healy et al. 2009). The images obtained from SEM were then analyzed for the same bothridial characteristics as were light microscope slides. After analysis, specimens were separated into preliminary categories based on combinations of their morphological features and each grouping was considered in the context of host locality and host specificity using collection data from the Global Cestode Database (Caira & Jensen 2013).

RESULTS

Following analysis, eight categories were erected to organize the diversity within Rhinebothriinae New Genus 3, with each grouping representing a potential species or morpho- type. Groupings were organized into a dichotomous key (Figure 2) in which bothridial features were used to distinguish each group and illustrate the diversity within the genus.

DISCUSSION

It is clear that a high degree of morphological diversity exists within Rhinebothriinae New Genus 3, a conclusion that aligns with previous taxonomic assessments of these marine tapeworms (Healy et al. 2009). Figure 2 illustrates this diversity in terms of the ~eight morphological species that were encountered. This demonstrates that several distinct species exist within this genus that remain undescribed at this time. Preliminary organizations of tapeworm genera, such as the one produced from this study, can provide support to future species description efforts. They also have the potential to serve as useful tools in examinations of coevolution and host specificity (e.g. Caira & Jensen 2001), both of which are vital components to understanding trophic relationships within this marine system. Concrete species descriptions are necessary to obtain a true understanding of parasite-host relationships within this community. Hence, for the next step in this process, we will formally describe one of these species from within this diverse genus.

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Figure 2. A Dichotomous key representation of the morphological diversity that exists within Rhinebothriidea New Genus 3 (Class: Cestoda) using line drawings of individual bothridia and SEM images of scoleces to represent individual species or morpho-types.

CONCLUSION

This study demonstrates that a great amount of morphological species diversity awaits discovery in the world. The eight new species illustrated here are just one small component of the biological diversity represented by tapeworms and other parasites. Efforts must continue to document this biodiversity.

ACKNOWLEDGEMENTS

Our many thanks to Dr. Janine Caira (University of Connecticut, Storrs, Connecticut), Dr. Kirsten Jensen (University of Kansas, Lawrence, Kansas), and their colleagues for conducting the fieldwork and providing the specimens for this study. Also thanks to previous Biological Field Station Intern and SUNY Oneonta alumna Danielle Willsey. This study was partially supported with funds from NSF BS&I grants (DEB 0103640, 0118882, 0542846, 0542941), an NSF PEET grant (0818696), and an NSF PBI collaborative grant (0818823), as

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well as by an NSF FSML Grant to W.N. Harman (DBI 1034744), and also by two SUNY Oneonta Research Foundation grants (Student Research Grants awarded to D. W., and to K. H. and R. R.).

REFERENCES

Caira, J.N. & K. Jensen. 2001. An investigation of the co-evolutionary relationships between onchobothriid tapeworms and their elasmobranch hosts. International Journal for Parasitology 31: 960-975.

Caira, J.N. & K. Jensen. 2013. Planetary Biodiversity Inventory: A Survey of Tapeworms from Vertebrate Bowels of the Earth (online). Accessed 09 August 2013 at https://sites.google.com/site/tapewormpbi/ and http://tapewormdb.uconn.edu/.

Caira, J.N. & F.B. Reyda. 2005. Eucestoda (true tapeworms). In Marine Parasitology, K. Rohde, Ed. CSIRO Publishing, Collingwood, Australia, pp. 92-104. (Invited book chapter).

Eyring, K.L., C.J. Healy, & F.B. Reyda. 2012. A new genus of cestode (Rhinebothriidea) from Mobula kuhlii Rajiformes: Mobulidae) from Malaysian Borneo. Journal of Parasitology 98(3): 584-91.

Healy, C.J., J.N. Caira, K. Jensen, B.J. Webster, & D.T.J. Littlewood, 2009. Proposal for a new tapeworm order, Rhinebothriidea. International Journal for Parasitology 39: 497-511.

Marcogliese, D.J. 2005. Parasites of the superorganism: Are they indicators of ecosystem health? International Journal for Parasitology 35:705-716.

Price, P. W. 1980. Evolutionary Biology of Parasites. Princeton University Press, Princeton.

Reyda, F.B. & J.N. Caira. 2006. Five new species of Acanthobothrium (Cestoda: Tetraphyllidea) from Himantura uarnacoides (Myliobatiformes: Dasyatidae) in Malaysian Borneo. Comparative Parasitology 73(1): 49-71.

Reyda, F.B. & P.D. Olson. 2003. Cestodes of cestodes of Peruvian freshwater stingrays. Journal of Parasitology 89(5): 1018-1024.

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Cestodes of the fishes of Otsego Lake and nearby waters

Amanda Sendkewitz1, Illari Delgado1, and Florian Reyda2

INTRODUCTION

This study of fish cestodes (i.e., tapeworms) is part of a survey of the intestinal parasites of fishes of Otsego Lake and its tributaries (Cooperstown, New York) from 2008 to 2014. The survey included a total of 27 fish species, consisting of six centrarchid species, one ictalurid species, eleven cyprinid species, three percid species, three salmonid species, one catostomid species, one clupeid species, and one esocid species. This is really one of the first studies on cestodes in the area, although one of the first descriptions of cestodes was done on the Proteocephalus species Proteocephalus ambloplitis by Joseph Leidy in Lake George, NY in 1887; it was originally named Taenia ambloplitis.

Parasite diversity is a large component of biodiversity, and is often indicative of the health and stature of a particular ecosystem. The presence of parasitic worms in fish of Otsego County, NY has been investigated over the course of a multi-year survey, with the intention of observing, identifying, and recording the diversity of cestode (tapeworm) species present in its many fish species. The majority of the fish species examined harbored cestodes, representing three different orders: Caryophyllidea, Proteocephalidea, and Bothriocephalidea.

METHODS

The fish utilized in this survey were collected through hook and line, gill net, electroshock, or seining methods throughout the year from 2008-2014. Cestodes were collected in sixteen sites throughout Otsego County. These sites included Beaver Pond at Rum Hill, the Big Pond at Thayer Farm, Canadarago Lake, a pond at College Camp, the Delaware River, Hayden Creek, LaPilusa Pond, Mike Schallart’s Pond in Schenevus, Moe Pond, a pond in Morris, NY, Oaks Creek, Paradise Pond, Shadow Brook, the Susquehanna River, the Wastewater Treatment Wetland (Cooperstown), and of course Otsego Lake. The specimens were then dissected and analyzed at the SUNY Oneonta Biological Field Station. The main organ examined was the intestine. However, other organs such as the stomach, pyloric cecae, heart, liver as well as the body cavity were also examined.

Cestode specimens collected were preserved either in 4% neutral-buffered formalin or in 100% ethanol. They were stained with Delafield’s Hematoxylin, dehydrated using a graded ethanol series, mounted in Canada balsam, and preserved as whole mount slides. The parasites were then studied with a light microscope and identified with the aid of a plethora of published scientific literature (i.e., all those listed in the References section). Representative specimens were also photographed with a digital camera. Electron microscopy was also used to aid in the identification of the cestodes. ______

¹ SUNY Oneonta undergraduate student, Biology Department, SUNY Oneonta 2Assistant Professor of Invertebrate Zoology and Researcher, Biology Department and Biological Field Station, SUNY Oneonta

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RESULTS

Among the fish examined, ~120 of the 400+ fish that were examined were infected with either adult or juvenile cestodes. To date, adult specimens of eight species, representing four genera and three orders of cestodes, were encountered in Otsego Lake or in nearby water bodies. The caryophyllid cestode Glaridacris catostomi (Figure 1) occurred in Catostomus commersoni and in Hypentelium nigricans. At least four species of Proteocephalus were also encountered. Proteocephalus cf ambloplitis (Figure 2) occurred in Micropterus dolomieu, Micropterus salmoides, and A. rupestris. Proteocephalus cf longicollis (Figure 3) occurred in Coregonus clupeaformis. Proteocephalus cf pinguis (Figure 4) occurred in Esox niger. Proteocephalus cf pearsei (Figure 5) occurred in Perca flavescens. Additional specimens of Proteocephalus that were encountered but not identified to species were found in the fish species Lepomis gibbosus, Catostomus commersonii, Coregonus clupeaformis, Salvelinus nemaycush, Cottus cognatus, and Noturus insignis. Each Proteocephalus species had distinctive scolex and bothrial features in combination with features of the strobila, but preliminary morphological examinations did not enable species-specific identification, owing to the lack of adequately mature specimens and the limitations in the reference taxonomic literature. Hence, the “cf” designation was given for each of the three Proteocephalus species encountered. The bothriocephalid cestode Bothriocephalus cuspidatus (Figure 6) occurred in Perca flavescens and in Sander vitreus. Another bothriocephalid cestode, Bothriocephalus acheilognathi (Figure 7), was found in Semotilus atromaculatus and in Notemigonus crysoleucas in a beaver pond at the base of Rum Hill. Additional specimens of Bothriocephalus that were not identified to species–i.e., they may or may not be conspecific with those mentioned above–were found in Micropterus salmoides, Lepomis gibbosus, Lepomis macrochirus, Ambloplites rupestris, Perca flavescens and Notropis heterodon. These specimens were limited in number and were generally not sexually mature, precluding specific identifications. Finally, a species of Eubothrium was found in Salvelinus nemaycush, but additional specimens of that taxon are needed to make a specific identification possible.

The juvenile cestodes encountered represented two genera of different cestode orders, Proteocephalus and Bothriocephalus. Unknown cestodes awaiting identification were also found in Ameiurus nebulosus, and Pimephales promelas.

DISCUSSION

Out of all the parasitic worms encountered during the survey, the cestodes were observed the least frequently. Among those cestodes encountered, many were sexually immature. This is a problem because it limited our ability to make firm identifications. For example, our identifications of the species of Proteocephalus encountered in this survey were based on scolex morphology. Fully confirmed identifications would, however, require us to examine proglottid morphology in addition to scolex morphology. This was not an option owing to the generally lack of sexually mature specimens. In this study, species of Proteocephalus were therefore only tentatively identified to species, hence designated “cf”. This abbreviation is for the Latin word “confer”, meaning compare to the form in this taxonomic context. More work is needed to properly identify the species. Another limitation with identifying species of Proteocephalus is a lack of complete and useful reference in the taxonomic literature.

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At least two species of Bothriocephalus were observed in this study, Bothriocephalus cuspidatus and Bothriocephalus achelognathi. The latter of the two is an invasive species of tapeworm, it was only found in one pond however, this finding is a first for New York state; B. cuspidatus has previously only been reported elsewhere in North America. Additional specimens of the species of Eubothrium that was encountered in Lake trout are needed to identify that tapeworm to species.

CONCLUSION

Furthering our understanding of the cestode species of Otsego Lake and nearby water bodies will require the collection of additional specimens of Proteocephalus and Eubothrium that are sexually mature, to facilitate specific identifications.

ACKNOWLEDGMENTS We would like to thank the students of Dr. Reyda’s Lab, the numerous summer interns from SUNY Oneonta, Steve Stowell, (SUNY Cobleskill) and Tim Pokorny (NYS DEC), for their help in fish and snail collections. We thank Anindo Choudhury (St. Norbert College) for his helpful comment on species identifications. Funding for this research was provided in part an NSF FSML grant to W. Harman (DBI 1034744). Also, special thanks to Dr. Bill Harman, Matt Albright, and Holly Waterfield of the Biological Field Station, and Dr. Janine Caira of the University of Connecticut. SUNY Biological Field Station provided the necessary equipment and work space to carry out this project. This work was done as an independent study by A.S. and I.D. over the course of three semesters. REFERENCES

Andersen, K. 1979. Studies on the Scolex Morphology of Eubothrium spp. with emphasis on Characters Usable in the Species Discrimination and with Brief References on the Scolices of Bothriocephalus sp. And Triaenophorus spp. (Cestoda; Pseudophyllidea). Z. Parasitenkd. 60:147-156.

Hanzelová, V., T. Scholz, D. Gerdeaux and R. Kuchta. 2002. A comparative study of Eubothrium salvelini and E. crassum (Cestoda: Pseudophyllidea) parasites of Artic char and brown trout in alpine lakes. Environmental Biology of Fishes. 64:245-256.

Kuchta, R., T. Scholz, J. Brabec and R.A. Bray. 2008. Suppression of the tapeworm order Pseudophyllidea (Platyhelminthes: Eucestoda) and the proposal of two new orders, Bothriocephalidea and Diphyllobothriidea. International Journal for Parasitology. 38:49- 55.

Mackiewicz, J.S. 1965. Redescription and Distribution of Glaridacris catostomi Cooper, 1920 (Cestoidea: Caryophyllaeidae)." Journal of Parasitology. 51(4): 554-560.

Scholz, T. 1997. “A revision of the species of Bothriocephalus Rudolphi, 1808 (Cestoda: Pseudophyllidea) parasitic in American freshwater fishes. Systematic Parasitology. 36: 85-107.

Scholz, T. and A. de Chambrier. 2003. “Taxonomy and biology of proteocephalidean cestodes: current state and perspectives.” Helmnithologia. 40(2): 65-77.

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Forest bryophyte reproduction and dispersal: An update

Alexander Lawrence1 and Sean C. Robinson 2

During the summer of 2012 we, along with Rebekah Obenauer, established a long-term study investigating forest bryophyte dispersal and colonization patterns at three SUNY Oneonta properties including: Greenwoods Conservancy (Figure 1), Thayer Farm (Figure 2), and Rum Hill (Figure 3) (Robinson et al. 2013). Three sites at each of these three locations (nine sites total) were selected, based on an initial survey of bryophyte diversity and forest composition. Two of these nine sites were intensively sampled for bryophytes including one at Thayer Farm and one at Rum Hill. For each bryophyte patch we encountered, a sample was collected for later identification, and the following information was recorded: substrate type, patch location, presence/absence of sporophytes, site name, date, and time. This work resulted in the mapping and identification of 572 bryophyte patches, representing 50 species.

During the summer of 2013, we continued this work with the help of Mathew Dami, a SUNY Oneonta graduate student. Two additional sites were sampled including one at Thayer Farm and one at Greenwoods Conservancy, resulting in a total of 824 bryophyte patches that have now been mapped and identified and an additional 901 bryophyte patches that have been mapped but still remain to be identified. Our 2012 report included a comparison of data collected at one Rum Hill site and one Thayer Farm site. Here we present a comparison between those two sites and the first site established at Greenwoods Conservancy.

The Greenwoods Conservancy site was the second-most species rich with 37 species encountered (25 and 40 species were encountered at Thayer Farm and Rum Hill, respectively) (Figure 4). Two measures of site similarity based on species composition and abundances showed that each of the three sites harbors a relatively unique assemblage of bryophytes (Table 1). This justifies the need for additional sites at each location for adequate replication. Ultimately, however, this will result in a more robust dataset by capturing a greater diversity of species.

Overall sporophyte production was highest at the Rum Hill site (44% of material collected) compared to Thayer Farm (35% of material collected) and Greenwoods Conservancy (31%). As seen in the 2012 data, there was a strong correlation, across all sites between sporophyte production and species abundances (r = 0.93, p = 0.0000), indicating a greater dependence on spores for dispersal and colonization. Five asexually reproducing species that showed high abundances at Rum Hill and Thayer Farm, however, were also found to be among the most abundant species at Greenwoods Conservancy. This continued result appears to support current work that suggests that asexual propagules play an equally important role in the dissemination of bryophytes beyond the immediate vicinity of a given patch.

1 Biological Field Station Intern, summer 2013.Environmental Science Major, SUNY-Oneonta 2 Assistant Professor of Biology, Biology Department, SUNY-Oneonta

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Figure 1. Map of Greenwoods Conservancy, Burlington, NY, showing sampling sites (SITE 1, SITE 2 and SITE 3).

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Figure 2. Map of Thayer Farm, Springfield, NY showing sampling sites (SITE 1, SITE 2 and SITE 3).

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Figure 3. Map of Rum Hill, Springfield, NY, showing sampling sites (SITE 1, SITE 2 and SITE 3).

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Figure 4. Total number of collections made (black bars), and number of collections with sporophytes present (gray bars) for each species encountered at the Greenwoods Conservancy site.

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Table 1. Bryophyte community similarity between sampling sites at Thayer Farm, Rum Hill, and Greenwoods Conservancy. Similarity values are based on species composition and abundances.

Site Comparison Bray-Curtis Similarity Percent Similarity in Species Index Value Composition and Abundance Thayer Farm vs. Rum Hill 0.73 32% Thayer Farm vs. Greenwoods 0.78 38% Greenwoods vs. Rum Hill 0.76 54%

REFERENCES

Robinson, S., A. Lawrence and R. Obenauer. 2013. Bryophyte reproduction and dispersal in a mixed hardwood forest. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Monitoring the effectiveness of the Cooperstown wastewater treatment wetland, 20131

M.F. Albright

BACKGROUND (from Albright and Waterfield 2011)

In 2002, the US Army Corp of Engineers (ACE) initiated a 1.6 million dollar Upper Susquehanna River Watershed-Cooperstown Area Ecosystem Restoration Feasibility Study And Integrated Environmental Assessment. Authorized by the U.S. Congress, the pilot program was to “use wetland restoration, soil and water conservation practices, and non-structural measures to…improve water quality and wildlife habitat…in the Upper Susquehanna River Basin…” (ACE 2001). Initially identified were eight Field Assessed Benefit and Design Strategy sites (FABADS) in Otsego County. During 2003, the SUNY Oneonta Biological Field Station (BFS) monitored two restored sites which receive agricultural runoff as well as a local “pristine reference site”. Comparisons were made between inflows and outflows, and between the wetlands, of concentrations of different nutrient fractions, suspended sediments and fecal coliform bacteria. This short term study did indicate water quality improvements when nutrient levels at the inflow were elevated (Fickbohm 2005), though it is probable that not enough time had elapsed to allow these systems to naturalize to the point where treatment potential was realized.

A third ACE restoration wetland was sited in the outskirts of the Village of Cooperstown adjacent to the municipal sewage treatment facility (Figure 1). The primary function of this 3 acre wetland was phosphorus and nitrogen removal, potentially by converting this site into a treatment wetland for the Village’s municipal effluent. However, at that time, funds to deliver the effluent to the wetland were lacking. The wetland design, provided by Ducks Unlimited, did not necessarily follow that generally utilized for treatment wetlands.

In 2009, funding was provided by the Village of Cooperstown’s Sewer Reserve Fund to hire the services of Lamont Engineering to evaluate alternatives to address nutrient reduction from the wastewater treatment plant (Jackson 2009). A more restrictive SPDES permit by the NYSDEC regarding nutrient loading to the Susquehanna River is consistent with New York State being cosignatory with the Chesapeake Bay Nutrient Reduction Strategy. The engineering report evaluated approaches to reducing phosphorus and nitrogen introduced into the Susquehanna River, their capital and annual operational costs, and expected nutrient reductions. In conclusion, it was recommended that utilizing the existing wetland for tertiary treatment would likely meet the nutrient reduction goals while costing substantially less than other approaches (i.e., addition of chemical coagulants, modification to the treatment plant, etc.).

1 Funding for this project was provided by the Village of Cooperstown.

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Figure 1. Bathymetric map of the wastewater treatment wetland, Cooperstown, NY (modified from Robb 2012).

Rationale for monitoring

Wetlands have been used as water treatment cells for a number of years, but, until recently, only on a very limited basis. Since the mid 1990s, however, the number of constructed wetlands, having a broad range of system configurations and treatment applications, has increased markedly (Kadlec and Wallace 2009). When associated with municipal sewage outfalls, the parameters that are most often targeted for reduction are phosphorus, various nitrogenous compounds (ammonia, nitrate, total nitrogen), suspended solids and biological oxygen demand. The demonstrated effectiveness of the removal of these constituents has been promising, though quite variable, as design and site characteristics are, in practically every case, unique. Because of this, every time a treatment wetland is utilized, the opportunity exists to collect meaningful data which can aid in the design of future systems. More directly, data collection at some level is necessary to evaluate whether or not the goals of the treatment wetland, and the regulated limits of the parameters, are met.

For the Cooperstown WWT wetland, the concentration limit requirements of total phosphorus, total nitrogen, ammonia and nitrates are not more stringent under the new SPDES/Chesapeake Bay Nutrient Reduction Strategy than they had been. It is not likely that concentrations leaving the wetland will be any higher than those entering it (excepting some short- term, meteorologically driven events). Ongoing studies of the monitoring of nutrient reduction

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dynamics could provide insight into, among other things, modification of the wetland’s flow patterns, depth, substrate composition, plant assemblages, etc. which might enhance performance.

METHODS

Early efforts to quantify flow from the wetland were frustrated by two factors. A V-notch weir was constructed of doubled ¾” treated plywood (glued and screwed together). The notch was beveled ~30o (with the bevel facing downstream), and a metal 90o edge was fastened to the upstream face, to reduce friction. A scaled gauge was attached to upstream face, off to the side, so that water level could be recorded. An adhesive-backed gasket was used to line the edges of the downstream face of the weir, and the bottom edge, to ensure that no leakage occurred around or under the device. The top four boards were removed from the flood control device and the weir was inserted into the channels. A Solinst® Levelogger, a pressure-transducer depth recorder, was purchased and attached to the upstream face of the weir, the intention being to collect a continuous log of water heights above the V-notch for flow determination. However, the flow values logged in this manner exhibited such a discrepancy with the metered flow of the wastewater treatment plant that they were considered unreliable and were not used. Efforts in 2011 focused on direct reading of the gauge on the weir face. Robb (2012) mounted a programmable Reconyx® trail camera so that it would capture images of the gauge at 15-minute intervals. In the absence of moderate rainfall (> ~ 1 cm/24 hr), the mean daily inflow from the wastewater treatment plant nearly equaled the outflow from the wetland. During wetter periods, a small stream entering the west side of the wetland can contribute enough flow so that the outflow exceeds the input from the treatment plant for short periods of time. Efforts to accurately estimate the flow at the outlet structure were further compounded over 2011 by activities of muskrat, which plugged the surface outflow structure, and beaver, which were believed to plug the subsurface pipe leading to the outflow structure. Eventually, efforts to gauge the flow were abandoned and the flow of effluent into the wetland was assumed to equal the flow out of it. Sampling did not coincide with runoff events in order to minimize their influence.

During summer and fall of 2013, activities by muskrat were leading to excessive blockage on the outfall device to the point where levels far exceeded those for which the wetland was designed and considerable effort was required by the village treatment plant operators to clear the device. Screening the exterior cage was ineffective as the animals were presumably moving into the outflow device through the drain pipe (which proved to be much larger and nearer to the shore than had been expected). To address this issue, several boards were removed, draining the wetland surface to about 1 m below “normal” height in early November. Efforts to reduce muskrat numbers commenced with the onset of NYS trapping season.

Sampling began in February 2010, and was done monthly through May 2010 to evaluate nutrient conditions prior to the diversion of effluent to the wetland (which commenced on 17 June 2010). Thereafter, samples were collected two to four times a month from the wastewater treatment plant (effluent), the wetland’s outlet and the stream feeding the wetland (to evaluate contributions from this source). This report summarizes results through December 2013. Samples were processed according to automated methods using a Lachat QuikChem FIA+ Water Analyzer. Samples were analyzed for total phosphorus using ascorbic acid following persulfate digestion

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(Liao and Martin 2001), total nitrogen using the cadmium reduction method (Pritzlaff 2003) following peroxodisulfate digestion Ebina et.al (1983), ammonia using the phenolate method (Liao 2001), and for nitrate+nitrite nitrogen using the cadmium reduction method (Pritzlaff 2003). Missing values were approximated by interpolating existing data.

RESULTS

Prior to the wetland receiving effluent, the outflowing concentrations of ammonia were below detection, nitrate averaged 0.28 mg/l, total nitrogen averaged 0.72 mg/l and total phosphorus averaged 0.043 mg/l. For the tributary inflow to the wetland from February 2010 through December 2013, mean nutrient concentrations were 0.01 mg/l (SE= 0.01) for ammonia, 0.26 mg/l (SE= 0.04) for nitrite+nitrate, 0.44 mg/l (SE= 0.04) for total nitrogen and 0.040 mg/l (SE= 0.004) for total phosphorus. The typical low flows and low nutrient concentrations of this tributary indicate that its influence on calculating nutrient retention rates, and investigating nutrient transformations, is minimal.

Summaries of the annual retention, as both net volumes (kg) as a percent of the inputs, of ammonia, nitrite+nitrate, total nitrogen and total phosphorus following the diversion of effluent to the wetland are provided in Table 1. Tables 2-5 provide mean monthly concentrations of the wastewater effluent and wetland’s outfall, as are total monthly nutrient volumes (kg), the volume of nutrients retained (kg) and the mean retention rate (%).

Between June 2010 and December 2013, the total amount of nutrients retained by the treatment wetland included 1,230 kg of ammonia-N, 7,446 kg of nitrate-N, 10,904 kg of total nitrogen-N and 1124 kg of total phosphorus-P. The monthly rates of retention of ammonia varied much more so than did other nutrients, which seemed mainly due to high variability of its concentration in the treatment plant effluent (mean= 1.50 mg/l, SE= 0.24); concentrations were lower and less variable at the wetland’s outlet (mean= 1.01 mg/l, SE= 0.13). This temporal variation led to calculated negative retention rates (or release) of ammonia in some months. Overall, the mean retention declined from about 42% in 2010 to 27.4% in 2011 and 24% in 2012. Over 2013, it had increased to 44.7%. However, this is not believed to reflect varying treatment by the wetland, so much as the fact that the ammonia concentration of the effluent flowing to the wetland has been variable over the course of the study (2.46 mg/l in 2010, 1.67 mg/l in 2011, 0.93 mg/l in 2012 and 1.83 in 2013).

Retention of both nitrate and total nitrogen was higher in 2012 (46% and 42%, respectively) and 2013 (35 % and 43%) than in 2010 and 2011 (between 28 and 30% for both parameters during both years). The retention of total phosphorus was highest in 2010 at 36% and lowest in 2011 at about 15% . It was 22% in both 2012 and 2013. In 2011, retention was lowest in summer months whereas that was when it was highest in 2010 (at the onset of the treatment wetland’s use). Phosphorus removal would be expected to decline if the main mechanism for removal is sediment binding (as the sediments become saturated) as opposed to biological uptake (Kadlec and Wallace 2009).

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Two anomalous situations over the 2013 monitoring effort are noteworthy. The first was the near record rainfall from mid-May through June. The second involved water levels, which were occasionally much higher (~1 m) than normal from August through October due to plugging at the outlet by muskrat, then the subsequent drawdown of the wetland by about 1 m below normal from early November through the end of the year to deal with that issue. This would have reduced the volume of the wetland by more than 50%. Interestingly, these issues did not meaningfully affect the nutrient removal efficiency of the system.

Given that this wetland was designed more for waterfowl habitat than for water quality improvement, the nutrient removal capacity seems promising. As vegetation densities increase, so should nutrient reduction, both directly though vegetative uptake and enhanced microbial denitrification due to increased microsites (Kadlec and Wallace. 2009). Investigations into phosphorus uptake by rooted plants at the Cooperstown wetland provided conflicting results. Olsen (2011) found elevated phosphorus content in the leaf tissue of reed canary grass (Phalaris arundinacea) within the wetland than that of plants in nearby areas not influenced by the treatment wetland. However, similar investigations in 2011 on reed canary grass and cattail (Typha sp.) did not show meaningful differences in phosphorus uptake (Gazzetti 2012).

Table 1. Summary of ammonia, nitrate, total nitrogen and total phosphorus retention by the Cooperstown treatment wetland, 2010 through 2012.

Ammonia retention Nitrate retention T. Nit. retention T. Phos. retention Kg % Kg % Kg % Kg % 2010 (17 June-Dec) 234.8 41.9 796.9 28.0 1148.8 29.9 252.3 36.2 2011 365.6 27.4 2016.7 28.4 2684.8 28.1 251.0 15.3 2012 116.3 24.4 2636.8 46.4 3302.0 42.0 306.2 21.9 2013 514.0 44.7 1995.4 22.6 3768.6 43.4 314.8 22.4 Total (mean) 1230.7 (34.6) 7445.8 (34.4) 10904.2 (35.8) 1124.4 (24.0)

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Table 2. Mean monthly concentrations of ammonia in the wastewater effluent and wetland’s outfall (mg/l), total monthly ammonia volumes (kg) entering and leaving the wetland, the volume of ammonia retained (kg) and the mean retention rate (%). (Projected).

Month Eff flow NH4 (kg) EFF CM/day (mg/l) Out (mg/l) EFF (kg) OUT (kg) RET. (kg) % RET. Jun-10* 1728 4.10 1.41 63.8 21.9 41.8 65.6 Jul-10 1692 0.31 1.03 16.1 54.0 -37.9 -235.5 Aug-10 1526 5.35 2.46 252.9 116.2 136.8 54.1 Sep-10 1186 2.19 1.17 77.9 41.6 36.3 46.6 Oct-10 1476 2.19 1.17 100.2 53.5 46.7 46.6 Nov-10 1447 0.60 0.45 26.0 19.7 6.3 24.3 Dec-10 1330 0.60 0.48 23.9 19.1 4.9 20.3 2010 560.9 326.1 234.8 41.9 Jan-11 1222 0.68 0.478 24.9 17.5 7.4 29.6 Feb-11 1319 2.68 2.488 98.8 91.8 6.9 7.0 Mar-11 2707 4.30 1.995 360.4 167.4 193.0 53.6 Apr-11 2824 1.96 1.883 166.2 159.5 6.7 4.0 May-11 2816 0.68 1.134 59.6 99.0 -39.4 -66.0 Jun-11 2495 2.15 1.816 161.1 135.9 25.2 15.6 Jul-11 1862 0.54 0.953 31.1 55.0 -23.9 -77.0 Aug-11 1859 3.99 2.073 230.0 119.4 110.6 48.1 Sep-11 2532 0.87 0.758 66.0 57.5 8.5 12.9 Oct-11 2120 0.71 0.341 45.2 21.7 23.5 52.0 Nov-11 1896 0.75 0.461 42.5 26.2 16.3 38.3 Dec-11 1961 0.77 0.262 46.8 15.9 30.9 66.0 2011 1332.6 967.0 365.6 27.4 Jan-12 1972 0.80 0.45 47.3 26.6 20.7 43.8 Feb-12 1722 0.69 0.44 33.3 21.2 12.1 36.2 Mar-12 1832 0.55 0.44 31.2 25.0 6.2 20.0 Apr-12 1427 0.55 0.44 23.5 18.8 4.7 20.0 May-12 1718 0.41 0.14 21.8 7.5 14.4 65.9 Jun-12 1442 0.68 0.61 29.4 26.4 3.0 10.3 Jul-12 1340 2.46 2.28 102.2 94.7 7.5 7.3 Aug-12 1344 1.39 1.50 57.9 62.5 -4.6 -7.9 Sep-12 1124 0.63 0.55 21.2 18.5 2.7 12.7 Oct-12 1196 0.52 0.43 18.7 15.4 3.2 17.3 Nov-12 1128 0.77 0.86 26.1 29.1 -3.0 -11.7 Dec-12 1245 1.68 0.40 64.9 15.4 49.4 76.2 2012 477.5 361.2 116.3 24.4 Jan-13 1431 0.28 0.25 12.4 11.3 1.1 9.1 Feb-13 1321 1.21 0.46 44.9 17.1 27.8 61.9 Mar-13 1473 1.55 1.00 70.5 45.7 24.8 35.2 Apr-13 1893 4.08 2.36 231.7 134.0 97.7 42.2 May-13 1522 0.81 0.81 38.2 38.3 -0.1 -0.2 Jun-13 2192 2.26 0.89 148.6 58.5 90.1 60.6 Jul-13 2332 1.58 1.13 113.9 81.5 32.3 28.4 Aug-13 1703 3.96 1.58 209.1 83.2 125.9 60.2 Sep-13 1647 2.77 1.08 137.0 53.6 83.4 60.9 Oct-13 1348 1.32 0.92 55.0 38.3 16.7 30.4 Nov-13 1242 0.92 0.87 34.3 32.4 1.9 5.4 Dec-13 1427 1.21 0.93 53.5 41.1 12.4 23.1 2013 1149.0 635.0 514.0 44.7 To date 3520.0 2289.3 1230.7 34.6

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Table 3. Mean monthly concentrations of nitrite+nitrate in the wastewater effluent and wetland’s outfall (mg/l), total monthly nitrite+nitrate volumes (kg) entering and leaving the wetland, the volume of nitrite+nitrate retained (kg) and the mean retention rate (%).

NO2+NO3 Month Eff flow (kg) EFF CM/day (mg/l) Out (mg/l) EFF (kg) OUT (kg) RET. (kg) % RET. Jun-10* 1728 11.85 8.05 184.3 125.2 59.1 32.1 Jul-10 1692 9.37 7.65 491.2 401.0 90.2 18.4 Aug-10 1526 9.13 5.75 432.2 272.1 160.1 37.0 Sep-10 1186 10.85 5.80 385.8 206.2 179.6 46.6 Oct-10 1476 10.70 6.43 489.6 294.2 195.4 39.9 Nov-10 1447 11.38 8.33 493.9 361.4 132.4 26.8 Dec-10 1330 9.15 9.65 365.1 385.0 -20.0 -5.5 2010 2842.0 2045.1 796.9 28.0 Jan-11 1222 13.35 13.150 489.5 482.2 7.3 1.5 Feb-11 1319 12.70 11.280 468.9 416.5 52.4 11.2 Mar-11 2707 5.31 4.100 445.5 344.0 101.5 22.8 Apr-11 2824 6.75 3.707 571.6 314.0 257.5 45.1 May-11 2816 8.83 6.665 770.9 581.9 189.0 24.5 Jun-11 2495 10.48 5.575 783.9 417.2 366.7 46.8 Jul-11 1862 9.44 7.850 545.0 453.2 91.8 16.8 Aug-11 1859 12.83 8.095 739.3 466.4 272.9 36.9 Sep-11 2532 9.12 5.075 692.8 385.5 307.3 44.4 Oct-11 2120 7.81 6.350 496.6 403.8 92.8 18.7 Nov-11 1896 8.45 5.585 480.8 317.8 163.0 33.9 Dec-11 1961 10.30 8.420 626.1 511.8 114.3 18.3 2011 7111.0 5094.3 2016.7 28.4 Jan-12 1972 8.47 7.94 501.1 469.7 31.4 6.3 Feb-12 1722 9.55 7.81 460.5 376.6 83.9 18.2 Mar-12 1832 9.75 7.82 553.7 444.1 109.6 19.8 Apr-12 1427 12.12 9.02 518.8 386.1 132.7 25.6 May-12 1718 9.52 4.39 507.1 233.9 273.3 53.9 Jun-12 1442 12.92 3.71 559.0 160.5 398.4 71.3 Jul-12 1340 10.10 3.08 419.5 127.9 291.6 69.5 Aug-12 1344 7.80 2.75 324.9 114.5 210.4 64.7 Sep-12 1124 8.56 1.96 288.7 66.1 222.6 77.1 Oct-12 1196 10.20 2.73 366.0 98.0 268.0 73.2 Nov-12 1128 15.70 8.65 531.3 292.7 238.6 44.9 Dec-12 1245 17.00 7.25 656.3 279.9 376.4 57.4 2012 5686.8 3050.0 2636.8 46.4 Jan-13 1431 10.50 9.43 465.8 418.1 47.7 10.2 Feb-13 1321 11.18 8.43 413.4 311.6 101.7 24.6 Mar-13 1473 7.03 4.42 320.7 201.5 119.1 37.2 Apr-13 1893 7.47 5.12 424.2 290.7 133.4 31.5 May-13 1522 10.90 4.65 514.2 219.4 294.8 57.3 Jun-13 2192 12.10 12.90 795.6 848.2 -52.6 -6.6 Jul-13 2332 9.44 3.83 682.0 276.9 405.2 59.4 Aug-13 1703 9.20 2.61 485.8 137.6 348.3 71.7 Sep-13 1647 9.46 1.50 467.4 74.0 393.5 84.2 Oct-13 1348 7.19 6.00 300.4 250.7 49.7 16.6 Nov-13 1242 9.19 10.20 342.3 379.9 -37.6 -11.0 Dec-13 1427 11.80 7.46 522.0 329.8 192.2 36.8 2013 5733.7 3738.3 1995.4 34.8 To date 21373.6 13927.8 7445.8 34.4

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Table 4. Mean monthly concentrations of total nitrogen in the wastewater effluent and wetland’s outfall (mg/l), total monthly total nitrogen volumes (kg) entering and leaving the wetland, the volume of total nitrogen retained (kg) and the mean retention rate (%).

Month Eff flow TN (kg) EFF CM/day (mg/l) Out (mg/l) EFF (kg) OUT (kg) RET. (kg) % RET. Jun-10* 1728 17.30 9.51 269.0 147.9 121.2 45.0 Jul-10 1692 14.10 12.50 739.6 655.7 83.9 11.3 Aug-10 1526 16.03 9.38 758.3 443.6 314.7 41.5 Sep-10 1186 13.47 7.32 479.0 260.2 218.7 45.7 Oct-10 1476 12.40 7.29 567.4 333.3 234.0 41.3 Nov-10 1447 13.65 10.05 592.6 436.3 156.3 26.4 Dec-10 1330 10.90 10.40 434.9 415.0 20.0 4.6 2010 3840.8 2692.0 1148.8 29.9 Jan-11 1222 16.48 16.375 604.1 600.4 3.7 0.6 Feb-11 1319 19.28 14.875 711.7 549.2 162.5 22.8 Mar-11 2707 10.20 6.775 855.8 568.4 287.4 33.6 Apr-11 2824 9.38 5.942 794.9 503.4 291.6 36.7 May-11 2816 11.45 9.075 999.7 792.3 207.4 20.7 Jun-11 2495 17.58 8.338 1315.3 624.0 691.3 52.6 Jul-11 1862 14.46 11.038 835.0 637.2 197.7 23.7 Aug-11 1859 11.11 12.350 639.8 711.6 -71.7 -11.2 Sep-11 2532 11.65 6.275 885.1 476.7 408.4 46.1 Oct-11 2120 9.83 7.720 625.1 491.0 134.2 21.5 Nov-11 1896 9.97 6.365 567.2 362.1 205.1 36.2 Dec-11 1961 11.80 9.045 717.3 549.8 167.5 23.3 2011 9551.0 6866.2 2684.8 28.1 Jan-12 1972 10.20 9.01 603.4 533.0 70.4 11.7 Feb-12 1722 11.20 8.84 540.1 426.3 113.8 21.1 Mar-12 1832 13.23 12.28 751.3 697.4 54.0 7.2 Apr-12 1427 19.08 12.30 816.8 526.5 290.2 35.5 May-12 1718 13.98 6.35 744.7 338.3 406.5 54.6 Jun-12 1442 14.40 6.04 623.0 261.3 361.7 58.1 Jul-12 1340 17.00 8.05 706.1 334.4 371.8 52.6 Aug-12 1344 13.55 7.00 564.4 291.6 272.8 48.3 Sep-12 1124 14.88 4.27 501.8 144.0 357.8 71.3 Oct-12 1196 17.20 5.48 617.2 196.6 420.5 68.1 Nov-12 1128 18.28 9.45 618.6 319.8 298.8 48.3 Dec-12 1245 20.30 12.95 783.6 499.9 283.7 36.2 2012 7871.0 4569.1 3302.0 42.0 Jan-13 1431 14.78 13.03 655.4 577.8 77.6 11.8 Feb-13 1321 16.70 11.23 617.7 415.2 202.5 32.8 Mar-13 1473 11.73 8.03 535.2 366.3 168.9 31.6 Apr-13 1893 21.46 12.19 1218.5 692.2 526.4 43.2 May-13 1522 15.00 6.69 707.6 315.6 392.0 55.4 Jun-13 2192 14.45 6.47 950.1 425.4 524.7 55.2 Jul-13 2332 12.80 5.75 925.3 415.3 510.0 55.1 Aug-13 1703 13.41 4.99 707.9 263.6 444.3 62.8 Sep-13 1647 14.34 3.88 708.2 191.9 516.3 72.9 Oct-13 1348 11.80 9.33 493.0 389.6 103.4 21.0 Nov-13 1242 12.50 10.80 465.6 402.3 63.3 13.6 Dec-13 1427 15.95 10.55 705.6 466.5 239.1 33.9 2013 8690.1 4921.6 3768.6 43.4 To date 29952.9 19048.8 10904.2 35.8

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Table 5. Mean monthly concentrations of total phosphorus in the wastewater effluent and wetland’s outfall (mg/l), total monthly total phosphorus volumes (kg) entering and leaving the wetland, the volume of total phosphorus retained (kg) and the mean retention rate (%).

Month Eff flow TP (kg) EFF CM/day (mg/l) Out (mg/l) EFF (kg) OUT (kg) RET. (kg) % RET. Jun-10* 1728 4.36 1.49 67.7 23.2 44.5 65.7 Jul-10 1692 1.49 0.81 78.3 42.6 35.7 45.6 Aug-10 1526 3.20 2.15 151.3 101.6 49.7 32.8 Sep-10 1186 3.39 2.32 120.6 82.5 38.1 31.6 Oct-10 1476 2.41 1.56 110.2 71.4 38.8 35.2 Nov-10 1447 2.28 1.52 99.2 66.2 33.0 33.3 Dec-10 1330 1.76 1.45 70.2 57.7 12.6 17.9 2010 697.4 445.1 252.3 36.2 Jan-11 1222 2.205 2.030 80.8 74.4 6.4 7.9 Feb-11 1319 2.225 1.865 82.2 68.9 13.3 16.2 Mar-11 2707 1.168 0.701 98.0 58.8 39.1 39.9 Apr-11 2824 1.088 0.700 92.2 59.3 32.9 35.7 May-11 2816 1.645 1.160 143.6 101.3 42.3 29.5 Jun-11 2495 3.092 2.402 231.4 179.7 51.6 22.3 Jul-11 1862 2.807 2.870 162.1 165.7 -3.6 -2.2 Aug-11 1859 3.700 4.125 213.2 237.7 -24.5 -11.5 Sep-11 2532 1.419 1.308 107.8 99.3 8.5 7.8 Oct-11 2120 4.400 3.950 279.8 251.2 28.6 10.2 Nov-11 1896 1.480 0.681 84.2 38.8 45.4 54.0 Dec-11 1961 0.996 0.818 60.6 49.7 10.9 17.9 2011 1635.8 1384.8 251.0 15.3 Jan-12 1972 0.941 0.810 55.7 47.9 7.7 13.9 Feb-12 1722 1.230 1.237 59.3 59.6 -0.3 -0.6 Mar-12 1832 2.020 1.462 114.7 83.0 31.7 27.6 Apr-12 1427 3.100 2.193 132.7 93.9 38.8 29.3 May-12 1718 2.535 1.437 135.0 76.5 58.5 43.3 Jun-12 1442 3.340 2.578 144.5 111.5 33.0 22.8 Jul-12 1340 3.052 3.152 126.8 130.9 -4.2 -3.3 Aug-12 1344 3.360 2.645 140.0 110.2 29.8 21.3 Sep-12 1124 4.060 3.345 136.9 112.8 24.1 17.6 Oct-12 1196 2.930 1.985 105.1 71.2 33.9 32.3 Nov-12 1128 3.367 2.502 113.9 84.7 29.3 25.7 Dec-12 1245 3.480 2.860 134.3 110.4 23.9 17.8 2012 1399.0 1092.8 306.2 21.9 Jan-13 1431 1.170 1.183 51.9 52.5 -0.6 -1.1 Feb-13 1321 2.453 1.645 90.7 60.9 29.9 32.9 Mar-13 1473 1.610 1.148 73.5 52.4 21.1 28.7 Apr-13 1893 2.835 2.475 161.0 140.5 20.4 12.7 May-13 1522 1.880 1.380 88.7 65.1 23.6 26.6 Jun-13 2192 1.376 1.115 90.5 73.3 17.2 19.0 Jul-13 2332 3.093 2.315 223.5 167.3 56.2 25.1 Aug-13 1703 2.990 1.989 157.9 105.0 52.9 33.5 Sep-13 1647 2.569 1.928 126.9 95.2 31.7 25.0 Oct-13 1348 2.838 2.393 118.5 99.9 18.6 15.7 Nov-13 1242 3.335 2.340 124.2 87.2 37.1 29.8 Dec-13 1427 2.230 2.075 98.7 91.8 6.9 7.0 2013 1406.0 1091.1 314.8 22.4 To date 5138.2 4013.8 1124.4 24.0

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CONCLUSION

The bathymetry of the wetland (see Figure 1) implies that much of the wetland is considerably deeper than that recommended for maximum nutrient removal; shallower systems allow for the colonization of suitable plants, preferably emergents (Kadlec and Wallace 2009). Also, work by Robb (2012), considering nutrient concentrations across the standing water and by using fluorescent tracers, indicate that the most suitable portion of the system regarding depth (the southeast arm) is ineffective since it is not in the flow path of the effluent. It may be worth considering maintaining the water level lower than the original design, as it was through the later months of 2013 to alleviate plugging issues, in order to encourage plant growth.

REFERENCES

ACE. 2001. Upper Susquehanna River Watershed-Cooperstown Area Ecosystem Restoration Feasibilty Study and Integrated Environmental Assessment. Project management Plan. United States Army Corps of Engineers, Planning Division. Baltimore, MD.

Albright, M.F. and H.A.Waterfield. 2011. Monitoring the effectiveness of the Cooperstown wastewater treatment wetland. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Ebina, J., T. Tsutsi, and T. Shirai. 1983. Simultaneous determination of total nitrogen and total phosphorus in water using peroxodisulfate oxidation. Water Res.7 (12):1721-1726.

Fickbohm, S.S. 2005. Upper Susquehanna River Watershed- Cooperstown Area Ecosystem Restoration Feasibility Study And Integrated Environmental Assessment: Post- restoration water quality and wildlife analysis of the FABADS sites (2003-2004). In 37th Ann. Rept. (2004). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Gazzetti, E. 2012. Efficacy of emergent plants as a means of phosphorus removal in a treatment wetland, Cooperstown, New York. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Jackson, M.H. 2009. Wastewater treatment facility modifications engineering report. Lamont Engineers, Cobleskill, NY.

Kadlec, R.H and S.D. Wallace. 2009. Treatment wetlands (second ed.). CRC Press, Boca Raton.

Liao, N. 2001. Determination of ammonia by flow injection analysis. QuikChem ® Method 10-107-06-1-J. Lachat Instruments, Loveland, CO.

Liao, N. and S. Marten. 2001. Determination of total phosphorus by flow injection analysis colorimetry (acid persulfate digestion method). QuikChem ® Method 10-115-01-1- F. Lachat Instruments, Loveland, CO.

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Olsen, B. 2011. Phosphorus content in reed canary grass (Phalaris arundinacea) in a treatment wetland, Cooperstown, NY. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Pritzlaff, D. 2003. Determination of nitrate/nitrite in surface and wastewaters by flow injection analysis. QuikChem ® Method 10-107-04-1-C. Lachat Instruments, Loveland, CO.

Robb, T. 2012. Insight into a complex system: Cooperstown wastewater treatment wetland, 2011. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Groundwater flow and geochemistry at Greenwoods Conservancy

Myles Moore1 and Les Hasbargen2

INTRODUCTION

Greenwoods Conservancy is located on the northern edge of the Appalachian plateau in central New York (latitude 42.719, longitude -75.098). Well-developed stream networks etch the region, which is underlain by gently southwest dipping sedimentary Devonian age rocks. Glaciers repeatedly advanced over this region during the Pleistocene epoch, and they scoured rock basins into the uplands, draped till sheets over the landscape, and mounded moraines in trunk valleys. Cranberry Bog rests in one of these rock basins. Fractures in bedrock are visible in a few places in and around Greenwoods, and linear topographic features appear to follow the trend of these fractures. One such fracture runs right through the east side of Cranberry Bog. This project reports on the orientation of fractures exposed in bedrock around Greenwoods, and whether groundwater geochemistry is distinctive around fractures. One might hypothesize that deeper water is rising toward the surface along vertical fractures, and thus would have a measurably different chemistry from near surface waters. Several shallow dug wells (1-10 m depth) and 3 drilled deep wells (>40 m deep) were sampled extensively as a means of testing the hypothesis (Figure 1).

The main geologic layers in this region are, from youngest to oldest: stream and lake deposits; glacial drift; the Moscow formation (mostly shale and siltstone); the Panther Mountain formation (mostly shale and siltstone); the Marcellus formation (black shale, shale and limestone); and the Onondaga formation (mostly limestone). See Figure 2 below for a geologic cross section under Greenwoods. The Greenwoods Conservancy study deals mainly with the Moscow formation and the glacial till that lies above it, and water draining from those units.

1 Peterson Family charitable Trust Intern, summer 2013. Water Resources Major, Earth & Atmospheric Sciences, SUNY Oneonta, BFS Summer Internship. 2Assistant Professor, Earth & Atmospheric Sciences, SUNY Oneonta, BFS Summer Fellowship.

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Figure 1. Topographic map displaying sampling locations, the regional fracture, and the locations where strike and dips were taken. The cluster of dots at the top of the map and just south of Cranberry Bog represent where the strike and dips were taken.

Figure 2. Geologic formations below Greenwoods Conservancy. (NYS Museum, 1999)

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The Moscow formation represents a marine environment, ranging from deeper water where black shale accumulated, to shallower depths where thin ripple-marked sandstone beds and occasional marine fossils were deposited. The upper portion of the Moscow formation grades into terrestrial deposits. It underlies the Unadilla formation. The Panther Mountain formation consists mostly of shale layers, with lesser amounts of siltstone and sandstone beds. It underlies the Moscow formation (NYS Museum, 1999).

The Marcellus formation covers a wide region, from New York to Ohio, West Virginia, Pennsylvania and Maryland. It consists of black shale and occasional beds of medium-gray shale and limestone nodules or beds of dark gray to black limestone. The Onondaga formation consists of limestone, dolostone, and shale beds. The Onondaga was deposited in a shallow warm tropical sea. Waters draining from the Onondaga usually have high amounts of dissolved calcium, sulfur, and high electrical conductivity (NYS Museum, 1999).

Figures 3 and 4 illustrate the bedrock geology and topography of the area around Cranberry Bog in Greenwoods Conservancy, respectively.

Figure 3. A map displaying the bedrock geology for the Greenwoods area and the locations of the wells sites and strike and dip sites. Elevation scale appears at the left side of the map.

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Figure 4. Topographic map displaying the water sampling locations. Scale is provided in the bottom right.

Hydrologic Setting: The topography of the study area was caused by long term dissection of the Appalachian plateau by streams, and the action of glaciers advancing and receding over the area several times in the last few million years. This caused etching and burying of the pre-existing landscape. Several rock basins have been scoured into the uplands by the glaciers, forming the hollows and hills observed throughout Greenwoods. Cranberry Bog and other smaller water bodies in Greenwoods occupy these basins. One might expect that local groundwater flow is controlled by topography; flow moves from higher ridges into the valleys, and then into streams. However, this prediction of flow could be altered if vertical fractures are present, which would allow groundwater to move vertically along the fracture. The hypothesis that groundwater could be moving up from deeper aquifers to shallower aquifers and surface water forms the core of this study. Water derived from deeper geologic units, such as the Marcellus and Onondaga, should have distinctly different chemical signatures. This study evaluates these signatures by measuring

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elemental concentrations and bulk parameters (electrical conductivity, total dissolved solids, salinity and pH) in surface water, shallow and deep aquifers and groundwater across Greenwoods Conservancy. Tables 1 and 2 summarize and describe the sampling locations.

Table 1. Summary of the location and elevation at each of the sampling sites.

Location Latitude Longitude Elevation (masl) Greenwoods 1 42.71967 -75.08913 567.3 Greenwoods 2 42.71963 -75.08998 572.6 Greenwoods 4 42.71651 -75.08779 552.8 Greenwoods 5 42.71308 -75.10094 567.1 Well at Greenwoods 5 42.71249 -75.10022 577.2 Greenwoods 6 42.71256 -75.10045 579.3 Greenwoods 7 42.71254 -75.10981 567.8 Greenwoods 8 42.71117 -75.10586 569.5 Greenwoods 14 42.71104 -75.09547 555.0 Cranberry Bog South 42.71167 -75.09778 556.0 Cranberry Bog North 42.71875 -75.09604 551.3

Table 2. Attributes of the sampling sites.

Dug Wells Drilled Wells Water Bodies

Greenwoods 2 (GW2) Greenwoods 1 (GW1) Cranberry Bog, South End (CBSE)

Greenwoods 4 (GW4) Greenwoods 8 (GW8) Cranberry Bog, North End (CBNE)

Greenwoods 6 (GW6) Greenwoods 5 (GW5)

Greenwoods 7 (GW7)

Greenwoods 14 (GW14)

Goals for the study: To identify fractures from linear trends on topographic maps and on the ground; to measure the water chemistry of shallow dug wells, deeper drilled wells and the bog; to identify spatial trends in water chemistry and characterize the bedrock associated with that water chemistry; to generate shallow groundwater flow path maps and to test the hypothesis that water is upwelling from deeper aquifers along regional fractures.

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METHODS

Dug well sampling collection method: Materials: ● Sampling pole and bottle ● 3 sample bottles

● HNO3 , used as a preservative for cation sampling ● 0.22 micron filter paper (See Figure 7) ● Vacuum filter, including hand pump, rubber hose that attaches hand pump to collection bottle, glass Buchner funnel that sample water is poured into to be filtered (see Figure 7) ● PCSTestr 35 Multi-Parameter Probe for temperature, pH, electrical conductivity ● Bucket ● Deionized (DI) water for rinsing the sampling bottles ● Duct tape/bottle labeling tape ● Sharpie ● Write-in-rain field guide book to record data ● Gas-powered pump to dewater wells, hose and gasoline ● Water-level indicator to determine depth to water table in wells

Steps for water sampling: At each location, the sampling pole bottle, the top of the sampling pole, and the bucket were rinsed with deionized (DI) water. Water was collected from the site and used to rinse the bucket three times. The bucket was then filled with sample water, and tested with the multi parameter probe for electrical conductivity, total dissolved solids, salinity and pH (bulk chemical parameters.) After this, water was filtered through a vacuum filtration system (Buchner funnel and filter paper), and collected in a collection bottle (See Figures 5 and 6).

The vacuum filtered water was used to rinse the three sample bottles three times, filtered water was added to the sample bottle, along with several drops of HNO3 to a pH < 2, which acts as a preservative to prevent microbial growth. Each bottle was labeled with date, time, location, presence of HNO3 , samplers, water body type, and unique identifier. Typically, 500 ml was collected for cation chemical analyses, and 500 ml collected for anions (sulfate, chloride) and nutrients. These samples were analyzed at the BFS lab for calcium, chloride, and alkalinity. Table 3 summarizes the methodologies for these, and all other, analyses.

Steps for pumping of dug wells: Many of the wells in the monitoring program were shallow dug wells. While it seemed likely that groundwater was flowing through the well, to verify that the water was fresh from the surrounding aquifer, the drilled wells were purged. The following steps were followed: ● Extract a water sample, following procedures above. ● Measure the ground-to-water table depth using the water-level indicator tape measure. ● Insert hose pump all the way down the dug well and begin pumping.

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● Every 2 minutes re-measure and record the ground to water depth using the water-level indicator. Drain dug well completely. ● Allow at least 24 hours before measuring groundwater level. Extract a water sample, following procedures above.

Drilled well sampling collection method: Materials ● At least 3 sample bottles ● Bottle of HNO3 preservative ● 0.22 micron Filter paper (See Figure 5) ● Filtration set-up, including hand pump, rubber hose that attaches hand pump to collection bottle, glass Buchner funnel that sample water is poured into to be filtered (see Figure 5) ● PCSTestr 35 Multi-Parameter Probe ● Bucket ● Gallon jug filled with deionized (DI) water ● Duct tape/bottle labeling tape ● Sharpie ● Write-in-rain field guide book to record data

Steps for collecting drilled well water samples: The drilled wells were all located where a sink inside a cabin, or a faucet located outside, can be tested to get the parameters of the drilled well. Rinse bucket, multi-meter and vacuum filtration set-up materials with DI water.

The bucket that was used for testing was filled and rinsed three times with the sink/faucet water. The multimeter probe was placed into bucket with the conductivity/temperature setting on. The conductivity was recorded every minute until the conductivity and temperature began to stabilize. This typically took 10 to 15 minutes. The goal was to test the well water and not the water that has stagnantly been sitting in the pipes and could give unrepresentative results.

Once the temperature and conductivity stabilized, the bucket was emptied and allowed to refill. The multi-parameter probe was used to get the bulk parameters of conductivity, total dissolved solids, salinity, pH and temperature. This was done immediately, before the temperature of the water would rise. After this, the vacuum filtration was set up (displayed in Figure 5 and Figure 6). The vacuum was used to filtered water to rinse out the three sample bottles each three times. After rinsing, a water sample was collected in one bottle until it was half full. Then 11 drops of HNO3 was added as a preservative. The bottle was capped, shaken and then filled the rest of the way with filtered water. The bottle was labeled “cations, 11 drops of HNO3 preservative” and the designation of the well site, the date, the time, the samplers, and indicating it is a drilled well.

The remaining bottles were filled with the filtered water and labeled as “anions, general” along with the other labels stated above. These samples were analyzed at the BFS for calcium, chloride, and alkalinity (see Table 3).

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Figure 5. Photo displaying, starting from top left and working towards bottom right; filter paper, Buchner funnel, filter, hand pump, rubber hose, collection bottle, pole sampling bottle.

Figure 6. Display of the vacuum filtration set-up with sampling bottles also shown.

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Table 3. Summary of analytical chemistry methods.

Parameter Preservation Method Reference

Electrical None, Measured Oakton PCS Testr Oakton Conductivity in field 35 Multiparameter Instruments; (µS/cm.) Eutech Instruments

Total Dissolved None, Measured Oakton PCS Testr Oakton Solids (ppm.) in field 35 Multiparameter Instruments; Eutech Instruments

Salinity (ppm.) None, Measured Oakton PCS Testr Oakton in field 35 Multiparameter Instruments; Eutech Instruments

pH None, Measured Oakton PCS Testr Oakton in field 35 Multiparameter Instruments; Eutech Instruments

Temperature (°C) None, Measured Oakton PCS Testr Oakton in field 35 Multiparameter Instruments;

Calcium Store at 4 °C EDTA titrimetric APHA 1989 (mg/L.) method, 3500- Ca. D. Standard methods

Chloride Store at 4 °C Titrimetric APHA 1989 (mg/L.) mercuric nitrate method, 4500-Cl. C. Standard Methods

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Table 3 (cont.). Summary of analytical chemistry methods.

Alkalinity Store at 4 °C Titrimetric sulfuric HACH Company

(CaCO3 mg/L.) Acid Method, 820 HACW Test Kit- Model AL-DT

Alkalinity Store at 4 °C Titrimetric Method APHA 1989 (CaCO3 mg/L.) 2320 B. Standard Methods

Total H2SO4 to pH < 2 Persulfate Liao and Marten Phosphorous digestion followed 2001 by single reagent

ascorbic acid

Total Nitrogen H SO to pH < 2 Cadmium Pritzlaff 2003; 2 4 reduction method Ebina et al. 1983 following peroxodisulfate digestion

Nitrate+ Nitrite H2SO4 to pH < 2 Cadmium Pritzlaff 2003 reduction method

Ag, Al, As, B, Ba, HNO3 to pH < 2 ICP-MS Activation Be, Bi, Ca, Ce, Laboratories Ltd. Co, Cd, Cr, Cs,

Cu, Dy, Er, Eu,

Fe, Ga, Ge, Hf, Ho, In, K, La, Li, Lu, Mg, Mn, Mo, Na, Nb, Ni, Pb, Pr, Rb, Re, Sb,

Se, Sm, Sn, Sr,

Ta, Tb, Te, Th, Ti, Tl, Tm, U, V, W, Y, Yb, Zn, Zr

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RESULTS AND DISCUSSION

Geology: In order to study the regional fractures in the area, the orientation of the fractures were measured at two locations: at a bedrock outcrop of the Moscow formation located along Highway 80 north of Greenwoods (latitude: 42.72928 and longitude: -75.095779; displayed in Figure 9 as site 1) and along a stream where bedrock of the Moscow Formation was exposed (latitude: 42.71025 and longitude: -75.09826; displayed in Figure 9 as site 2 located just south of Cranberry Bog). These were the only two outcrops where fractures were readily visible and were suitable for accurate orientation measurements. The Moscow formation outcrop located along Highway 80 had a southside outcrop and a northside outcrop each situated on opposite sides of the road about 15 meters away from each other. Only fractures which were through-going, that is, cut through the entire section of rock, were measured.

We plotted the orientation data in a program called Open Stereo (Figures 7 and 8). This program creates projections called stereonets that show the orientations of planes in geographical space. Two directions are measured in the field which characterize the direction a plane is dipping and the magnitude of the dip. Directions are measured around the outside of the circle from 0 (North) to 360 in a clockwise fashion. The stereonet for the Highway 80 outcrop shows that the main regional fracture runs nearly north-south, with a second set of fractures that runs northeast to southwest (about 40-220 degrees) (Figure 7) . The fractures along the stream appear to have the same orientation (Figure 8).

Figure 7. Display of a stereonet for the strike and dip values at the North End of Greenwoods, this Moscow formation outcrop is located right by highway 80. This represents site 1 in Figure 9.

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Figure 8. Display of a stereonet for the strike and dip values along a stream, where the bedrock outcrops were located South of Cranberry Bog. This represents site 2 in Figure 9.

Cranberry Bog lies directly between the two fracture measurement locations, and aligns nearly perfectly with the north-south set of fractures. This suggests that erosional processes, such as rock quarrying by glaciers and preferential plucking of fractured blocks by streams, have taken advantage of the regional fracture system over geologic time to hollow out the depression in which Cranberry Bog rests.

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Site 1; see Figure 7

Site 2; see Figure 8

Figure 9. Overview of the map locations for where the fracture measurements were taken. Site 1 represents the location of the highway 80 bedrock outcrop (see Figure 7).Site 2 represents the location of the stream bedrock outcrop (see Figure 8). The other dots indicate the location of wells and other water bodies tested.

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Hydrology: Since this project hinged on sampling surface and shallow groundwater, a significant question arises to what hydrologic conditions were like during the sampling period. Figure 10 below shows that April and May were a bit drier than normal, but June was substantially wetter, as reflected in the discharge in the Unadilla River, downstream from the Greenwoods watershed area. The data shows higher gage heights and larger discharge rates that were occurring while sampling for this project was taking place.

Figure 10. Seasonal river flow compared to long term records on the Unadilla River at Rockdale, NY. Data and chart courtesy USGS http://waterdata.usgs.gov/ny/nwis/uv?site_no=01502500.

One method to test the hypothesis of water upwelling from deeper aquifers along the regional fracture is to plot distance from regional fracture against the water chemistry parameters we tested for to see if there is a correlation. We measured electrical conductivity, pH, total dissolved solids, alkalinity and temperature. We also analyzed for a number of major and trace elements. Higher concentrations of barium and strontium in the water samples that are nearer to the fracture could indicate signs of deeper waters mixing with shallow groundwater as well.

Figures 11 through 19 plot the various parameters evaluated against the distance of the site to the regional fracture to identify potential correlation, which could indicate deep-water upwelling. Shallow groundwater flowlines, based upon contours, are proposed in Figure 20. Conductivity is presented in a map in Figure 21.

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Electrical Conductivity:

Figure 11. Conductivity versus distance from regional fracture. There is almost no correlation as the R^2 is 0.0001

Total Dissolved Solids:

Figure 12. T.D.S versus Distance from regional fracture. There is almost no correlation as the R^2 is 0.0097

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Salinity:

Figure 13. Salinity versus distance from regional fracture. There is almost no correlation as R^2 is 0.0173.

Salinity is very important to evaluate when testing the hypothesis of deeper aquifer waters entering into shallow aquifers via a large regional fracture. Due to the Moscow formation and other shale formations in this region of New York forming in a deep ocean environment, they tend to exhibit high salinity concentrations because they form deep aquifer brines. This means that if we were to see higher salinity values for water bodies closer to the regional fracture, then the deep aquifer brines high in salinity have been making their way to shallower aquifers via the distance from the regional fracture. There being little correlation, we can deduct that the regional fracture is having little influence on deeper high in saline brines make their way to shallower waters. pH:

Figure 14. pH versus Distance from regional fracture. There is almost no correlation as R^2 is 0.0251.

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Calcium:

Figure 15. Calcium vs. Distance from regional fracture. There is little to no correlation as R^2 is 0.1243

Alkalinity:

Figure 16. Alkalinity vs Distance from regional fracture. Almost no correlation is found as the R^2 value is 0.047.

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Temperature:

Figure 17. Temperature vs Distance from regional fracture. The correlation is very low with an R^2 value of 0.2699. Though this is a low correlation, it is higher than any of the other parameters tested in this study. As you can see the water samples taken further away from the regional fracture appear to be lower in temperature, but this is due to the fact that surface and groundwater data are included in the above diagram, and the highest temperatures are in Cranberry Bog during the height of summer.

Barium:

Figure 18. Barium vs Distance from the regional Fracture, the correlation is almost none as the R^2 value is 0.0368.

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Strontium:

Figure 19. Strontium vs Distance from the regional Fracture, the correlation is little to none as the R^2 value is 0.0069.

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Figure 20. This map displays shallow groundwater flow paths. This map was generated in Google Fusion with 40 meter contours.

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Figure 21. Map summarizing electrical conductivity. The small dots are representative of sample sites with electrical conductivity (E.C) of 0 to 100 μS/cm. The squares represent an E.C of 100 to 200 μS/cm, the triangles represent an E.C. of 200 to 300 μS/cm, and the circular symbol with a dot inside it represents 300 to 400 μS/cm. The line going through the middle of the map represents the regional fracture in Greenwoods. This helps to disprove the theory of the regional fracture allowing deeper aquifer waters reaching the surface, because deeper aquifer waters tend to have higher electrical conductivity. This means if the theory was true, we would see higher electrical conductivity along the regional fracture line. This does not appear to the case as 3 to 4 of the small dots representing E.C.s of 0 to 100 fall along this line.

Hydrology and Geology: This section is a comparison of the sampled water bodies that are in contact with the glacial till (such as the dug wells, streams and bog) and the sampled water bodies that are in contact with the Moscow formations.

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Upon viewing Figure 21, one can conclude that the water-rock relations of the Moscow formation are creating environments that allow for higher elemental concentrations than those in the glacial till water-rock relations. The glacial till water bodies are all at lesser depths than the Moscow formation water bodies. Most of the Moscow formation water bodies are drilled wells that penetrate to about 50 meters depth, whereas the glacial till water bodies are all within 10 meters depth as the dug wells do not go as deep. Upon viewing the visual above it appears (with the exception of chloride, which was in very small concentrations) that the higher elemental concentrations remain in the lower aquifer as these values were repeatedly taken from the same well sites. It also appears that the lower conductivity of the glacial till is allowing water to move quicker not allowing stagnant water to accumulate higher element concentrations. The Moscow formation water is not moving as fast and higher elemental concentrations are existing in these waters.

Lastly, Figure 23 compares the elemental concentrations and electrical conductivities in lakes, streams, and wells at Greenwoods Conservancy. The stagnant waters in the wells appears to be in higher elemental concentrations and higher in electrical conductivity than the quicker moving and lower water residence times of the stream and lake.

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Figure 21. A display of the concentrations of strontium, barium, calcium, chloride, and electrical conductivity for water bodies located in glacial till vs the Moscow formation.

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Figure 23. These are charts comparing the elemental concentrations and electrical conductivities in lakes, streams, and wells.

CONCLUSIONS

We find no correlation between well proximity to regional fractures and well chemistry (Figures 14 to 22.) Presence of ponds and bogs in upland regions implies low conductivity and/or high water tables. Zones of deep groundwater discharge at the surface are highly unlikely, especially in upland regions which serve as recharge zones. Water chemistry of Cranberry Bog is consistent with local shallow water wells and rules out groundwater discharge into the bog from deeper groundwater flowing up along a regional fracture zone.

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Comparing the flowlines data with the electrical conductivity map and the glacial till elemental concentrations with the Moscow formation elemental concentrations, one can conclude that the flowlines do not have as great of an influence on elemental concentrations in water bodies than the water rock relations do (Figures 20 and 21.) The water rock relations have a much greater influence because the Moscow formation water rock relations were higher in most elemental concentrations (Figure 22)

REFERENCES

APHA, AWWA, WPCF. 1989. Standard methods for the examination of water and wastewater, 17th ed. American Public Health Association. Washington, DC.

Ebina, J., T. Tsutsi, and T. Shirai. 1983. Simultaneous determination of total nitrogen and total phosphorus in water using peroxodisulfate oxidation. Water Res. 17(12):1721-1726.

Liao, N. and S. Marten. 2001. Determination of total phosphorus by flow injection analysis colorimetry (acid persulfate digestion method). QuikChem ® Method 10-115-01-1-F. Lachat Instruments, Loveland, CO.

NYS Museum, NYS Geological Survey, NYS Museum Technology Center, 1999, 1:250,000 Bedrock geology of NYS, http://www.nysm.nysed.gov/gis.html.

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Utilizing environmental DNA to identify aquatic invasive species

Lisa Newton1

INTRODUCTION

This project was designed to develop methods to assess the presence and extent of incursion of several selected aquatic invasive species in the Croton and Delaware watersheds for New York City. There is a need to determine techniques to assess whether invasive species are present and the level of invasion that are cost-effective, rapid and accurate, especially when the species are present in the reservoirs at low densities. Physical surveys and environmental DNA (eDNA) techniques will be used to identify the presence of invasive species in the Ashokan and Rondout reservoirs of the Delaware Watershed and the West Branch, New Croton and Kensico reservoirs of the Croton Watershed.

Environmental DNA is genomic DNA that is released into the environment by an organism in a variety of ways (Pilliod et al. 2013). These include epithelial tissue, feces, gametes and mucous, as well as cells that are released from dead organisms (Pilliod et al. 2013). The use of eDNA can allow a more efficient and relatively inexpensive manner to examine presence/absence of invasive species (Pilliod et al. 2013). Techniques involving eDNA may be especially useful for the early detection of unwanted non-native species, monitoring the levels of native species after invasion of non-native species, and determining the effectiveness of management techniques (Wilcox et al. 2013, Pilliod et al. 2013). A last important benefit of eDNA techniques is that they can replace the traditional survey methods used to assess the levels of invasive species in an area at a much lower cost and with a drastic decrease in overall expended effort required to determine the levels of invasive species (Wilcox et al. 2013).

While the use of environmental DNA has its advantages, it also possesses limitations. DNA obtained from the environment is often very dilute with the DNA of the target species contributing only a small percentage of the total DNA obtained from the eDNA sample, and the non-target DNA possibly being made up of closely related species (Wilcox et al. 2013). Species- specific molecular markers, or other means of differentiating the target species from genetically similar species that may be present, must therefore be developed.

The species selected for this project include Cipangopaludina chinesis, Corbicula fluminea, Dreissena polymorpha, Hydrilla verticillata, Orconectes rusticus and Myriophyllum spicatum. Each has been identified as a known invasive species of New York State and considered of interest to the NYC Department of Environmental Protection in order to determine their presence, as well as abundance, in the selected reservoirs.

Hydrilla verticillata is an aquatic plant of the family Hydrocharitacea. Hydrilla possess slim, branching stems that can grow up to 7.6 m long. Its leaves are small and narrow with a pointed tip, and grow around the stem in whorls, ranging from four to eight leaves in each whorl

1 Biological Field Station Intern, summer 2013. Funding provided by NYC DEP.

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(Ramey & Peichel 2001a). Its native range is thought to include India, Africa, Australia and portions of Asia. Hydrilla verticulata was most likely introduced to the United States through aquarium trade in Florida in the 1950s and 1960s, as well as through mail-order water lilies. Hydrilla forms dense stands of vegetation in almost any freshwater habitat (Ramey & Peichel, 2001a). This plant can grow in both lotic and lentic water, including lakes, rivers, marshes and ditches, and it can grow in a few inches of water or in water up to 6.1 m deep (Ramey & Peichel, 2001a). Hydrilla verticillata is also capable of thriving in a variety of conditions, ranging from eutrophic to oligotrophic, as well as being able to grow in a mere 1% of full sunlight, making it highly competitive (Ramey & Peichel 2001a).

Myriophyllum spicatum, also known as Eurasian water-milfoil, is a member of the Haloragaceae family (Ramey & Peichel 2001b). It has thin, branched stems that typically grow 1.8-2.7 m in length, although they can reach up to 6.1 m long (Ramey & Peichel 2001b). The leaves of M. spicatum are olive-green colored and feather-like in appearance (Ramey & Peichel 2001b). The leaves are arranged in whorls, with 3-6 leaves in each whorl, and each leaf less than 5 cm in length (Ramey & Peichel 2001b). Eurasian water-milfoil is thought to be native to Europe, Asia and northern Africa (NBII & IUCN/SSC ISSG 2006). It is suspected that this plant was introduced intentionally to the United States or was introduced in the 1800s through the release of ship ballast and was first documented in the U.S. in Washington, D.C. in 1942 (Ramey & Peichel 2001b). Myriophyllum spicatum prefers habitats which possess slow moving water, such as lakes, ponds and sluggish rivers (Ramey & Peichel 2001b). While its preference is for water movement that is slow, Eurasian water-milfoil can also grow in fast moving water (Ramey & Peichel 2001b). Like Hydrilla, M. spicatum can tolerate a wide range of habitat conditions. This plant can prosper in spring water, as well as brackish water (Ramey & Peichel 2001b). It also possess a great deal of temperature tolerance, being able to withstand the winter conditions of frozen lakes in northern areas of the U.S. and to flourish in over-heated bays in southern areas of the U.S. (Ramey & Peichel 2001b).

Dreissena polymorpha, commonly known as the zebra mussel, is a small, striped, shellfish that can grow up to 50 mm in size (Benson et al. 2013). Dreissena polymorpha will attach itself to any stable substrate in the water body, both natural and artificial, including rock, cement, boat props and other organisms living in the water. Zebra mussels are considered native to the Azov, Black and Caspian Seas (Benson et al. 2013). Dreissena polymorpha was initially documented in the United States in 1988, where it was found to be present in the Great Lakes (Benson et al. 2013). Introduction is thought to have occurred through the release of the larval stage of the zebra mussels from a ballast exchange of a commercial ship, which had travelled from the Black Sea to the Great Lakes (Benson et al. 2013). Dreissena polymorpha can withstand a variety of temperature variations and has been known to grow in temperatures ranging from 3-30˚C, with optimal growth occurring in the 20-25˚C range (Benson et al. 2013).

Orconectes rusticus, or the rusty crayfish, is a large crayfish characterized by an overall greenish blue color, large, tough claws and prominent rust colored spots on either side of its carapace (National Biological Information Infrastructure (NBII & ISSG 2010). The rusty crayfish has been known to grow up to 10 cm in length (NBII & ISSG 2010). Orconectes rusticus is native to portions of the U.S., including areas in Indiana, Illinois, Kentucky, Michigan and Ohio (NBII & ISSG 2010). Introduction of Orconectes rusticus to other areas in the United

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States has occurred through the use of the crayfish as bait by anglers, leading to its classification as an invasive species in many areas (NBII & ISSG 2010). The rusty crayfish can be found in lakes, ponds and streams, failing to display a predilection for lotic or lentic environments (NBII & ISSG 2010). A variety of substrates are acceptable for habitation of O. rusticus, including sand, silt, clay and rock (NBII & ISSG 2010). One habitat requirement for the rusty crayfish are areas that contain a variety of debris, such as rocks and submerged wood fragments that can be used to cover and protection (NBII & ISSG 2010). Often, areas with large amounts of debris are found to be preferred by the crayfish. Orconectes rusticus is often found in water bodies that have a depth of less than 1 m, but they have also been sampled in water with depths up to almost 15 m (NBII & ISSG 2010). Water that contains high levels of oxygenation is preferred by O. rusticus, and it thrives in a temperature range of 20-25˚C (NBII & ISSG 2010). While this range of temperature is preferred, the rusty crayfish can survive in temperatures as low as 0˚C and as high as 39˚C (NBII & ISSG 2010).

Corbicula fluminea, commonly known as the Asian clam, is a freshwater mollusk, with shell coloration ranging from a yellow-brown shade to black (National Biological Information Infrastructure (NBII) & IUCN/SSC Invasive Species Specialist Group (ISSG) 2005). The Asian clam can grow up to sizes of 50 mm in length and possesses uniformly spaced concentric rings on the surface of its shell (NBII & ISSG 2005). The clam is native to southeastern areas in China and Russia, as well as Korea and the Ussuri Basin (NBII & ISSG, 2005). Corbicula fluminea was first discovered in the United States in the Columbia River in Washington in 1938 (Foster et al. 2013). It was first thought to have entered the U.S. through Chinese immigrants importing the clam for consumption (Foster et al. 2013). The Asian clam is now present in at least 38 states and the District of Columbia (NBII & ISSG 2005). This magnitude of dispersal is thought to be aided by usage as bait, aquarium trade and release of ship ballast water (NBII & ISSG 2005). Corbicula fluminea can be found in lakes, streams and estuaries which can contain a range of substrates, including silt, sand and gravel (NBII & ISSG 2005). Its preference of substrate is fine clay or sand, and coarse sand that allows for burrowing (NBII & ISSG 2005). High levels of oxygenated water are required by Asian clams for suitable habitation and it can survive in temperatures that vary from 2-30˚C (NBII & ISSG 2005).

Cipangopaludina chinesis, also called the Chinese mystery snail, is a freshwater mollusk that has a shell that can be brown, reddish or greenish brown or an olive green color and can grow up to 70 mm in length (National Biological Information Infrastructure (NBII) & IUCN/SSC Invasive Species Specialist Group (ISSG) 2011). The thick shell is convex and typically contains 6 to 7 whorls (NBII & ISSG 2011). Cipangopaludina chinensis is considered native to China, Korea, Japan, Java, Myanmar, and the Philippines (NBII & ISSG 2011). It is also native to Eastern Russia, Taiwan and Vietnam (NBII & ISSG 2011). Introduction to the United States is thought to have occurred through sale in the Chinese food market in the late 1800s in San Francisco (NBII & ISSG 2011). It was first documented in Boston in 1914 and was suspected of being released sometime between 1931 and 1942 in the Niagara River (Kipp et al. 2013). The Chinese mystery snail may also have been introduced to non-native areas in a variety of other methods, such as aquarium trade, attachment to recreational boat hulls, ornamental plants shipped into the U.S. and stocked as a food source for catfish in Lake Erie (NBII & ISSG 2011). Cipangopaludina chinesis inhabits both slow-moving lotic and lentic water systems that have a substrate consisting of mud or silt (NBII & ISSG 2011). Favorable habitats include

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streams, lakes, rivers, canals and ditches (NBII & ISSG 2011). Cipangopaludina chinesis has been found to be highly resistant to desiccation, facilitating its distribution to other non-native areas through transport on boats and can also tolerate stagnant water conditions (NBII & ISSG 2011). Discovered at depths ranging from 0.2 to 3 m, the Chinese mystery snail can tolerate a temperature range of 0-30˚C and a pH of 6.5-8.4 (NBII & ISSG 2011).

METHODOLOGY

This project will involve the utilization of environmental DNA to identify the six selected invasive species of New York through the collection of water samples from five different reservoirs of the Croton and Delaware watersheds. To complete this research, several steps must be accomplished to ensure accuracy and reliability of the techniques developed. DNA extraction directly from each target organism will occur to determine whether the techniques developed to identify each selected species will produce results. To determine whether the target species is present in the selected reservoir, species-specific primers will be utilized when they are obtainable. Other further techniques will be used to determine specific species when species- specific primers are not attainable. Following the direct DNA extraction, extraction of eDNA from an artificial aquatic environment containing the organism will occur. This artificial environment will consist of a tank containing a known quantity of water harboring the target species. Following this, serial dilutions of water from the artificial environments will occur to determine the lowest density of target species at which presence can be detected. Once these levels have been determined, the techniques will be tested on water collected from water bodies that are known to be inhabited by the target species. If successful, this process will then ultimately lead to the testing of the developed techniques on the selected NYC reservoirs.

Molecular identification of each species will occur using a variety of manners. Cipangopaludina chinesis and Corbicula fluminea will be identified through the development of species-specific microsatellite markers. Extracted DNA from both species will be sent to Cornell where it will be sequenced and microsatellite markers that are specific to each species will be identified. Family specific primers designed by Theriault et al. (2003) will be utilized for identification of Dreissena polymorpha. These primers amplify the 28S rRNA gene. One limitation of the use of these primers is that they fail to distinguish between zebra mussels and another member of the dreissenid family, quagga mussels. This is not considered to be significant due to the fact that both species are considered invasive to NYS. Hydrilla verticillata will be identified through the use of species-specific primers obtained from Rybicki et al. (in press). These primers will be able to distinguish Hydrilla from other morphologically similar waterweeds, including Egeria densa, Elodea nuttallii, and Elodea canadensis. Orconectes rusticus will be identified through the use of family-specific primers developed by Taylor & Knouft (2006) which amplify Cytochrome Oxidase I. The utilization of restriction fragment length polymorphisms (RFLPs) will assist in differentiating between the different species of the family cambaridae. RFLPs use restriction enzymes that cut at known locations on the DNA creating specific lengths of fragments based on the species. The restriction enzymes that were selected for O. rusticus include NsiI, BsaHI and BsrGI. Molecular identification of Myriophyllum spicatum will also involve the use of RFLPs. Moody & Les (2002) developed primers that were effective on M. spicatum, but also were effective on other members of the

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Myriophyllum genus. To identify our target species, the restriction enzymes BsaHI and Hpy99I will be utilized.

REFERENCES Benson, A.J., D. Raikow, J. Larson and A. Fusaro. 2013. Dreissena polymorpha. USGS Nonindigenous Aquatic Species Database, Gainesville, FL. Retrieved from http://nas.er.usgs.gov/queries/factsheet.aspx?speciesid=5.

Foster, A.M., P. Fuller, A. Benson, S. Constant, D. Raikow, J. Larson and A. Fusaro. 2013. Corbicula fluminea. USGS Nonindigenous Aquatic Species Database, Gainesville, FL. Retrieved from http://nas.er.usgs.gov/queries/FactSheet.aspx?speciesID=92.

Kipp, R.M., A. J. Benson, J. Larson and A. Fusaro. 2013. Cipangopaludina chinensis malleata. USGS Nonindigenous Aquatic Species Database, Gainesville, FL. http://nas.er.usgs.gov/queries/factsheet.aspx?SpeciesID=1045

Moody, M.L. and D.H. Les. 2002. Evidence of hybridity in invasive watermilfoil (Myriophyllum) populations. Proceedings of the National Academy of Sciences, 99(23), 14867-14871.

National Biological Information Infrastructure (NBII) and IUCN/SSC Invasive Species Specialist Group (ISSG). 2005. Corbicula fluminea. Global Invasive Species Database. Retrieved from http://www.issg.org/database/species/ecology.asp?fr=1&si=278.

National Biological Information Infrastructure (NBII) and IUCN/SSC Invasive Species Specialist Group (ISSG). 2006. Myriophyllum spicatum. Reviewed by J. Clayton, Global Invasive Species Database. Retrieved from http://www.issg.org/database/species/ecology.asp?fr=1&si=278.

National Biological Information Infrastructure (NBII) and IUCN/SSC Invasive Species Specialist Group (ISSG). 2010. Orconectes rusticus. Reviewed by B. Hazlett, Global Invasive Species Database. Retrieved from http://www.issg.org/database/species/ecology.asp?si=217.

National Biological Information Infrastructure (NBII) and IUCN/SSC Invasive Species Specialist Group (ISSG). 2011. Bellamya chinensis. Reviewed by J. Olden, Global Invasive Species Database. Retrieved from http://www.issg.org/database/species/ecology.asp?si=1812&fr=1&sts=&lang=EN.

Pilliod, D.S., C.S. Goldberg, M.B. Laramie and L.P. Waits. 2013. Application of environmental DNA for inventory and monitoring of aquatic species: U.S. Geological Survey Fact Sheet 2012-3146, 4 p.

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Ramey, V. and B. Peichel. 2001a. Hydrilla verticillata. Center for Aquatic and Invasive Plants, University of Florida, IFAS, Retrieved from http://plants.ifas.ufl.edu/node/183.

Ramey, V. and B. Peichel. (001b. Eurasian water-milfoil. Center for Aquatic and Invasive Plants, University of Florida, IFAS, Retrieved form http://plants.ifas.ufl.edu/node/278.

Rybicki, N.B., J.D. Kirshtein and M.A. Voytek. In Press. Molecular techniques to distinguish morphologically similar Hydrilla verticillata, Egeria densa, Elodea nuttallii, and Elodea canadensis.

Taylor, C.A. and J.H. Knouft. 2006. Historical influences on genital morphology among sympatric species: Gonopod evolution and reproductive isolation in the crayfish genus Orconectes (Cambaridae). Biological Journal of the Linnean Society, 89, 1-12.

Therriault, T.W., M.F. Docker, M.I. Orlova, D.D. Heath and H.J. MacIsaac. 2004. Molecular resolution of the family Dreissenidae (Mollusca: Bivalvia) with emphasis on Ponto- Caspian species, including first report of Mytilopsis leucophaeata in the Black Sea basin. Moulecular Phylogenetics and Evolution, 30, 479-489.

Wilcox, T.M., K.S. McKelvey, M.K. Young, S.F. Jane, W.H. Lowe, A.R. Whiteley and M.K. Schwartz. 2013. Robust detection of rare species using environmental DNA: The importance of primer specificity. PLoS ONE, 8(3), e59520.

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A biosurvey of Allen’s Lake, Richfield Springs, NY1

Paul H. Lord

INTRODUCTION

Allen’s Lake serves as the water supply for Richfield Springs and given the lengthy underground pipe connecting the reservoir to the water treatment plant, there are concerns regarding the potential for invasive species e.g., zebra mussels (Dreissena polymorpha) blocking the pipe system. We performed a quick check to ascertain if adult zebra mussels were evident in Allen’s Lake and, while doing so, we checked for the presence of aquatic plants, macroalgae, and pearly mussels.

METHODS

I entered the lake, in the vicinity of the water intake, with SCUBA equipment, on 19 May 2013 and surveyed depths from 0 -5 m using a generalized u-shaped search pattern. I collected a sample of every plant and macroalgae species that I noted. Subsequent to the dive, we separated the plants and macroalgae and confirmed identifications.

RESULTS

We identified 13 plants and one macroalgae in our survey (Table 1).

Table 1. Plants and macroalgae in Allen’s Lake Reservoir.

1 Funding provided by Otsego County SWCD dive fund.

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DISCUSSION

The presence of water marigold and northern watermilfoil attests to the protections provided Allen’s Lake. The one exotic noted was curly leaved pondweed which was likely introduced by the Canadian geese (Branta canadensis) using the reservoir.

Additionally, I noted a healthy population of eastern floater (Pyganodon cataracta) pearly mussels and no zebra mussels (Dreissena polymorpha) or quagga mussels (Dreissena rostriformsi bugensis).

Given the shallow morphology of Allen’s Lake and the proximity of waters holding zebra mussels and water thyme (Hydrilla verticillata), we recommend increased vigilance to unauthorized uses of Allen’s Lake.

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Chronological field observations at various BFS sites, 2013

William L. Butts

Date Location Notes V.1.2013 Greenwoods/Thayer Greenwoods; Canada Goose (10), Hooded Merganser♂,♀;Mallard ♂ Thayer; Ring-necked duck 7♂,2♀-Pond below UIC, Wild Turkey(7)no beards, red on wattles, all feeding, no obvious interaction. V.2.2013 Thayer Pond below UIC-Mallard ♂,♀, Ring-necked duck 5♂,4♀ V.15.2013 Greenwood/Thayer Greenwoods; Canada Goose(3); Merganser ( prob. Red-Breasted); Mallard(3♂). Thayer-No birds on pond below UIC V.15.2013 Greenwoods/Thayer Greenwoods-No birds Lower Mill Pond; Canada Goose & 3 goslings; Hooded Merganser ♂,♀ on Upper Mill Pond. Thayer- Crow (4 lg) in field across road from UIC V.22.2013 Thayer Checked ground pools below pond edge, Pond edge both sides boat launch, puddles in road, water levels low - all impoundments, upper step pond. No larvae. Sampled outer edge off embankment of Pond below UIC, Pond edge both sides boat launch – no larvae. Step ponds – no larvae. V.28.2013 Thayer Checked ground pools around pond below UIC, pond edge both sides boat dock, puddles in path, blow- down hole – no larvae any site. V.29.2013 Impoundments None VI.5.2013 Greenwood/ Thayer Greenwoods –No birds any impoundment Thayer-UIC –both sides from boat launch, no larvae, small Ephemeropteran naiads. Damage from Pileated Woodpecker activity. VI.12.2013 Thayer/ Greenwoods Thayer-UIC pond edge both sides boat launch – no larvae, low area below pond berm, standing water, water nearly gone < 2± inches deep –heavily silted,1 small pupa in blow-down hole by road VI.13.2013 Thayer Drove through to gate in heavy rain. No birds on pond. Wild Turkey (8 lg) on road – fled to woods. VI.19.2013 Thayer Check ground pools below pond edge/ pond edge both sides boat launch, puddles in road, upper step pond. No larvae any site.

VI.25.13 Greenwoods No waterfowl in any impoundment; Upper Mill Pond largely covered w/ algal mat. Set

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trap in low area beyond pond drainage; 2 mosquitoes approached – one attempted to feed. VII.2.2013 Upper Site, Greenwoods Upper Site: Set light trap at site beside path to boat launch, steady light rain falling – no mosquitoes approached. Drove through Greenwoods – Canada Goose(14) on upper Mill Pond. VII.9.2013 Upper Site Checked pond edge and trail for larva, none collected. Blow down hole only site with sufficient water to support larvae. No mosquitoes approached. VII.16.2013 Upper Site Set light trap at edge of upper step pond. VII.17.2013 Upper Site / Greenwoods No catch in light traps – No birds any impoundment VII.30.2013 Upper Site / Greenwoods Set trap at step pond site. Canada Goose (2) on upper mill pond at Greenwoods. Large area of both ponds with vegetative cover. VII.31.2013 Upper Site Picked up light trap – very little catch. VIII.6.2013 Greenwoods Drove through impoundment behind grange drained, covered with herbaceous growth, lower mill pond nearly covered with vegetative growth with some erect emergent plants. Upper Mill pond nearly covered ( ~ 80%) with vegetative growth. Virtually no open water in pond above cross road. Significant change in habitat. No waterfowl. Most water surface with vegetative cover. VIII.14.2013 Upper Site / Greenwoods Walked trail to lower trap site with side trip to boat launch. No mosquitoes approached. No waterfowl on any impoundment. Apparent increase in extent of floating (some rooted) vegetation. VIII.16.2013 Upper Site / Greenwoods No waterfowl. On pond below Interpretive Center. Very little standing water at any site below pond edge. Some sites might support Anopheles spp., cattail stand appears to be enlarging, wood duck eclipse ♂ (3) on pond. Drove through Greenwoods. No waterfowl observed. Available open water greatly reduced. VIII.20.2013 Upper Site / Greenwoods Walked to boat launch and lower trap site with right arm fully exposed. No mosquitoes approached. Drove through Greenwoods: no birds on any impoundment. Vegetative cover appears to be expanding. Not much open water. VIII.26.2013 Upper Site / Greenwoods No waterfowl visible form road at either site IX.4.2013 Upper Site / Greenwoods No waterfowl on pond below Interpretive Center. Checked pond edge both sides boat launch. No larvae. Low area below pond beam with no standing water. Most of ground surface not notable damp. Greenwoods: No waterfowl. Increased vegetative growth.

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IX.10.2013 Upper Site / Greenwoods No waterfowl on pond below Interpretive Center. Greenwoods: Black Duck and 3 smaller ( probably Gadwall) IX.12.2013 Upper Site / Greenwoods No waterfowl on pond below Interpretive Center. Whitetail deer 7-2 antlered, one with rack well above ears. Greenwoods: 2 ducks on lower mill pond, Canada Goose (6) on upper mill pond. IX.26.2013 Heavy Ground fog. No sightings X.3.2013 Upper Site / Greenwoods No Sightings X.4.2013 Upper Site / Greenwoods No Sightings X.7.2013 Upper Site / Greenwoods No Sightings X.10.2013 Upper Site / Greenwoods No Sightings

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Dynamics of Galerucella spp. and purple loosestrife (Lythrum salicaria) in Goodyear Swamp Sanctuary, summer 2013 update

H.A. Waterfield

INTRODUCTION

The distribution and effectiveness of Galerucella spp. populations as a biocontrol agent of purple loosestrife (Lythrum salicaria) were monitored within Goodyear Swamp Sanctuary as part of an ongoing monitoring regime that began in 1997. Annual spring monitoring of the impact of Galerucella spp. on purple loosestrife is updated in this report. Details of the history of this study can be found in Albright et al. (2004); the concise summary below was presented in Albright 2013.

L. salicaria is an emergent semi-aquatic plant that was introduced into the United States from Eurasia in the early 19th century (Thomson 1987). It is an aggressive and highly adaptive invasive species which inhabits wetlands, flood plains, estuaries and irrigation systems. Once established, purple loosestrife often creates monospecific stands, displacing native species including cattails (Typha spp.), sedges (Carex spp.), bulrushes (Scirpus spp.), willows (Salix spp.) and horsetails (Equisetum spp.). Recent efforts, which include both chemical application and the use of biocontrol methods, have focused on controlling L. salicaria where stands impede well-diversified wetland communities (Thomson 1987).

In June 1997, 50 adults each of Galerucella calmariensis and G. pusilla were introduced into Goodyear Swamp Sanctuary (N42°48.6’ W74°53.9), located at the northeastern end of Otsego Lake (Austin 1998). The beetles were initially released in cages from sites 1 and 2 (Figure 1). In 1998, sites 3-5 were introduced into the study in order to monitor the distribution of Galerucella over time to other stands of purple loosestrife (Austin 1999). Sampling sites were established to monitor the qualitative and quantitative effects of the beetles on purple loosestrife and also to examine the extent of any recovery by the native flora (Austin 1998). It was expected that these beetles would lessen the competitive ability of purple loosestrife by feeding upon their meristematic regions, resulting in defoliation, impaired growth, decreased seed production, and increased mortality (Blossey et al. 1994).

METHODS

Spring monitoring was performed according to protocols established by Blossey et al. (1997). Observations of the insects and plants were made within the five 1m2 quadrats, marked by four visible stakes (Figure 1).

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Figure 1. Map of Goodyear Swamp Sanctuary showing sampling sites. Sites 1 and 2 are 1997 Galerucella spp. stocking sites; sites 3-5 were established to evaluate the spread of Galerucella spp. within the Sanctuary over time.

Spring monitoring was completed on 23 May 2013. This first assessment is typically completed within 2-3 weeks after overwintering adults appear (Blossey 1997). Galerucella spp. abundance was estimated in each life stage (egg, larva, adult) according to the established abundance categories (Table 1). The number of stems of L. salicaria within each quadrat were counted, and the five tallest were measured. The percent cover of L. salicaria and the percent damage attributable to Galerucella spp. were both estimated according to established frequency categories. Fall monitoring, which was completed on 21 August 2013, consisted of the same metrics measured in the spring monitoring along with measurements to gauge the vigor of L. salicaria plants, including the number of inflorescences per plant and per quadrat, as well as the number of flowers per inflorescence.

Table 1. Categories prescribed by Blossey’s (1997) protocol for reporting abundance and frequency categories.

Abundance Categories Frequency Categories Number category range category mid point 0 1 0% A 0% 1-9 2 1-5% B 2.5% 10-49 3 5-25% C 15% 50-99 4 25-50% D 37.5% 100-499 5 50-75% E 62.5% 500-1000 6 75-100% F 87.5% >1000 7 100% G 100%

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RESULTS & DISCUSSION

Most monitoring data are represented by abundance and frequency categories, defined in Table 1. Changes between these frequency categories from year-to-year or plot-to-plot can represent a substantial change in abundance due to the broad ranges covered by each category (Albright 2004). It should be noted that the actual number of L. salicaria stems are presented in the following results, while all other metrics are categorical. Variation in the number of stems between years or plots may not correspond with a shift in percent cover category, due to the above-stated lack of sensitivity that is inherent in a categorical classification scheme.

Spring Monitoring (23 May 2013) Eggs of the Galerucella beetle were present in four of the five of the quadrats at moderate densities (Figure 2). Larvae were not found in any quadrat, which is consistent with typical conditions since 1998 but is in contrast with larval abundances observed in 2012 (Figure 3). Adult beetles were found in 2 of 5 quadrats at moderate densities (Figure 4).

6

5

4

3

2 Abundance category 1 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5

Figure 2. Comparison of Galerucella spp. egg abundance from yearly spring samplings. Abundance categories taken from Table 1. 6

5

4

3

2

Abundance category 1 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5

Figure 3. Comparison of Galerucella spp. larval abundance from yearly spring samplings. Abundance categories taken from Table 1.

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6

5

4

3

2

1 Abundance category 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5

Figure 4. Comparison of Galerucella spp. adult abundance from yearly spring samplings. Abundance categories taken from Table 1.

Lythrum salicaria abundance, based on the number of stems in the quadrat, was at the lowest level since monitoring began in 1998 (Figure 5). Estimated percent cover was also as low as has ever been recorded, and for the second time was not present in two of the five quadrats (Figure 6). The loosestrife that was in the quadrats was damaged by herbivory (Figure 7).

100 90 80 70 60 50 40 30

Number Number Stemsof 20 10 0 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5

Figure 5. Comparison of the number of purple loosestrife stems from yearly spring sampling observations.

70

- point 60 50 40 30 20 10

Frequency Category Mid 0 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5 Figure 6. Comparison of percent cover estimates by purple loosestrife from yearly spring samplings. Frequency category mid points derived from Table 1.

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70

- point 60 50 40 30 20 10

Frequency Category Mid 0 1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5

Figure 7. Comparison of percent damage estimates to purple loosestrife leaves from yearly spring samplings. Frequency category mid points derived from Table 1.

Fall Monitoring (21 August 2013)

The number of L. salicaria stems and estimated percent cover was low again in fall 2013, marginally higher than in 2012 (Figures 8 and 9, respectively). No inflorescences (flower clusters) were recorded in the quadrats, though individual stems of L. salicaria were in bloom elsewhere in the swamp.

Galerucella are host-specific and as such feed exclusively on purple loosestrife. This characteristic results in a beetle population that is directly dependent upon loosestrife densities within the swamp. Abundance patterns observed within the swamp since 1998 illustrate the population dynamics of host-specific organisms and their dependency upon host populations (Fagan et al. 2002). It appears that once Galerucella spp. became established in the Sanctuary in about 2001, control of purple loosestrife has been effective, with abundance at a fraction of what it had been prior to the onset of biocontrol efforts.

120

100

80

60

40

20 NA

Number Number Stemsof NA 0 1997 2000 2001 2002 2003 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5

Figure 8. Number of purple loosestrife stems per quadrat during fall monitoring, 1997, 2000- 2013. Flooding in fall 2011 precluded sampling.

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100 90 point - 80 70 60 50 40 30 20 10 NANA Frequency Category Mid Category Frequency 0 1997 2000 2001 2002 2003 2005 2006 2007 2008 2009 2010 2011 2012 2013

quadrat 1 quadrat 2 quadrat 3 quadrat 4 quadrat 5 Figure 9. Estimated percent cover (category midpoints) of purple loosestrife during fall monitoring, 1997, 2000-2012. Categories as presented in Table 1. Flooding in fall 2011 precluded sampling.

CONCLUSIONS

Spring 2013 monitoring indicated that L. salicaria abundance continues to be less (based on percent cover and number of stems) than in most year since monitoring began in 1997, while enough exists to sustain a Galerucella population, as evidenced by the moderate abundance of both eggs and larvae. Fall monitoring reveals that Galerucella spp. are effective at controlling not only the abundance of L. salicaria, but also the overall vigor and fitness based on reduced plant height and failure to produce flowering bodies. Observations related to the presence of Galerucella spp. at sites outside of Goodyear Swamp Sanctuary (i.e., Lydon 2008) indicate that the dispersal of Galerucella spp. continues from the original site and it shows promising potential as a biological agent against the invasive plant.

REFERENCES

Albright, M.F. 2013. An update on the dynamics of Galerucella spp. and purple loosestrife (Lythrum salicaria) in Goodyear Swamp Sanctuary, summer 2012. In: 45th Ann. Rept. (2012) SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Albright, M.F., W.N. Harman. S.S. Fickbohm, H.A. Meehan, S. Groff and T. Austin. 2004. Recovery of native flora and behavior responses by Gallerucella spp. following biocontrol of purple loosestrife. Am. Midl. Nat. 152:248-254.

Austin, T. 1998. Biological control of purple loosestrife in Goodyear Swamp Sanctuary using Galerucella spp., summer 1997. In 30th Ann. Rept. (1997). SUNY Oneonta. Biol. Fld. Sta., SUNY Oneonta.

Austin, T. 1999. Biological control of purple loosestrife in Goodyear Swamp Sanctuary using Galerucella spp., summer 1998. In 31st Ann. Rept. (1998). SUNY Oneonta. Biol. Fld. Sta., SUNY Oneonta.

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Blossey, B. 1997. Purple loosestrife monitoring protocol, 2nd draft. Unpublished document. Dept. of Natural Resources, Cornell University.

Blossey, B., D. Schroeder, S.D. Hight and R.A. Malecki. 1994. Host specificity and environmental impact of two leaf beetles (Galerucella calmariensis and G. pusilla) for the biological control of purple loosestrife (Lythrum salicaria). Weed Science. 42:134-140

Fagan, W.F., M.A. Lewis, M.G. Neubert, P. van den Driessche. 2002. Invasion theory and biological control. Ecology Letters 5(1) 148.

Groff, S. 2001. Biological control of purple loosestrife (Lythrum salicaria) in Goodyear Swamp Sanctuary using leaf-eating beetles (Galerucella spp.), summer 2000. In 34th Annual Report (2000). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Lydon, J.C. 2008. Monitoring the dynamics of Galerucella spp. and purple loosestrife (Lythrum salicaria) in the Goodyear Swamp Sanctuary and along the shorelines of Otsego, Weaver and Youngs Lakes, summer 2007). In 40th Ann. Rept. (2007). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Meehan, H.A. 2006. Biological control of purple loosestrife (Lythrum salicaria) in Goodyear Swamp Sanctuary using leaf-eating beetles (Galerucella spp.), summer 2005. In 38th Annual Report (2005). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Rubenstein, M. 2010. Monitoring the dynamics of Galerucella spp. And purple loosestrife (Lythrum salicaria) in the Goodyear Swamp Sanctuary, summer 2009. In 42nd Ann. Rept. (2009). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Snyder, C.M. 2007. Monitoring the dynamics of Galerucella spp. and purple loosestrife (Lythrum salicaria) in the Goodyear Swamp Sanctuary and along the Otsego Lake shoreline, summer 2006. In 39th Annual Report (2006). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Thompson, Daniel Q., R.L. Stuckey, E. B. Thompson. 1987. Spread, Impact, and Control of Purple Loosestrife (Lythrum salicaria) in North American Wetlands. U.S. Fish and Wildlife Service. 55 pages. Jamestown, ND: Northern Prairie Wildlife Research Center Online. http://www.npwrc.usgs.gov/resource/plants/loosstrf/loosstrf.htm (04JUN99).

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Walleye (Sander vitreus) movement and depth utilization in response to changes in alewife (Alosa pseudoharengus) abundance in Otsego Lake

John R. Foster1 & Daniel J. Drake2

Abstract: The goal of this study was to determine if walleye movement and habitat utilization differed between years of high and low alewife abundance. Walleye locations were recorded over 24 hour periods using ultrasonic telemetry transmitters implanted in 9 fish from 2007-2008 (high alewife abundance) and 6 fish from 2011-2012 (low alewife abundance). Walleyes utilized the deeper open water sections of the lake and moved greater distances when alewives were abundant. Years when alewives were scarce, walleye spent nearly 50% of their time in shallow, weedy inshore areas where they had little movement. Changes in walleye movement and habitat utilization demonstrate a behavioral response to forage availability and location that has not been documented before.

INTRODUCTION

There have been numerous studies of habitat utilization and movements of walleye in lakes and reservoirs using radio and ultrasonic telemetry (Foust and Haynes 2007, Holt et al. 1977, Palmer et al. 2005, Hanson 2006). With the exception of studies carried out in Otsego Lake (Stich et al. 2008, Byrne et. al. 2009, Potter et al. 2010) these earlier studies have primarily been focused on shallow, weedy, warm-water reservoirs (Ager 1976, Ross and Winter 1975, Schlagenhaft and Murphy 1985, Johnson et al. 1988, Munger 2002) and lakes (Holt et al. 1977, Nate et al. 2003, Foust and Haynes 2007). From these studies, differences have been reported in walleye habitat utilization and movements. Even when the same stocked Oneida Lake strain of walleye have been studied in Honoeye Lake (Foust and Haynes 2007) and nearby Otsego Lake (Byrne et al. 2009) substantial differences in behavior have been observed. In Honoeye Lake, the Oneida strain of walleye seldom moved and was confined to shallow, weedy littoral waters. However, in Otsego Lake, stocked Oneida strain walleye made use of virtually all lake habitats, and made substantial movements into deep open water.

While observed differences in walleye behavior may be primarily due to differences in lake morphometrics and limnology, another possibility is that they may be due to differences in the forage base. Alewives became the principle forage species in Otsego Lake within a few years after their introduction in 1986 (Foster 1990) and were the primary forage of walleye in Otsego Lake (McBride and Cornwell 2008). However, recently the abundance of alewives in Otsego Lake has decreased considerably (Waterfield and Cornwell 2013). Alewife density was close to 4,000 fish per ha in 2007-2008, but had dropped to 11 - 58 fish per hectare in 2011- 2012. In this study we will examine the hypothesis that when alewives were abundant walleye exhibit pelagic feeding behavior, moving considerable distances in open water to locate and feed on alewives. However, with low alewife abundance in recent years, walleye would cease to be an active, open water predator and would become more sedentary in shallow, near-shore weedy areas as described by Foust and Haynes (2007).

1 Professor & Chair, Fisheries, Wildlife & Environmental Science Department, SUNY Cobleskill, Cobleskill, NY. 2 Fisheries & Aquaculture Student, SUNY Cobleskill, Cobleskill, NY.

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In order to test the hypothesis that the reduction in alewife population resulted in reductions in walleye movements and utilization of deep, open waters, walleye were tracked in 2011-2012 and compared to data from 2007-2008. It was hypothesized that with the reduction in pelagic forage in 2011-2012, walleyes would be more sedentary and make greater utilization of shallow, weedy near-shore habitats than they did in 2007-2008.

MATERIALS & METHODS

Otsego Lake (42°40’ N, 74°55’ W) is located in Otsego County, New York at an elevation of 1190 ft. The lake has a surface area of 1,711 ha, the physical traits of an oligotrophic lake, but the water chemistry of a mesotrophic lake. It is a long narrow lake, with a length of 13.28 km and a mean width of 1.28 km (Figure 1). It is deep and steep sided with a maximum depth of 50.5 m and a mean depth of 24.9 m (Harman et al. 1997).

Figure 1. Bathymetric map of Otsego Lake, Otsego County New York.

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In Otsego Lake, recent studies using hourly diel sampling of walleye positions have provide a much more accurate characterization of walleye habitat utilization and movements than previous studies based on daily or weekly sampling (Stich et al. 2008; Byrne et al. 2009). Therefore, in this study, only fish position data collected over a 24 hour period were used. Fish position data collected from previous studies (Stich et al. 2008, Byrne et al. 2009) were used as well as data from fish implanted with transmitters in 2009, 2010 and 2012 (Table 1). The diel data set for 2007-2008 consisted of 371 hourly fish positions, in fifteen 24-hour sampling periods for nine individual walleye (Table 1). The 2011-2012 diel data set consisted of 329 fish positions, in fourteen 24-hour sampling periods for six different fish.

As part of this study, walleye were tagged in April 2012 with Sonotronics© DT-97-L transmitters using methods adapted from Paragamian (1989). Walleye were tracked by boat and located using a Sonotronics© DH-4 directional hydrophone and Sonotronics© USR-96 receiver. Fish position was determined when the hydrophone could be turned 360° without attenuation of signal strength. Coordinates of fish positions were determined with a Garmin® GPS unit. ArcMap 10 was used to find water depth at each location. MS Excel was used to analyze data.

Table 1. Tagging information (year tracked, year tagged, carrier frequency, pulse interval, fish length and fish sex) for diel tracks used in this study.

Year Year Tagged Carrier Pulse Length Sex Tracked Frequency Interval 2007, 2008 2006 73 910 NA Female 2008 2006 76 1040 NA Female 2007 2006 71 1010 NA Male 2007 2006 69 970 NA Male 2008 2007 74 960 NA Female 2007, 2008 2007 70 860 NA Female 2007, 2008 2007 77 1230 NA Female 2008 2008 69 1210 NA Male 2008 2008 71 1230 NA Male 2011 2009 74 1020 NA Female 2011, 2012 2010 77 990 561 Male 2011, 2012 2010 81 1030 586 Male 2012 2012 76 1080 597 Female 2012 2012 71 1030 431 Female 2011, 2012 2012 73 890 632 Female

RESULTS

Walleye utilized different areas of the lake when alewives were abundant than when they were scarce (Figure 2). In 2007-2008 during high alewife abundance, walleyes were located in the deepest portion of the lake (150-168 ft) 47% of the time. However, in 2011-2012 during periods of very low alewife abundance walleyes were located in the shallowest portion of the lake (0-25 ft) 49% of the time (chi square test, P < .005).

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60% High Alewife Abundance (2007-2008) 50% Low Alewife Abundance (2011-2012)

40%

30%

20% Percent Occurrence Percent 10%

0% 0-25 25-50 50-75 75-100 100-125 125-150 150-168 Water Depth (ft)

Figure 2. Walleye depth utilization during years of high and low alewife abundance.

Walleye distances travelled were also considerably different between the periods of high and low alewife abundance (Figure 3). On average walleye moved 4.9 m/minute when alewives were abundant, but only 1.9 m/min. when alewives were scarce (T-test, P < .005). Walleye were stationary during 1 hour observation periods twice as often when alewives were scarce than when alewives were abundant. Walleye were also 3 times as likely to move more than 4 m/min when alewives were abundant (28%) than when alewives were scarce (9%)

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45% High Alewife Abundance (2007-2008) 40% Low Alewife Abundance (2011-2012) 35% 30% 25% 20% 15% 10% Percent Occurrence 5% 0% 0-1 1-2 2-3 3-4 4-5 5-6 6-7 7-8 8-9 9-10 >10 Movement (m/min)

Figure 3. Distances walleye travelled during periods of high and low alewife abundance.

DISCUSSION

The hypothesized behavioral response of walleye to forage availability and location was as predicted. During low alewife abundance walleye movement was significantly reduced. Walleye stayed in the shallowest part of the lake nearly half the time when alewives were scarce. However, when alewives were abundant they spent nearly half their time in the deepest waters of the Otsego Lake. While walleye behavior has been studied in Otsego Lake as well as other lakes and reservoirs, none of these earlier studies have documented a change in activity and habitat utilization in response to reductions in a forage species abundance.

Optimum forage theory suggests that walleyes in the lake would expend the least amount of energy possible to make the greatest possible gains in nutrition (Beck 1993). When the number of alewives decline in open water, it would be a waste of energy for walleye to actively forage in those waters. Staying more sedentary in shallow water closer to shore would require less energy expenditure in order to obtain food. A follow-up study should be conducted to determine if walleye have indeed changed their diet to littoral zone species.

Walleye have been reported to confine themselves to shallow, weedy littoral waters in some lakes (Foust and Haynes 2007), while utilizing deeper open waters in other lakes and reservoirs (Ager 1976; Festa et al. 1987; Palmer et al. 2005). Perhaps as suggested in this study, these differences in habitat utilization and movement have more to do with the density of the forage base than lake morphometric or limnological characteristics. Walleye behavioral responses to a change in their main forage species indicate that they are an adaptable and resilient species.

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The lake trout population in Otsego Lake must also be impacted from the reduction in alewives and further studies should examine their feeding response, diet and population dynamics. While walleyes have shown that they can adapt to a change in prey species, lake trout cannot feed in the shallow weedy areas during summer when the lake becomes thermally stratified. Reduction of the alewife population may therefore have a much greater impact on the lake trout population of Otsego Lake, than on the walleye population.

Acknowledgements

Data utilized in this study were collected by numerous SUNY Cobleskill’s Fisheries & Aquaculture students: Dan Stich, John Byrne, Justin Potter, Ben German and Justin Hulbert. Henry Whitbeck was a great help in aiding with GIS application.

REFERENCES

Ager, L.M. 1976. A biotelemetry study of the movement of walleye in Center Hill Reservoir, Tennessee. Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies. 30:311-323.

Beck, J.E. 1993. Foraging theory and piscivorous fish: Are forage fish just big zooplankton? Transactions of the American Fisheries Society 122(5): 902-911.

Byrne, J.M., D.S. Stich, and J.R. Foster. 2009. Diel movements and habitat utilization of walleye (Sander vitreus) in Otsego Lake. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Stat., SUNY Oneonta.

Cornwell, M.D. and N.D. McBride. 2008. Walleye (Sander vitreus) reintroduction update: Walleye stocking, gill netting and diet analysis 2007. In 40th Ann. Rept. (2007). SUNY Oneonta Bio. Fld. Stat., SUNY Oneonta.

Festa, P.J., J.L. Forney, and R.T. Colesente. 1987. Walleye management in New York State. NYSDEC Bureau of Fisheries. 104p.

Foster, J. R. 1990. Introduction of the alewife (Alosa pseudoharengus) in Otsego Lake. In 22nd Ann. Rept.(1989) SUNY Oneonta Bio Fld. Stat., SUNY Oneonta.

Foust, J.C. and J.M. Haynes. 2007. Failure of walleye recruitment in a lake with little suitable spawning habitat is probably exacerbated by restricted home ranges. Journal of Freshwater Ecology 22(2):297-304.

Hanson, J.R. 2006. Seasonal Movement Patterns of Walleye (Sander vitreus) in Muskegon River and Muskegon Lake, Michigan. School of Natural Resources and Environment: The University of Michigan. 1-36.

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Harman, W.N., L.P. Sohacki, M.F. Albright and D.L. Rosen. 1997. The state of Otsego Lake 1936-1996. Occass. Pap. No. 30. SUNY Oneonta Biol. Fld. Stat., SUNY Oneonta.

Holt, C.S., G.S.D. Grant, G.P. Oberstar, C.C. Oakes, and D.W. Bradt. 1977. Movement of walleye, Stizostedion vitreum, in Lake Benmidji, Minnesota as determined by radio- biotelemetry. Transactions of the American Fisheries Society 106(2):163-169.

Johnson, B.L., D.L. Smith, and R.F. Carline. 1988. Habitat preferences, survival, growth, foods and harvests of walleyes and walleye x sanger hybrids. North American Journal of Fisheries Management. 8: 292-304.

Munger, C. 2002. A review of walleye management in Texas with emphasis on Meridith Reservoir. North American Journal of Fisheries Management. 22: 1064-1075.

Nate, N.A., M.A. Bozek, M.J. Hansen, C.W. Ramm, M.T. Bremigan and S.W. Hewett. 2003. Predicting the occurrence and success of walleye populations from physical and biological features of northern Wisconsin Lakes. North American Journal of Fisheries Management. 23: 1207-1214.

Palmer, G.C., B.R. Murphy and E.M. Hallerman. 2005. Movements of walleyes in Claytor Lake and the Upper New River, Virginia, indicate distinct lake and river populations. North American Journal of Fisheries Management 25(4):1448-1455.

Paragamian, V.L. 1989. Seasonal habitat use by walleye in a warmwater river system, as determined by biotelemetry. North American Journal of Fisheries Management 9(4):392- 401.

Potter, J., J. M. Byrne, D.S. Stich and J.R. Foster. 2010. Walleye (Sander vitreus) Seasonal Activity And Habitat Utilization In Otsego Lake, New York. In 43rd Ann. Rept. SUNY Oneonta Biol. Fld. Stat., SUNY Oneonta.

Ross, M.J. and J.D. Winter. 1981. Winter movements of four fish species near a thermal plume in Northern Minnesota. Trans. American Fisheries Society. 110:14-18.

Schlagehaft, T.W. and B. R. Murphy 1985. Habitat use and overlap between adult largemouth bass and walleye in a West Texas Reservoir. North Am. J. of Fisheries Management 5(3) 465-470.

Stich, D.S., B. Decker, J. Lydon, J. Byrne and J.R. Foster. 2008. Summer diel habitat utilization of walleye in Otsego Lake, NY. In 40th Ann. Rept. (2007). SUNY Oneonta Bio. Fld. Stat., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2013. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2012. In 45th Ann. Rept, (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Hydroacoustic survey of Otsego Lake’s pelagic fish community, spring 20131

Holly A. Waterfield2 and Mark Cornwell3

INTRODUCTION

A hydroacoustic survey was conducted in June 2013 to estimate pelagic fish abundance in Otsego Lake (Otsego County, NY). Historically, hydroacoustic surveys were conducted to assess alewife abundance. Following their introduction in 1986, alewife (Alosa pseudoharengus) quickly became the most abundant forage fish in Otsego Lake (Foster 1990) and had major impacts on trophic relationships, chlorophyll a, water clarity, and hypolimnetic oxygen depletion rates (Warner 1999, Harman et al. 2002). Walleye (Sander vitreus) stocking resumed in 2000 (previously stocked through 1934) to establish a recreational fishery while at the same time introducing a predator which would potentially control the alewife population. The population of spawning adult walleye has steadily increased since 2008; in 2012 the population was estimated at 10,520 fish (Willson et al. 2013). Increased predation by this growing population in combination with winter predation by lake trout have drastically reduced the alewife population and led to changes in lake conditions that suggest reversal of the alewife-induced trends previously described. However, the establishment of zebra mussels in Otsego Lake in 2007 has confounded efforts to directly attribute recovery of certain trophic measures directly to decreased alewife abundance. The acoustic survey detailed in this report is part of an ongoing effort to document changes in the pelagic fish community and lake trophic condition. Comparisons are made to the results of past hydroacoustic surveys (1996 to 2007) which were summarized by Brooking and Cornwell (2008) and those from 2009 to 2013 (Waterfield and Cornwell 2011, 2012, 2013). Previous reports were focused on estimating alewife abundance, as this was the dominant forage fish in the open water community; 2012 and 2013 results are not species- specific due to uncertainty associated with the composition of acoustic targets, though consistent criteria have been used in the analysis to produce comparable results.

METHODS

A single acoustic survey was conducted on the night of 05 June 2013 according to standard procedures (Parker-Stetter 2009). Beginning 1-2 hours after dark, the survey was conducted from north to south following a zig-zag pattern from shore to shore (Figure 1). Approximately east-west transects were connected with a diagonal “zig” to yield two sets of parallel transects for additional analysis of methods and statistical approaches. Both down- looking and side-looking data were collected using a BioSonics DtX echosounder with 123kHz 7.5o beam and 120kHz 7.8o beam transducers, respectively; data collection settings for each transducer are listed in Table 1. Performance of each transducer was checked against a standard tungsten carbide sphere; no calibration offsets were used.

1 Made possible by 2007 NSF grant entitled Acquisition of hydroacoustic and associated instrumentation for fisheries research (DBI: 0722764). 2 Research Support Specialist, SUNY Oneonta Biological Field Station, Cooperstown, NY. 3 Assistant Professor, SUNY Cobleskill Department of Fisheries and Wildlife, Cobleskill, NY.

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Figure 1. Bathymetric map of Otsego Lake, NY with acoustic transects surveyed 05 June 2013.

Table 1. Data collection settings for 05 June acoustic survey conducted on Otsego Lake, NY.

Transducer Number of Frequency Average Survey Pulse Duration Ping Rate Orientation transects (kHz) Speed (m/s) (ms) (pps) Vertical - Down 12 123 1.9 0.4 3 Horizontal 12 120 1.9 0.4 3

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Hydroacoustic data were analyzed using the Sonar5-Pro post processing system (Sonar5) following conversion of the raw acoustic files to Sonar5-compatible formats (Balk and Lindem 2007). Surface and bottom exclusion zones were established on each echogram to isolate open water areas for analysis. The surface exclusion zone was established from the surface to 2.0m to eliminate data from the transducer’s near-field and surface noise. The bottom boundary included a margin of approximately 0.5 m above the actual detected bottom to avoid inclusion of bottom echoes in abundance estimates. Echograms were visually inspected to remove echoes from targets other than fish (i.e., submerged vegetation). Passive noise, calculated from passive listening data collected prior to the survey, was subtracted from all files before analysis. Fish density (#/ha) was calculated using Sv/TS scaling method in Sonar5 (Balk and Lindem 2007) based on a minimum in situ target strength threshold of -55 dB. In past analyses the TS threshold was -61dB; the higher thresholds of -55dB was used to exclude an abundance of small targets from the alewife abundance estimate (Figure 2). Target strengths of -61 to -55 dB have been associated with alewife ranging in length from 1.5 to 2.0 cm (Warner 2002), a size class that would not be present at the time of the 2013 survey. Each transect was divided into layers for analysis based on areas of relatively homogeneous fish distribution. Abundance estimates were determined for predator-sized (TS >-35 dB) and prey-sized (TS -55dB to -36dB) targets in each layer. Total fish abundance and predator abundance were each based on the proportion of targets in each TS range; forage fish abundance was based on estimates from the surface through the bottom of the metalimnion. Layers used in the analysis were 2-14m and 14m+.

Figure 2. Target strength distribution (in dB) for targets detected during the 05 June 2013 hydroacoustic survey of Otsego Lake. Variable small mesh experimental gill nets were set concurrent with each survey. Nets were 21 meters long by 6 meters deep and composed of seven 3-meter wide panels of different mesh sizes (6.2, 8, 10, 12.5, 15, 18.7, and 25mm bar mesh). Three nets were set on the night of 05 June 2013 for about 12 hours, set just before dark, pulled early morning. Fish were tallied by panel and vertical position (top, middle, or bottom) in the net. Species, length, and weight were recorded for all fish caught.

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RESULTS AND DISCUSSION

Lake-wide epilimnetic fish density in June was estimated to be 25 fish/ha (95% CI +/- 15.7 fish/ha) based on the 12 transects (Table 2); this is lower than most recent estimates of alewife-only abundance (Table 3, Figure 3). 2013 spring survey results are not alewife-specific due to uncertainty related to the composition of targets contributing to forage fish abundance estimates. Table 2 contains 2013 estimates of epilimnetic fish (all fish and forage-only) and lake- wide predators within each transect. Figure 3 illustrates spring and fall alewife abundance estimates since 1996. Estimates of forage fish varied among transects ranging from 0 fish/ha to 73 fish/ha; this range of density estimates is less than those reported in 2012 (Waterfield and Cornwell 2013). A high percentage of targets were of low-intensity (-80 to -60 dB); this range is often associated with fry of smelt, alewife, coregonids, and percids. Though the presence of fry was not confirmed during the survey, percid fry were collected in the weeks prior (Waterfield and Albright 2014b).

Gill netting efforts yielded multiple lake trout in the net fishing 8-14m depth, with no fish caught in the nets fishing from 0-6m and 4-10m. If this catch is representative of lake-wide alewife distribution, it is possible that the acoustic-derived abundance is an over-estimate of the actual alewife abundance in the water column. Increased effort to ground-truth the acoustic data is necessary in order to identify the organisms contributing to spring abundance estimates. A fall survey was not conducted in 2013, though a summary of historical fall survey data is presented below for reference (Table 4) and is included in Figure 3. The decrease in estimated alewife abundance agrees with other data collected since 2008, including gill net catch for 2011-2013 surveys, as well as results documented in other ongoing and past studies, including increased mean alewife size concurrent with decreased trap net catch rate (Potter 2010, German 2012), only 2 individual alewife being recorded in 2012 (none recorded in 2013, Stowell 2014) and the past prevalence of alewife in stomachs of captured walleye (McBride and Cornwell 2008). Other trophic indicators measured in and calculated for Otsego Lake also point toward changes that are directly related to the decreased alewife abundance including increased mean size of cladoceran zooplankton (Tanner and Albright 2014) and reduced areal hypolimnetic oxygen depletion rates (Waterfield and Albright 2014a). Definitive conclusions regarding the “success” of the walleye stocking program in balancing the lake’s trophic condition have been precluded by the establishment of zebra mussels in Otsego Lake and the resultant trophic changes, however, it is undeniable that the walleye have an impact on the alewife population. Additional changes observed in lake trout and walleye reflect the marked decrease in alewife abundance. Fitness and condition of lake trout decreased in 2012, as documented by Region 4 DEC Fisheries gill net survey (Wells 2013). Angler reports indicate uncharacteristic foraging by lake trout in shallow waters during the late fall and winter, with stomach contents including crayfish, pickerel, and walleye fingerlings. Anecdotal observations of stomach contents indicate that walleye are targeting species other than alewife as abundance remains low.

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Table 2. Fish abundance estimates of epilimnetic forage fish and predator-sized targets throughout the water column for transects surveyed 05 June 2013, including lake-wide mean, standard error (SE) and 95% confidence intervals.

epilimnetic whole-lake No. of all fish forage predator Transect SEDs (fish/ha) (fish/ha) (fish/ha) 1 1 21 1 20 2 0 0 0 0 3 2 47 35 13 4 2 12 6 7 5 2 22 11 10 6 10 82 73 8 7 4 31 31 0 8 3 21 21 0 9 2 14 14 6 10 0 0 0 0 11 5 52 52 5 12 0 0 0 0 Mean 25 21 5.9 Standard Error 7.1 6.8 1.8 Standard Deviation 24.7 23.6 6.4 Confidence Level (95.0%) 15.7 15.0 4.0

Table 3. Forage fish abundance estimates (fish/ha) for spring surveys conducted between 2004 and 2012 with number of transects, standard deviation (SD) and 95% Confidence Intervals (CI) Prior to 2012, forage fish abundance estimates were specific to alewife; since 2012 forage fish estimate is not species-specific.

Year Fish/ha # transects SD 95% CI 2004 907 9 175 114 2005 236 9 214 137 2006 2522 10 1463 907 2007 1330 9 611 399 2010 174 14 236 137 2011 78 8 52 43 2012 901 11 921 619 2013 25 12 24 16

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Table 4. Abundance estimates (fish/ha) from fall acoustic surveys conducted between 1996 and 2012. Modified from Brooking and Cornwell (2008). Forage fish estimates are assumed to be representative of alewife abundance.

Fall Forage Fish Abundance Fall Predator Abundance Year #/ha # transects SD 95% SE #/ha # transects SD 95% SE 1996 5170 7 1434 1063 7.5 7 4.2 3.1 1997 2053 9 798 521 3.3 9 3.4 2.2 2000 1382 8 925 774 2001 8562 9 3811 2490 35.2 9 13.9 9.1 2002 10901 16 4886 2394 15.2 16 10.7 5.2 2003 3851 16 2901 1421 1.2 16 1.5 0.7 2004 2418 9 1571 1026 3.5 9 4.7 3.1 2005 9562 9 3555 2322 8.6 9 8.8 5.7 2006 1631 7 2713 2010 19.4 7 25.6 19 2007 3921 11 2524 1492 6.5 11 5.7 3.4 2009 369 18 334 166 6.6 18 6.4 3.17 2010 275 16 464 247 2.9 16 4.3 2.3 2011 58 11 85 57 4.1 11 6.9 3.3 2012 11 10 16 11 2.7 10 6.5 4.6

12000

10000

8000

6000

Fish Density Density (#/ha)Fish 4000

2000

0

spring fall

Figure 3. Forage fish abundance estimates based on analysis of spring and fall acoustic surveys from 1996 through 2013. Prior to 2012, all values were specific to alewife abundance.

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REFERENCES

Balk, H. and T. Lindem. 2007. Sonar4 and Sonar5 post processing systems, operator manual version 5.9.7, 420p. Lindem Data Acquisition Humleveien 4b. 0870 Oslo Norway.

Brooking, T.E. and M.D. Cornwell. 2008. Hydroacoustic surveys of Otsego Lake, 2007. In 40th Ann Rept. (2007). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Cornwell, M.D. and N.D. McBride. 2009. Walleye (Sander vitreus) reintroduction update: Walleye stocking, gill netting 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

German, B.P. 2012. Summer 2011 trap net monitoring of fish communities utilizing the weedy littoral zone at Rat Cove and rocky littoral zone Brookwood Point, Otsego Lake. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Foster, J.R. 1990. Introduction of the alewife (Alosa pseudoharengus) into Otsego Lake. In 22nd Ann. Rept. (1989). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Harman, W.N., M.F. Albright, and D.M. Warner. 2002. Trophic changes in Otsego Lake, NY following the introduction of the alewife (Alosa psuedoharengus). Lake and Reservoir Management. 18(3)215-226

Parker-Stetter, S.L., Rudstam, L.G., Sullivan, P.J., and Warner, D.M. 2009. Standard operating procedures for fisheries acoustic surveys in the Great Lakes. Great Lakes Fish. Comm. Spec. Pub. 09-01.

Potter, J. 2010. Littoral fish community survey of Rat Cove & Brookwood Point, summer 2009. In 42nd Ann. Rept. (2009). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Tanner, C. and M.F. Albright. 2014. A survey of Otsego Lake’s zooplankton community, summer 2013. In: 45th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Warner, D.M. 1999. Alewives in Otsego Lake, NY: a comparison of their direct and indirect mechanisms of impact on transparency and chlorophyll a. Occas. Pap. No. 32. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Warner, D.M., Rudstam, L.G., and Klumb, R.A. 2002. In situ target strength of alewives in freshwater. Transactions of the American Fisheries Society. 131(2): 212-223.

Waterfield, H.A. and M.F. Albright. 2014a. Otsego Lake limnological monitoring 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.F. Albright. 2014b. Otsego Lake fry sampling, 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Waterfield, H.A. and M.D. Cornwell. 2011. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2010. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2012. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2011. In 44th Ann. Rept. (2011). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A. and M.D. Cornwell. 2013. Hydroacoustic surveys of Otsego Lake’s pelagic fish community, 2012. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Wells, S.M. 2013 Personal Communication.

Willson, J., J.R. Foster, M.D. Cornwell. 2013. 2012 population estimate of walleye (Sander vitreus) in Otsego Lake. In 45th Ann. Rept. (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Natural recruitment of lake trout (Salvelinus namaycush) in Otsego Lake

Nicholas M. Sawick1 & John R. Foster2

Abstract: This study was conducted to determine if natural recruitment of lake trout was still occurring in Otsego Lake after zebra mussels have carpeted their spawning shoals. Emergent fry traps were placed on traditional lake trout spawning shoals at Bissel Point to capture lake trout fry as they emerge from the substrate. Thirteen fry were captured in 2013, which was significantly less than the number captured in 2003 and 2004, before zebra mussels were introduced, but more than captured in 2011 and 2012. More studies have to be conducted to determine the factors impacting the drop in natural lake trout recruitment in Otsego Lake.

INTRODUCTION

Lake trout spawning shoals are made up of angular, rocky substrates with deep interstitial spaces (Beauchamp et al. 1992, Nester & Poe 1987). During spawning, lake trout eggs drop into the interstitial spaces between the rocks, where they are protected from current, wave, ice scouring and wind action (Edsall et al. 1989; Dorr et al. 1981), as well as from a variety of fish and invertebrate egg predators (Claramunt et al. 2005, Marsden 1997, Tibbits 2007). Zebra mussels, which were first discovered in Otsego Lake in 2007 (Waterfield 2009) now cover almost every hard surface in the lake, including historic lake trout (Salvelinus namaycush) spawning shoals. When lake trout spawning beds are covered by large colonies of zebra mussels, eggs cannot drop into the interstitial spaces between the rocks, potentially resulting in higher egg mortality and lower recruitment from natural reproduction.

Although Otsego Lake is stocked with hatchery reared lake trout, recruitment of lake trout into the Otsego Lake fishery is largely dependent on natural reproduction ( McBride and Sanford 1997, Tibbits 2007). The most recent documentation of successful lake trout recruitment was made before zebra mussels had colonized Otsego Lake (Tibbits 2007). More recent studies in 2011 and 2012 following the introduction of zebra mussels were unable to document successful lake trout recruitment from the same Bissel Point shoals studied by Tibbits (J.R. Foster pers. com.). In this study, natural lake trout recruitment was again examined at Bissel Point.

The goal of this study was to determine if and to what extent natural lake trout recruitment was still occurring on the traditional lake trout spawning shoal at Bissel Point studied by Tibbits (2007). In order to meet this goal, fry traps were deployed in early spring to capture and enumerate the lake trout fry emerging from the Bissel Point spawning shoal. Unsuccessful or lower lake trout fry emergence may provide strong evidence that zebra mussels are impacting lake trout recruitment in Lake Otsego.

1 Fisheries & Aquaculture Student, SUNY Cobleskill, Cobleskill, NY. 2 Professor & Chair, Fisheries, Wildlife & Environmental. Science Department, SUNY Cobleskill, Cobleskill, NY.

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MATERIALS & METHODS

This study was conducted just off Bissel Point, Otsego Lake (W74° 54.141; N42° 45.550, Town of Otsego, Otsego County, New York), at the same site used in Tibbits’ (2007) study (Figure 1). Fry traps were set near shore along a transect line perpendicular to the shoreline out to approximately 1 meter in depth. Three traps were used in each of the 4 transect lines. The traps were centered at depths of 30, 60 and 90 cm along each transect.

Figure 1. Study location, Otsego Lake, Otsego County, NY.

The fry traps used in this study followed the design of Chotkowski et al. (2002) and were the same emergent fry traps used by Tibbits in his 2003-2004 study. Each circular trap had a diameter of 81 cm and covered a bottom area of .52 m2. In order to anchor the traps to the substrate, rocks were placed on the bottom ring. Foam blocks with a hole drilled in the center were placed under the jar for extra floatation. As they fry swim out of the substrate to gulp air, they swim up into the bottle and are trapped inside (Figure 2).

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Figure 2. Soft mesh fry emergence traps used in this study.

The study began immediately after the ice came off the lake in the spring of 2013. The traps were set on 23 April and removed on 21 May. This was the same time frame used by Tibbits (2007) and corresponded to the natural emergence of lake trout fry in Otsego Lake. Ice- out on Otsego Lake was late in 2013 due to the harsh winter.

The traps were checked 3 times a week either by boat or shore access. The collection bottles at the top of the traps were checked for fry and other fish. All fry were captured unharmed and released back into the lake. The weather conditions, water temperature and clarity were measured each time the traps were checked.

RESULTS

Thirteen lake trout fry were captured in this study. Most of the fry (7) were captured at 30 cm deep traps (Figure 3). Five fry were captured in 60 cm of water and only 1 fry was captured in 90 cm of water.

While emergence traps were set for 29 days in this study, lake trout fry only emerged between 23 April and 7 May (Figure 4). In earlier studies, fry emergence occurred over a longer period. In 2003 fry emergence occurred from 25 April to 15 May and in 2004 from 25 April to 19 May.

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60.0

50.0

40.0

30.0

% of Emergent % of Emergent Fry 20.0

10.0

0.0 30 60 90 Water Depth in cm

Figure 3. Percentage of 2013 lake trout emergent fry captured at three water depths.

In 2003 fry emergence had a nice bell shaped curve with a peak occurring on 5 May (Figure 4). The emergence pattern for 2004 and 2013 was much more sporadic.

60

2013 50 2003 2004 40

30

20

# Fry Caught in Emergence Emergence FryTraps Caught in # 10

0

5/1 5/2 5/3 5/4 5/5 5/6 5/7 5/8 5/9 4/25 4/26 4/27 4/28 4/29 4/30 5/10 5/11 5/12 5/13 5/14 5/15 5/16 5/17 5/18 5/19 5/20 5/21 Figure 3: Lake trout fry emergence by date inDate Otsego Lake for 2003, 2004 and 2013.

Figure 4. The number and date emerging lake trout fry were captured in 2003, 2004 and 2013 off the Bissel Point, Otsego Lake.

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In 2003 and 2004, significantly more lake trout fry emerged than in 2013 (chi square test, P < .001). In 2003, 173 lake trout fry were captured and in 2004, 43 lake trout fry were captured, compared to only 13 fry in 2013. The number of fry captured per m² of substrate per day provides a better comparison, particularly when the data is limited to the length of the season fry were captured. In 2013, 1.59 fry were captured per m² per day, a much lower capture rate than 3.44 fry/ m²/day captured in 2003 and the 3.96 fry/ m²/day captured in 2004.

DISCUSSION

Emerging lake trout fry were successfully captured during this study. Therefore, natural reproduction of lake trout occurred on their historic Bissel Point shoal in 2012 and their eggs and fry survived to swim-up in 2013. Natural lake trout recruitment did occur even though zebra mussels covered the substrate. However, lake trout fry were captured in much higher numbers in 2003 and 2004 before zebra mussels colonized the lake (Tibbits 2007).

The difference in recruitment does not appear to be due to low numbers of lake trout spawners. NYSDEC gill net catches (NYSDEC 2013) of lake trout spawners (fish 21 inches or larger) was 3 fish per net in 2002 (spawning season for fry to emerge in 2003) compared to 5 fish per net in 2012, (the spawning time for fry to emerge in 2013).

The low numbers of emerging lake trout fry in 2013 may indicate that zebra mussels are impacting lake trout recruitment. The presence of zebra mussels on the spawning shoals has been shown to interfere with the deposition of eggs, as well as their survival. The presence of zebra mussels reduces egg deposition by discouraging adult lake trout from spawning. Further zebra mussels increases damage to lake trout eggs (Marsden and Chotkowski 2001), as well as vulnerability of eggs to predators (Claramunt et al. 2005, Marsden 1997, Tibbits 2007).

Low recruitment in 2011-2013 could be due to a variety of factors other than the zebra mussels. There were significant yearly variations in recruitment that occur in Tibbits (2007) and other studies (Marsden and Chotkowski 2001). By spawning in very shallow water at Bissel Point, lake trout eggs would be very vulnerable to ice scour and wave action (Edwards et al. 1990), which would differentially impact recruitment from year to year.

Another factor in this study was vandalism. Thirty-nine trap days were lost because traps were pulled out of the water and thrown on shore. The traps were back in the water during the dates of peak fry emergence in 2003 and 2004, however, vandalism was a major setback to the research and could have negatively impacted the results.

The ultimate cause for lack of lake trout recruitment at Bissel Point in 2011 and 2012, and the low recruitment in 2013 is still in question. More studies need to be conducted to determine the ultimate cause of low lake trout recruitment in Otsego Lake. If the downward trend in natural recruitment continues, than additional stocking would be necessary to prevent a substantial decline in Otsego Lake’s lake trout fishery.

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ACKNOWLEDGEMENTS

The SUNY Oneonta Biological field station provided boats and traps and Matthew Albright provided guidance and assistance to this study. SUNY Cobleskill students Corey Sullivan, Eric Malone, Nick Madderom, Justin Sheward, Shaun Story and John Dooly helped set and check the traps. The Schoharie County Conservation association provided the senior author with a research grant to carry out this study.

REFERENCES

Beauchamp D. A. , B. C. Allen, R. C. Richards , W. A. Wurtsbaugh and C. R. Goldman. 1992. Lake Trout spawning in Lake Tahoe: egg incubation in deepwater macrophyte beds. North American Journal of Fisheries Management 12:442-449.

Chotkowski M.A., E.J. Marsden and B. Ellrot. 2002. An inexpensive modified emergent fry trap. N. Amer. J. Fish. Manage. 22: 621-624.

Claramunt, R. M., J. L. Jonas, J. D. Fitzsimons and J. E. Marsden. 2005. Influences of spawning habitat characteristics and interstitial predators on Lake Trout egg deposition and mortality. Transactions of the American Fisheries Society 134:1048-1057.

Dorr, J. A., D. V. O’Connor, N. R. Foster and D. J. Jude. 1981. Substrate conditions and abundance of Lake Trout Eggs in a traditional spawning area in southeastern Lake Michigan. North American Journal of Fisheries Management 1:165-172.

Edwards, C. J., R. A. Ryder, and T. R. Marshall. Using lake trout as a surrogate of ecosystem health for oligotrophic waters of the Great Lakes. 1. Great Lakes Res. 16(4):591-608.

Marsden, J. E. 1997. Common carp diet includes zebra mussels and lake trout eggs. Journal of Freshwater Ecology 12: 491-492.

Marsden, J. Ellen & Michael A. Chotkowski. 2001. Lake trout spawning on artificial reefs and the effect of zebra mussels: fatal attraction? Journal of Great Lakes Research 27:1 33– 43.

McBride, N.D. and D.K. Sanford. 1997. An interim fish management plan for Otsego Lake. New York State Department of Environmental Conservation, Region 4 Fisheries Office, Stamford, New York: 112 pp.

Nester, R. T. and T. P. Poe. 1987. Visual observations of historical Lake Trout spawning grounds in western Lake Huron. North American Journal of Fisheries Management 7:418-424.

NYSDEC. 2013. Summary of salmonid gill net catches in Otsego Lake from 1969-2012. Unpublished data from the New York Dept. of Env. Consvn. Bureau of Fisheries, Region 4 Stamford, NY 12167 USA.

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Tibbits, W. T. 2007. The behavior of lake trout, Salvelinus namaycush, in Otsego Lake: A documentation of the strains, movements and the natural reproduction of lake trout under present conditions. Occas. Pap. #42. SUNY Oneonta Biological Field Station, SUNY Oneonta.

Waterfield, H.A. 2009. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake, 2008. In 41st Ann. Rept. (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Monitoring the Moe Pond ecosystem and population estimates of largemouth bass (Micropterus salmoides) post unauthorized introduction Stephen G. Stowell1 ABSTRACT A study of Moe Pond, located in Otsego County, NY, at N42°43.00’W74°56.75’ (Albright et. al 2004) was conducted to evaluate the water quality, estimate a population for largemouth bass (Micropterus salmoides) through mark and recapture, and analyze the stomach contents of the largemouth bass within the pond. Bass abundance was also estimated using seining techniques, as has historically been conducted at Moe Pond. Water temperature average was 20.7°C, average conductivity was 0.068 (mS/cm), pH averaged 7.6, average oxidation-reduction potential (ORP) was 64.67, average DO (%) was 67.43, DO (mg/L) average was 6.35, and the average Secchi disk reading was 2.33m. The estimated population of largemouth bass within Moe Pond was 4,205, down from 2012’s estimate of 6,480 bass (VanDerKrake 2013). The stomach contents of 40 largemouth bass showed mostly Daphnia spp., damselfly larvae, and amphipods.

INTRODUCTION As part of ongoing research through the SUNY Oneonta Biological Field Station (BFS), Moe pond in Cooperstown, NY (Figure 1), was monitored for several weeks. Water quality, turbidity, chlorophyll a, and fish abundance, diet and growth were studied. At the start of the study, data collected from 1972, 1994, 2000-2008, and 2012 had been previously compiled. Continuing the study in 2013 enables the BFS to add to the long-running data set for Moe Pond and notice any trends. The goal of the study was to compare current water quality with past as well as compare the fish community and abundance of largemouth bass to prior findings. Prior to 1998, seining in Moe Pond revealed that the only fish species in the water body were brown bullhead (Ictalurus nebulosus) and golden shiner (Notemigonus crysoleucas) (McCoy et al. 2001). In 1998 or 1999, largemouth bass (Micropterus salmoides) and smallmouth bass (M. dolomieu) were illegally introduced. The disappearance of the golden shiner population led to an increase in zooplankton mean size and abundance and increased algal grazing (Albright et al. 2004). Monitoring since has revealed conditions varying between dominance by algae and rooted macrophytes, presumably driven by trophic changes.

1 Robert C. MacWatters Internship in the Aquatic Sciences, summer 2013. Present affiliation: Department of Fisheries and Wildlife Technology, SUNY Agriculture and Technical College, Cobleskill, NY.

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Figure 1. Topographic map showing the location of Moe pond and Otsego County, NY.

METHODS AND MATERIALS On a weekly basis from 4 June 2013 to 3 July 2013, water quality was assessed at Moe Pond’s deepest point (Figure 2). A YSI® multiprobe was used to record temperature, conductivity, pH, oxidation-reduction potential (ORP), dissolved oxygen (%), and dissolved oxygen (mg/L) in profile. A Secchi disk reading was also taken each week. Water samples were collected and taken back to the lab for nutrient and chlorophyll a analysis. The fish community was evaluated via haul seine and electrofishing, as described below. A 200ft haul seine was deployed using a john boat in a teardrop shape with the bag at the top of the teardrop, furthest from shore. The net was pulled to shore, and the bag was lifted out of the water and the fish were transferred to totes. The study only focused on largemouth bass, and brown bullhead (Ameiurus nebulosus, the only other species collected) were returned to the pond immediately. The length of each largemouth bass was recorded. The first 10 fish over 150mm had a gastric lavage performed. A syringe with a 4in piece of aquarium hosing attached was filled with water. The hose was gently placed down the throat of the fish until resistance was felt. The fish was held inverted over the opening of a Whirl-Pak. The water was pushed from the syringe into the fish’s stomach. This caused the fish to regurgitate any forage in to the Whirl- Pak. The bag was labeled with the date and fish’s length, preserved with ethanol, and stored in the cooler until it was processed. Prior to being released, each fish received a caudal fin clip to identify it as having been collected. This was partly done to mark fish for an upcoming recapture effort to calculate an abundance estimate.

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Figure 2. Water quality sampling location at Moe Pond, Otsego County, NY (modified from Sohaki 1972).

A mark and recapture analysis was also performed at Moe Pond to further evaluate largemouth bass abundance. Using an electrofishing boat, courtesy of SUNY Cobleskill and Mark Cornwell, the entire perimeter of the pond was shocked on 14 July 2013 and all largemouth bass were netted using scap nets. The fish lengths were measured and recorded, a partial pelvic fin clip was performed and the fish were released back into the pond. On 24 July 2013 the same team of people shocked the entire perimeter of the pond again for the recapture for the same amount of seconds and the same number of amps and D/C volts. All largemouth bass were netted and put into the live well. The lengths were recorded and each fish was examined for the partial pelvic fin clip that was used to mark fish in the previous survey. A data sheet containing numbers of clipped and non-clipped fish was kept; data were used in the following formula (N = MC/R) to derive an estimate of the largemouth bass population within Moe Pond, where M = number of fish originally marked, C = the sample size at the time of recapture, R = number of marked fish at the time of recapture (Cornwell 2013).

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RESULTS AND DISCUSSION

The water quality values from the YSI from 4 June 2013 to 3 July 2013 are shown in Table 1. Data show that the temperature steadily increased from the beginning to June to the beginning of July. Conductivity and pH stayed fairly constant at around 0.068 mS/cm and 7.6, respectively. Dissolved oxygen near the bottom occasional was <1.0 mg/l, likely following calm periods during which mixing was not entraining oxygen as fast as it was being lost through respiration. In Table 2, results from the analyses of nutrients, chlorides, calcium and chlorophyll a are shown. All parameters are in line with values of recent years. The Secchi disk readings were all greater than 2 meters. Compared to previous years of studies conducted on Moe Pond, transparency has greatly increased since 1992 (Table 3). This was likely a direct result of the absence of golden shiners. Larger zooplankton, such as daphnia, effectively graze on algae. Therefore, prevalence of planktivorous fish in the waterbody can greatly influence algae, which in turn effects transparency. Initially, golden shiner reduced daphnia abundance, while more recently daphnia have been a major food item in young largemouth bass in Moe Pond (i.e., VanDerKrake 2013).

Table 1. Water quality parameters of Moe pond between 4 June 2013 and 3 July 2013.

Date Depth Temp. Sp. Cond. pH ORP DO DO TURB. Secchi (oC) (us/cm) (% sat.) (mg/l) (NTU) (m) 6/4/2013 0 21.39 0.053 8.75 91 101.1 8.91 3 N/A 1 21.38 0.053 8.29 90.4 100.2 8.85 3.1 2 16.47 0.056 8.26 87.1 92.3 9.01 4.9 2.5 15.32 0.143 7.27 -177.1 26.8 2.68 4 6/12/2013 0 18.93 0.056 7.77 154.1 87.1 8.08 3.7 2.3 1 18.72 0.056 7.61 164.6 84.1 7.86 3.6 2 18.34 0.056 7.54 167.3 85.6 8.04 3.7 2.5 18.46 0.055 7.36 128.5 84.9 9.57 3.9 6/20/2013 0 20.95 0.055 8.15 152.6 98.1 8.75 N/A 2.3 1 20.05 0.054 7.63 165.8 94.4 8.57 N/A 2 19.71 0.054 7.43 137.8 89.8 8.27 N/A 2.3 19.54 2056 6.97 131.4 68.4 6.17 N/A 6/27/2013 0 25.73 0.055 7.32 33.4 91.2 7.52 0.6 2.5 1 24.45 0.055 8.18 83.3 94.5 7.7 0.4 2 22.18 0.064 7.5 102.7 15.3 1.26 27.2 2.7 20.08 0.151 6.99 -194 4.2 0.36 5.7 7/3/2013 0 24.66 0.051 8.03 106.9 73.6 6.11 N/A 2.3 1 23.59 0.054 7.34 124.4 51.2 4.3 N/A 2 21.87 0.083 6.65 -84.4 3.2 N/A N/A 2.3 21.39 0.098 6.41 -106.6 2.6 0.23 N/A

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Table 2. Nutrient Data of Moe pond between 4 June 2013 and 27 June 2013. Total Date Nitrite+nitrate Total nitrogen phosphorus Chlorophyll a (mg/l) (mg/l) (µg/l) (µg/l) 6/4/2013 < 0.02 0.32 22 6/12/2013 < 0.02 0.30 21 6/20/2013 < 0.02 0.27 17 2.16 6/27/2013 < 0.02 0.36 20 6.85 7/3/2013 < 0.02 0.23 23 2 7/12/2013 < 0.02 0.36 46 7.77

Table 3. Average values of Secchi depth, total phosphorus, nitrate+nitrite, and chlorophyll a (bd=Below detectable limits) (Modified from VanDerKrake 2013).

1972 1994 2000 2001 2002 2003 2004 2005 2006 2007 2008 2012 2013 Secchi Depth (m) NA 0.85 1.2 1.1 2.2 2.33 1.26 1.26 2.2 2.62 1.35 2.24 2.33 Total Phosphorus (µg/l) 40-70 36.7 NA NA 26.4 29.05 42.29 56.64 26.91 20.5 28.95 26.33 20 Nitrate+nitrite (mg/L) NA <.05 NA NA 0.14 0.11 0.1 0.01 0.01 <.01 0.003 bd bd Chlorophyll a (ppb) NA 2.2 0.2 8.1 2.4 2.49 4.4 2.41 19.4 2.75 3.94 4.99 4.69

The stomach contents of 40 largemouth bass collected with the haul seine were analyzed in the lab (Table 4). The most common forage found in the stomachs was daphnia. This could suggest that the pond’s Secchi disk readings would start to decrease as more bass forage on daphnia. Other popular forage for the bass was damselfly larvae and pupae as well as amphipods. Three stomachs on 20 June 2013 contained brown bullhead fry (Table 4). Brown bullhead spawn in late spring early summer and incubate for roughly 20 days before hatching (Werner 2004). Because bass are piscivorous, one would expect to encounter more bass containing other fish in the stomach. Overpopulation of the pond likely plays a major role in such omnivorous diets for the bass of Moe Pond.

- 232 - Table 5. Stomach Contents of 40 Largemouth Bass from Moe pond between 4 June 2013 and 27 June 2013 .

4-Jun-13 Length (mm) Brown Bullhead fry Crayfish (Pieces) Copepod Daphnia Midge Amphipod Aquatic Worm Mayfly Larvae Dragonfly Larvae Beetle Damselfly Pupae Damselfly Larvae Grasshopper Fly Backswimmer

215 - 1 6 1 ------214 - - 2 - 4 1 ------226 - - - - 1 2 2 ------110 - - - 182 2 1 ------204 - - - - - 6 - 1 ------219 - - 2 2 - 1 ------269 - - - - 1 - - - 1 ------276 - - 1 ------1 - - --- 264 - - - - 1 - - - - - 1 1 --- 262 - - 1 2 1 - - - 1 ------12-Jun-13 Length (mm) Brown Bullhead fry Crayfish (Pieces) Copepod Daphnia Midge Amphipod Aquatic Worm Mayfly Larvae Dragonfly Larvae Beetle Damselfly Pupae Damselfly Larvae Grasshopper Fly Backswimmer 250 - 1 - - 1 - - - 1 1 1 3 --- 223 - - - 1 1 - - - 1 - - 1 --- 212 - - 2 ------3 - --- 211 ------8 2 --- 237 - - - - 2 ------2 --- 251 - - 1 - 1 - - - - - 3 - --- 239 ------2 - - 12 ---

- 233 218 - - 1 - 2 - - - 1 - - 1 1 - - 171 ------184 ------1 - --- 20-Jun-13 Length (mm) Brown Bullhead fry Crayfish (Pieces) Copepod Daphnia Midge Amphipod Aquatic Worm Mayfly Larvae Dragonfly Larvae Beetle Damselfly Pupae Damselfly Larvae Grasshopper Fly Backswimmer 151 - - - - 4 63 - - - - 6 - --- 189 - - - 31 ------2 - - 2 - 192 - - - 213 -- - - 3 - 6 - --- 179 1 ------1 ---

264 13 - - - 5 - - - - - 3 3 --- 46th AnnualReportoftheBiological Field Station 219 - - - 67 - 3 - - - - - 1 --- 206 1 - - 113 4 - - - - - 3 - --- 221 - - - 11 ------169 - - - 6 ------3 - --- 187 - - - 7 - 9 - - - - 4 3 --- 27-Jun-13 Length (mm) Brown Bullhead fry Crayfish (Pieces) Copepod Daphnia Midge Amphipod Aquatic Worm Mayfly Larvae Dragonfly Larvae Beetle Damselfly Pupae Damselfly Larvae Grasshopper Fly Backswimmer 210 ------2 --- 212 - - - 6 1 ------114 ------135 - - - 3 5 - - - - - 3 - --- 189 ------2 --- 147 - - - 13 3 ------132 - - --- 10 - - 1 - - - - - 1 209 - - - - - 3 - - - - - 2 --- 147 ------1 134 ------1 ---

Totals 40 15 2 16 658 39 99 2 1 11 2 47 37 1 2 2 46th Annual Report of the Biological Field Station

In past years, largemouth bass abundance was estimated through the area extrapolation method. The area seined was estimated to be 300 m². The number of bass caught per seine was divided by the area seined, and that number of fish per m² was then multiplied by 155,800m², the area of Moe Pond (Reinicke 2006). Though this method is not considered to accurately estimate abundance, it was considered a proxy of abundance (Lopata 2004). For this study, a comparison between the area extrapolation method and the mark and recapture method was evaluated to see how the two methods compared. Table 5 shows that after finding the number of bass in Moe Pond based on each haul seine, an average was taken and the population estimate was 13,560 Largemouth bass in Moe Pond.

Table 5. Area extrapolation methods for haul seines on Moe Pond between 4 June 2013 and 27 June 2013.

Date 4-Jun 12-Jun 20-Jun 24-Jun 24-Jun 24-Jun 25-Jun 25-Jun 27-Jun Total bass caught in Seine 81 22 36 12 19 15 13 24 13 Area of Seine (m2) 300 300 300 300 300 300 300 300 300

Fish per m2 0.27 0.07 0.12 0.04 0.06 0.05 0.04 0.08 0.04 Area of Moe Pond (m2) 155800 155800 155800 155800 155800 155800 155800 155800 155800 # of bass in Moe Pond/seine 42066 11425 18696 6232 9867 7790 6751 12464 6751 Average # of Bass in Moe Pond 13560

A second population estimate used for comparison was mark and recapture. Ninety six largemouth bass received a caudal fin clip after being caught in the haul seine. From the 96 fish clipped, only 2 were recaptured while using the electrofishing boat. The Peterson mark and recapture formula N = MC/R was used to estimate a population of 21,600 largemouth bass in Moe pond (Table 6). This number was about 7000 fish higher than the area extrapolation number. However, the low number of recaptured fish (about 2%) provides results having low precision and high potential error (Quinn and Deriso 1999).

Table 6. Population estimate of Moe pond through the Peterson mark and recapture method, using the haul seine (mark) and electrofishing boat (recapture).

Peterson Mark and Recapture (Haul Seine)

# of fish originally marked(M) 96 Sample size taken at time of Recapture(C) 450 # of marked fish at time of recapture(R) 2

Estimate of population in Moe Pond 21600

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The third method used to derive a population estimate of Moe pond was the use of the electrofishing boat to both mark and recapture fish. On the first night of shocking, 14 July 2013, 450 largemouth bass were shocked, given a pelvic fin clip, and released back to the water. The second night shocking, 24 July 2013, focused on recaptures. The same route around the pond was taken, but shocking was done for only 2180 seconds, and a total of 271 fish were shocked. Of those 271, 29 had a pelvic fin clip from the previous week’s shocking. Using the same Peterson mark and recapture formula, N = MC/R, a population estimate of ~4205 largemouth bass was calculated for Moe pond (Table 7).

Table 7. Population estimate of Moe pond through the Peterson mark and recapture method, using the electrofishing boat only.

Peterson Mark and Recapture (E-Boat)

# of fish originally marked(M) 450 Sample size taken at time of Recapture(C) 271 # of marked fish at time of recapture(R) 29

Estimate of population in Moe Pond 4205

The total fish population of Moe Pond has been estimated for several years. Table 8 shows that golden shiners were extirpated around 2001 and smallmouth bass around 2006. Largemouth bass populations grew from 1999 to around 2007 and then decreased significantly. This could be due to the high competition and lack of forage. Further studies with determine if the population of largemouth bass continues to decrease.

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Table 8. Populations of golden shiner, largemouth bass, and smallmouth bass for 1994, 1999- 2008, and 2012-2013.

Year golden shiner largemouth bass smallmouth bass (Notemigonus (Micropterus (Micropterus crysoleucas) salmoides) dolomieu) 1994 (McCoy et al., 2000) 7,154: +12,701;-6,356 0 0 1999 (Wilson et al., 2000) 3,210+/-1760 1,588+/-650 958+/-454 2000 (Tibbits, 2001) 381+/-296 2,536+/-1,177 945+/-296 2001 (Wojnar, 2002) 1,708+/-1,693 3,724+/-3,447 504+/-473 2002 (Hamway, 2003) 3 206 20 2003 (Hamway, 2004) 2 318 1 2004 (Lopata, 2005) 0 6,924+/-2,912 0 2005 (Dresser, 2006) 0 12,019+/-3,577 223+/-257 2006 (Reinicke & Walters, 2007) 0 11,555.17+/- 0 2007 (Underwood, 2008) 0 13,373+/-249 0 2008 (Finger, 2009) 0 46,740+/-13,220 0 2012 (VanDerKrake 2013) 0 6,480+/-1,533 0 2013 (current, seining) 0 13,560 0 2013 (current, mark-recap) 0 4205 0

CONCLUSION Through comparisons of long running data sets, 2013 Moe Pond water quality was fairly consistent with 2012 averages. Phosphorus and chlorophyll a were lower than 2012, hinting that there has been less algal growth in 2013. The large mats of Elodea sp. could also be taking up the nutrients first and leaving less in the water samples tested. Related to decreased algae, Secchi disk readings increased from 2012, as expected with the greater abundance of zooplankton resulting from less predation. The fish of Moe Pond have changed significantly since studies of the pond began in 1994. The pond was dominated by golden shiners and brown bullhead until the illegal introduction of largemouth and smallmouth bass somewhere around 1999. As the bass populations took hold, the golden shiner population decreased to all but extinct in just 3 years. The largemouth bass have since out-competed the smallmouth bass and now dominate the pond, with few brown bullhead trolling the bottom. Since the bass have taken over the pond, their diet has switched from mostly forage fish to primarily zooplankton and invertebrates. Only 3 out of 40 stomachs had fish in them, showing that the bass aren’t consuming fish for forage, and seem to be stunted as a result.

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REFERENCES Albright, M.F., W.N. Harman, W.T Tibbits, M.S. Gray, D.M. Warner, and R.J. Hamway. 2004. Biomanipulation: A classic example in a shallow eutrophic pond. Lake and Reserve Management. 20(04):263-269. Cornwell, M.D. 2013. Personal communication. Assistant professor, SUNY Cobleskill Dept. of Fisheries and Wildlife, Cobleskill, NY.

Dresser, K. 2006. Continued monitoring of the Moe Pond ecosystem following the introduction of smallmouth and largemouth bass (Micropetrus dolomieu and M. salmoides, respectively). 38th Annual Report (2005). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Finger, K. M. 2009. Continued monitoring of the Moe Pond ecosystem following the introduction of smallmouth and largemouth bass (Micropetrus dolomieu and M. salmoides, respectively). 41st Annual Report (2008). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. Hamway, R. J. 2003. Continued monitoring of Moe Pond following the unauthorized stocking of smallmouth and largemouth bass. In 35th Annual Report (2002). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. Hamway, R. J. 2004. Continued monitoring of Moe Pond following the unauthorized stocking of smallmouth and largemouth bass. In 36th Annual Report (2003). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. Lopata, K. 2004. Fifth annual report on the status of Moe Pond following the stocking of Micropterous salmaoides and M. dolomieui. 37th Ann. Rept. (2004). SUNY Oneonta Bio. Fld. Sta.,SUNY Oneonta. McCoy, M.C. III, C.P. Madenjian, J.V. Adams and W.N. Harman. 2001. The fish community of a small impoundment in upstate New York. J. of Freshwater Ecology. 16(3):389-394. Quinn, T.J.III and R.B. Deriso. 1999. Quantitative Fish Dynamics. Oxford University Press, Oxford. Reinicke E. and Walters G.M. 2006. Continued monitoring of fish community dynamics and abiotic factors influencing Moe Pond, summer 2006. 39th Annual Report (2006). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta Sohacki, L. P.1972. Limnological studies on Moe Pond. 5th Annual Report (1972). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. Tibbits, W.T. 2001. Conseuences and management strategies concerning the unauthorized stocking of smallmouth and largemouth bass in Moe Pond. In 33rd Ann. Rept. (2000). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Underwood , E.S. 2008. Continued monitoring of the ecosystem dynamics of Moe Pond following the introduction of largemouth bass (Micropterus salmoides) and smallmouth bass (M. dolomieu). In 40th Annual Report (2007). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. VanDerKrake, A.J. 2012. Monitoring of the Moe Pond ecosystem and largemouth bass (micropterus salmoides) population before considering biomanipulation options. 45th Annual Report (2012). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta 126-136. Werner R.G. 2004. A Field guide: Freshwater Fishes of the Northeastern United States. 1(1) 157. Wilson, B.J., D.M. Warner and M. Gray. 1999. An evaluation of Moe Pond following the unauthorized introduction of smallmouth and largemouth bass. In 32nd Ann. Rept. (1998). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta. Wojnar, 2002. The continued evaluation of Moe Pond following the unauthorized stocking of smallmouth and largemouth bass. In 34th Annual Report (2001). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

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Otsego Lake, NY ice phenology 1843-2014

Holly A. Waterfield, CLM

Otsego Lake’s ice phenology, including date of ice-on (freeze) and ice-off (break-up), and duration of ice cover is summarized in this report (Table 1, Figure 1), providing an update to a 2005 report which included data for most years from 1843-2005 (Altman 2005). Missing data points have been filled by collaboration with K. Addington (Pers. Comm.). Biological Field Station faculty and staff observe lake conditions and maintain a record of annual data.

Table 1. Otsego Lake, NY freeze and thaw dates and duration of ice cover (days) 1843-2014.

Break-up Ice Duration Break-up Ice Duration Winter of Freeze Date Date (days) Winter of Freeze Date Date (days) 1842-43 4/26/1843 1872-73 12/24/1872 5/4/1873 131

1843-44 4/13/1844 1873-74 1/26/1874 5/5/1874 99

1844-45 4/1/1845 1874-75 12/31/1874 5/7/1875 128

1845-46 4/7/1846 1875-76 1/13/1876 4/26/1876 104

1846-47 4/25/1847 1876-77 12/17/1876 4/27/1877 131

1847-48 4/10/1848 1877-78 1/29/1878 4/1/1878 62

1848-49 4/7/1849 1878-79 1/3/1879 4/30/1879 117

1849-50 02/01/1850 4/24/1850 83 1879-80 2/3/1880 4/7/1880 64

1850-51 12/30/1850 3/30/1851 90 1880-81 12/29/1880 4/25/1881 118 1851-52 12/24/1851 4/26/1852 124 1881-82 1/5/1882 4/6/1882 91 1852-53 1/17/1853 4/9/1853 82 1882-83 1/6/1883 4/26/1883 110 1853-54 1/24/1854 4/20/1854 86 1883-84 1/6/1884 4/22/1884 107 1854-55 12/20/1854 4/24/1855 125 1884-85 1/19/1885 4/26/1885 97 1855-56 1/5/1856 4/26/1856 112 1885-86 1/11/1886 4/13/1886 92 1856-57 12/18/1856 4/6/1857 109 1886-87 1/2/1887 5/1/1887 119 1857-58 2/5/1858 4/5/1858 59 1887-88 12/30/1887 4/30/1888 122 1858-59 1/9/1859 3/30/1859 89 1888-89 1/19/1889 4/11/1889 82 1859-60 12/28/1859 4/7/1860 101 1889-90 2/22/1890 4/8/1890 45 1860-61 1/8/1861 4/14/1861 96 1890-91 12/25/1890 4/15/1891 111 1861-62 1/5/1862 4/22/1862 107 1891-92 1/20/1892 4/5/1892 76 1862-63 1/17/1863 4/23/1863 96 1892-93 12/28/1892 4/18/1893 111 1863-64 1/3/1864 4/21/1864 109 1893-94 1/13/1894 3/22/1894 63 1864-65 1/8/1865 4/5/1865 87 1894-95 12/29/1894 4/20/1895 112 1865-66 1/7/1866 4/14/1866 97 1895-96 1/6/1896 4/20/1896 91 1866-67 1/3/1867 4/15/1867 102 1896-97 1/18/1897 4/21/1897 92 1867-68 1/6/1868 4/16/1868 101 1897-98 1/28/1898 3/18/1898 44 1868-69 12/27/1868 4/21/1869 115 1898-99 1/10/1899 4/24/1899 104 1869-70 1/8/1870 4/26/1870 108 1899-00 1/9/1900 4/19/1900 101 1870-71 1/4/1871 3/17/1871 72 1900-01 1/19/1901 4/17/1901 88 1871-72 12/21/1871 4/27/1872 128 1901-02 1/17/1902 3/29/1902 71

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Table 1. Cont’d.

Ice Ice Freeze Break-up Duration Freeze Break-up Duration Winter of Date Date (days) Winter of Date Date (days) 1902-03 1/19/1903 3/17/1903 57 1942-43 12/20/1942 4/12/1943 113 1903-04 12/29/1903 4/24/1904 117 1943-44 12/24/1943 4/20/1944 118 1904-05 1/23/1905 4/19/1905 86 1944-45 12/28/1944 3/26/1945 88 1905-06 2/3/1906 4/14/1906 70 1945-46 1/20/1946 3/24/1946 63 1906-07 12/26/1906 4/4/1907 99 1946-47 2/8/1947 4/11/1947 62 1907-08 1/12/1908 4/8/1908 87 1947-48 1/5/1948 3/27/1948 82 1908-09 1/8/1909 4/9/1909 91 1948-49 2/6/1949 3/26/1949 48 1909-10 12/30/1909 3/28/1910 88 1949-50 2/8/1950 4/17/1950 68 1910-11 1/4/1911 4/23/1911 109 1950-51 1/28/1951 4/9/1951 71 1911-12 1/7/1912 4/19/1912 103 1951-52 1/8/1952 4/10/1952 93 1912-13 2/18/1913 3/21/1913 31 1952-53 2/2/1953 3/23/1953 49 1913-14 1/14/1914 4/19/1914 95 1953-54 1/14/1954 3/9/1954 54 1914-15 12/26/1914 4/19/1915 114 1954-55 1/21/1955 4/11/1955 80 1915-16 1/8/1916 4/17/1916 100 1955-56 12/23/1955 4/8/1956 107 1916-17 2/12/1917 4/18/1917 65 1956-57 1/11/1957 4/16/1957 95 1917-18 12/13/1917 4/14/1918 122 1957-58 2/1/1958 4/18/1958 76 1918-19 2/8/1919 3/28/1919 48 1958-59 12/29/1958 4/18/1959 110 1919-20 12/18/1919 4/17/1920 121 1959-60 1/12/1960 4/17/1960 96 1920-21 1/18/1921 3/19/1921 60 1960-61 1/3/1961 4/26/1961 113 1921-22 1/3/1922 4/11/1922 98 1961-62 1/18/1962 4/18/1962 90 1922-23 1/6/1923 4/11/1923 95 1962-63 1/16/1963 4/17/1963 91 1923-24 1/24/1924 4/17/1924 84 1963-64 1/12/1964 4/15/1964 94 1924-25 12/28/1924 3/27/1925 89 1964-65 1/15/1965 4/25/1965 100 1925-26 1/13/1926 4/23/1926 100 1965-66 1/15/1966 4/19/1966 94 1926-27 12/19/1926 4/10/1927 112 1966-67 1/19/1967 4/9/1967 80 1927-28 1/27/1928 4/7/1928 71 1967-68 1/2/1968 4/5/1968 94 1928-29 1/15/1929 3/29/1929 73 1968-69 12/26/1968 4/15/1969 110 1929-30 1/23/1930 4/7/1930 74 1969-70 1/2/1970 4/24/1970 112 1930-31 1/14/1931 4/10/1931 86 1970-71 1/9/1971 4/30/1971 111 1931-32 2/25/1932 4/22/1932 57 1971-72 1/17/1972 4/25/1972 99 1932-33 2/25/1933 4/17/1933 51 1972-73 1/8/1973 3/18/1973 69 1933-34 12/29/1933 4/11/1934 103 1973-74 1/12/1974 4/14/1974 92 1934-35 1/12/1935 4/13/1935 91 1974-75 1/21/1975 4/20/1975 89 1935-36 1/27/1936 3/30/1936 63 1975-76 2/29/1976 3/29/1976 29 1936-37 1/28/1937 4/20/1937 82 1976-77 12/27/1976 4/12/1977 106 1937-38 1/10/1938 3/23/1938 72 1977-78 1/15/1978 4/17/1978 92 1938-39 1/18/1939 4/24/1939 96 1978-79 1/10/1979 4/6/1979 86 1939-40 12/28/1939 4/27/1940 121 1979-80 1/25/1980 4/11/1980 77 1940-41 1/7/1941 4/17/1941 100 1980-81 12/25/1980 4/13/1981 109 1941-42 1/16/1942 4/8/1942 82 1981-82 12/25/1981 4/20/1982 116

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Table 1. Cont’d. Break-up Ice Duration Break-up Ice Duration Winter of Freeze Date Date (days) Winter of Freeze Date Date (days) 1982-83 1/20/1983 4/1/1983 71 1999-00 1/19/2000 3/25/2000 66 1983-84 1/11/1984 4/14/1984 94 2000-01 12/29/2000 4/21/2001 113 1984-85 1/30/1985 4/5/1985 65 2001-02 did not freeze 0 1985-86 1/7/1986 3/30/1986 82 2002-03 1/14/2003 4/17/2003 93 1986-87 1/23/1987 4/4/1987 71 2003-04 1/10/2004 4/13/2004 94 1987-88 1/7/1988 4/1/1988 85 2004-05 1/21/2005 4/11/2005 80 1988-89 1/5/1989 4/1/1989 86 2005-06 2/8/2006 2/17/2006

1989-90 12/22/1989 3/21/1990 89 2005-06 2/26/2006 4/1/2006 43 1990-91 1/22/1991 3/28/1991 65 2006-07 2/12/2007 4/23/2007 71 1991-92 1/25/1992 4/17/1992 83 2007-08 1/27/2008 4/9/2008 75 1992-93 1/19/1993 3/14/1993 87 2008-09 1/10/2009 4/4/2009 84 1993-94 1/6/1994 4/20/1994 104 2009-10 1/23/2010 3/29/2010 65 1994-95 2/7/1995 3/30/1995 51 2010-11 1/14/2011 4/12/2011 88 1995-96 1/4/1996 4/15/1996 102 2011-12 did not freeze 0 1996-97 1/19/1997 4/6/1997 77 2012-13 1/24/2013 4/13/2013 79 1997-98 2/16/1998 3/31/1998 43 2013-14 1/22/2014 4/14/2014 82 1998-99 2/24/1999 4/6/1999 41

Figure 1. Duration of ice cover (days) on Otsego Lake, NY 1850-2014, by year and by 10 year mean. Linear trend lines and R2 values included for time periods 1850-2014 and 1960-2014.

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Long-term trends in the duration of ice cover follow those exhibited by many lakes throughout the northern hemisphere (Magnuson et al. 2000). In general, temperate lakes are freezing later and thawing earlier, with the greatest rates of change observed beginning around 1850 (Magnuson, et al. 2000). The rate of change in Otsego Lake’s length ice cover in more recent years (1960-2014) is greater than that seen from 1850-2014, based on decade mean ice cover duration (Figure 1). The ecological effects of such changes are currently a focus of researchers around the world.

REFERENCES

Addington, K. Personal communication.

Altman, M. 2005. History of ice on/off dates for Otsego Lake, NY. In. 38th Ann. Rept. (2005). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Magnuson, J.J., D.M. Robertson, B.J. Benson, R.H. Whynne, D.M. Livingstone, T. Arai, R.A. Assel, R.G. Barry, V. Card, E. Kuusisto, N.G. Granin, T.D. Prowse, K.M. Stewart, V.S. Vuglinski. 2000. Historical trends in lake and river ice cover in the northern hemisphere. Science. 289: 1743-1746.

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Presence of mercury and comparison to other metals in lakes, rivers, and streams in central New York

M. Moore1 and D. Castendyk2

INTRODUCTION This research project has two major parts to it: the water sampling in 2012 and the soil sampling in 2013. To help differentiate between the two parts the first part of this project when water samples were collected in 2012 will be referred to as water sampling. The second part of this project involving collecting soil samples in 2013 will be referred to as soil sampling.

Water sampling In 2004, a study was released by the USGS indicating that mercury concentrations in NY were below drinking water limits (Krabbenhoft 2004). However, in 2012 a Department of Environmental Conservation (DEC) fish advisories report, several streams and rivers in central NY were classified as having high mercury concentrations in fish (Table 1). Measured mercury concentrations in fish were above the limit dictated by the DEC (NYS DEC 2012). The DEC routinely tests rivers and streams. The purpose of this study was to test three lakes (Wilber, Goodyear, and Otsego Lake) and two rivers (Unadilla and Susquehanna River) for mercury in central New York. The water bodies were tested for methyl and total mercury. Part of the study included testing the drinking water source for Oneonta, NY (Wilbur Lake). All five bodies of water were found to have traces of total mercury and four out of five had traces of methyl mercury. Otsego Lake was the only site that had methyl mercury concentrations below detection limits. (The detection limit for total mercury is 0.50 ng/L and for methyl mercury is 0.050 ng/L). Although our results indicate concentrations under drinking water state and EPA limits, it is still a concern for human health in regards to fish consumption.

1 Peterson Family Conservation Trust Intern, summer 2013. Present affiliation: SUNY Oneonta. 2 Associate Professor, Earth Science Department, SUNY Oneonta.

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Table 1. Summary of some of the fish warnings released in the DEC fish advisories report of 2012. Females of child bearing age are informed to never eat a meal of fish from any of these water bodies, and others are instructed to eat only one meal a month. The ones highlighted in gray are sites that we tested and that have mercury contamination issues for fish (NYS DEC 2012).

Soil sampling The next part of the research was to look for a source of mercury that were found in the water samples. We were interested in seeing if the mercury in the water was coming from atmospheric loading or from baseflow in a body of water through the soil. We chose to look into the soil. The soil samples were analyzed for other metals that provide a screening indicator of industrial discharge where elevated mercury may be expected. An additional part of the research was to examine the impact of the watershed location (how far upstream or downstream the sites were located on the rivers) on chemical signatures of trace and heavy elements (Cu, Pb, Zn and Ni) in stream sediments. Eleven sediment samples were collected from Susquehanna drainage system and analyzed for acid and soluble extractible elements, and basic soil parameters such as organic matter (OM), cation exchange capacity (CEC) and soluble salts. Results showed that concentration of elements are log normally distributed. Concentration of elements demonstrated spatial variability in stream sediments with a decreasing order of abundance: Zn>Pb>Cu>Ni. Most of the element concentrations increase from upstream to downstream where there is a larger volume and possible longer contact time between the elements in solution and soil sediments. The downstream samples are the ones that have a larger volume of water entering this area and longer contact time as the water in the downstream samples travels through the watershed for a longer time and has more water and soil interactions. A positive correlation analysis of elemental concentrations increases as you go downstream is suggestive of similar source.

METHODS

Water Sampling The accepted method for collecting water samples for mercury concentrations in water is very specific because mercury concentrations in water samples are in such low concentrations. EPA method 1669 (Novak 2011), known as “Clean Hands Dirty Hands”, is used to ensure accuracy in collecting water samples analyzed for mercury. It is explained out step by step in the appendix. We used this method to collect 10 water samples at five locations (two at each site: one for total mercury and one for methyl mercury).

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Soil sampling Locations for testing soils were partially based on the water sampling portion of this project and to get more testing locations along the Susquehanna and Unadilla to see spatial changes along with the chemical concentration changes. The first thing needed to be done was to collect soil samples to be analyzed. This was done with a black Teflon scooping spoon. It was Teflon to try and limit trace metals from entering soil samples through sampling equipment. We had at least 20 Ziploc bags for collecting the soil samples. Areas having metals or a large amount of leaves were avoided, as they can skew soil results. The spoon was used to get a soil sample so that the bag was filled at least halfway, attempting to get soil samples that were low in rocks and pebbles. The spoon was rinsed in the surface water after finishing testing at a site. It was then rinsed again with the next sites water before collecting the next sample. After collecting the samples, they were laid out in the sun to dry. This was done for about 3 days on and off when the sun was out to ensure dry soil. After the samples were dried they were then filtered with a 0.22 millimeter sifter to get just the soil in the samples and no sticks or pebbles. Acid digestion was then performed on the dried soil samples by modified EPA method 3052. Modified EPA Soil Acid Digestion method 3052 (EPA, December 1996): Materials:

• Safety gloves, safety goggles, and lab coat • Hydrochloric Acid • Nitric Acid • Pipettes • Acid digestion microwave • Soil Samples • Scale • Centrifuge tubes that can hold at least 15 to 25 milliliters of sample. • Pen, notebook, sharpie, tape to label and keep track of samples • Deionized Water (DI water)

1. Weigh out 0.5 grams of each soil sample and put into separate centrifuge tubes and label each with its separate ID location. 2. Using a pipette add to each centrifuge tube nine ml. of nitric acid. 3. Using a pipette add an additional three ml. of hydrochloric acid (this is a modification; the original EPA Method 3052 uses three ml. of hydrofluoric acid. Hydrofluoric acid is much more dangerous than hydrochloric acid). 4. Allow the samples to sit loosely capped for at least 24 hour for the acid to begin digesting the soil sample.

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5. Insert centrifuge tubes in acid digestion microwave and put microwave at 100 ◦C. (This is a modification, the original EPA Method 3052 uses 180 ◦C. The lab technician did not allow us to turn the microwave up to 180◦C because he was worried that the centrifuge tubes would have melted.) 6. The microwave is left on for 10 minutes once it reaches the temperature of 100◦C. 7. Pipette one ml. of the newly made soil and acid mixture into new centrifuge tubes for each sample. 8. Add nine ml of DI water to the centrifuge tubes that now contain one ml of the acid and soil mixture. This dilution helps to ensure the soil samples are completely dissolved in liquid for inductively coupled plasma spectrometry (ICP-AES) Label each tube with site ID. 9. The samples are then sent to Virginia Tech Soil Laboratory to be analyzed with ICP- AES on the Acid Digested soil samples.

RESULTS/DISCUSSION Water sampling Figure 1 is a map displaying the locations of the water samples taken and shows methyl mercury concentrations (grey dots) and shows the total mercury concentrations (white dots) graphically. Table 2 summarizes the data in tabular form. Soil sampling Figure 2 and Table 3 indicate the sites sampled for soil analysis. Samples, from headwater to downstream locations, were LOC-4, LOC-2, LOC-3, LOC-1, LOC-7, LOC-8, and LOC-9 along the Susquehanna River. Samples LOC-11, LOC-10 and LOC-9 were all taken along the Unadilla. LOC-9 is a sample location where the Unadilla River and Susquehanna River meet. Also the Susquehanna River runs through Goodyear Lake. Points in Figure 2 that have a grey circle around them represent where both a water and soil sample were taken from. Table 4 summarizes concentrations of heavy and trace metals (mg/kg). pH is a measure of the concentration of hydronium ions, Organic Matter (OM) was measured as a percentage, and Cation Exchange Capacity (CEC) is measured in cmolc/kg. The shaded boxes indicate concentration values below the detection limit of the ICP-AES at Virginia Tech Soil Laboratory (Tilley 2013). All of the values for mercury and all the values but one for arsenic were below the detection limit. This is why they are left out of correlation analysis provided in Table 5.

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Figure 1. Map displaying the locations of the water samples taken. The grey dots show methyl mercury concentrations and the white dots represent total mercury concentrations.

Table 2. Concentrations of total and methyl mercury water samples for the five sampling locations and the days they were taken. Analyzed at Frontier Global Sciences (Siska 2011).

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Figure 2. Location of soil samples collected along the Susquehanna and Unadilla Rivers.

Table 3. Locations of the soil sampling sites and the site descriptions.

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Table 4. Summary of concentrations of heavy and trace metals (mg/kg). pH is measured in concentration of hydronium ions, Organic Matter (OM) was measured in percentage, Cation Exchange Capacity (CEC) is measured in cmolc/kg. The shaded boxes indicate concentration values below the detection limit of the ICP-AES at Virginia Tech Soil Laboratory (Tilley 2013).

Table 5. The negative and positive correlations between trace and heavy metals along with other important variables relating to soil such as pH, Organic Matter, Cation Exchange Capacity. The values were all put into log form and correlated using the Microsoft Excel Correlation Analysis Toolpak.

When comparing the concentrations throughout all eleven of the soil sample sites taken, there is a positive correlation between iron and nickel, copper and lead, copper and organic matter, and lead and zinc. A negative correlation is shown between iron and lead, iron and organic matter, and nickel and lead.

Figure 3 plots concentrations of zinc, lead and copper in the Susquehanna River against distance from its source, Otsego Lake (see Table 3 for site locations). Table 6 displays these concentrations in tabular form.

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Figure 3. Concentrations of zinc, lead and copper in the Susquehanna River sediments plotted against distance from its source, Otsego Lake. See Table 3 for site locations.

Table 6. Concentrations of zinc, lead and copper in sediments along the Susquehanna River.

LOC-3 is the location of Goodyear Lake after the dam, which has historically been used for hydro power. We hypothesized that elemental concentrations would increase while going downstream because of the longer contact time and volume of water increasing as you travel down the Susquehanna River. However, this location had the highest concentration of zinc and the second highest concentrations of lead and copper. This could be caused by the turbines that are located at LOC-3. These turbines are aged and could be contributing to the increased metal concentrations seen at Goodyear Lake right after the dam.

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Figure 4 plots concentrations of zinc, lead and copper in the Unadilla River against distance from the most upstream sample (see Table 3 for site locations). Table7 displays these concentrations in tabular form. The results from the Unadilla River fell more in-line with the hypothesis that concentrations in the metals increase as you travel downstream. LOC-9 had the highest concentrations in metals except for lead, where LOC-10 had the highest concentration. This difference, though, was quite small and could have been caused by data skews in sampling or EPA acid digestion modifications.

Figure 4. Concentrations of zinc, lead and copper in the Unadilla River sediments plotted against distance. See Table 3 for site locations.

Table 7. Concentrations of zinc, lead and copper in sediments along the Unadilla River.

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CONCLUSION Water sampling The drinking water maximum concentration limit (MCL) for mercury as mandated by the EPA is 200 ng/L (EPA 2012.) Therefore, all of the total and methyl mercury concentration limits in the water samples conducted in the water sampling phase of research of this study were far below the MCL for mercury in drinking water. This does not rule out mercury concentrations in fish, though, as this compound can bioaccumulate in fish (Chen et al. 2014). Further work on this project will be conducted to receive total concentrations of mercury in fish samples. Soil sampling The hypothesis that zinc, lead, and copper concentrations increase as one travels downstream holds true, with the exception of LOC-3. Concentrations were higher there, despite that site being fairly high in the watershed, just below the Goodyear Lake dam. There is a slight increase of metal concentrations as one travels down the Unadilla and Susquehanna Rivers. This supports the hypothesis of further downstream samples, with higher residence time and greater volumes of water, will have higher metal concentrations. Yet, since it is only a slight increase, more data needs to be completed to substantiate this hypothesis. We also can conclude that some other source from the hydroelectric dam, such as the turbines or transformers for electric usage, could be introducing metal concentrations to LOC-3.

ACKNOWLEDGMENTS I would like to thank the SUNY Oneonta grants department for funding this project. I would also like to thank Biological Field Station, in particular Matt Albright for allowing me usage of lab equipment. I would like to thank Colleen Parker, Dave Snyder, and Stephen Dechon for helping in collection. I would also like to thank Oyewumi Oluyinka for his countless guidance throughout the entire soil sampling portion of this project.

REFERENCES Chen, C.Y., M.E. Borsuk, D.M. Bugge, T. Hollweg, P.H. Bolcom, D.M. Wward, J. Williams and R.P. Mason. 2014. Benthic and pelagic pathways of methylmercury bioaccumulation in estuarine food webs of the northeast United States. PLos ONE 9(2):e89305.doi:10.1371/journal.pone.0089305. EPA, 21 May, 2012, Basic Information about Mercury (inorganic) in Drinking Water. http://water.epa.gov/drink/contaminants/basicinformation/mercury.cfm EPA. 1996. Method 3052- Microwave Assisted Acid Digestion Of Siliceous And Organically Based Matrices, http://www.epa.gov/osw/hazard/testmethods/sw846/pdfs/3052.pdf Goodyear Lake Association. Oct. 2011, Goodyear Lake Association Inc. http://www.goodyearlakeny.org/index.html

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Krabbenhoft, D. August, 2004, The mercury experiment to assess atmospheric loadings in Canada and the US (METALLICUS) project. http://diseasemaps.usgs.gov/mercuryworkshop/presentations.html Novak M. July 2011, Personal communication. Chief of the Statewide Waters Monitoring Section of DEC. NYS DEC. 2012. Fish Health Advisories. http://www.dec.ny.gov/docs/fish_marine_pdf/fishguide12ha.pdf NYS DEC. 2011. The Mercury Cycle, NY. http://www.dec.ny.gov/docs/wildlife_pdf/hgfish.pdf Siska. 2011. Frontier Global Sciences. Bothell, WA. Tilley, A.M. 2013. Personal communication. ICP-AES at Virginia Tech Soil Laboratory.

Appendix Basic Procedure for “Clean Hands/Dirty Hands” Sampling in a Waded Stream 1. Assess site for potential sources of trace metal contamination, adjust sampling location to be out of influence of contamination if possible, note any potential contamination sources on field sheets 2. Don waders and any other site specific safety gear short of gloves and other “clean” items 3. Designate “clean hands”(CH) and “dirty hands” (DH) members of the team, preserve roles through the entire sample collection day 4. Carry Hg cooler and PPE (Personal Protective Equipment) storage tub down to water’s edge and place in a stable area, away from potential contamination sources. 5. Don PPE 1. Gloves will be single bagged according to size and purpose and stored in a plastic storage tub with a sealing lid. 2. DH opens the storage tub and holds lid while CH opens bags and dons inner glovers (elbow length) and outer gloves (standard cuff) 3. CH removes 2 pairs of the appropriate gloves for DH, seals all bags 4. DH gets double-bagged sample bottle out of dedicated Hg cooler 5. DH unseals outer bag, holds open for CH 6. CH opens inner bag (if possible, do not remove from outer bag), and removes sample bottle (inner bag is folded over and stuffed back into outer bag without resealing)

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Note: From the time the sample bottle is removed from the inner bag, until the time it is returned, CH should keep the container away from his or her body and clothing. Both samplers should also avoid any direct breathing on the sample and/or sample container. 7. DH reseals outer bag, places bags back into cooler

8. CH wades out to sampling location, approaching the sample point from downstream and upwind of possible contamination sources if possible (downstream approach takes precedence)

9. Facing and reaching upstream, CH plunges the still capped, empty sampling container underwater to the desired depth (4-6” below the surface), taking care to avoid any surface scum or sheens.

10. Holding the bottle away from their body, with the opening facing upstream, CH uncaps the bottle underwater, allows it to fill, and re-caps the bottle underwater (keep cap underwater during filling).

11. CH wades back to the bank and staging area

12. DH removes bags from cooler and holds open outer bag for CH

13. CH places sample bottle in the inner bag and re-seals

14. DH re-seals outer bag.

All samples after collection were kept chilled. All of the samples were analyzed at Frontier Global Sciences, Bothell, WA 98011, United States of America using a cold-vapor atomic fluorescence spectrophotometer. (CV-AFS)

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The effects of zebra mussels on benthic macroinvertebrates in Otsego Lake

Jennifer M. Vanassche1, Wai Hing1,2*, Willard N. Harman2, and Matthew F. Albright2

ABSTRACT

The zebra mussel, Dreissena polymorpha (Pallas 1771), was first documented in North America in the 1980s. Zebra mussels were first documented in Otsego Lake in 2007 and were considered abundant of 2010. Surveys of macroinvertebrates were performed during and prior to 1997 and in 2008. In 2013, surveys of benthos in deep and shallow sites in the lake were repeated and taxonomic composition was described to assess the impacts of D. polymorpha on the benthic community.

INTRODUCTION

Dreissenid mussels including the zebra mussel (Dreissena polymorpha Pallas, 1771), originating from the Ponto-Caspian area (Black, Asov, and Caspian Sea), and the quagga mussel (Dreissena rostriformis bugensis Andrusov, 1897), originating from the mouths of the Rivers Southern Bug and Dnieper, are both species native to Eastern Europe (Van der Velde et al. 2010). Both zebra and quagga mussels were accidently introduced into the Laurentian Great Lakes in North America in the 1980s most likely in ballast water (Ludyanskiy et al. 1993; Carlton 2008; Van der Velde et al. 2010). It was first reported that the zebra mussel was found in North America in 1988 in Lake St. Clair (Hebert et al. 1989); however, a recent paper (Carlton 2008) provided convincing evidence that it was present as early as 1986 in Lake Erie. The first occurrence of the quagga mussel in North America was documented in 1989 in Lake Erie (Mills et al. 1993), but it was first identified as a separate species and given the common name “quagga” in a paper by May and Marsden (1992). This species was later identified from morphological and genetic material as Dreissena bugensis (Spidle et al. 1994, Rosenburg and Ludyanskiy 1994); however, a recent genetic comparison between D. bugensis and D. rostriformis indicated no distinct difference between the two taxa (Therriault et al. 2004).

Otsego Lake in Otsego County, New York is a mesotrophic, dimictic lake formed by glacial over-deepening of the Susquehanna River Valley (Harman 1997). Zebra mussels were first documented in Otsego Lake in 2007 (Waterfield 2009) and were abundant by 2010 (Anonymous 2009; Anonymous 2010a, b). Dreissenid mussels initially have a minor impact on the ecosystem in which they were introduced (Wong et al. 2011); however, over time exponential increases in dreissenid populations drastically affect the ecosystem by changing microhabitats (Higgins and Zanden 2010). Zebra mussels have a positive impact on some macroinvertebrates while negatively impacting others; they affect the environment directly by altering the substrate they colonize or indirectly by increasing water clarity (Ward and Ricciardi 2010). The ability of zebra mussels to easily colonize hard substrates is especially detrimental to

1 Department of Biology, SUNY College at Oneonta 108 Ravine Pkwy Oneonta, NY. 2 Biological Field Station, SUNY College at Oneonta, 5838 State Highway 80, Cooperstown, NY. *Corresponding author

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native mussels in the family Unionidae. Unionids, which have been in decline since the mid- 1800s, compete with zebra mussels for food and space (Cope et al. 2003). Zebra mussels often colonize the shells of the latter, preventing them from eliminating wastes and feeding; this contributes to the decline of native mussels (Cope et al. 2003). Several other species are negatively impacted by zebra mussels, including the molluscan family Pisidiidae and the amphipod genus Diporeia (Nalepa et al. 2009; Ward and Ricciardi 2010). Declines in populations of macroinvertebrates may contribute to declines in populations of fish that do not have flexible diets (Owens and Dittman 2003). Because zebra mussels are a low-energy alternative to the animals they displace, fish species that become dependent on zebra mussels for food are smaller and weigh less than fish of the same species that eat other macroinvertebrates (Owens and Dittman 2003). Dreissena polymorpha can have a positive effect on other macroinvertebrates; zebra mussel colonization increases the complexity of the substrate, providing more cover from predators (Ozersky et al. 2011; Nalepa et al. 2009; Ward and Ricciardi 2010). Feces and pseudofeces released by zebra mussels may also provide a food source for some macroinvertebrates (Hecky et al. 2004). Furthermore, dreissenids have indirect effects on macroinvertebrates by changing water clarity and phytoplankton production and by changing nutrient contents of lakes, including the removal of phosphorus in lakes (Higgins and Zanden 2010). As a result, the invasion of the zebra mussel to Otsego Lake is expected to have resulted in changes in the benthic community. Benthic macroinvertebrate surveys were performed in 2013 to evaluate that hypothesis.

METHODS

In 1968, Harman collected benthos at 53 shore stations and deep water benthos at sites along 6 transects at various depths from the surface of Otsego Lake (Harman 1994). The eulittoral and transect (littoral and profundal) studies were repeated in 1993 by Wheat (1994) and Hayes (1994), respectively. Harman (1994) calculated species richness at the eulittoral sites from records of taxa present between 1968 and 1988 and compared the data to collections made between 1989 and 1993.

Benthic samples were collected on Otsego Lake from the research vessel Anadontiodes in July 2008. An Ekman dredge was used to collect samples along three transects (Figure 1) at depths of 0m to 50m. Transects were sampled between 10 July 2008 and 21 July 2008. Samples were preserved in jars in 70% ethanol. In June and July 2013, rose bengal was added to each jar and macroinvertebrates were removed from the samples and sorted. If less than 50% of a single organism was present, the organism was discarded. Macroinvertebrates were identified to subclass, order, family, or genus. Nematodes were not identified further than phylum. Annelids were identified to subclass and Platyhelminthes were identified to class. All other taxa were identified to family or genus excluding two unknown arthropods. Morphological differences within these broader taxonomic groups were used to estimate more specific taxonomic richness, which was then calculated for each site (Peckarsky et al.1990; Merritt and Cummins 1996; Harman 1997).

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Figure 1. Transects and sites that were sampled in Otsego Lake, New York during July 2008 (Iannuzzi 1990). Contours are given in feet (1 m = 3.28 ft). Collections were made within the shaded areas.

Benthic samples were collected with an Ekman dredge on Otsego Lake along 7 transects from the research vessel Anadontiodes in June, July, and August 2013. Samples were preserved, sorted, and identified as in 2008. In August through November 2013, benthic macroinvertebrates were collected at 47 sampling stations along the shoreline of Otsego Lake. Most of the sampling stations are the same sites used by Harman (1994) except sites 610, 621-623, 640-642, 652, and 653 sampled in 2013 which were not sampled in 1997, and 613 and 646 sampled in 1997, which were not sampled in 2013. Sites at which samples were collected from in 2008 were sampled in nearby areas in 2013. Samples were preserved in 70% ethanol and identified in the same way as the deep samples.

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RESULTS

In the three transects sampled in 2008, 30 taxa were found compared to 19 in 2013 (Table 1). A similar reduction was found between 1997 and 2013; along all 7 transects sampled, 85 taxa were found in 1997 and 51 were found in 2013 (Table 2). The phyla Cnidaria, Ectoprocta, and Entoprocta and the family Unionidae were not found in 2008 or 2013. Plecopterans, which had not been found in Harman’s 1997 study or in 2008, were present in 2013. The reduction in richness in 2013 compared to 2008 along transects 2, 4, and 6 primarily includes Ephemeroptera, Odonata, and Trichoptera. Between 2008 and 2013, 4 genera of ephemeropterans disappeared while one new genus was identified. No families of Odonata and Trichoptera were identified in 2013 compared to 2 families each in 2008 along the three transects. A reduction in richness of ephemeropterans also occurred between 1997 and 2013. Hemipterans in Otsego Lake experienced decline; in 1997 6 families of hemipterans were found, whereas only one was found in 2013 along all 7 transects. Diversity of Odonata declined between 1997 and 2013. During 1997 7 taxa of odonates were found and 3 were found in 2013; however, the two years had no genera in common. The mollusk families Unionidae, Viviparidae, Pleuroceridae, and Succineidae were not found after 1997.

Table 1. Comparison of lake macrobenthic taxa of Otsego Lake between 2013 and 2008. Presence of a taxon is marked with an “x”. Taxon Taxon Presence Phylum Class Order Family Genus 2013 2008 Annelida Hirudinea x x Annelida Oligochaeta x x Arthropoda Branchiopoda Diplostraca (Suborder Cladocera) x Arthropoda Insecta Coleoptera Elmidae x Arthropoda Insecta Diptera Ceratopogonidae x x Arthropoda Insecta Diptera Chironomidae x x Arthropoda Insecta Diptera Tipulidae x Arthropoda Insecta Ephemeroptera Caenidae Brachycercus x Arthropoda Insecta Ephemeroptera Caenidae Caenis x Arthropoda Insecta Ephemeroptera Ephemeridae Hexagenia x x Arthropoda Insecta Ephemeroptera Heptageniidae Stenacron x Arthropoda Insecta Ephemeroptera Heptageniidae Stenonema x Arthropoda Insecta Ephemeroptera Leptophlebiidae Leptophlebia x Arthropoda Insecta Megaloptera Sialidae Sialis x Arthropoda Insecta Odonata Coenagrionidae Coenagrion x Arthropoda Insecta Odonata Corduliidae x Arthropoda Insecta Trichoptera Leptoceridae x Arthropoda Insecta Trichoptera Polycentropodidae x Arthropoda Arachnida Trombidiformes x x Arthropoda Entognatha Collembola x Arthropoda Malacostraca Amphipoda Gammaridae Gammarus x x Arthropoda Malacostraca Amphipoda Hyalellidae Hyalella x

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Arthropoda Malacostraca Isopoda Asellidae x x Mollusca Bivalvia Veneroida Dreissenidae Dreissena x x Mollusca Bivalvia Veneroida Sphaeriidae/Pisidiidae x x Mollusca Gastropoda Basommatophora Lymnaeidae x Mollusca Gastropoda Basommatophora Physidae x x Mollusca Gastropoda Basommatophora Planorbidae x x Mollusca Gastropoda Heterostropha Valvatidae x x Mollusca Gastropoda Neotaenioglossa Hydrobiidae x x Nematoda x x Platyhelminthes Turbellaria x x Porifera Demospongiae Haplosclerida Spongillidae Spongilla x

Table 2. Comparison of lake macrobenthic taxa of Otsego Lake between 2013 and 1997. Presence of a taxon is marked with an “x”. Taxon Taxon Presence Phylum Class Order Family Genus 2013 1997 Annelida Hirudinea x x Annelida Oligochaeta x x Arthropoda Branchiopoda Diplostraca (Suborder Cladocera) x x Arthropoda Insecta Coleoptera Chrysomelidae x Arthropoda Insecta Coleoptera Curculionidae x Arthropoda Insecta Coleoptera Dryopidae x Arthropoda Insecta Coleoptera Dytiscidae x x Arthropoda Insecta Coleoptera Elmidae x x Arthropoda Insecta Coleoptera Gyrinidae x Arthropoda Insecta Coleoptera Haliplidae x x Arthropoda Insecta Coleoptera Hydrophilidae x Arthropoda Insecta Coleoptera Psephenidae Ectropria x Arthropoda Insecta Coleoptera Psephenidae Psephenus x x Arthropoda Insecta Diptera Ceratopogonidae x x Arthropoda Insecta Diptera Chironomidae x x Arthropoda Insecta Diptera Culicidae x Arthropoda Insecta Diptera Sciomyzidae x Arthropoda Insecta Diptera Stratiomyidae x Arthropoda Insecta Diptera Tabanidae x x Arthropoda Insecta Diptera Tipulidae x Arthropoda Insecta Ephemeroptera Baetidae Baetis x x Arthropoda Insecta Ephemeroptera Baetidae Callibaetis x Arthropoda Insecta Ephemeroptera Baetidae Centroptilum x Arthropoda Insecta Ephemeroptera Baetidae Cloeon x Arthropoda Insecta Ephemeroptera Caenidae Brachycercus x Arthropoda Insecta Ephemeroptera Caenidae Caenis x x

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Arthropoda Insecta Ephemeroptera Ephemerellidae Ephemerella x Arthropoda Insecta Ephemeroptera Ephemerellidae Timpanoga x Arthropoda Insecta Ephemeroptera Ephemeridae Ephemera x x Arthropoda Insecta Ephemeroptera Ephemeridae Hexagenia x x Arthropoda Insecta Ephemeroptera Heptageniidae Epeorus x Arthropoda Insecta Ephemeroptera Heptageniidae Heptagenia x Arthropoda Insecta Ephemeroptera Heptageniidae Stenacron* x x Arthropoda Insecta Ephemeroptera Heptageniidae Stenonema x Habrophleboi Arthropoda Insecta Ephemeroptera Leptophlebiidae des x Arthropoda Insecta Ephemeroptera Leptophlebiidae Leptophlebia x x Paraleptophle Arthropoda Insecta Ephemeroptera Leptophlebiidae bia x Arthropoda Insecta Ephemeroptera Siphlonuridae x Arthropoda Insecta Hemiptera Aphididae Rhoalosiphum x Arthropoda Insecta Hemiptera Corixidae x Arthropoda Insecta Hemiptera Gerridae x Arthropoda Insecta Hemiptera Notonectidae x Arthropoda Insecta Hemiptera Pleidae Neoplea x x Arthropoda Insecta Hemiptera Sialidiae Saluda x Arthropoda Insecta Lepidoptera Pyralidae Acentria x Arthropoda Insecta Lepidoptera Pyralidae Nymphula x Arthropoda Insecta Lepidoptera Pyralidae Petrophila x Arthropoda Insecta Megaloptera Corydalidae Chauloides x Arthropoda Insecta Megaloptera Corydalidae Nigronia x x Arthropoda Insecta Megaloptera Sialidae Sialis x x Arthropoda Insecta Neuroptera Sisyridae Sisyra x Arthropoda Insecta Odonata Aeshnidae x Arthropoda Insecta Odonata Coenagrionidae Argia x Arthropoda Insecta Odonata Coenagrionidae Coenagrion x Arthropoda Insecta Odonata Coenagrionidae Enallagma x Arthropoda Insecta Odonata Coenagrionidae Ishnura x Arthropoda Insecta Odonata Corduliidae x Arthropoda Insecta Odonata Gomphidae Arigomphus x Arthropoda Insecta Odonata Gomphidae Gomphus x Arthropoda Insecta Odonata Lestidae Lestes x Arthropoda Insecta Odonata Libellulidae x Arthropoda Insecta Odonata Macromiidae Macromia x Arthropoda Insecta Plecoptera Nemouridae x Arthropoda Insecta Plecoptera Perlidae x Arthropoda Insecta Trichoptera Helicopsychidae x Arthropoda Insecta Trichoptera Hydropsychidae x Arthropoda Insecta Trichoptera Hydroptilidae x x

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Arthropoda Insecta Trichoptera Leptoceridae x x Arthropoda Insecta Trichoptera Limnephilidae x x Arthropoda Insecta Trichoptera Philopotamidae x Arthropoda Insecta Trichoptera Phryganeidae x Arthropoda Insecta Trichoptera Polycentropodidae x x Arthropoda Arachnida Trombidiformes x x Arthropoda Entognatha Collembola x x Crangony Arthropoda Malacostraca Amphipoda Crangonyctidae nx x Gammaru Arthropoda Malacostraca Amphipoda Gammaridae s x x Arthropoda Malacostraca Amphipoda Hyalellidae Hyalella x x Orconect Arthropoda Malacostraca Decapoda Cambaridae es x x Arthropoda Malacostraca Isopoda Asellidae x x Cnidaria Hydrozoa Anthoathecatae Hydridae x Ectoprocta Phylactolaemata Plumatellida Lophopodidae Pectinella x Fredrecel Ectoprocta Phylactolaemata Plumatellida Plumatellidae la x Entoprocta Phylactolaemata Plumatellida Lophopodidae Urnatella x Mollusca Bivalvia Unionoida Unionidae x Mollusca Bivalvia Veneroida Dreissenidae Dreissena x x Mollusca Bivalvia Veneroida Sphaeriidae/Pisidiidae x x Mollusca Gastropoda Architaenioglossa Viviparidae x Mollusca Gastropoda Basommatophora Ancylidae x Mollusca Gastropoda Basommatophora Lymnaeidae x x Mollusca Gastropoda Basommatophora Physidae x x Mollusca Gastropoda Basommatophora Planorbidae x x Mollusca Gastropoda Heterostropha Valvatidae x x Mollusca Gastropoda Neotaenioglossa Hydrobiidae x x Mollusca Gastropoda Neotaenioglossa Pleuroceridae x Mollusca Gastropoda Stylommatophora Succineidae x Nematoda x x Platyhelminthes Turbellaria x x Porifera Demospongiae Haplosclerida Spongillidae Spongilla x

*Includes MacCaffertium

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DISCUSSION

The introduction of Dreissena polymorpha to Otsego Lake drastically impacts the ecosystem and microenvironments within lakes. Sampling that took place prior to 2007 indicates the state of the benthic community had not yet been affected by zebra mussels. At the time of sampling in 2008, lake-wide zebra mussel abundance was low and their influence on the benthic community was assumed to be minimal. Changes in benthic macroinvertebrates in Otsego Lake have been seen as a result of other invasive species, such as Eurasian milfoil, Myriophyllum spicatum L. (Harman 1997). Unfortunately, the 2008 sampling sites were originally designed for water quality monitoring (Iannuzzi 1990) which makes it difficult to compare with earlier surveys of the benthos (Harman 1997, Vanassche et al. 2014); however, profundal communities in Otsego Lake historically have been homogenous throughout the lake on similar substrates (WN Harman, personal communications). Based on anecdotal observation, zebra mussels have become dominant in the benthic community of Otsego Lake (Anonymous 2009; Anonymous 2010a, b). A systematic survey on zebra mussel abundance and benthic community structure is needed to help determine how D. polymorpha has influenced the benthos.

With the introduction of zebra mussels, the benthic community may have drastically changed since 2008. Changes in the environment of the lake may benefit species that feed from the feces and pseudofeces deposited by the mussels (Atalah et al. 2010; Hecky et al. 2004). Increases in surface area, complexity, and heterogeneity of the substrate provide more refuges for macroinvertebrates to hide from predators and more habitat to live in (Horvath et al. 1999; Ozersky et al. 2011). The removal of phytoplankton and the subsequent increase in water clarity provides additional resources that may benefit some benthic organisms (Higgins and Zanden 2010). Of the taxa identified in 2013, 13 are new compared to 1997 and earlier while 3 are new compared to 2008. Despite reductions in overall taxonomic richness, 3 new genera of ephemeropterans were found and 3 new genera of odonates were found in 2013 compared to 1997. In addition, 2 new genera of pyralids were identified.

Zebra mussels, while beneficial to some macroinvertebrate populations, are detrimental to others. For example, declines in unionid mussel and other filter feeding populations (Cope et al. 2003, Higgins and Zanden 2010) may occur more rapidly from the colonization of zebra mussels. Similarly, declines in other macroinvertebrates such as Pisidiid clams in the Hudson River (Strayer et al. 1998) and some amphipods, such as those in the amphipod genus Diporeia in the Great Lakes, may experience population declines (Nalepa et al. 2009; Ward and Ricciardi 2010). The overall taxonomic richness of benthic macroinvertebrates has decreased since zebra mussels colonized the lake. Many families and genera that were present in 1997 and 2008 were absent in the 2013 survey. Notable families that experienced reductions in diversity included Ephemeroptera, Diptera, Hemiptera, Odonata, and Trichoptera. Lake ecosystems are also affected overall as a result of major changes inflicted by zebra mussels; changes in chlorophyll a and phytoplankton production may result as zebra mussels remove phosphorus from the lake (Wong et al. 2010, Higgins and Zanden 2010). The removal of phosphorus and phytoplankton may affect macrobenthos by altering the amounts of food available to benthos in the lake (Higgins and Zanden 2010). The reductions in diversity in Otsego Lake are likely a result of the introduction of zebra mussels. Further surveys of the lake including statistical analyses of

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changes in the benthic community should be performed to monitor the effects D. polymorpha has had in the lake.

ACKNOWLEDGEMENTS

We thank the sponsored undergraduate and high school interns at the Biological Field Station, and Emily Davidson from the Environmental Science Program, State University of New York at Oneonta who helped collecting benthic samples.

REFERENCES

Anonymous. 2009. Otsego Lake Zebra mussel update. SUNY Oneonta Biological Field Station Reporter, summer 2009, p.3. SUNY Oneonta Biological Field Station, Oneonta, NY.

Anonymous. 2010a. Otsego Lake Zebra mussel update. SUNY Oneonta Biological Field Station Reporter, winter 2010, p.2. SUNY Oneonta Biological Field Station, Oneonta, NY.

Anonymous. 2010b. Otsego Lake Zebra mussel update. SUNY Oneonta Biological Field Station Reporter, summer/fall 2010, p.3. SUNY Oneonta Biological Field Station, Oneonta, NY.

Atalah, J., M. Kelly-Quinn, K. Irvine and T.P. Crowe. 2010. Impacts of invasion by Dreissena polymorpha (Pallas, 1771) on the performance of macroinvertebrate assessment tools for eutrophication pressure in lakes. Hydrobiologia 654: 237-251.

Carlton, J.T. 2008. The zebra mussel Dreissena polymorpha found in North America in 1986 and 1987. Journal of Great Lakes Research 34(4): 770-773.

Cope, W.G., T.J. Newton and C.M. Gatenby. 2003. Review of techniques to prevent introduction of zebra mussels (Dreissena polymorpha) during native mussel (Unionoidea) conservation activities. Journal of Shellfish Research 22(1): 177-184.

González, M.J. and A. Downing. 1999. Mechanisms underlying amphipod responses to zebra mussel (Dreissena polymorpha) invasion and implications for fish-amphipod interactions. Canadian Journal of Fisheries and Aquatic Sciences 56(4): 679-685.

Harman, W.N. 1994. A comparison of the Otsego Lake macrobenthos communities between 1935 and 1993. SUNY Oneonta Biological Field Station (BFS). Annual Report No 26, 131-158.

Hayes, B. 1994. Analysis of littoral and profundal benthos from Otsego Lake, summer 1993. SUNY Oneonta Biological Field Station (BFS). Annual Report No 26, 127-130.

Haynes, J.M., N.A. Tisch, C.M. Mayer and R.S. Rhyne. 2005. Benthic macroinvertebrate communities in southwestern Lake Ontario following invasion of Dreissena and

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Echinogammarus: 1983 to 2000. Journal of the North American Benthological Society 24(1): 148-167.

Hebert, P.D.N., B.W. Muncaster and G.L. Mackie. 1989. Ecological and genetic studies on Dreissena polymorpha (Pallas) - a new mollusk in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 46: 1587-1591.

Hecky, R.E., R.E.H. Smith, D.R. Barton, S.J. Guildford, W.D. Taylor, M.N. Charlton and T. Howell. 2004. The nearshore phosphorus shunt: a consequence of ecosystem engineering by dreissenids in the Laurentian Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 61: 1285–1293.

Higgins, S.N. and M.J. Vander Zanden. 2010. What a difference a species makes: a meta– analysis of dreissenid mussel impacts on freshwater ecosystems. Ecological Monographs 80(2): 179-196.

Horvath, T.G., K.M. Martin and G.A. Lamberti. 1999. Effect of zebra mussels, Dreissena polymorpha, on macroinvertebrates in a lake-outlet stream. The American Midland Naturalist Journal 142: 340-347.

Iannuzzi, T.J. 1991. A model plan for the Otsego Lake watershed. Phase II: the chemical limnology and water quality of Otsego Lake. SUNY Oneonta Biological Field Station (BFS). Occasional Paper #23, Rept. No. 2a.

Ludyanskiy, M.L., D. McDonald and D. Macneill. 1993. Impact of the zebra mussel, a bivalve invader: Dreissena polymorpha is rapidly colonizing hard surfaces throughout waterways of the United States and Canada. Bioscience 43: 533-544.

May, B. and J.E. Marsden. 1992. Genetic identification and implications of another invasive species of dreissenid mussel in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 49: 1501-1506.

Mayer, C.M., R.A. Keats, L.G. Rudstam and E.L. Mills. 2002. Scale-dependent effects of zebra mussels on benthic invertebrates in a large eutrophic lake. Journal of the North American Benthological Society 21(4): 616-633.

Merritt, R.W. and K.W. Cummins (eds). 1996. An Introduction to the Aquatic Insects of North America, 3rd edition. Kendall/Hunt Publishing Company, Dubuque, Iowa, 862 pp.

Mills, E.L., R.M. Dermott, E.F. Roseman, D. Dustin, E. Mellina, D.B. Conn and A.P. Spidle. 1993. Colonization, ecology, and population structure of the quagga mussel (Bivalvia, Dreissenidae) in the lower Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 50: 2305-2314.

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Nalepa, T.F., S.A. Pothoven and D.L. Fanslow. 2009. Recent changes in benthic macroinvertebrate populations in Lake Huron and impact on the diet of lake whitefish (Coregonus clupeaformis). Aquatic Ecosystem Health & Management 12(1): 2-10.

Owens, R.W. and D.E. Dittman. 2003. Shifts in the diets of slimy sculpin (Cottus cognatus) and lake whitefish (Coregonus clupeaformis) in Lake Ontario following the collapse of the burrowing amphipod Diporeia. Aquatic Ecosystem Health & Management 6(3): 311-323.

Ozersky, T., D.R. Barton and D.O. Evans. 2011. Fourteen years of dreissenid presence in the rocky littoral zone of a large lake: effects on macroinvertebrate abundance and diversity. Journal of the North American Benthological Society 30(4): 913-922.

Peckarsky, B.L., P.R. Fraissinet, M.A. Penton and D.J. Conklin. 1990. Freshwater Macroinvertebrates of Northeastern North America. Cornell University Press, Ithaca, NY. 442 pp.

Rosenberg, G. and M.L. Ludyanskiy. 1994. A nomenclatural review of Dreissena (Bivalve, Dreissenidae), with identification of the quagga mussel as Dreissena bugensis. Canadian Journal of Fisheries and Aquatic Sciences 51: 1474-1484.

Spidle, A.P., J.E. Marsden and B. May. 1994. Identification of the Great Lakes quagga mussel as Dreissena bugensis from the Dnieper River, Ukraine, on the basis of allozyme variation. Canadian Journal of Fisheries and Aquatic Sciences 51: 1485-1489.

Strayer, D.L., L.C. Smith and D.C. Hunter. 1998. Effects of the zebra mussel (Dreissena polymorpha) invasion on the macrobenthos of the freshwater tidal Hudson River. Canadian Journal of Zoology 76(3): 419-425.

Therriault, T.W., M.F. Docker, M.I. Orlova, D.D. Heath and H.J. MacIsaaca. 2004. Molecular resolution of the family Dreissenidae (Mollusca: Bivalvia) with emphasis on Ponto- Caspian species, including first report of Mytilopsis leucophaeata in the Black Sea basin. Molecular Phylogenetics and Evolution 30: 479-489.

Vanassche, J.M., W.H. Wong, W.N. Harman and M.F. Albright. 2014. Zebra mussels and other benthic organisms in Otsego Lake, New York in 2008. SUNY Oneonta Biological Field Station (BFS).

Van der Velde, G., S. Rajagopal and A. Bij de Vaate. 2010. The Zebra Mussel in Europe. Backhuys Publishers, Leiden, The Netherlands, 490 pp.

Ward, J.M. and A. Ricciardi. 2010. Community-level effects of co-occurring native and exotic ecosystem engineers. Freshwater Biology 55: 1803-1817.

Waterfield, H.A. 2009. Update on zebra mussel (Dreissena polymorpha) invasion and establishment in Otsego Lake 2008. SUNY Oneonta Biological Field Station (BFS). Annual Report No 41.

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Wheat, E. 1994. A study of the macrobenthos of the eulittoral zone of Otsego Lake. SUNY Oneonta Biological Field Station (BFS). Annual Report No 26, 122-126.

Wong, W.H., T. Tietjen, S. Gerstenberger, G.C. Holdren, S. Mueting, E. Loomis, P. Roefer, B. Moore, K. Turner and I. Hannoun. 2010. Potential ecological consequences of invasion of the quagga mussel (Dreissena bugensis) into Lake Mead, Nevada-Arizona. Lake and Reservoir Management 26: 206-315.

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Control and eradication of water chestnut (Trapa natans, L.) in an Oneonta wetland, 2013 progress report

Holly A. Waterfield1 and Matthew F. Albright2

The marsh of concern is located within the city of Oneonta, Otsego County, NY. The majority of the shoreline (and site access point) is owned by Louis Blasetti with a number of other landowners along the northwestern shoreline behind Oneida Street. Water chestnut (Trapa natans L.) was first observed in the wetland in 2000, though it is likely that it was introduced in years prior but was not documented. A detailed description and history of the management efforts to control and eradicate the water chestnut within the marsh is given in a 2009 Master’s Thesis submitted by W. Eyres (2009) and subsequent report to the NYS Department of Environmental Conservation (Harman et al. 2012). In short, management activities have included a combination of herbicide applications and hand-harvesting of plants since 2006. The combination of chemical and manual control of plants was hugely effective in reducing the population from 2007 to 2010. Native floating-leaved plants were rebounding. Logistical complications resulted in a “missed” herbicide application during the 2010 growing season and subsequent rebound of the population. Following the 2011 herbicide application a second growth of plants was observed in mid-August. These plants were also producing fruits; a major hand-pulling event was held in mid-September, though growth was so prolific that effective harvest of all plants was not achievable. Grant funding for herbicide application expired in 2011. 2012 efforts consisted of hand harvesting on 2 dates; abundance was very low in comparison with past years and only a few plants, if any, remained following the harvesting effort (Waterfield and Albright 2013).

Water chestnut abundance was relatively high in 2013. Hand-harvesting was conducted on 3 dates in 2013, 18 July, 9 and 26 August, by BFS staff along with OCCA staff and volunteers from Headwater Youth Conservation Corps and SUNY Oneonta faculty and incoming freshmen (coordinated by OCCA). Approximately 23 canoe-loads of chestnut were removed, with only isolated individuals remaining, if any. All plants found were harvested. The greatest density of plants was found in the secluded north-eastern portion of the marsh. Harvesting in this area was completed on August 26, at which point some plants had already matured and falling apart and nuts had been released. It is likely that some nuts had already matured and were released. The increased water chestnut abundance in 2013 is evidence that hand-harvesting efforts will be necessary for the coming years, as a viable seed bed still remains.

HAND-HARVEST SUMMARY 2011: >12 canoe-loads harvested, with at least 12 remaining at the end of the season 2012: ~1 canoe-load harvested, “none” remaining at the end of the day. 2013: ~23 canoe-loads harvested, with only isolated individuals potentially remaining.

1 Research Support Specialist: SUNY College at Oneonta Biological Field Station, Cooperstown, NY. 2 Assistant to the Director: SUNY College at Oneonta Biological Field Station, Cooperstown, NY.

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REFERENCES

Eyres, W. 2009. Water Chestnut (Trapa natans L.) infestation in the Susquehanna River watershed: population assessment, control and effects. Master’s Thesis: SUNY College at Oneonta. Bio. Fld. Sta. OP No. 44.

Harman, W.N., H.A. Waterfield, M.F. Albright. 2012. DEC Invasive Species Eradication and Control Grant FINAL REPORT. In 44th Ann. Rept. (2011). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., and M.F. Albright. 2013. Control and eradication of water chestnut (Trapa natans, L.) in an Oneonta wetland, 2012 progress report. In 45th Ann. Rept. (2012). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., W.N. Harman, M.F. Albright. 2010. An update on the control and eradication of water chestnut (Trapa natans, L.) in an Oneonta wetland, 2009 summary report of activities. In.42nd Ann. Rept.(2009). SUNY Oneonta Bio. Fld. Sta., SUNY Oneonta.

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Aquatic macrophyte management plan facilitation, Lake Moraine, Madison County, NY 2013

Willard N. Harman and Matthew F. Albright

BACKGROUND (From Harman et al. 2010)

Located in Madison County NY, Moraine Lake (42o 50’ 47” N, 75o 31’ 39” W) was formed by a deposited glacial moraine damming a valley. The lake, which has been artificially raised, is divided into two basins separated by a causeway and interconnected by a submerged culvert. The north basin is approximately 79 acres, has a mean depth of 1.1m, and a maximum depth 3.7m. The south basin occupies 182 acres, has a mean depth of 5.4m, and a maximum depth of 13.7m. Most of the recreational activities such as fishing, boating and swimming take place in the south basin (Harman et al. 1997).

Moraine Lake has been regarded as meso-eutrophic due to the high productivity of algal and macrophytic plants, low transparency, and depleting levels of dissolved oxygen in the hypolimnion during summer stratification. Developments of lakeside residences and nearby agricultural activities are believed to have contributed to the current productivity status of the upper and lower basins (Anon. 1991). Nutrient loading as a result of faulty septic systems from the residences are believed to be a significant source of the problem in nutrient introduction (Harman et al. 1998). Many of the systems are out of date, undersized, and extremely close to the lake (Brown et al. 1983). Furthermore, soils surrounding the lake have poor percolation rates, steep slopes, shallow depths to bedrock, and fractured bedrock make the lake vulnerable to nutrient loading (Harman et al. 2008).

INTRODUCTION

The aquatic macrophyte communities of Moraine Lake have been monitored by the SUNY Oneonta Biological Field Station (BFS) since 1997. The purpose of monitoring these plant communities has historically been directed towards controlling Eurasian water-milfoil (Myriophyllum spicatum), though in recent years the expansion of the exotic starry stonewort (Nitellopsis obtusa) has been a matter of increased focus. Eurasian water-milfoil is an invasive species that grows rapidly and its extensive canopies cause problems for recreation and other species growth (Borman et al. 1999). Numerous methods of control have been applied to reduce the abundance of Eurasian water-milfoil (Harman et al. 2006). Since 1998, efforts have focused primarily on applications of Sonar®, which has been demonstrated to control Eurasian water- milfoil with some specificity. The goal of managing the Eurasian water-milfoil in the past has been to achieve a balance of species (Lembi 2000, Harman et al. 2008). Most recent activities have included a Sonar® application in the north basin in 2010 and in the south basin in 2011.

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MATERIALS AND METHODS

Sampling took place 20 June, 2 August and 26 September 2013. Five collection sites were sampled, two in the north basin and three from the south basin (Figure 1). The sampling method used was the Point Intercept Rake Toss Relative Abundance Method (PIRTRAM) (Lord and Johnson 2006). It was evaluated in 2008 by comparing the PIRTRAM and dry weight methods such that the rake toss method “could prove useful, if not too much value is placed on actual abundance estimates. …an adequate number of replicate samples could provide insight into species dominance and extent related to exotic nuisance species as well as efforts to control them” (Harman et al. 2008).

For this method two heads of garden rakes were welded together and connected to a 10m nylon cord. At each of the 5 sites, the rake was thrown out randomly 3 times. The rake was allowed to settle to the bottom of the lake and slowly pulled into the boat. Once in the boat, species were separated and each was assigned an abundance category. The 5 abundance categories are “no plants” (denoted by “Z”), “fingerful” (“T”= trace), “handful” (“S” = sparse), rakeful (“M” =medium), and “can’t bring into the boat” (“D” = dense). Table 1 provides biomass range estimates associated with each category. Each rake toss triplicate sample’s category was converted to its corresponding mid-point (Harman et al. 2008). The mid-points were averaged for each species at each site. These species averages were then summed together to look at overall biomass at each site.

In each basin at the deepest location, water quality parameters were measured with a YSI® multiprobe. From surface to substrate, temperature, dissolved oxygen, conductivity, pH, and ORP were measured. A water sample was taken from each basin and returned to the lab to be analyzed using the Lachat QuickChem FIA+Water Analyzer®. The ascorbic acid method following persulfate digestion (Liao and Marten 2001) was used to determine total phosphorus. For total nitrogen, the cadmium reduction method (Pritzlaff 2003) was used following peroxodisulfate digestion as described by Ebina et al. (1983). The phenolate method (Liao 2001) was used to measure ammonia and the cadmium reduction method (Pritzlaff 2003) for nitrate+nitrate-nitrogen. (Harman et al. 2008)

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Figure1. Bathymetric map of Moraine Lake, Madison County, NY. Contours in feet. WQ1 and WQ2 represent were water quality data were collected, sites 1-5 represent where plant biomass and rake toss methods were performed (Harman et al. 2008).

Table 1. Categories, field measurements, midpoint of each category (g/m2) and dry weight ranges applied for the rake toss method and used to generate Tables 2-6 (Harman et al. 2008).

Abundance Categories Field Measure Total Dry Weight (g/m^2) mid low high "Z" = no plants Nothing 0 0 0 0 "T" = trace plants Fingerful .0001 - 2.000 1 0.0001 2 "S" = sparse plants Handful 2.001 - 140.000 71 2.001 140 "M" = medium plants Rakeful 140.001 - 230.000 185 140.001 230 "D" = dense plants Can't bring in boat 230.001 - 450.000+ 340 230.001 450

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RESULTS

Plant Biomass

Tables 2-6 proved biomass estimates, by species, during 2013 for sites 1-5 at Moraine Lake. Eurasian water-milfoil density was generally low in the south basin sites throughout the summer (Tables 2-4). However, it was abundant in the north basin throughout the summer (Tables 5 and 6). Of particular note is the dominance by starry stonewort (Nitellopsis obtusa) at sites 1, 2 and 3; the biomass estimates given in Tables 2, 3 and 4 undoubtedly underestimate actual values because masses of this plant would collapse and fall off the rake as it was being pulled into the boat. Beds of this plant were often so thick that the perception was that a false bottom existed over 1 m from the actual bottom. 2013 was the first year that it was noted at site 2, and by summer’s end it dominated there.

Figures 2-6 graphically summarize the plant biomass contributed by starry stonewort (Nitellopsis obtusa), Eurasian milfoil (Myriophyllum spicatum) and other plant species between 2008 and 2013. While not the original focus of study, starry stonewort is highlighted along with milfoil because it is also an exotic nuisance species and management efforts ought to focus upon control both species. Also, the temporal variations implied by these figures may be misleading. The highest midpoint that can be assigned by any given species is 340 g/m2 (see Table 1). In some visits prior to 2012, more than one species was assigned this highest value, though the cumulative biomass value, in actuality, likely did not exceed the monocultural beds encountered in 2012 and 2013.

Table 2. Mean biomass (g/m2) category mid-points for each species found at Site 1 during 2013 sampling events.

Site 1 6/20/2013 8/2/2013 9/26/2013 Myriophyllum spicatum Megalodonta beckii Zosterella dubia 1.00 Najas spp. Ceratophyllum demersum Chara vulgaris 62.00 85.33 198.67 Vallisneria americana Elodea canadensis Ranunculus aquatilis Ranunculus trichophyllus Stuckenia pectinata 62.00 Potamogeton crispus 1.00 Potamogeton zosteriformis 0.33 Potamogeton pusillus Nitellopsis obtusa 113.33 227.00 61.67 Total 239.33 312.67 260.33

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Table 3. Mean biomass (g/m2) category mid-points for each species found at Site 2 during 2013 sampling events.

Site 2 6/20/2013 8/2/2013 9/26/2013 Myriophyllum spicatum Megalodonta beckii Zosterella dubia 0.67 Najas spp. Ceratophyllum demersum Chara vulgaris Vallisneria americana Elodea canadensis 0.33 0.67 Ranunculus aquatilis Ranunculus trichophyllus Stuckenia pectinata 62.00 Potamogeton crispus 24.33 Potamogeton zosteriformis 23.67 Potamogeton pusillus Nitellopsis obtusa 85.67 340.00 147.00 Total 196.33 340.33 147.67

Table 4. Mean biomass (g/m2) category mid-points for each species found at Site 3 during 2013 sampling events.

Site 3 6/20/2013 8/2/2013 9/26/2013 Myriophyllum spicatum 0.33 Megalodonta beckii Zosterella dubia Najas spp. Ceratophyllum demersum 23.67 23.67 Chara vulgaris Vallisneria americana Elodea canadensis Ranunculus aquatilis Ranunculus trichophyllus Stuckenia pectinata 0.67 Potamogeton crispus 47.67 Potamogeton zosteriformis 0.67 Potamogeton pusillus Nitellopsis obtusa 236.67 340.00 288.33 Total 309.33 363.67 288.67

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Table 5. Mean biomass (g/m2) category mid-points for each species found at Site 4 during 2013 sampling events.

Site 4 6/20/2013 8/2/2013 9/26/2013 Myriophyllum spicatum 185.00 340.00 288.33 Megalodonta beckii Zosterella dubia Najas spp. Ceratophyllum demersum 23.67 0.33 Chara vulgaris Vallisneria americana 0.33 Elodea canadensis Ranunculus aquatilis Ranunculus trichophyllus Stuckenia pectinata 62.33 Potamogeton crispus 24.00 Potamogeton zosteriformis 1.00 Potamogeton pusillus Nitellopsis obtusa Total 296.00 340.00 289.00

Table 6. Mean biomass (g/m2) category mid-points for each species found at Site 5 during 2013 sampling events.

Site 5 6/20/2013 8/2/2013 9/26/2013 Myriophyllum spicatum 236.67 198.67 236.67 Megalodonta beckii Zosterella dubia Najas spp. Ceratophyllum demersum 24.00 24.00 23.67 Chara vulgaris Vallisneria americana Elodea canadensis Ranunculus aquatilis Ranunculus trichophyllus Stuckenia pectinata 24.33 Potamogeton crispus 1.00 Potamogeton zosteriformis 23.67 Potamogeton pusillus Nitellopsis obtusa Total 309.67 222.67 260.33

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Site 1 Plant Community 1000 900 800

700 600 500 Nitellopsis obtusa 400 Myriophyllum spicatum 300 Other Dry Weight (g/m^2) 200 100 0

Figure 2. Comparison of biomass (g/m2) of starry stonewort (Nitellopsis obtusa), Eurasian milfoil (Myriophyllum spicatum) and other plant species present, 2008 (Harman et al. 2009), 2009 (Harman et al. 2010), 2010 (Harman et al 2011), 2011 (Harman et al. 2012), 2012 (Harman and Albright 2013) and 2013, Site 1 (see Figure 1 for site locations).

Site 2 Plant Community 1000 900 800

700 600 500 Nitellopsis obtusa 400 Myriophyllum spicatum 300 Other Dry Weight (g/m^2) 200 100 0

Figure 3. Comparison of biomass (g/m2) of starry stonewort (Nitellopsis obtusa), Eurasian milfoil (Myriophyllum spicatum) and other plant species present, 2008 (Harman et al. 2009), 2009 (Harman et al. 2010), 2010 (Harman et al 2011), 2011 (Harman et al. 2012), 2012 (Harman and Albright 2013) and 2013, Site 2 (see Figure 1 for site locations).

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Site 3 Plant Community 1000 900

800 700 600 500 Nitellopsis obtusa 400 Myriophyllum spicatum 300

Dry Weight (g/m^2) Other 200 100 0

Figure 4. Comparison of biomass (g/m2) of starry stonewort (Nitellopsis obtusa), Eurasian milfoil (Myriophyllum spicatum) and other plant species present, 2008 (Harman et al. 2009), 2009 (Harman et al. 2010), 2010 (Harman et al 2011), 2011 (Harman et al. 2012), 2012 (Harman and Albright 2013) and 2013, Site 3 (see Figure 1 for site locations).

Site 4 Plant Community 1000 900 800 700 Nitellopsis obtusa 600 Myriophyllum spicatum 500 400 Other 300

Dry Weight (g/m^2) 200 100 0

Figure 5. Comparison of biomass (g/m2) of starry stonewort (Nitellopsis obtusa), Eurasian milfoil (Myriophyllum spicatum) and other plant species present, 2008 (Harman et al. 2009), 2009 (Harman et al. 2010), 2010 (Harman et al 2011), 2011 (Harman et al. 2012), 2012 (Harman and Albright 2013) and 2013, Site 4 (see Figure 1 for site locations).

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Site 5 Plant Community 1000 900 800

700

600 Nitellopsis obtusa 500 Myriophyllum spicatum 400 300 Other Dry Weight (g/m^2) 200 100 0

Figure 6. Comparison of biomass (g/m2) of starry stonewort (Nitellopsis obtusa), Eurasian milfoil (Myriophyllum spicatum) and other plant species present, 2008 (Harman et al. 2009), 2009 (Harman et al. 2010), 2010 (Harman et al 2011), 2011 (Harman et al. 2012), 2012 (Harman and Albright 2013) and 2013, Site 5 (see Figure 1 for site locations).

Water Quality Analysis

Water quality parameters over summer 2013 were comparable to those of recent years. In the south basin, waters below 8m were essentially anoxic by the first sampling date (20 June). Transparency was 2.5m on 20 June, 3.5m on 2 August and 2.5m on 26 September. pH was typically between 6.8 and 8.5. In the shallower north basin, stratification was evident in the June visit, with bottom waters being anoxic. pH ranged from 7.0 to 8.8.

DISCUSSION

By 21 June, the north basin was practically covered in milfoil, in which mats of filamentous green algae algae was growing. Coontail was collected at moderate levels. Milfoil was essentially absent from collection sites in the south basin, though it was observed at low density in some areas, notably near the launch area. Curly leaf pondweed was common in the south basin on the June sampling date, but had died off by the next visit. Given the dominance by milfoil through the north basin in 2013, as it was in 2012, an herbicidal application seems appropriate.

For the fifth year, starry stonewort has been dominant where present (Sites 1 and 3, the south end of the south basin and the eastern embayment at the north end of the south basin (see Figures 2 and 4)). For the first year, stonewort was collected at site 2. At the August and

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September sampling dates, it was practically the only species collected. (It should be noted that its abundance is likely much higher than those figures imply due to difficulties related to measurement via the rake toss method. On a number of samples collected, masses of this species collapsed off the rake as the sample was lifted into the boat.) That it is beginning to spread in the lake is worrisome. It has become a serious recreational pest in many lakes (i.e., Pullman and Crawford 2010), and in Moraine Lake it likely will become increasingly common and problematic and may soon be considered the focus of management concerns. We are not aware of proven methods of selective control of this species.

Lastly of note, on the 26 September sampling effort, a bloom of cyanobacteria was observed which was denser than any we have observed on Moraine Lake since our monitoring began in 1997; most of the south basin was covered with a surface scum. We identified the dominant contributors to be Lingbya sp. and Coelosphaerium sp. (= Woronichinia sp.). Also present were Anabaena sp. and Microcystis sp. All of these blue-green algae have been associated with the presence of toxins. Recreational use of the lake should be discouraged if such conditions reoccur.

REFERENCES

Anon. 1988. Madison County septic system survey. Madison County Planning Department, Wampsville, NY 13163.0

Borman, S., R. Korth, and J. Tempte. 1999. Through the looking glass. A field guide to aquatic plants. Wisconsin Lakes Partnership.

Crow, G. E. and C. B. Hellquist. 2000a. Aquatic and wetland plants of Northeastern North America. V.1. Pteridophytes, gymnosperms, and angiosperms: dicotyledons. The University of Wisconsin Press.

Crow, G. E. and C. B. Hellquist. 2000b. Aquatic and wetland plants of Northeastern North America. V.2. Angiosperms: monocotyledons. The University of Wisconsin Press.

Fuller, R. 1997. Unpublished data. Colgate University, Hamilton, NY 13346.

Harman, W.N. and M.F. Albright and O. Zaengle. 2012. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison, N.Y. 2011. Tech. Rept. #30. SUNY Oneonta Bio. Fld. Sta., Oneonta, NY.

Harman, W.N. and M.F. Albright. 2013. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison, N.Y. 201. 45th Ann. Rept. (2012). SUNY Oneonta Bio. Fld. Sta., Oneonta, NY.

Harman, W.N. and M.F. Albright and T.F. Smith. 2011. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison, N.Y. 2010. Tech. Rept. #29. SUNY Oneonta Bio. Fld. Sta., Oneonta, NY.

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Harman, W.N., M.F. Albright, H.A. Waterfield and M. Rubenstein. 2010. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison, N.Y. 2009. Tech. Rept. #27. SUNY Oneonta Bio. Fld. Sta., Oneonta, NY.

Harman, W.N. and M.F. Albright and L. Zach. 2009. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison, N.Y., 2008. Tech. Rept. #26. SUNY Oneonta Bio. Fld. Sta., Oneonta, NY.

Harman, W. N. and M. F. Albright, and C. M. Snyder. 2008. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison County, NY., 2007 Tech. Rept. #25. SUNY Oneonta Bio. Fld. Sta., Oneonta, NY.

Harman, W. N. and M. F. Albright, and A. Scorzafava. 2006. Aquatic macrophyte management plan facilitation, Lake Moraine, Madison County, NY. Tech. Rept. #23. SUNY Oneonta Bio. Fld. Sta., Oneonta NY.

Harman, W. N. and M. F. Albright, P.H. Lord and M. E. Miller. 2002. Aquatic macrophyte management plan facilitation of Lake Moraine, Madison County. Tech. Rept. #13. SUNY Oneonta Bio. Fld. Sta., Oneonta NY.

Harman, W. N. and M. F. Albright, P.H. Lord and M. Miller. 2000. Aquatic macrophyte management plan facilitation of Lake Moraine, Madison County. Tech. Rept. #9. SUNY Oneonta Bio. Fld. Sta., Oneonta NY.

Harman, W. N. and M. F. Albright, P.H. Lord and D. King. 1998. Aquatic macrophyte management plan facilitation of Lake Moraine, Madison County. Tech. Rept. #5. SUNY Oneonta Bio. Fld. Sta., Oneonta NY.

Harman, W. N. and M. F. Albright. 1997. Aquatic macrophyte survey of Lake Moraine, Madison County, summer 1997, as related to management efforts by Sonar ® application. SUNY Oneonta Bio. Fld. Sta., Oneonta NY.

Lembi, C.A. 2000. Aquatic Plant Management. Purdue University, Cooperative Extension Service. West Lafayette, IN 47907.

Lord, P.H. and R.L. Johnson. 2006. Aquatic plant monitoring guidelines. http://www.dec.ny.gov/docs/water_pdf/aquatic06.pdf

NYS Department of Environmental Conservation, Water Division. A Primer on Aquatic Plant Management in NYS. http://www.dec.ny.gov/docs/water_pdf/ch6p2.pdf. 8.12.2009.

Pulman, G.D. and G. Crawford. 2010. A decade of starry stonewort in Michigan. Lakeline, summer 2010. North American Lake Management Society.

- 279 - Otsego Lake fry sampling, 2013

Holly A. Waterfield, CLM and Matthew F. Albright, CLM

INTRODUCTION

Walleye stocking began in 2000 to re-establish a population of the popular game-fish, which would take advantage of the abundant forage provided by alewife (Cornwell 2005). Alewife are known to prey heavily on walleye fry; given the low abundance of alewife in recent years (Waterfield and Cornwell 2014), it is plausible that natural recruitment of walleye is occurring in Otsego Lake. Walleye spawning has been thoroughly documented (i.e., Foster et al. 2011), though no work has been done to document their recruitment. The intent of this survey was to qualitatively assess the success of walleye spawning in the streams tributary to Otsego Lake. The presence of fry would indicate successful spawning by adult walleye, hatching of eggs, and migration of fry into open waters.

METHODS

On 28 and 29 May, thirteen five-minute tows were performed following guidance from T. Brooking (Pers. Comm.) for a total tow time of about 40 minutes; approximate locations are illustrated in Figure 1. A 1-meter diameter net with 1mm mesh and an attached sample cup was used to collect qualitative samples for fry presence. Tow depth was approximated, ranging from roughly 1 meter below the surface to a depth of 4 meters. No estimates of volume sampled were generated, though distance and tow track were recorded via GPS. Aqueous samples were processed within 24 hours; rose bengal was used to aid in the separation of fry from zooplankton. Zooplankton were discarded, fry were preserved with 70% ethanol and sent to T. Brooking at Cornell Biological Field Station for identification.

RESULTS & DISCUSSION

Samples were dominated by zooplankton, primarily Daphnia spp., though fish fry were collected from many tows. In total, 40 fry were collected (total tow time about 40 minutes), ranging from 6 to 10mm in length. Anecdotally, the greatest abundance of fry occurred in tows with lower zooplankton abundance. Fry were not positively identified to species level, though they were decidedly not walleye fry (Brooking, pers. comm.). NYS DEC staff have suggested that effort should be focused on further investigation of potential walleye recruitment in Otsego Lake; 2014 surveys should include use of a miller sampler for fry sampling.

Figure 1. May 2013 tows to assess presence or absence of walleye fry in Otsego Lake, NY.

REFERENCES

Brooking, T.E., Personal Communication. Cornell Biological Field Station at Shackelton Point, NY. Cornell University.

Cornwell, M.D. 2005. Re-introduction of walleye to Otsego Lake: Reestablishing a fishery and subsequent influences of a top predator. Occas. Pap. No. 40. SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Foster, J.R., J.C. Lydon, R.C. Hilsdorf and C.J. Tizzio.2011. Spawning site fidelity of walleye (Sander vitreus) in Otsego Lake, NY. In 43rd Ann. Rept. (2010). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Waterfield, H.A., and M.D. Cornwell. 2014. Hydroacoustic surveys of Otsego Lake, 2013. In 46th Ann. Rept. (2013). SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.

Willson, J., J.R. Foster, and M.D. Cornwell. 2013. 2012 population estimate of walleye (Sander vitreus) in Otsego Lake. In: 45th Ann. Rept. (2012) SUNY Oneonta Biol. Fld. Sta., SUNY Oneonta.