fisheries€¦ · fisheries • vol 38 no 9 • september 2013 • 389 in videos milton love, alena...

Meet Your New Executive DirectorAFS on Capitol HillDeficient Mining RegulationsFish Diseases in the Salish SeaTuna Spawning in CagesA Common LanguagePay Attention to the TwitchFirst Call for Papers: Québec City FAO Fisheries Reports

FisheriesAmerican Fisheries Society • www.fisheries.org

VOL 38 NO 9 SEPT 2013

03632415(2013)38(9)

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Fisheries • Vol 38 No 9 • September 2013 • www.fisheries.org 389

In VideosMilton Love, Alena Pribyl, and Daniel M. Pauly

In BlogsDerrick Ogle

In NewsNew fish species awaits AFS name approval.

MEMBER SPOTLIGHT 418

NEW AFS MEMBERS 419

427 Fisheries Events

CALENDAR

Contents

Fisheries VOL 38 NO 9 SEPTEMBER 2013

President’s Commentary391 A Call for Better Mining RegulationsThere is clearly room for improvement in North America for how governments develop environmental impact assessments and regulate those mines and wells believed appropriate for development.

Bob Hughes, AFS President

Policy393 AFS Talks Climate Change and Fisheries on Capitol HillThe Potomac Chapter of AFS, in conjunction with the AFS External Affairs Committee, convened a briefing on Climate Change and Fisheries this past May on Capitol Hill. Here are the highlights.

Ward Slacum and Lee Benaka

Guest Director’s Line396 Getting in the Swim of ThingsAFS executive director Doug Austen on his new role and vision for the Society.

John C. Bowzer and Jesse T. Trushenski

Fish Habitat Connections399 Connecting the Habitat Dots: Perceptions and ExpectationsAcross professional and academic fields, as well as among the general public, one challenge in the habitat world is to converse with a common language.

Thomas E. Bigford

The Cast400 The TwitchSometimes just believing in possibilities is all that’s needed in order to actually detect them.

Donald C. Jackson

COLUMNS

402 Infectious Diseases of Fishes in the Salish SeaDiseases of concern for wild fishes in the Puget Sound / Strait of Georgia region.

Paul Hershberger, Linda Rhodes, Gael Kurath, and James Winton

FEATURES

Cover: Juvenile Pacific Herring demonstrating external signs of VHSV, including exophthalmia, focal hemorrhaging around the eyes, mouth, and fin bases, and diffuse epithelial hemorrhaging along the flank. Photo credit: Paul Hershberger.

420 First Call for Papers: Québec City 2014

AFS ANNUAL MEETING 2014

423 New FAO Fisheries ReportsThe Fisheries and Aquaculture Department of the Food and Agriculture Organization of the United Nations (FAO) announces the release of three new publications: “Advances in Geographic Information Systems and Remote Sens-ing for Fisheries and Aquaculture,” “A Global Assessment of Potential for Offshore Mariculture Development from a Spatial Perspective,” and “National Aquaculture Sector Overview Map Collection—User Manual.”

José Aguilar-Manjarrez

ANNOUNCEMENT

396 New executive director Doug Austen holding a King Salmon caught on Oregon’s Columbia River. Photo credit: Jim Martin.

LETTERS TO THE EDITOR 392

410 Morphological and Genetic Identification of Spontaneously Spawned Larvae of Captive Bluefin Tuna in the Adriatic SeaBluefin Tuna spawning in cages.

Leon Grubišić, Tanja Šegvić-Bubić, Ivana Lepen Pleić, Krstina Mišlov-Jelavić, Vjeko Tičina, Ivan Katavić, and Ivona Mladineo

425 North American Journal of Fisheries Management, Volume 33, Number 4, August 2013

JOURNAL HIGHLIGHTS

Fisheries • Vol 38 No 9 • September 2013• www.fisheries.org 390

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The American Fisheries Society (AFS), founded in 1870, is the oldest and largest professional society representing fisheries scientists. The AFS promotes scientific research and enlightened management of aquatic resources for optimum use and enjoyment by the public. It also encourages comprehensive education of fisheries scientists and continuing on-the-job training.

Fisheries (ISSN 0363-2415) is published monthly by the American Fisheries Society; 5410 Grosvenor Lane, Suite 110; Bethesda, MD 20814-2199 © copyright 2013. Periodicals postage paid at Bethesda, Maryland, and at an additional mailing office. A copy of Fisheries Guide for Authors is available from the editor or the AFS website, www.fisheries.org. If requesting from the managing editor, please enclose a stamped, self-addressed envelope with your request. Republication or systematic or multiple reproduction of material in this publication is permitted only under consent or license from the American Fisheries Society. Postmaster: Send address changes to Fisheries, American Fisheries Society; 5410 Grosvenor Lane, Suite 110; Bethesda, MD 20814-2199. Fisheries is printed on 10% post-consumer recycled paper with soy-based printing inks.

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AFS OFFICERSPRESIDENTBob Hughes

PRESIDENT ELECTDonna L. Parrish

FIRST VICE PRESIDENTRon Essig

SECOND VICE PRESIDENTJoe Margraf

PAST PRESIDENTJohn Boreman

EXECUTIVE DIRECTORDoug Austen

FISHERIES STAFFSENIOR EDITORDoug Austen

DIRECTOR OF PUBLICATIONSAaron Lerner

MANAGING EDITORSarah Fox

EDITORSCHIEF SCIENCE EDITORJeff Schaeffer

SCIENCE EDITORSMarilyn “Guppy” Blair Jim BowkerMason BryantSteven R. ChippsSteven CookeKen CurrensAndy DanylchukMichael R. DonaldsonAndrew H. FayramStephen FriedLarry M. GigliottiMadeleine Hall-ArborAlf HaukenesJeffrey E. HillDeirdre M. Kimball

DUES AND FEES FOR 2013 ARE:$80 in North America ($95 elsewhere) for regular members, $20 in North America ($30 elsewhere) for student members, and $40 ($50 elsewhere) for retired members.

Fees include $19 for Fisheries subscription.

Nonmember and library subscription rates are $174.

Jeff KochJim LongDaniel McGarveyRoar SandoddenJesse TrushenskiUsha Varanasi Jack E. WilliamsJeffrey Williams

BOOK REVIEW EDITORFrancis Juanes

ABSTRACT TRANSLATIONPablo del Monte Luna


Mineral mining and oil and gas drilling are essential indus-tries for modern industrial and technological economies. How-ever, mining can negatively alter water bodies and the biota and fisheries that they support (Figure 1). Even at concentrations less than 5 µg/L, heavy metals can alter fish physiology, im-munology, behavior, disease resistance, and growth—leading to reduced population sizes. Ions leaching from coal and metal mines can produce toxic levels of conductivity, alkalinity, and acid that can eliminate aquatic biota across extensive stream reaches. The drilling and transporting of petroleum products have produced devastating spills in aquatic ecosystems, as have failures at mine tailings sites. Fracking for oil and gas can con-taminate surface and groundwater. The burning of fossil fuels has altered the atmosphere and oceans, leading to global climate change, threatening cold water and marine fisheries. Near- and in-stream gravel mining can increase channel erosion, reduce riparian woody vegetation, and degrade aquatic habitats.

Despite these continued threats to aquatic ecosystems and fisheries, mineral exploitation is underregulated. Oil and gas wells in the United States are exempted from regulation by the Clean Water Act and the Safe Drinking Water Act (Hal-liburton Loophole). In the United States, the General Mining

Law of 1872 shifted mineral wealth from federal lands to min-ing firms and billions of dollars in cleanup costs to taxpayers. Wa-ters in Canada that do not support a fishery are not protected by the Fisheries Act of 2012, under which the depo-sition of mining wastes is regulated. Mexico’s PEMEX IXTOC 1 well exploded and spilled oil in 1979, resulting in fisheries closures and loss of estuarine and lagoon species; but, being a federal corporation, PEMEX was immune from fines.

Approximately 10.8% of U.S. streams have a mine within their network, and 152,272 km (2.7%) of U.S. streams have mines in their local catchment. These numbers were calculated based on the National Fish Habitat Partnership mines data set used in the mines figure, aggregated National Fish Habitat Part-nership catchments, and the NHDPlus (v1) (National Hydrogra-phy Dataset) reach information.

It would seem wise to mine and drill only in locations where the public benefits exceed their public costs, and where minerals can be exploited in an environmentally responsible and fish- and fisheries-friendly manner. There is clearly room for improvement in how North American governments develop environmental impact assessments and regulate those mines and wells believed appropriate for development. For an example of an improved environmental assessment regarding a major min-ing district, see www2.epa.gov/bristolbay and reviews thereof by the American Fisheries Society (AFS; Boreman 2013) and the Western Division of the AFS (Swanson 2013). The AFS Re-source Policy Committee is also developing a mining policy that will revise or replace AFS Policy Statement #13: Effects of Surface Mining on Aquatic Resources in North America; see fisheries.org/docs/policy_statements/policy_13f.pdf.

REFERENCES

Boreman, J. 2013. Comments on USEPA’s 30 April 2013 draft as-sessment of potential mining impacts on salmon ecosystems of Bristol Bay, Alaska. American Fisheries Society, Docket # EPA-HQ-ORD-2013-0189, Bethesda, Maryland.

Grabarkiewicz, J. D., and W. S. Davis. 2008. An introduction to freshwater fishes as biological indicators. Environmental Pro-tection Agency, Office of Environmental Information, EPA-260-R-08-016. U.S., Washington, D.C.

COLUMNPresident’s Commentary

AFS President Bob Hughes can be contacted at: [email protected]

A Call for Better Mining RegulationsBob Hughes, AFS President

Figure 1. Percentage of generally intolerant fish individuals as a func-tion of mine density for the conterminous United States (n = 33,538). Mines include coalmine and supporting activities (U.S. Geological Survey 2012), hard rock and aggregate mines (U.S. Geological Survey 2009), and uranium mines (U.S. Environmental Protection Agency 2006). Intolerant fish species are from Grabarkiewicz and Davis (2008) and Whittier et al. (2007). Fish data were provided by the National Fish Habitat Partnership (W. M. Daniel et al., Department of Fisheries & Wildlife, Michigan State University, unpublished data).

Oil and gas wells in the United States are exempted from regulation by the Clean Water Act and the Safe Drinking Water Act (Halliburton Loophole).

Continued on page 426


LETTERS TO THE EDITOR

COMMON FISH NAMES

The 2013 edition of Common and Scientific Names of Fishes from the United States, Canada, and Mexico is out. Generally, this is a time of reflection as once favorite scien-tific names (i.e., the alliterative Pimelometopon pulchrum) give way to more scientifically accurate ones (i.e., Semicossyphus pulcher). In general, I take these changes with something ap-proaching good grace, because, as noted above, there is at least a soupçon of scientific rationale behind them.

Imagine my horror and chagrin, though, when I read in the new edition “Common names in English shall be capitalized.” No explanation, no apology, nothing to ameliorate the angst, just a fiat from the names cabal overturning many decades of tradition.

So, to be precise, we are now expected to write “The most abundant fishes on Richardson’s Reef were the Kelp Bass, Blacksmith, Walleye Perch, and Kelp Perch.”

Words cannot express how profoundly dorky that looks.

Really, when I see Kelp Bass or Walleye Perch, all those capital letters just give me the willies. It looks like those e-mails I get from fourth-graders: “Dear Mr. Milton, for my science project I have to write about a fish. I have picked the Yellow Bearded Rock Sucker. Please send me everything you know about the Yellow Bearded Rock Sucker and please send it to me today because the Yellow Bearded Rock Sucker project is due tomorrow.”

“But,” you say, “ornithologists have been capitalizing the common names of birds forever.”

Uh-huh. So, just because a bunch of chinless birders do something, does that mean you have to do it? If ornithologists decided to jump off a cliff would you jump off a cliff?

You would? Okay, I’ll get back to you.

Milton Love [email protected]

Response from the Editors:Your letter was very timely. Look for an article on this subject in the near future.

PROPER PHOTOS

Too often, electrofishing photos are shown with people not wearing insulated gloves; such was the case with the cover photo of the April 2013 issue of Fisheries. Electrofishing with-out shock protection is not a safe practice and the AFS should not inadvertently support it by publishing such photos. I suggest that if such images are received by contributors, they should be politely rejected with a request for a proper photo. Doing so will (1) make the contributors think about the safety of their practices and (2) send the right message to readers from AFS.

Jim [email protected]

Response from the Editors:The photo in question was taken outside the U.S. where safety standards may have differed, but your point is well taken and we agree. In the future we will look at photographs more closely to be sure they show correct safety and operating procedures.


COLUMNPolicyAFS Talks Climate Change and Fisheries

on Capitol HillWard SlacumVersar, Columbia, MD

Lee BenakaNational Oceanic and Atmospheric Administration Fisheries, Office of Science and Technology, Silver Spring, MD 20910. E-mail: [email protected]

The Potomac Chapter of the American Fisheries Society (AFS), in conjunction with the AFS External Affairs Commit-tee, convened a briefing on Climate Change and Fisheries on May 9, 2013, in the Rayburn House Office Building on Capitol Hill. This was the first Capitol Hill briefing sponsored by the Potomac Chapter since a July 2001 briefing on Aquatic Nui-sance Species and Fisheries.

The briefing drew over 40 people, including staffers from the offices of seven members of Congress, as well as profession-als from the Food and Agriculture Organization of the United Nations, the Natural Resources Defense Council, Oceana, and the Center for American Progress. AFS past-president John Boreman opened the briefing by describing the AFS Climate Change Policy Statement (AFS 2010). President Boreman also described the contents of his January 2013 letter to President Obama regarding climate change.

The briefing included four presentations on a variety of as-pects of climate change and fisheries. Cora Campbell, Commis-sioner of the Alaska Department of Fish and Game, discussed how the North Pacific Fishery Management Council has been taking a precautionary, adaptive management approach in re-sponse to climate change; for example, by approving in 2009 a Fishery Management Plan for Fish Resources of the Arctic Management Area (Arctic FMP). The Arctic FMP (Federal Reg-ister Office 2009) declared that all federal waters of the U.S. Arctic will be closed to commercial fishing for any species of finfish, mollusks, crustaceans, and all other forms of marine animal and plant life. Commissioner Campbell also expressed concern regarding several recent attempts to list corals and seals under the Endangered Species Act based on possible future ef-fects of climate change. According to Commissioner Campbell, it makes more sense to allocate scarce resources for research and monitoring rather than for responses to Endangered Species Act petitions.

Malin Pinsky of Princeton University focused his presenta-tion on the concept of climate velocity and how fish and fisheries respond to changing temperatures across seascapes. According to Dr. Pinsky, fish stocks shift their distributions at much the same rate, and in the same direction, as climate velocity. For ex-ample, many species expanded their range northward as waters warmed. Fisheries usually follow these shifting fish stocks, but do so more slowly than the rate of the migrating fish. Fisheries managers can help fishing communities and fish stocks adapt to these changes by ensuring that fish habitat remains intact,

Cora Campbell—Commissioner of the Alaska Department of Fish and Game.

Then-president John Boreman opens the session.

by setting sustainable fishing quotas, and by incorporating an ecosystem focus into management measures.

Jon Hare of the National Oceanic and Atmospheric Admin-istration (NOAA) Fisheries’ Northeast Fisheries Science Center discussed the need to consider habitat and climate variables in the science enterprise. Dr. Hare noted that climate is variable and also changing in the Northeast, and impacts to fisheries will occur causing “winners” (e.g., Atlantic Croaker) and “losers”


(Atlantic Cod) as species abundance and distributions shift to adapt to warming ocean temperatures Dr. Hare concluded that the current scientific enterprise should be augmented by adding climate variables, reaching out to management and stakehold-ers, and completing climate vulnerability analysis to evaluate biological productivity and fishery sustainability in the context of future environmental conditions.

The final presenter, Leaf Hillman of the Karuk Tribe of California, discussed the cultural effects of climate change on his fisheries-dependent tribe. Mr. Hillman described the threats to fisheries in the Klamath River basin, including mining, com-mercial fishing, irrigated agriculture, mismanaged landscapes, and dams that restrict fish from colder waters as available river and stream waters warm due to climate change. As fish have become less available, members of the tribe have been less able to pass down fishing traditions and have experienced higher rates of heart disease and diabetes. Mr. Hillman recommended increasing river flows, improving habitat, and increasing access to coldwater refugia as solutions to his tribe’s climate change challenges.

This successful Capitol Hill briefing showcased the AFS Potomac Chapter’s ability to facilitate the communication of AFS messages to leaders in Washington, D.C. The AFS planned this meeting in conjunction with the NOAA Fisheries’ Manag-ing our Nation’s Fisheries III conference, which saved travel expenses for some speakers during this time of tight travel bud-gets. The National Sea Grant College Program’s Knauss Fel-lows, some of whom serve in offices of Capitol Hill, were also important to this briefing’s success through securing a meeting room and publicizing the briefing. The AFS Potomac Chapter looks forward to continuing to help the AFS communicate its positions and value to members of Congress and other leaders in Washington, D.C.

REFERENCES

AFS (American Fisheries Society). 2010. Policy statement on climate change. Available: http://fisheries.org/docs/policy_statements/policy_33f.pdf. (June 2013).

Federal Register Office. 2009. Fisheries of the United States Exclusive Economic Zone off Alaska; Fisheries of the Arctic Management Area; Bering Sea Subarea, Final Rule. 74 Federal Register 211 (November 3, 2009), pp. 56734–56746.

Ward Slacum, acting President of the AFS Potomac Chapter, introduces Malin Pinsky of Princeton University.

Jon Hare—NOAA Fisheries’ Northeast Fisheries Science.

Leaf Hillman of the Karuk Tribe of California.

The meeting took place in a packed conference room at the Rayburn House Office Building on Capitol Hill.


John C. BowzerCenter for Fisheries Aquaculture and Aquatic Sciences, Southern Illinois University Carbondale, Carbondale, IL 62901. E-mail: [email protected]

Jesse T. TrushenskiSouthern Illinois University Carbondale, Carbondale, IL

We caught Doug Austen coming back from a jaunt to the Conodoguinet Creek with his two English Springer Spaniels—Cooper and Bailey—and managed to interview him (with the occasional “the dog just stepped on my leg” sentence interjected here and there). He was relaxed, on vacation with his family (wife, Lezli, and their fourteen-year-old twins, Ally and Zach), and getting ready to come on board as the new AFS executive director. But consider this. AFS is in a huge transition period; it’s managed to get through the thick of the economic crisis and remain in the black, shore up its web presence, turn Fisheries magazine into a high-impact factor journal cum membership magazine—but now there’s a need to put more focus on our newer members who live in a vastly different world than the members who signed on twenty (or even ten) years ago, all while helping them in their vocations, and still continuing to maintain our solid reputation and keeping with the objectives in the AFS Constitution. That’s a tall order. But Doug comes

COLUMNGuest Director’s Line

Getting in the Swim of ThingsAFS executive director Doug Austen on his new role and vision for the Society

with energy and a healthy respect for this need for change. And, maybe most important of all, he’s a fish guy. So how does he see his role in this new process of securing the Society’s cur-rent and future relevance in the fisheries community and in broader contexts? Read our interview with Doug and see why past-president Don Jackson said, “Doug is a heck of a fine guy. We’re lucky.”

Bowzer/Trushenski: Growing up in the suburbs of Chicago, what were your early fishing experiences?

Doug: I’m not sure that my Dad really was interested in fishing but he took me because I really wanted to fish. We went fishing on family vacations and an occasional trip to Midwest locations like Lake Michigan, Wisconsin’s Wolf River, and Michigan’s St. Joseph River. Most of my local fishing was in pretty unglam-orous highway borrow pits and area ponds, and about the only thing I caught was stunted bluegill, carp, and maybe a rare bass, but I was hooked.

Bowzer/Trushenski: In high school you knew you wanted to be a fisheries biologist, but did you even know what that might entail?

Doug: It’s funny, but back then you would ask guidance coun-selors about fisheries biology and they would look at you with this dumbfounded expression. They just had no idea what that meant. Now they probably still don’t know, but can do an In-ternet search and figure it out.

Bowzer/Trushenski: Your fisheries career has been broad—geographically and experientially—so how do you see the world of fisheries having changed from then to now?

Doug: People are not necessarily getting into the fisheries field for the same reasons that they used to. Fisheries used to attract primarily anglers and others actively using the resource, but in today’s classrooms and agency rosters, you’re more likely to find people who were drawn to fisheries by more general inter-ests in conservation, ecology, or environmental issues.

Bowzer/Trushenski: Do you believe this transition will help drive positive changes in our profession?

Doug: Yes. It will create greater diversity in terms of perspec-tives, expertise, and values, as well as gender, race, and ethnic-ity—and it will affect our Society in the same positive way. Our diverse pool of expertise provides the Society with tools needed

Doug Austen holding a King Salmon caught on Oregon’s Columbia River. Photo credit: Jim Martin.

Doug earned degrees in fisheries sciences at South Dakota State University (B.S.), Virginia Tech (M.S.), and Iowa State University (Ph.D.). After completing his formal education, he held positions with the Illinois Natural History Survey, the Illinois Department of Natural Resources, and served for six years as the Executive Direc-tor of the Pennsylvania Fish and Boat Commission. Prior to coming to AFS, Doug was the national coordinator of Landscape Conserva-tion Cooperatives for the U.S. Fish and Wildlife Service.


to tackle contemporary and future challenges as they arise. Our role, as a Society, is to pull together the state of the science, explore how complete or incomplete it is, and give voice to our members’ perspectives. Many of these issues are too com-plex to be resolved in the short-term, but the very exercise of wrestling with the issues—which the Society can do in a neutral way—is valuable in understanding and moving the dialogue for-ward without devolving into taking sides.

Bowzer/Trushenski: You’re a self-described “fish guy.” That’s important. You’re well positioned to communicate the value of fisheries and AFS to the public, to strengthen existing lines of internal and external communication, develop new vehicles for education and outreach, and empower our members to do the same. What are the priorities on Doug’s to-do list?

Doug: Certainly, my career started out as being a fishery biolo-gist and that was the focus for many years, but it also included work on large collaborative efforts such as MARIS (the Multi-state Aquatic Resources Information System), the National Fish Habitat Action Plan, and recently the Landscape Conservation Cooperatives (LCC). Along with time in some leadership roles and involvement in state and federal capital politics, I hope that I can bring some helpful perspectives and experiences to AFS. People see AFS as conveying an unbiased, impartial message that is based on science. We can continue to do that through Fisheries and our other peer-reviewed publications, our web-site, briefings, and by developing partnerships with mainstream media and other groups with common interests. Organizations interested in aquatic conservation—and there are many—look to us to provide credible, science-based information. By partner-ship and by retooling our existing modes of communication, we can create a broader circle of influence.

Bowzer/Trushenski: You sound committed to challenging your-self and all fisheries professionals to remember what brought us to fisheries in the first place, what our connection to the resource is, and to give some of our best back to the profession and the Society. What do you mean by “best?”

Doug: Is your best doing science, mentoring students, and pub-lishing in our journals? Then do that and do it at the highest level you can. Is it being the best person at the front of the shocking boat or doing field work? Then do that, and maybe consider showing someone else what you know along the way. It’s then the Society’s responsibility to create a receiving en-vironment, make members aware of opportunities available to them, give them room to be their best, and help them realize a real return on their investment. We must be able to engage groups via dialogues and teams… create a series of events and conversations where we accomplish something.

Bowzer/Trushenski: What is your philosophy about students?Doug: As far as I’m concerned, students aren’t the AFS of the

future; they’re the AFS of right now. Students provide different perspectives, ways to communicate, and philosophies that need to be part of the conversation. There’s always some tension be-tween students and younger members and the old guard at AFS. That happens in virtually every organization. It’s a little bit of a generational thing, along with changes that have taken place over time. But this can be a very healthy issue. These younger members want to know what AFS is going to do to open doors for them that will allow them to make their life better or richer. They’re less enthralled with the fact that we’ve been around for hundreds of years. They want to be with organizations that can make a positive impact on the resource—and there are many good groups doing excellent work out there. For example, look at all the information that people can get now via blogs, elec-tronic newsletters, dynamic web sites, and much more. How much have we done to address this for the younger generation or, for that matter, for all members? I’m not sure we’ve done nearly what we can or should. We may not be connected in what their needs are. It’s different than it was when I was a younger guy. When I talk to them, they tell me they haven’t heard from AFS in years. They get nothing. “I let my membership expire,” they say. What should we be doing to really help them in their career that fits in with the realty of their situation? What is it that we should be doing that will help them? I’m not sure we really know, but it is my priority to find out.

Bowzer/Trushenski: You will have to be quite a leader to get this done. How do you lead? What are your challenges?

Doug: My favorite definition for leadership is “making change happen at a rate that people can accept.” Don’t shy away from it, but also don’t cram it down people’s throats, unless, of course, there is something so important that the society will die if we don’t do it. We need to continuously and regularly press forward and do things, in ways that won’t break the bank or cause people to think subversive thoughts. Part of our job, particularly my job, will be to help create the conversations at the right time with the right people and target those issues and get the people to buy into the solutions with them. As for my challenges? There are a lot of things I could strengthen about myself. I’ve got weaknesses all over the place. But what I’ve been able to do is to find people to work with me and be com-petent with those things.

Bowzer/Trushenski: We understand that during the interview process you described the concept of “re-setting” the conversa-tion with the membership. What does that mean?

Doug: Like any organization, AFS needs to ensure that it is relevant and of value to its members. There is a great deal of competition for the time, attention, and money of our members: new journals to read and publish in, many other groups that are increasingly effective in carrying the ball for aquatic resources and science, and the general busyness of all of us, etc. What is it about AFS that is unique, that attracts their attention, which responds to a need? What makes someone want to become a member of AFS and contribute to the success of the AFS mis-sion? We need to make sure that we know the answer to these

As far as I’m concerned, students aren’t the AFS of the future; they’re the AFS of right now.


questions—not what the answers used to be, but what they are now and in the near future. We need to understand what is missing in the professional life of our members and how we can address that vacuum. We need to know the kind of information that members need, the best way to deliver it, and the timeli-ness of that delivery. We need to know how to better provide for the social and professional interaction that is so important but is also clearly and increasingly not supported due to policy issues, budget cuts, and politics. In essence, we need to re-think how we do our job, to re-set a new conversation about the role of AFS. Likely, such a conversation will validate many cur-rent practices, and should also challenge us and demand new approaches.

Bowzer/Trushenski: Tell us how you will go about doing this re-setting?

Doug: First of all, a substantial part of my job is to bring alive the vision of the AFS leadership. The Society’s elected lead-ership will be a key part of ensuring that we have a vision of the future that is on-target, energetic, and moves us in the right direction. So I need to understand their vision and help them—in whatever way I can—craft the vision, based upon the best information possible, so we can make it come to life. We will do this, together, through a mixture of spending old-fashioned, face-to-face time with members just talking, identifying needs, and debating options. I would love to visit as many chapters, sections, and divisions as possible in the next year to create as many opportunities as possible for these dialogues. We will also use whatever media we can to engage the many members who won’t be able to be a part of these in-person conservations. Our staff has some great ideas for how we can vastly expand our tools to address this need, and I hope to be able to employ those that will be most effective.

Bowzer/Trushenski: We know that one of your core interests has always been professional development of staff. Back in Illinois you invested a great deal of time in creating continu-ing education programs for the Illinois chapter, you worked on leadership development at the Fish and Boat Commission and actively encouraged staff, including yourself, to be involved in the National Conservation Leadership Institute. How do you see this playing out as AFS executive director?

Doug: Continuing education in the Illinois chapter of AFS was a direct response to a need of the members. In most state agen-cies, those hired come in with a Masters degree and are well educated in the academic sense. They then learn on the job but, unfortunately, that is often the extent of the training provided. States simply don’t have the resources or expertise for exten-sive training programs for agencies as specialized and relatively small as the natural resources agencies. What training the states provide is usually generalized and difficult to apply to the situa-tions that our members find themselves in. The federal agencies are, for the most part, substantially different. Their size and budgets provide for a much richer opportunity for training. The obvious example is the tremendous resource afforded by the Na-tional Conservation Training Center, the leadership training that

most federal agencies offer, and the tremendous opportunity for short term “details” or other vehicles to provide staff with new experiences. AFS can and does provide an invaluable service in organizing continuing education at multiple levels, but we can and must strive to do more. We need to broaden the opportuni-ties for those with limited access to have training available and be supported in taking part in that training. We need to create more opportunities for members to have new experiences that will enrich them professionally and find ways to support these opportunities. This is critically important if we are to address the challenges we face as our aquatic resources become increas-ingly stressed.

Bowzer/Trushenski: What exactly is the job of executive director?

Doug: Members should feel confident that someone is there that knows enough about fisheries to be an effective advocate for the resource and the science that is a critical part of AFS. It’s important to also recognize that fisheries and aquatic resources management and science are increasingly diverse and special-ized. We need to be able to draw upon the AFS membership to engage and benefit from that expertise in order to effectively address the challenges that we face and I hope to be able to do that. Members should get good gains for their investment in the Society. I’ve been involved in enough national activities that I can be the person for AFS who can go nose to nose with the people in Congress and be able to argue persuasively. I’m an advocate for the resource, and I bring people together who can find solutions to our fisheries resource problems. In this job, I will help to orchestrate opportunities to protect fisheries and aquatic resources. In fact, the first objective in the AFS Consti-tution is to “promote the conservation, development, and wise use of fisheries.” So one of my main functions is to create an environment that will help to ensure that we make the mission and vision of the Society come to life. This includes a wide va-riety of roles such as finances, effective and creative meetings, excellence in publications and communications, advocating for development and use of sound science, and much more.

Bowzer/Trushenski: Any closing thoughts?

Doug: To me, AFS was how I grew and matured in the profes-sion. AFS has been the source of many of my lifelong friends and continues to be a strong part of my social network, almost like an extended family. AFS has also been a place that creates and facilitates opportunities to work for the resource in ways above and beyond what we can do in our day-to-day jobs or in support of those jobs. AFS is a place that I am thrilled to be working for and I look forward to the years ahead. I hope to see many of you in Little Rock, and for those who can’t make it, or haven’t found the time to come meet me, please keep this conversation going through the Talk AFS blog at www.talkafs.wordpress.com. Please share your thoughts, concerns, and sug-gestions with me about anything related to AFS. Or just come say hello. Hearing from our members will always be my first priority.

You can also contact Doug directly at [email protected].


It is reassuring that there are solu-tions or at least rea-sonable options. To catch up with other fields, the fish habitat community must accelerate its efforts and then persist. The gaps deserving our immediate attention will be the subject of a future column, but hints are everywhere. Because the habitat world lags behind population dynamics and hydrological modeling (and others) on the knowledge curve, I often feel that our work includes equal measures of explora-tion, discovery, and learning. With each approach we must con-vey knowledge to citizens and professionals alike. Sometimes the gap is within our own organization or is manifested as dis-agreement between groups or perhaps directly with the public. Because of the daunting information gaps, learning for habitat professionals and sharing with colleagues and the general public is a career-long obligation. The same is true for teaching, the obverse of learning. The combination represents the increased awareness so deserving of fish habitat.

Fisheries is not your typical magazine, so it’s safe to assume that readers of this column have a solid academic background plus coursework in at least one advanced field of study—social sciences, engineering, natural sciences, law, communications, or the like. And the majority are likely to be professional fish folk of some kind. Across those fields, one challenge in the habitat world and with the public generally is to converse with a common language. Our words and messages must connect with our audience of academic leaders, agency representatives, sector associations, aspiring students, adoring family members, and the semi-interested hordes who rely on mass media for mes-sages. We must speak of fish, not ichthyology, bottom habitat rather than benthic rugosity, quality of life instead of ecosystem services, environmental change, not ecogeomorphic feedback. That may sound condescending, but I’ve been to many meetings where strong fish messages accompanied by statistical punc-tuation points were easily trumped by simple messages about jobs and the local economy. We need deeper knowledge about habitat and then we need to translate that for each audience. Continued learning, coupled with careful communications, will entice our many publics to lean toward the importance of sound natural resource management. As the old proverb says, “It’s not easy to teach an old dog new tricks.” Fortunately, most of you are younger than me and may even be dog whisperers.

Welcome to the fifth column in this series on fish habitat. In May we introduced plans to use this forum to elevate habitat awareness. The June column explained how habitat connects American Fisheries Society units and members. In July, I fo-cused on the need for urgent and sustained action. The August column explained how we must think and act on an ecosys-tem scale. This month we’ll cover the importance of education, awareness, and overall creativity, for both habitat aficionados and the many other fisheries professionals and students who are vital to our success.

Beginning with subtle messages in elementary school, continuing beyond college graduation, and finally reflected in our professional life, we learn and apply valuable lessons about habitat. One common theme is change. Though not unique to habitat, the shifts, twists, and surprises do seem endless and often inspire us to be more creative than the typical fish person. Happily, the unforeseen often delivers bona fide opportunities. Though the habitat arena has been the malnourished aspect of aquatic resource management, it is now primed to be a key player. It’s up to us to seize the moment and learn on the fly.

Habitat professionals usually greet new opportunities with unbridled enthusiasm. I know that many of us view landscape-scale efforts, research initiatives on ecosystem connections, cli-mate change, and other opportunities as the next best chance to educate, to convince, to advance our agenda. New roles challenge us to learn and then communicate better. While we craft precise messages, we are reminded that our knowledge is limited. To tackle climate issues, we need to know how rivers morph over the course of centuries. To mitigate the effects of dams and to meet angler demands, we need to design reservoirs and passage facilities to last beyond the typical lifetime. To de-cide how to manage wetlands, we need to understand the jux-taposition of habitat protection and coastal development. With education, we learn, we make better decisions, and our society benefits.

COLUMNFish Habitat ConnectionsConnecting the Habitat Dots: Perceptions

and Expectations Thomas E. BigfordOffice of Habitat Conservation, NOAA/National Marine Fisheries Service, Silver Spring, MD 20910. E-mail: [email protected]

Continued on page 426

“We have demonstrated the value of place-based, collab-orative approaches to habitat conservation. They stimulate innovative thinking, uncover previously unrecognized col-laboration, and create a contagious energy and enthusiasm to work together to achieve common objectives.”

From Eric Schwaab, formerly director of National Oceanic and Atmospheric Administration/National Marine Fisheries Service, now senior vice president and chief conservation officer of the National Aquarium in Baltimore.


Although I live in a coastal state (Mississippi), I don’t do a lot of saltwater fishing. At best I make the 4-hour drive down to the coast two or three times a year and fish a day or two on each trip. Most recently,

this past June, I was fishing for Red Snapper with my brother and my daughter. The trip was my daughter’s high school gradu-ation present. We fished for 2 days.

On the second day the fishing was somewhat slow. We’d caught a few Snappers, a few King Mackerel, and several At-lantic Sharp-Nose Sharks. The day was very hot and we began to worry about our supply of beer in the boat’s coolers. Waves gently rocked the boat. The air was thick. We daydreamed. Con-versation was minimal. It was hard to stay focused on the rods in our hands, the line, and, in fact, the whole fishing enterprise. And the beer was running out.

I was standing beside my brother and happened to notice a slight twitch on his line. Since we were using circle hooks, technique demands that there be no jerk to set the hook. The hook sets itself if the fisher simply tightens up. My brother did this and nearly got yanked out of the boat. He was fast on to something very big—actually, not big but absolutely huge! He fought the thing for about 20 minutes before we got a glimpse of what he’d tied into. It was a monster Hammerhead Shark about 7 feet long with an estimated weight of over 200 pounds.

After about a half an hour of fighting the beast, my brother was ready to hand off the rod to somebody else. Everybody on the boat got to feel the weight, the incredible strength, and the unyielding determination of that shark. We didn’t want to actually catch it; and we certainly did not want it in the boat, thrashing and snapping and wreaking havoc. It was a fish that belonged in the ocean. It was our privilege to be attached to it—just for a while. Finally, we broke off and the shark settled back into the dark depths of the Gulf.

It all started with a twitch, a barely noticeable twitch, one almost imperceptible. But the results were glorious and made much more so by my brother’s willingness to share with us all the source of that twitch.

COLUMNThe Cast The Twitch

Donald C. JacksonDepartment of Wildlife, Fisheries, and Aquaculture, College of Forest Resources, Mississippi State Univer-sity, Mississippi State, MS 39762. E-mail: [email protected]

Fast-forward 3 weeks. I met the dawn the next morning on my pond. It is a small pond, a shade larger than an acre in size. It’s located in a remote corner of my 50-acre farm, which itself is located in a remote corner of Mississippi way down at the very end of a narrow and innocuous gravel road. The dawn was full of hush, a hush made more intense—almost overpowering—by the flutelike calls of a wood thrush down in the shadows of the forest below my pond. A wood duck flushed from the pond at my approach. Prothonotary warblers (“swamp canaries”) flitted and flashed yellow and blue among the willows.

I went to my little fishing boat that I keep out there on the pond’s edge and loaded my dog and my fly-fishing equipment. My assignment was to collect the basic ingredients for a family fish fry scheduled for that evening, those basic ingredients being fat Bluegill that I knew would be cruising just outside the littoral zone of the pond.

There was no wind. The surface of the pond was like a mir-ror. From time to time I could see swirls as a Bluegill came to the surface to snatch an insect. It was a perfect morning to fish a dry fly. So I selected a small popping bug, a fly that has a cork body, rubber legs, a few hackles, and a tail made of feathers. It is supposed to mimic a grasshopper.

My rod is a custom-built rod, designed specifically for Bluegill fishing in ponds. For those who care about such things and might be interested, it is a two-weight, 7-foot rod. The length gives me casting distance. The light weight allows me to feel every fin twitch when I’m playing a hooked fish. It is the perfect tool for the job—almost a magic wand.

I was soon rewarded with several nice Bluegills with lengths ranging from 8 to 13 inches. The fish would come to the surface and hit that bug with a variety of approaches. Some would slash. Some would slurp. Some would swirl at it and then take it under the surface. I didn’t hook fish on every strike. In fact, I missed as many as I hooked and I also lost some that were hooked but pulled free while I was trying to bring them to the boat. But the beautiful thing about fishing on the surface is that I can see what’s going on (or think I can). Even when I miss a strike there’s still the thrill of the splash.

Eventually the fish stopped hitting the surface bug, so I switched to a wet fly called a Brim Killer. It looks like a black spider with white legs. I cast it, let it sink a little, twitch it some, then retrieve. When a fish strikes this underwater fly it is harder to detect the strike. There’s no splash and dash. At most there’s just a slight twitch in the line. Sometimes the twitch isn’t there at all but rather the end of the line goes almost imperceptibly slack for only a fraction of a moment. I’m still using the same

We didn’t want to actually catch it; and we certainly did not want it in the boat, thrashing and snapping and wreaking havoc. It was a fish that belonged in the ocean.


the same tool, and the different sorts of responses I might get depending on how I use it under different conditions.

But most of all that morning I thought about the power of the “twitch”—that virtually undetectable small signal that something of significance is happening—out of sight, but still within the realm of the senses—or intuition. The twitches are easy to overlook. If there are ripples under the “surface” of an experiment, you may not see the twitches at all, but still they are there. Sometimes just believing in their possibility is all that’s needed in order to actually detect them. A few times I wasn’t sure I’d had a strike but reacted as if I had. Something sublimi-nal had alerted me and I was rewarded with a fish on the end of my line.

The most successful anglers are those who think that they are going to catch fish. The most successful scientists are those who believe that they will discern truth. Focus, faith, and sensi-tivity. Pay attention to the twitch.

rod, the same line, the same leader as earlier, but with the dif-ferent fly it is a very different game than fishing on the surface. Under the right conditions the different technique can yield phenomenal results. That morning, the fish I caught below the surface were on average much larger than those I caught on the surface. They filled my heart.

The key to both techniques is paying attention to what’s going on and recognizing the differences in potential responses to treatments. I was trying to catch the same species of fish and I needed fish with particular characteristics (in this case, ap-propriate size for eating) to meet the ultimate objective of my enterprise—an evening fish fry for my family.

As a scientist I couldn’t help but recognize the metaphor from fishing—how it relates to my professional endeavors in research, the design and analyses of my experiments, the tools I use to detect responses to treatments, the different ways I use


FEATURE

Infectious Diseases of Fishes in the Salish Sea

Enfermedades infecciosas en los peces del mar de SalishRESUMEN: Tal como sucede en las regiones marinas de otras partes del mundo, los peces del mar de Salish hospedan cantidad de patógenos, incluyendo nematodos, tremátodos, protozoarios, protistas, bacterias, virus y crus-táceos. En este trabajo se hace una revisión de algunas de las enfermedades infecciosas mejor documentadas que muy posiblemente contribuyen a la pérdida significativa de peces en el mar de Salish y se discuten los factores am-bientales y ecológicos que pueden influenciar los impactos de la enfermedad a nivel poblacional. La demostración de estas enfermedades y sus consecuencias en recursos críti-cos y amenazados, justifica expandir la vigilancia sobre patógenos e incorporar un pronóstico de enfermedades y herramientas de mitigación en los esfuerzos de restaura-ción de ecosistemas.

Paul HershbergerU.S. Geological Survey, Western Fisheries Research Center, Marrowstone Marine Field Station, 616 Marrowstone Point Road, Nordland, WA 98358. E-mail: [email protected]

Linda RhodesNorthwest Fisheries Science Center, National Marine Fisheries Service, National Oceanic and Atmospheric Administration, Seattle, WA

Gael Kurath and James WintonU.S. Geological Survey, Western Fisheries Research Center, Seattle, WA

ABSTRACT: As in marine regions throughout other areas of the world, fishes in the Salish Sea serve as hosts for many pathogens, including nematodes, trematodes, protozoans, protists, bacteria, viruses, and crustaceans. Here, we review some of the better-documented infectious diseases that likely contribute to significant losses among free-ranging fishes in the Salish Sea and discuss the environmental and ecological factors that may affect the population-level impacts of disease. Demonstration of these diseases and their impacts to critical and endangered resources provides justification to expand pathogen surveillance efforts and to incorporate disease fore-casting and mitigation tools into ecosystem restoration efforts.

INTRODUCTION

The Salish Sea encompasses the nearshore marine waters of western Washington and British Columbia, including Puget Sound, the Strait of Juan de Fuca, and the Strait of Georgia (Figure 1). The region was previously referred to as the Puget Sound–Georgia Basin; however, the official name was recently changed to pay homage to numerous groups of indigenous peo-ples, collectively referred to as the Coast Salish, who continue to inhabit coastal areas throughout the region.

Recent restoration and research efforts for the Salish Sea have been championed by the Puget Sound Partnership, a Wash-ington State agency charged with coordinating federal, state, local, tribal, and private resources and creating an action agenda, a strategy for cleaning up, restoring, and protecting Puget Sound by 2020 (Puget Sound Partnership 2013). The objec-tives of the action agenda are heavily weighted toward achiev-ing tangible habitat and restoration goals through the cleanup or remediation of environmental contaminants and the reduction of nutrient loadings. Further, numerous salmon recovery plans, the Georgia Basin Action Plan, and other restoration efforts are intended to bridge national and international jurisdictional boundaries in the Salish Sea (Gaydos et al. 2008); however, none have incorporated the effects of disease into their resto-ration plans for marine fishes or other marine animals. These oversights are unfortunate, because massive disease epizootics repeatedly cause drastic population declines and extirpations in

wild animal populations (Smith et al. 2009), including popula-tions of marine fishes (e.g., Jones et al. 1997; McVicar 1999). The impacts of these disease-mediated population declines can cascade throughout the food web and indirectly affect sympatric species (Dann et al. 2000).

Here we introduce some of the infectious and parasitic dis-eases affecting marine and anadromous fishes in the Salish Sea. As in marine regions throughout the world, fishes in the Sal-ish Sea serve as hosts to a broad range of pathogens, including bacteria (e.g., Renibacterium sp., Listonella (Vibrio) spp., and Tenacibaculum spp.), viruses (e.g., infectious hematopoietic ne-crosis virus, viral hemorrhagic septicemia virus, erythrocytic necrosis virus, infectious pancreatic necrosis virus), myxozo-ans (e.g., Parvicapsula, Henneguya, and Kudoa spp.), coccid-ians (e.g., Goussia spp.), monogenean and digenean trematodes (e.g., Gyrodactylus and Nanophyetus spp.), protists (e.g., Ich-thyophonus sp.), and parasitic crustaceans that are collectively referred to as “sea lice” (e.g., Lepeophtheirus, Caligus, and Argulus spp.). The impacts of these and other pathogens on free-ranging fishes remain largely underinvestigated, to a great extent because diseases are very difficult to visualize in marine systems, where dead and moribund hosts sink to the bottom and are consumed by predators. Further, endemic pathogens typi-cally exist chronically and covertly, with host- and population-level impacts occurring only periodically when host, pathogen, and/or environmental conditions shift in favor of the disease. The diseases described below do not constitute a comprehensive list, nor are they reviewed in detail; rather, we begin with those that have been studied more thoroughly and are generally ac-cepted to cause population-level losses. We end by discussing emerging threats, including the introduction of novel pathogens


and the synergistic effects of multiple stressors imposed by infectious diseases, habitat alteration, environmental contami-nants, and climate change on the health and survival of fishes in the Salish Sea.

DISEASES POTENTIALLY HAVING POPULATION-LEVEL IMPACTS

Ichthyophoniasis

Ichthyophonus sp. is an internal protistan (Mesomyceto-zoean) parasite, primarily of marine fishes, with an extremely broad host range that includes more than 80 known fish species (McVicar 1999). The fungal-like parasite occurs in wild and cultured fishes throughout the world, including the Salish Sea. Though numerous hypotheses have been proposed, the complete life cycle of Ichthyophonus has not yet been fully described, with the most obvious gap involving route(s) of transmission for planktivorous fishes like Atlantic/Pacific Herring (Clupea har-rengus/pallasii) and American Shad (Alosa sapidissima). This information gap has largely hindered progress in developing an understanding of host and environmental conditions that preface outbreaks of Ichthyophonus disease (ichthyophoniasis) in wild fishes.

In the North Pacific Ocean, mortality from diseases, in-cluding ichthyophoniasis, represents a leading hypothesis ac-counting for the decline and failed recovery of Pacific Herring populations in Prince William Sound (Marty et al. 2010). In other clupeids, an epizootic of ichthyophoniasis occurred among American Shad in the Columbia River, with infection prevalence reaching 72% near the peak in shad abundance and then decreasing with host population declines (Hershberger et al. 2010c). Among non-clupeids, an ichthyophoniasis epizootic occurred in adult Chinook Salmon (Oncorhynchus tshawyts-cha) in the Yukon River in Alaska, where infection prevalence exceeded 40% (Kocan et al. 2004). Recently, as with Ameri-can Shad, the infection prevalence in Yukon River Chinook Salmon decreased as the population biomass declined (Zuray et al. 2012). In addition to causing host- and population-level impacts through direct mortality and decreased swimming per-formance (Kocan et al. 2006), ichthyophoniasis can result in economic losses by causing macroscopic lesions in the skeletal muscle of infected hosts that negatively affect the marketability of diseased fillets.

Throughout the Salish Sea and the northeast Pacific Ocean, Ichthyophonus is ubiquitous in populations of Pacific Herring (Hershberger et al. 2002); however, the parasite also occurs in juvenile American Shad, Pacific Tom Cod (Microgadus proxi-mus), Speckled Sand Dab (Cithanichthys stigmaeus), Surf Smelt (Hypomesus pretiosus), Puget Sound Rockfish (Sebastes emphaeus), Copper Rockfish (S. caurinus), and likely many other species (Rasmussen et al. 2010). Of all of these hosts, the epizootiological relationships have been most thoroughly examined in Pacific Herring, where the prevalence of infection increases with host size and age (Hershberger et al. 2002). In-fection prevalence as high as 48% in populations of adult Her-ring from the Salish Sea (P. Hershberger, unpublished data) are concerning because Ichthyophonus can be highly pathogenic to Pacific Herring (Figure 2), with laboratory exposures often resulting in rapid mortality (Kocan et al. 1999). The combina-tion of high pathogenicity and increasing infection prevalence with age indicates that Ichthyophonus may be an important fac-tor influencing recent zoographic trends in Pacific Herring in Puget Sound, where estimates of annual natural mortality (ex-clusive of commercial fishing) increased from 20% to 40% in the 1970s to 68% in the years since 1990 (Stick and Lindquist 2009). Most of this mortality has occurred among the oldest age cohorts having the highest prevalence of Ichthyophonus infec-tion. As a result of this age-biased mortality, the median age of Pacific Herring in Puget Sound has decreased in recent years, and newly recruited cohorts (age 2–3 years) currently represent the majority of the spawning stock biomass.

Viral Hemorrhagic Septicemia

Viral hemorrhagic septicemia virus (VHSV) is a fish rhab-dovirus that is best known as an important pathogen affecting Rainbow Trout (Oncorhynchus mykiss) in European aquacul-ture. However, in the late 1980s, isolations of the virus were made from apparently healthy adult salmonids returning from the Salish Sea to spawn at state, tribal, and federal hatcheries

Figure 1. Map of the Salish Sea and surrounding basin. Included with permission of the cartographer, Stefan Freeland, Western Washington University, 2009.


(Meyers and Winton 1995). Because VHSV is listed as report-able to the World Organization for Animal Health (OIE), ex-tensive hatchery fallowing and disinfection procedures were immediately implemented in an effort to eradicate the virus. Additional isolations of VHSV from a range of marine and anadromous species indicated that the virus was broadly pres-ent in the North Pacific Ocean and genetic analysis indicated that these isolates subscribed to a novel genotype (commonly referred to as the North American strain or Genogroup IV) that is distinct from European types (designated Genogroups I–III). More recently, VHSV emerged in the Laurentian Great Lakes during the mid-2000s and massive epizootics occurred in cer-tain fish species; this freshwater virus, a sublineage of the North

American strain designated subgroup IVb, is genetically distinct from the type occurring in the Salish Sea (and greater northeast Pacific), which is now designated as subgroup IVa.

In the Salish Sea and the northeast Pa-cific Ocean, VHSV has a broad host range, occurring in Pacific Herring (Figure 3), Pa-cific Sardines (Sardinops sagax), Pacific Sandlance (Ammodytes hexapterus), and other marine fishes (Hedrick et al. 2003). Among free-ranging fishes in the Salish Sea, large-scale disease outbreaks accompa-nied with high mortality are most common in populations of Pacific Herring and Pacific Sardines (Traxler et al. 1999), and in Alaska, the disease has also been reported to affect Pacific Hake (Merluccius productus) and Walleye Pollock (Theragra chalcogramma; Meyers et al. 1999). Mortality from VHSV (along with ichthyophoniasis and other dis-eases) is believed to be a compounding factor associated with the decline and failed recov-ery of Pacific Herring populations in Prince William Sound, Alaska (Marty et al. 2010).

Laboratory experiments demonstrate that Pacific Herring are highly susceptible to in-fection with VHSV and experience explosive outbreaks with high mortality in which viral shedding from infected individuals can reach 5.0 × 108 infectious units/Herring/day (Her-shberger et al. 2010a). These observations and reports of large-scale VHSV epizootics in populations of wild Pacific Herring (Garver et al. 2013) suggest the potential for VHSV to cause disease outbreaks or fish kills whenever susceptible populations of Pacific Herring are exposed to the virus. However, VHSV com-monly persists in apparently healthy Pacific Herring for extended periods, with disease and mortality occurring only when host and environmental conditions permit. For ex-ample, the capture and confinement of seem-

ingly healthy wild Pacific Herring in laboratory tanks or net pens often results in the progression of VHSV and other disease epizootics (Hershberger et al. 2006). A partial explanation of this apparent paradox was provided by studies demonstrating that chronic and persistent VHSV infections can occur in Pa-cific Herring (Hershberger et al. 2010b). Although these VHSV survivors become refractory to the disease, they remain capable of replicating VHSV upon subsequent reexposure (Hershberger et al. 2010b; Lovy et al. 2012). Therefore, chronic and persis-tent infections with VHSV are thought to occur among a small percentage of individuals in a population, and these chronic infections can ignite population-level epizootic cascades when host and environmental conditions shift in favor of the disease (e.g., host resistance declines due to colder water temperatures,

Figure 2. Pacific Herring demonstrating external lesions of ichthyophoniasis. “Sandpaper skin” and black ulcers on the flank are characteristic of the disease in juvenile Pacific Her-ring; external signs such as these are infrequently observed in older age cohorts, where the disease typically presents as white nodular lesions on the heart and other internal organs. Photo credit: Paul Hershberger.

Figure 3. Juvenile Pacific Herring demonstrating external signs of VHSV, including exophthal-mia, focal hemorrhaging around the eyes, mouth, and fin bases and diffuse epithelial hemor-rhaging along the flank. Photo credit: Paul Hershberger.


infection pressure increases due to limited water exchange or elevated fish densities, and/or herd immunity decreases due to the removal of refractory individuals or the in-flux of susceptible cohorts to the population).

Infectious Hematopoietic Necrosis

Infectious hematopoietic necrosis is a disease of salmonid fishes caused by an-other fish rhabdovirus, infectious hematopoi-etic necrosis virus (IHNV). Geographically, IHNV is enzootic in nearly all coastal wa-tersheds of Western North America, ranging from Alaska to California, and in major river systems including the Fraser River drainage of British Columbia; the Columbia River basin in Washington, Oregon, and Idaho; and the Sacramento River of California (Bootland and Leong 1999). During the 1970–1980s, the virus also spread to Asia and Europe via the shipment of infected fish or fish eggs.

Losses due to IHNV are most notable in Rainbow Trout reared in the commercial aquaculture industry, as well as in Chinook Salmon, steelhead (an anadromous form of Rainbow Trout), and Sockeye Salmon (O. nerka) reared at federal, pro-vincial, state, and tribal or First Nation facilities throughout the region. This disease is most common in juvenile fish in freshwa-ter, where infection can result in extensive external and internal hemorrhaging (Figure 4), with mortality rates up to 90% due to destruction of the kidney and spleen. Older fish are typically more resistant to disease but can be infected and become virus carriers. Transmission of IHNV occurs when an infected fish sheds virus into the surrounding water or from parent to off-spring at spawning. Outbreaks of IHNV have also been docu-mented in wild Sockeye Salmon (O. nerka) and kokanee (its lacustrine life history variant) in lakes, but these are relatively infrequent.

In the Salish Sea, limited surveillance for IHNV occurs among wild marine fishes; however, the virus has been detected in returning adult Sockeye Salmon in seawater (Traxler et al. 1997) and in 7 of 436 adult Sockeye during a broader survey of fishes from marine areas adjacent to the Salish Sea (Kent et al. 1998). In addition, IHNV is enzootic in several Sockeye Salmon populations spawning in rivers that flow into the Sal-ish Sea, including the Fraser River in British Columbia and the Skagit, Cedar, and Baker river systems in Washington State. Disease outbreaks have occurred in juvenile Sockeye Salmon from several of these watersheds, including Sockeye Salmon from Baker Lake, which have experienced several epizootics in the artificial spawning beaches used to hatch and rear fry. Among wild or free-ranging fish, two natural IHNV disease out-breaks have been reported in Salish Sea watersheds: in 1973 in Sockeye fry in Chilco Lake, on a tributary of the Fraser River in British Columbia (Williams and Amend 1976), and in 1986 in feral 2-year-old kokanee in a lake on Vancouver Island (Traxler

1986). Thus, it is evident that wild Sockeye Salmon of the Sal-ish Sea are affected by IHNV, but population-level impacts are difficult to estimate. Sporadic outbreaks of IHNV resulting in significant mortality have also occurred among Atlantic Salmon (Salmo salar) reared in commercial net-pens in the Salish Sea during 1992–1996, 2001–2003, and in 2012. Genetic typing indicated that the source of the virus was from native Pacific salmon, most likely from infrequent but significant virus spill-over events from free-ranging Sockeye.

Our understanding of the ecology of IHNV among both hatchery and wild populations of Pacific salmon in Western North America has been greatly improved by the genetic analy-sis of over 2,000 virus isolates. This has revealed three genetic subgroups of IHNV that show some host specificity, designated U, M, and L, because they occur in the upper, middle, and lower regions of the West Coast range of IHNV, respectively (Kurath et al. 2003). In the Salish Sea, all isolates of IHNV to date have been members of the U genogroup, which has general host spec-ificity for Sockeye Salmon. The M subgroup of IHNV has gen-eral host specificity for Rainbow Trout and steelhead and has historically been limited the Columbia River basin. However, since 2007 the M subgroup of IHNV has emerged to affect steel-head in Washington coastal rivers of the Olympic Peninsula, where it has spread northward as far as the Bogachiel River (Breyta et al. 2013). Steelhead within the Salish Sea are as yet naïve to this subgroup of IHNV, and the salmon co-managers of Washington State are actively pursuing strategies to prevent the further geographic spread of the M group viruses.

Thus, the impact of IHNV in the Salish Sea involves the constant presence of U group virus in Sockeye Salmon, asso-ciated with disease impacts that have also remained relatively constant over the last 35 years. Factors that could influence this situation include climate or environmental changes that neg-atively impact the general health of salmonids, making them more susceptible to disease caused by the endemic U genogroup of the virus or by the invasion of the M or L strains of IHNV, which are more pathogenic for steelhead or Chinook Salmon, respectively.

Figure 4. Juvenile Sockeye Salmon after laboratory exposure to IHNV. External signs of disease include exophthalmia and hemorrhaging around eyes, mouth, fins, and ventral surface. Photo credit: Gael Kurath.


Viral Erythrocytic Necrosis

Viral erythyocytic necrosis (VEN) is a disease that can lead to severe anemia in many marine fish species worldwide. The disease, caused by a fish iridovirus, occurs in the Salish Sea, where it affects populations of Pacific Herring (Hershberger et al. 2009) and the seawater stages of Pink (O. gorbuscha) and Chum (O. keta) Salmon (Evelyn and Traxler 1978). The extent and severity of VEN in the Salish Sea are likely underestimated due to the absence of a coordinated marine fish health surveil-lance program in the region and lack of sensitive or standard-ized diagnostic procedures for the disease. The etiological agent of VEN, generally referred to as erythrocytic necrosis virus, is easily visualized within affected cells by electron microscopy; however, the virus has yet to be isolated on established fish cell lines, resulting in difficulties in characterizing the causative agent or in developing tools needed to understand the epizooti-ology of the disease. From a diagnostic perspective, cytoplas-mic inclusion bodies, consisting of viral nucleic acid and viral proteins, can be easily detected by examining fixed and stained blood films (Figure 5); however, routine fish health examina-tions by fish health professionals throughout the Salish Sea re-gion often do not include the preparation and examination of stained blood films.

Viral erythrocytic necrosis typically occurs at low inten-sity and prevalence among populations of Pacific Herring in the Salish Sea; however, poorly understood host and environ-mental factors can produce an increase in severity leading to large outbreaks accompanied by fish kills. Epizootics are re-ported most frequently in juvenile Herring cohorts (Meyers et al. 1986; Hershberger et al. 2009); however, low-intensity in-fections are often seen in adults (Hershberger et al. 2009, and unpublished data). Correlations with environmental variables during periods of epizootics led to the hypothesis that cool tem-peratures—possibly accompanied by stress from exposure to low salinity water—were associated with the increased severity of VEN (Meyers et al. 1986). Additionally, the proximity of im-munologically naïve juvenile cohorts to fish carrying low-level infections also represents a likely risk factor (Hershberger et al. 2009). Epizootics can be easily induced after confinement of

free-ranging Herring into net pens or laboratory tanks (Traxler and Bell 1988; Hershberger et al. 2006), likely as a result of viral shedding from a few infected individuals that, when com-bined with limited water exchange in confined situations, results in increased infection pressure among susceptible cohorts.

Although most species of Pacific salmonids are susceptible to VEN after experimental exposures, the disease in free-rang-ing salmonids is generally limited to Pink and Chum Salmon (Evelyn and Traxler 1978). Epizootics can be induced upon confinement of these species into net-pens (Evelyn and Traxler 1978). Most available data indicate that the disease is of marine origin; however, an apparent progression of the disease in adult Pink Salmon during their freshwater migration up the Fraser River British Columbia indicates that reduced salinities may be an important factor influencing the progression of the disease (Bell and Traxler 1985).

Bacterial Kidney Disease

Bacterial kidney disease (BKD) is generally regarded as one of the most important bacterial diseases affecting wild and cultured salmonids in the Pacific Northwest (Weins and Kaat-tari 1999). The causative agent, Renibacterium salmoninarum, occurs in salmonid populations worldwide and is endemic to the Salish Sea. Infection prevalence in wild or feral stocks of salmon ranges up to 40%, strongly supporting the designation of R. salmoninarum as an endemic pathogen for anadromous salmonids in the Salish Sea (Banner et al. 1986; Rhodes et al. 2011). Contrary to a widely held assumption, BKD can occur at moderate to high infection intensities among free-ranging sal-monids in the Salish Sea and greater Pacific Northwest (Banner et al. 1986). Outside the Pacific Northwest, significant BKD epizootics have occurred in Chinook Salmon following their introduction to the Laurentian Great Lakes (Holey et al. 1998), where host–pathogen coevolution during the past several de-cades has selected less susceptible hosts (Purcell et al. 2008).

Our understanding of epizootiological relationships sur-rounding BKD remains limited, primarily because controlled laboratory studies are confounded by difficult bacterial isola-tion, slow bacterial growth in vivo, and slow disease progres-sion. The disease occurs in both acute and chronic forms in freshwater and marine-phase salmonids (Austin and Rayment 1985), and host mortality can occur either directly from the acute disease or indirectly as a result of behavioral changes stemming from the chronic form of the disease (Mesa et al. 1999). Host susceptibility is species-specific, with Sockeye, Chinook, and Chum Salmon demonstrating high susceptibility and Lake Trout (Salvelinus namaycush) and Bull Trout (S. con-fluentus) demonstrating low susceptibility (Jones et al. 2007). Asymptomatic fish and non-salmonid fish species can serve as long-term R. salmoninarum reservoirs (Rhodes et al. 2011). Transmission from an infected individual can occur either verti-cally through intraovum transfer (Figure 6) or horizontally by exposure to shed waterborne bacteria (McKibben and Pascho 1999) or by fecal–oral routes (Balfry et al. 1996). The likelihood of horizontal transmission is greater under conditions of higher

Figure 5. Giemsa-stained blood cells from Pacific Herring. (A) Normal herring erythrocytes, with dark-staining nuclei. (B) Erythrocytes from a herring with VEN, demonstrating viral inclusion bodies in the cytoplasm of affected cells. Photo credit: Paul Hershberger.


fish density (Rhodes et al. 2011), which can be influenced by schooling and spawning behaviors, loss of critical habitat, and episodic releases of hatchery salmonids. The likelihood of in-fection or disease-related mortality in free-ranging salmonids can be influenced by a range of host and environmental condi-tions, including temperature (Jones et al. 2007; Rhodes et al. 2011), smoltification (Mesa et al. 1999), and hatchery practices (Fenichel et al. 2009).

DISEASE AND THE HEALTH OF FISH POPULATIONS IN THE SALISH SEA

Our understanding of the effects of pathogens, contami-nants, and other factors that affect aquatic animal health is heav-ily weighted toward cultured stocks or a few model systems. For example, the Salish Sea and its associated river systems are home to a large number of salmonid hatcheries and captive rearing facilities, in addition to extensive runs of native, free-ranging populations of Pacific salmon. Hatchery-reared salmon are routinely examined for pathogens at different life stages, in-cluding as juveniles prior to transportation to another facility, as prespawn adults returning to the hatcheries, and whenever clini-cal disease signs are observed in the hatcheries; however, few efforts have been made to assess the health of their wild cohorts. Thus, though information is available about the infectious dis-eases affecting hatchery-reared salmonids, less is known about the health status of wild salmonids or other marine fishes of the Salish Sea. Further, much less is known about the range of in-fectious diseases or contaminants affecting the health of marine fish that have important commercial or recreational value or that serve as critical components of the aquatic food chain.

Movement of Fish and Pathogens

Among the many threats to the health of wild fish in the Sal-ish Sea is the introduction of novel pathogens via the activities of man. Increased global trade provides greater opportunities for the introduction of nonnative species, including pathogens, via ballast water or by international shipments of ornamental fish (Hedrick 1996). Also important is the movement of live

fish or eggs and their associated pathogens by public or private aquaculture practices. For example, several massive epizootics swept through populations of wild pilchards in south Australia, presumably as the result of a herpesvirus that was introduced through feed imports used in the tuna mariculture industry (Jones et al. 1997). From a prophylactic disease management perspective, the largest importation concern in the Salish Sea involves those diseases listed by World Organization for Ani-mal Health (OIE). Concerns surfaced after the recent molecular identification of an infectious salmon anemia-like virus in the Canadian portion of the Salish Sea. Although the virus has not yet been isolated in cell culture, and its pathogenicity to en-demic fishes remains uncertain, expanded and coordinated sur-veillance efforts have been implemented. A related risk occurs from changes in the host range or virulence of endemic patho-gens, like IHNV, that are associated with intensive aquaculture (Kurath and Winton 2011).

In addition to the ecosystem impacts of nonnative or inva-sive fish species, an important but sometimes overlooked fac-tor is the range of pathogens that accompany their introduction and spread. Not only can nonnative species introduce exotic pathogens, they can also amplify endemic pathogens (Hersh-berger et al. 2010c) and serve as long-term reservoirs of infec-tion (Groocock et al. 2007).

Synergistic Effects of Contaminants and Climate Change

The presence or severity of infectious diseases in fish is controlled by factors present in the host, the pathogen, and the environment. As poikilothermic aquatic vertebrates, fish are uniquely susceptible to environmental conditions such as tem-perature that control most aspects of their physiology(including the immune response) and to the effects of contaminants and toxins present in the aquatic environment in which they con-tinuously reside. The state of Washington has been monitoring levels of contaminants in marine biota, including fish, since 1989 through the Puget Sound Assessment and Monitoring Pro-gram. In addition to identifying high prevalence of neoplasms in flatfishes from urban areas, these surveillances indicate that Chinook Salmon from Puget Sound are three times more con-taminated with polychlorinated biphenyls than are those from six other West Coast populations (Puget Sound Assessment and Monitoring Program 2007). There is a critical need to better un-derstand the impacts of confounding stressors, such as contami-nants and climate change (Harvell et al. 2002) on infectious and parasitic diseases throughout the Salish Sea. Such an integrated understanding would provide valuable information to managers who are currently tasked with managing resources in a poorly understood and dynamic system.

CURRENT NEEDS FOR INCORPORATING DISEASE INTO SALISH SEA ECOSYSTEM MANAGEMENT

Anecdotal observations and a limited number of field studies indicate that numerous other fish pathogens are present within the region, and some—such as Nanophyetus salmonicola and

Figure 6. Photomicrograph of Chinook Salmon ovary infected with Reni-bacterium salmoninarum. (A) Hematoxylin and eosin stain. (B) Immunos-tain with fluoroscein-conjugated anti–R. salmoninarum antibody. Arrows, bacterial clusters labeled with antibody. Photo credit: Anne Baxter.


Listonella (Vibrio) spp.—may have population-level impacts. Unfortunately, as presented here, most of these pathogens have been underinvestigated, and a true understanding of their im-pacts on affected host populations remains uncertain. Coordi-nated surveillance activities are required to define the pathogens present in fishes from the Salish Sea, assess their prevalence and intensity of infection, and determine their host and geographic ranges. In addition, controlled empirical studies, intended to de-termine cause-and-effect relationships, are necessary to under-stand the factors that control the ecology of infectious diseases in the region.

Ambitious habitat restoration efforts are also envisioned for the Salish Sea, and recovery strategies are needed for popula-tions of forage fishes (e.g., herring) as well as recreationally, culturally, and commercially important species (e.g., salmon) and the highly charismatic predators like orcas (Orcinus orca) that depend on them (Ford et al. 1998). These recovery efforts require knowledge about the sources of natural mortality (in-cluding infectious diseases and contaminants) that affect these free-ranging populations. Although a better understanding of the natural factors that have historically governed diseases in wild fish populations is necessary, current and future efforts must also determine the impacts of a wide range of emerging anthropogenic stressors, including exotic pathogens or species, habitat loss, novel contaminants, and climate change, as well as the synergistic effects of these stressors on the severity of diseases among wild fish in the Salish Sea.

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FEATURE

Morphological and Genetic Identification of Spontaneously Spawned Larvae of Captive Bluefin Tuna in the Adriatic Sea

Identificación morfológica y genética de larvas espontáneamente desovadas de atún aleta azul cultivado en el mar AdriáticoRESUMEN: a través de jaulas, se observó el comporta-miento de desove del atún aleta azul (Thunnus thynnus) en cautiverio en el mar Adriático durante el verano de 2011. Se colectaron aproximadamente 20,000 huevos a partir de jaulas y se transfirieron a un criadero en tierra, con un éxito de fertilización del 80%. Los huevos pelágicos fueron esféricos con un diámetro de 1,035.06 ± 20.16 µm y un solo glóbulo de aceite (227.63 ± 8.07 µm). Las larvas recién eclosionadas (3.08 ± 0.14 mm longitud total) siguieron el desarrollo larval propio de los escómbridos: melanóforos dispersos en el cuerpo, cabeza, notocorda y vitelo, excepto en el pliegue de la aleta; se observó pigmentación ocular a los 1.5 días después de la eclosión, comenzaron a aparecer las aletas pectorales y se reabsorbieron dos tercios del saco vitelino; a los 2.5 días de la eclosión, se desarrolló la boca y la larva comenzó a alimentarse de rotíferos enriquecidos. Después de haber ocurrido la mortalidad en el criadero (5 días después de la eclosión) se extrajo una muestra de ADN de las larvas. Se amplificaron y secuenciaron 890 pares de bases de una región mitocondrial control con el fin de corroborar genéticamente la identidad de la especie. Sobre la base de una alineación secuencial múltiple, las secuencias de las larvas no mostraron ambigüedades en la región control mitocondrial de T. thynnus, por lo tanto se confirmó el desove espontáneo de la especie en cautiverio. Si bien tal evento ya se había documentado previamente en el Mediterráneo, este es el primer reporte basado en datos morfológicos y genéticos.

Leon Grubišić, Tanja Šegvić-Bubić, and Ivana Lepen PleićInstitute of Oceanography & Fisheries, Split, Croatia

Krstina Mišlov-JelavićKali Tuna d.o.o., Island of Ugljan, Croatia

Vjeko Tičina and Ivan KatavićInstitute of Oceanography & Fisheries, Split, Croatia

Ivona MladineoInstitute of Oceanography & Fisheries, Šetalište Ivana Meštrovića 63, 21000 Split, Croatia. E-mail: [email protected]

ABSTRACT: The spawning behavior of captive Bluefin Tuna (Thunnus thynnus) was observed in cages at an Adriatic facil-ity in summer 2011. Approximately 20,000 eggs were collected from cages and transferred to a land-based nursery, with an estimated fertilization success rate of 80%. Eggs were spheri-cal with a diameter of 1,035.06 ± 20.16 µm and were pelagic with a single oil globule (227.63 ± 8.07 µm). Newly hatched larvae (3.08 ± 0.14 mm total length) followed scombrid larval development: melanophores were scattered over the body, head, notochord, and yolk, except finfold; eye pigmentation was ob-served 1.5 days posthatch, pectoral fins started to appear, and two-thirds of the yolk sac were absorbed; the mouth developed at 2.5 days posthatch, and larvae began feeding upon enriched rotifers. After mortalities occurred in the nursery (5 days post-hatch), DNA was extracted from a sample of larvae. An 890 base pair fragment of the mitochondrial partial control region was amplified and sequenced to genetically confirm fish spe-cies identity. Based on multiple sequence alignment, larval se-quences showed no ambiguities to the T. thynnus mitochondrial control region, thereby confirming spontaneous spawning in captivity. Although such an event has previously been reported in the Mediterranean, this is the first report supported by both morphological and genetic data.

INTRODUCTION

The captive Bluefin Tuna (Thunnus thynnus) fattening in-dustry as a capture-based practice was introduced in the Medi-terranean Sea in early 1990s, with the goal of obtaining a fresh product rich in muscle fat content, a prerequisite for obtain-ing a high price on the demanding Japanese sushi and sashimi markets (Ottolenghi 2008). Consequently, growing industry fishing pressure and the need for live Bluefin Tuna for the fat-tening process stimulated the establishment of the International Commission for the Conservation of Atlantic Tuna (ICCAT), which established fishing quotas for the Eastern Atlantic stock. Despite various management and conservation measures intro-

duced by ICCAT, the decline of the Bluefin Tuna population due to continuous overfishing was not halted (ICCAT 2008), starting a wide debate over the enlisting of Tuna on the International Union for Conservation of Nature Red List of endangered spe-cies (IUCN 2013). As a consequence, this would limit fishing and international trading activities (Atlantic Bluefin Tuna Status Review Team 2011; Colette et al. 2011). In 2006, the ICCAT management plan to rebuild the stock to BMYS (biomass of the maximum sustainable yield) by 2022 with 50% or greater prob-ability of success was revised in response to the emergence of various issues related to its implementation (ICCAT 2011a). The plan incorporated a number of conservation measures: country-specific total allowable catches, minimum size limits, closed fishing seasons, management controls of fishing and farm-ing capacity, as well as monitoring, control, and surveillance


measures (vessel registers, vessel monitoring systems, observer programs, transshipment prohibitions, weekly catch reporting). In addition, in order to improve data transparency and avail-ability and the fundamental knowledge on Bluefin Tuna biology and behavior, ICCAT established a research program jointly run by the European Community (80%), Canada, Croatia, Japan, Libya, Morocco, Norway, Turkey, the United States, Chinese Taipei, and the ICCAT Secretariat, with a total budget of up to €19 million for a 6-year period (ICCAT 2011b). Lastly, ICCAT established a total allowable catches of 12,900 tons for 2011 for the eastern Atlantic and Mediterranean stock, which is esti-mated to have a 60% probability of allowing the stock to rebuild to a maximum sustainable yield level by 2022 (ICCAT 2011a).

Seeing a need to establish hatchery production technology for Bluefin Tuna larvae for the eastern Atlantic and Mediter-ranean stock, as was being developed for Thunnus orientalis (Miyashita et al. 2001; Sawada et al. 2005), Thunnus albacares (Wexler et al. 2003), and Thunnus maccoyii (Hutchinson 2009), the European Union secured funding of large Framework Pro-gram projects that produced the first hatchery-reared Bluefin Tuna larvae using hormonal implants to stimulate gamete matu-ration and spawning of captive adults and obtaining the first hatchery-reared Bluefin larvae (see Mylonas et al. 2007, 2010). Starting in 2001, the European Union projects alone amounted to a budget of approximately €6.6 million and resulted in ap-proximately 16 publications in peer-review and non-peer-re-view journals through 2009 (Reproduction of the Bluefin Tuna in Captivity 2006; Self-sustained Aquaculture and Domestica-tion of Bluefin Tuna Thunnus thynnus 2008, 2009). Although hormonally induced spawning represented enormous success for the European scientific community and ignited great hopes in the industry, larval survival was poor during the first 10 days posthatching due to buoyancy and weaning issues, cannibalism, and collisions with the tank or net walls (De Metrio et al. 2010). Meanwhile, the first reports of spontaneous spawn-ing without hormonal induction began to emerge from different farms throughout the Mediterranean but were based only on morphological identification of larvae (Gordoa et al. 2009; Gordoa 2010; de la Gándara et al. 2011).

In Croatia, Bluefin Tuna farming is based on catches of small (usually 8–15 kg) juvenile Bluefin Tuna that are transported from the southern Adriatic Sea to a facility where they are kept at least 1.5 years to reach desired harvesting weight (>30 kg; ICCAT 2011a). Large Bluefin Tuna may sometimes be kept for several months before harvesting them. In sum-mer 2011, the first of what appeared to be sponta-neously spawned Bluefin Tuna larvae from offshore Adriatic farm cages were collected and subjected to genetic and morphological identification using a molecular marker to infer species identity. Captive spawning populations of other Tuna species have been monitored using mitochondrial DNA (mtDNA) markers to estimate spawning frequencies, spawning period, and number of females spawning each day

(Masuma et al. 2003; Niwa et al. 2003; Nakadate et al. 2011). Such data contribute to both aquaculture operations and stock enhancement efforts, allowing genetic variation of offspring to be determined and potential genetic impact of fry releases on wild stocks to be considered (Niwa et al. 2003). Therefore, we used the control region of mtDNA to infer the genetic identity of the larvae hatched from eggs collected in Tuna cages and also compared them to wild Tuna sampled from the Adriatic Sea.

MATERIALS AND METHODS

Larval Collection, Incubation, and Development

The captive Bluefin Tuna broodstock we used for our study was composed of 70 fish with an estimated mean body weight of 120 kg and unknown numbers of males and females. They were kept in captivity for 4 years in a floating cage at an Adri-atic Tuna farm situated 200 m from the Island of Ugljan (Figure 1). In July 2011, the net cage was equipped with a plastic foil egg collector surrounding the top 2 m of the net sidewall to pre-vent eggs from flowing out of the cage. Seawater temperature was measured daily at 1 m below the water surface inside the cage (Figure 2). Spawning behavior patterns, as described by Masuma et al. (2006) were observed at dusk on July 18. The following morning, zooplankton was collected from the surface inside the cage using a 700-µm mesh size net at a measured sea water temperature of 23.9°C. The zooplankton was transferred into a tank in the land-based nursery, kept in a fine mesh net at 24°C, and later transferred to a Petri dish for examination under a stereomicroscope. Fish eggs with the morphological features of scombrid eggs were isolated (~20,000 eggs), transferred into a 400-L rearing tank, and, each hour, observed under an Olym-pus BX40 light microscope at 20–100× magnification and pho-tographed with an Olympus C0404 (Olympus, Croatia) digital camera. No larvae were found in the zooplankton sample. The

Figure 1. Geographic locations of the Bluefin Tuna sampling sites in the eastern Adriatic Sea, 2011. RL = reared larvae (44°2′18″ N; 15°10′55″ E); WL = wild larvae (43°5′0″ N; 15°27′0″ E); RA = captive adults (43°17′16″ N; 16°28′56″ E).


diameter of 30 eggs was measured with an ocular micrometer attached to a microscope at an accuracy of 0.01 mm. In order to calculate fertilization rate, a random subsample of about 200 eggs was taken and the percentage of fertilized eggs was esti-mated using the light microscope.

Hatched larvae from the land-based nursery were anesthe-tized with benzocaine (5 mgL−1) and measured using the ocular micrometer. For analysis of larval morphology, 15 anesthetized larvae were randomly sampled and measured for total length (distance along midline of body from tip of mouth to end of caudal fin), notochord length (distance along midline of body from tip of mouth to end of the notochord), preanal distance (distance along midline of body from tip of snout to vent), body depth (perpendicular depth of trunk at anus), head length (dis-tance between tip of upper jaw and cleithrum), longest yolk sac diameter, longest yolk sac oil globule diameter, horizontal eye diameter, and pectoral fin length. The time of yolk sac resorp-tion and opening of the mouth were also recorded.

Rearing conditions were based on clear water methods, with a constant daily exchange of 100–200% filtrated seawater, artificial photoperiod of 8 h of dark and 16 h of light at 500 lux, and a constant water temperature of 24 ± 0.1°C. A surface skim-mer was installed on the second day of posthatch to maintain a clean water surface that would facilitate swim bladder inflation. Feeding was based on rotifers (5–7 individuals/mL) enriched with DHA Protein Selco (Artemia Systems SA, Ghent, Bel-gium), beginning at 2 days and terminating at 5 days posthatch due to massive larvae mortality. Water quality measures (mean ± SE) were 8.13 ± 0.08 pH, 37.6 ± 0.2 ppt salinity, and 96.5 ± 2.6% oxygen saturation for the duration of the experiment.

DNA Extraction and Polymerase Chain Reaction Amplification

To confirm that eggs collected from the cages were Bluefin Tuna, 8 larvae were sampled postmortem from the land-based nursery tank for mtDNA control region analysis (Boustany et al. 2008). Additional samples for mtDNA control region analysis were obtained from 23 larval Bluefin Tuna collected in the Adri-atic Sea off the Island of Jabuka (43°05′67″ N, 15°28′63″ E) and fin clips from nine adults held in facility operated by Sardina on the Island of Brač.

DNA was isolated using the QIAGEN DNeasy Blood and Tissue Kit (Qiagen), and the quantity and purity of isolated DNA and amplified fragments were checked using a BioPho-tometer 6131 (Eppendorf, Hamburg, Germany). The mtDNA control region locus was amplified by polymerase chain reac-tion (PCR) using 0.8 µM of each degenerate primers, forward primer 5′ CTA CCC CTA ACT CCC AAA GC 3′ and reverse primer 5′ GCT TTA GTT AAG CTA CG 3′ to amplify ~1 kilo-base pair of the Bluefin Tuna mtDNA control region. The rest of the reaction mix consisted of 2.5 mM of MgCl2, 2 mM of each dNTP, 1.25 U of Platinum Taq polymerase (Invitrogene, California, USA) and 3 ng/µL of template. The amplification profile consisted of initial denaturation for 2 min at 94°C, 35 cycles of denaturation for 30 s each at 94°C, annealing at 45°C for 30 s, and elongation for 60 s at 72°C with a final extension of 7 min at 72°C. Products were loaded on a 1% agarose gel and visualized by adding SYBR Safe (1%) (Invitrogen, California, USA) directly into the gel.

Fragment Sequencing

PCR products were purified using a QIAquick PCR Puri-fication Kit (Qiagen) and sequenced on an ABI 3100 automatic DNA sequencer (Applied Biosystems, California, USA), using the ABI PRISM BigDye Terminator Cycle Sequencing Kit, in

Figure 2. Mean sea surface temperature measured in the cage of an Adriatic Tuna farm, situated 200 m from the Island of Ugljan, used to hold Bluefin Tuna for a spontaneous spawning study. Left panel shows mean monthly temperatures from March to August 2011. Left panel arrow indicates month of spontaneous spawning, July, and points to right panel, which shows daily July temperature variations. The temperature on the egg collection date, July 19, is circled.


both directions. Sequences were aligned with other scombrids in GenBank (North Atlantic T. thynnus DQ855113, Mediterranean T. thynnus DQ087594, T. maccoyii HQ630707, T. albacares AF301184, T. alalunga JN007599, T. orientalis HQ630709) by Clustal X, implemented in the MEGA 5 software using default parameters. The same tool was used to perform neighbor-joining analysis, based on p-distance. Reliabilities of phylogenetic rela-tionships were evaluated using nonparametric bootstrap analy-sis (Felsenstein 1985) with 2,000 replicates for neighbor-joining analysis. Bootstrap values exceeding 70 were considered well supported (Hills and Bull 1993). Sequences were submitted to GenBank, and the accession numbers of sequences obtained in this study are JN620217–JN620224 (reared Tuna larvae), JX273444–JX273467 (wild Tuna larvae), and JX273435–JX273443 (reared adult Tuna).

Statistical Analysis

Molecular diversity among the three Adriatic groups of samples (reared larvae, wild larvae, and adult reared Tuna) was measured using Dnasp 5.0 (Librado and Rozas 2009) and Arlequin 3.0 (Excoffier et al. 2005). Values for the number of haplotypes, polymorphic sites, haplotype diversity (Nei 1987), nucleotide diversity (Nei 1987), and the average numbers of pairwise nucleotide differences (Tajima 1983) were estimated. Pairwise and overall distances among haplotype sequences were calculated in MEGA 5.0 (Tamura et al. 2007). Pairwise genetic differentiation between populations was estimated using the fixation index (FST) and statistical significances were tested with 10,000 permutations. FST calculation was performed in Arlequin 3.0.

RESULTS AND DISCUSSION

It is known that the main spawning trigger for the Bluefin Tuna is a sudden and drastic rise in temperature to 25°C (Mylo-nas et al. 2007), which was also observed in this study. Though the exact time of fertilization could not be established, embryo morphology and comparisons to previous studies’ (Mylonas et al. 2007; De Metrio et al. 2010) approximations suggested that it occurred 5 days after the temperature rise.

Approximately 20,000 eggs were collected the next day on July 19, and about 80% were estimated to have been fertil-ized. The 30 eggs measured were spherical with a diameter of 1,035.06 ± 20.16 µm (mean ± SD), pelagic, contained a single oil globule with a diameter of 227.63 ± 8.07 µm (mean ± SD), and seemed to be similar to earlier descriptions of eggs from Bluefin Tuna held in other Mediterranean facilities (Ghysen et al. 2010). At the time the eggs were examined in the land-based nursery (1500 hours), the embryo form was detectable, encir-cling the yolk by 180° with a typical central yolk constriction and tail bud still attached to the yolk. A maximum of five early somites were visible along the frontal part of the notochord and on Kupffer’s vesicle (Figure 3a). The exact age of the embryos was not known, because the time of spontaneous spawning was not precisely determined, but, according to descriptions by De Metrio et al. (2010) and Miyashita (2002), the eggs were esti-

mated to be around 13–15 h postfertilization, confirming a clear pattern of nocturnal spawning from 0300 to 0500 hours (Gordoa et al. 2009). Between 20 and 23 h postfertilization, the appear-ance of melanophores and a heartbeat were observed (Figure 3b).

Hatching occurred 32 h after the estimated spawning time at a constant seawater temperature of 24°C. According to De Metrio et al. (2010), the incubation period for eggs obtained from hormonally treated Bluefin Tuna in captivity was 30 h at water temperature of 25–26°C or, as we found in our study, 32 h at 24°C.

Figure 3. Development of the Bluefin Tuna: (a) embryo formed, appear-ance of Kupffer’s vesicle; (b) embryo 3 h before hatching; (c) yolk sac larvae 3 h after hatching; (d) 1.5 days posthatch yolk sac larvae; (e) 2.5 days posthatch larvae, mouth developed, first feeding; (f) 4 days post-hatch larvae. Scale bars = 0.5 mm.


The newly hatched larvae were 3.08 ± 0.14 mm (mean ± SD) in total length and had small melanophores that were scat-tered over the body, head, notochord, and yolk but not over the fin fold (Figure 3c; 3 h posthatch). During the first 5 days post-hatch, larvae length, body depth, and eye diameter increased, whereas yolk sac and oil globule diameter decreased (Table 1). At 1.5 days posthatch, eye pigmentation was observed and pec-toral fins started to appear, and two-thirds of the yolk sac was absorbed (Figure 3d). At 2.5 days posthatch, the mouth was de-veloped and larvae began feeding on enriched rotifers (Figure 3e). The yolk was visible until 3 days posthatch. From hatching to 5 days posthatch (Figure 3f), the growth rate was 0.21 mm/day. Massive mortality occurred between 3 and 5 days post-hatch, with less than 1% overall survival at 6 days posthatch, likely related to unsuccessful initial feeding and larvae sinking to the tank bottom.

An 890 base pair partial mtDNA control region sequence was obtained from the isolated DNA samples. Based on results of multiple sequence alignment, all isolates likely belonged to Bluefin Tuna. The mtDNA control region has been documented to be particularly sensitive in detecting genetic diversity and population genetic structure of marine fish and has been used for analysis of populations in various fish (Bremer et al. 1996; Iguchi et al. 1999; Tabata and Taniguchi 2000; Ishikawa et al. 2001). In the 40 Tuna samples we analyzed (eight reared larvae; nine captive adults; 23 wild larvae), we found 90 variable sites and 30 haplotypes (Table 2). Sequence divergence (Kimura dis-tance) among mtDNA control region haplotypes ranged from 0.01% to 0.39%, with an average of 0.15%. Among the 90

polymorphic sites, 47 were singleton variable sites and 43 were parsimony informative. Among the 30 haplotypes, most (83%) were unique and represented by a single individual. Two haplo-types were found in the eight reared larvae examined, and one of the haplotypes was shared by seven of these larvae (Table 3). Nine haplotypes were found in the nine captive adults examined, none of which were shared. Nineteen haplotypes were found in the 23 wild tuna larvae examined, 15 of which were unique. None of the 30 haplotypes examined were shared among the three groups of samples, which suggests that these haplotypes have no use for Bluefin Tuna stock identification in the Adriatic.

Overall mtDNA control region diversity of 90 Bluefin Tuna samples was 0.968 ± 0.019 (mean ± SD) for haplotypes and nucleotide diversity of 0.01318 ± 0.00113 (mean ± SD) for nucleotides (Table 2). The high level of mtDNA polymor-phism, indicated by high nucleotide and haplotype diversity in wild larvae and captive adults, has been reported to occur in other Thunnus species (Viñas et al. 2004; Alvarado-Bremer et al. 2005; Durand et al. 2005; Ely et al. 2005; Martinez et al. 2006; Boustany et al. 2008; Nakadate et al. 2011). In contrast, the low haplotype and nucleotide diversity observed in reared larvae might be due to the small number of larvae examined and the small number of females that spawned in the cage. Un-fortunately, it was not possible to use a larger number of reared larvae for mtDNA analysis due to financial and technical limita-tions.

Significant (P < 0.05) genetic differences were found be-tween reared larvae and captive adults (FST = 0.3624) as well

Table 1. Changes in length and shape of Thunnus thynnus yolk sac larvae (n = 15) during the first 5 days posthatch at a seawater temperature of 24 ± 0.1°C (mean ± SD).

Hours posthatching

Total length (mm)

Standard length (mm)

Preanal length (mm)

Head length (mm)

Body depth (mm)

Longest yolk sac diameter (mm)

Oil globule diameter (mm)

Eye diameter (mm)

0 3.08 ± 0.14 3.03 ± 0.13 1.32 ± 0.09 0.36 ± 0.03 0.61 ± 0.06 1.40 ± 0.10 0.31 ± 0.04 0.23 ± 0.01

+12 3.38 ± 0.16 3.30 ± 0.12 1.55 ± 0.11 0.43 ± 0.02 0.65 ± 0.05 1.24 ± 0.08 0.28 ± 0.03 0.24 ± 0.01

+24 3.69 ± 0.20 3.63 ± 0.17 1.46 ± 0.11 0.44 ± 0.04 0.72 ± 0.07 0.63 ± 0.04 0.23 ± 0.01 0.26 ± 0.02

+48 3.79 ± 0.19 3.68 ± 0.21 1.87 ± 0.14 0.73 ± 0.03 0.85 ± 0.07 0.33 ± 0.09 0.17 ± 0.01 0.34 ± 0.01

+72 3.99 ± 0.22 3.76 ± 0.19 1.99 ± 0.18 0.85 ± 0.04 0.87 ± 0.10 — — 0.39 ± 0.03

+96 4.15 ± 0.23 4.00 ± 0.25 2.11 ± 0.18 0.92 ± 0.05 0.91 ± 0.12 — — 0.45 ± 0.02

Table 2. Genetic diversity statistics for Thunnus thynnus sampled from the Adriatic Sea based on mtDNA control region sequence data. N = sample size; H = number of haplo-types; S = number of segregating sites; h = haplotype diversity (mean ± S.D.); π = nucleo-tide diversity (mean ± SD); k = pairwise difference (mean ± SD).

Fish origin N H S h π k

Reared larvae 8 2 8 0.250 ± 0.180 0.00253 ± 0.00182 2.00000 ± 1.256177

Reared adults 9 9 57 1.000 ± 0.052 0.02038 ± 0.00254 16.13889 ± 8.062041

Wild larvae 23 19 53 0.984 ± 0.017 0.01148 ± 0.00117 9.08300 ± 4.680749

Total samples 40 30 90 0.968 ± 0.019 0.01318 ± 0.00113 10.42821 ± 5.103292


as between reared larvae and wild larvae (FST = 0.3036) but not between captive adults and wild larvae (FST = 0.0084). This is likely due to a very high diversity of haplotypes where the two populations do not share a single common haplotype, result-ing in a low frequency of each haplotype and constrained FST (Hedrick 1999).

Neighbor-joining analysis generated an unrooted tree resulting from 2,000 bootstrap replicates with interspersed sequences of the three groups of Adriatic Bluefin Tuna samples we analyzed in addi-tion to reference specimens from the North Atlantic (T. thynnus NA or DQ855113) and Mediterranean (T. thynnus Med or DQ087594; Figure 4). The great haplotype diversity observed within most of the populations was fully supported by the tree topology, which showed scattered sequences through the clades and no distinct clustering.

In the small sample of reared larval, two haplotypes were iden-tified, suggesting that at least two females contributed to spawning. Future genetic analysis should be based on more extensive samples of larvae from captive spawning events, as well as samples from cap-tive adults having known spawning grounds (Carlsson et al. 2007).

Although the eastern stock of Bluefin Tuna is known to spawn south of Malta and in the Balearic and Tyrrhenian seas (West Mediterranean spawning ground) as well as in the Io-nian and Levantine seas (eastern spawning ground), Fromentin and Powers (2005) have suggested the existence of additional spawning grounds. Though genetic confirmation of larvae re-sulting from Bluefin Tuna spawning in holding cages has not previously been available, Gordoa et al. (2009) reported that Bluefin Tuna caught on their spawning ground spawned within

Table 3. Number of mtDNA control region haplotypes obtained from each sample group and related pair base positions. RL = reared larvae; CA = captive adults; WL = wild larvae.

Haplotypes RL CA WL Base pair position

10; 27; 45; 56; 64; 65; 66; 87; 89; 94; 105; 114; 117; 137; 139; 140; 144; 145; 150; 156; 157; 158; 164; 168; 170; 175; 190; 194; 197; 201; 219; 228

239; 242; 247; 253; 259; 261; 264; 269; 280; 287; 291; 303; 324; 339; 341; 349; 366; 384; 385; 386; 387; 390; 395; 396; 409; 412; 417; 422; 429

2 9 19 444; 465; 467; 475; 515; 537; 541; 550; 566; 575; 580; 581; 587; 592; 596; 610; 615; 618; 621; 627; 628; 629; 637; 687; 733; 745; 775; 777; 788

Hap1 7 ATTTATATTCTAGCAAACGGTCTGCACATTTTGACCCCTGATTTAATTTAGCCGATCCCAGGCGCCGCATCTATACCAAAGAGCGTTTGG

Hap2 1 ..................AA................................T..C.T.......T.......C.........T......

Hap3 1 .......A.........................................G..T.......A..............T.........G....

Hap4 1 ..................AA...A......C.....................T..C....A.....A....C.CC....CC...C.....

Hap5 1 .......A.G..A.....AA..........C.......G.............T..C....A.....A...GC..TT..........AC.A

Hap6 1 G....CGA.GC.......AA...A......C.....................T..C....A.....A.......................

Hap7 1 ..C..........T..GTAA............A..T...........C....T..C....A.......G...G.........A.......

Hap8 1 ..................AA.......G.......................TT....T..A...........................A.

Hap9 1 ..........C.......AA...A......C.....AAG.C.CC....C...T..CT.TCA.G...A..C..................A.

Hap10 1 G.........C.......AA...........C....................T..C....A....T........................

Hap11 1 .............................................G...GA.T.......A.............................

Hap12 1 ..................AA...........C....................T..C....A.............................

Hap13 1 ....G...........G.AA..........C..........C..........T.......A.....A.......................

Hap14 2 ...C.C....C.......AA...A......C.....................T..C....A...T.A.......................

Hap15 1 ...............G..AAC...............................TA.C....A.............................

Hap16 1 ...........G......AA................................T..C.T.......T.......C.........T......

Hap17 2 ..................AA....T..............A.........G..TA.C.......A..........................

Hap18 2 ..................AA...............................TT....T..A.................G...........

Hap19 1 ..C..........T..G.AAC...........A..............C....T.......A.......G.......T.............

Hap20 1 ..................AA...................A............T..C..................................

Hap21 1 ..................AA.......G.......................TT....T..A.............................

Hap22 1 ..................AA................................T.......A.............................

Hap23 2 ..................AA...................A......C.....T..C....A.............................

Hap24 1 ..........C.......AA...................A......C.....T..C....A....................G........

Hap25 1 ................G.AA...A...............A.........G..T..C....A.............................

Hap26 1 .C................AA......T..C.C.G.T........G.......T.GC....A......T......................

Hap27 1 ..C..........T..G.AAC...........A..............C....T.......A.......G.....................

Hap28 1 ..............G...AA..CAT....C....T.................T..C....A.............................

Hap29 1 .....C....C.......AA...A......C.....................T..C....A.....A.......................

Hap30 1 ........C.........AA.T.A.G..CC..............G....G..T.......AA...............C............


the transport cage being towed to a fattening facility, and de la Gándara et al. (2011) and Gordoa (2010) made similar observa-tions of spawning in rearing cages. Our study provided the first genetic evidence supporting spontaneous spawning of captive Bluefin Tuna in the Adriatic Sea. Though our study suggests that suitable spawning conditions occur in this area, no wild Bluefin Tuna spawning activity has ever been documented in the Adriatic Sea, although its potential as a spawning site has been recognized.

European Union Member States have made strong efforts to closely control the rearing cycle of Bluefin Tuna and have achieved promising results with the hormonal induction of spawning (Mylonas et al. 2007; De Metrio et al. 2010). Grow-ing evidence of spontaneous spawning of captive Bluefin Tuna throughout the Mediterranean suggests that experiments focus-ing on large-scale larval rearing should be undertaken to domes-ticate this species as well as to help rebuild wild populations.

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Bremer, J. R. A., J. Mejuto, T. W. Greig, and B. Ely. 1996. Global popu-lation structure of the Swordfish (Xiphias gladius L.) as revealed by analysis of the mitochondrial DNA control region. Journal of Experimental Marine Biology and Ecology 197:295–310.

Carlsson, J., J. R. McDowell, J. E. L. Carlsson, and J. E. Graves. 2007. Genetic identity of YOY bluefin tuna from the eastern and west-ern Atlantic spawning areas. Journal of Heredity 98:23–28.

Collette, B., A. F. Amorim, A. Boustany, K. E. Carpenter, N. de Oliveira Leite Jr, A. Di Natale, D. Die, W. Fox, F. L. Fredou, J. Graves, F. H. Viera Hazin, M. Hinton, M. Juan Jorda, O. Kada, C. Minte Vera, N. Miyabe, R. Nelson, H. Oxenford, D. Pollard, V. Restrepo, J. Schratwieser, R. P. Teixeira Lessa, P. E. Pires Ferreira Travassos, and Y. Uozumi. 2011. Thunnus thynnus in IUCN 2011. IUCN Red List of Threatened Species. Version 2011.2. Available: www.iucnredlist.org. (March 2012).

de la Gándara, F., A. Ortega, A. Belmonte, and C. R. Mylonas. 2011. Spontaneous spawning of Atlantic Bluefin Tuna Thunnus thynnus kept in captivity. In Mediterranean Aquaculture 2020, Aquacul-ture Europe 11, Rhodes, Greece, October 18–21, 2011. European Aquaculture Society.

De Metrio, G., C. R. Bridges, C. C. Mylonas, M. Caggiano, M. De-florio, N. Santamaria, R. Zupa, C. Pousis, R. Vassallo-Agius, H. Gordin, and A. Corriero. 2010. Spawning induction and large-scale collection of fertilized eggs in captive Atlantic Bluefin Tuna (Thunnus thynnus L.) and the first larval rearing efforts. Journal of Applied Ichthyology 26:596–599.

Durand, J. D., A. Collet, S. Chow, B. Guinand, and P. Borsa. 2005. Nuclear and mitochondrial DNA markers indicate unidirectional gene flow of Indo-Pacific to Atlantic Bigeye Tuna (Thunnus obe-sus) populations, and their admixture off southern Africa. Marine Biology 147:313–322.

Ely, B., J. Viñas, J. R. Alvarado Bremer, D. Black, L. Lucas, K. Cov-ello, A. V. Labrie, and E. Thelen. 2005. Consequences of the historical demography on the global population structure of two highly migratory cosmopolitan marine fishes: the Yellowfin Tuna (Thunnus albacares) and the Skipjack Tuna (Katsuwonus pela-mis). BMC Evolutionary Biology 5:19.

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Ghysen, A., K. Schuster, D. Coves, F. de la Gándara, N. Papandroula-kis, and A. Ortega. 2010. Development of the posterior lateral line system in Thunnus thynnus, the Atlantic Blue-fin Tuna, and in its close relative Sarda sarda. International Journal of Developmen-tal Biology 54:1317–1322.

Gordoa, A. 2010. The Atlantic Bluefin Tuna: study of the temporal pattern of spawning in the Western Mediterranean region and re-productive capacity in captivity. Collective Volumes of Scientific Papers ICCAT 65:837–847.

Figure 4. Unrooted phylogenetic tree based on the Thunnus sp. fragment (792 base pair) of mtDNA control region locus, inferred by neighbor-join-ing analysis. RL = reared larvae; RA = captive adult; WL = wild larvae sequences.


Gordoa, A., M. P. Olivar, R. Arevalo, J. Viñas, B. Moli, and X. Illas. 2009. Determination of Atlantic Bluefin Tuna (Thunnus thynnus) spawning time within a transport cage in the western Mediterra-nean. ICES Journal of Marine Science 66:2205–2210.

Hedrick, P. W. 1999. Perspective: highly variable loci and their inter-pretation on evolution and conservation. Evolution 53:313–318.

Hills, D. M., and J. J. Bull. 1993. An empirical test of bootstrapping as a method for assessing confidence in phylogenetic analysis. Systematic Biology 42:182–192.

Hutchinson, W. 2009. Southern Bluefin Tuna (Thunnus maccoyii) lar-val rearing advances at the South Australian Research and Devel-opment Institute and collaborating institutions. Pages 38 –42 in G. Allan, M. Booth, G. Mair, S. Clarke, and A. Biswas, editors. Pro-ceedings of the 2nd Global COE Program Symposium of Kinki University, 2009, sustainable aquaculture of the Bluefin and Yel-lowfin Tuna—closing the life cycle for commercial production. Kinki University, Osaka, Japan.

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———. 2011b. 22nd Regular Meeting of ICCAT, Istanbul, Turkey, 19th November 2011. Press release. ICCAT, Madrid, Spain. Avail-able: www.iccat.es/Documents/Meetings/COMM2011/PRESS-REL2011_ENG.pdf. (March 2012).

Iguchi, K., Y. Tanimura, H. Takeshima, and M. Nishida. 1999. Genetic variation and geographic population structure of amphidromous Ayu Plecoglossus altivelis as examined by mitochondrial DNA sequencing. Fisheries Science 65:63–67.

Ishikawa, S., J. Aoyama, K. Tsukamoto, and M. Nishida. 2001. Popula-tion structure of the Japanese Eel Anguilla japonica as examined by mitochondrial DNA sequencing. Fisheries Science 67:246–253.

IUCN (International Union for Conservation of Nature and Natural Resources). 2013. IUCN Red List of Threatened Species. Version 2013.1. Available: www.iucnredlist.org. (August 2013).

Librado, P., and J. Rózas. 2009. DnaSP v5: a software for compre-hensive analysis of DNA polymorphism data. Bioinformatics 25:1451–1452.

Martinez, P., E. G. Gonzalez, R. Castilho, and R. Zardoya. 2006. Genetic diversity and historical demography of Atlantic Bigeye Tuna (Thunnus obesus). Molecular Phylogenetics and Evolution 39:404–416.

Masuma, S., N. Tezuka, M. Koiso, T. Jinbo, T. Takebe, H. Yamazaki, H. Obana, K. Ide, H. Nikaido, and H. Imaizumi. 2006. Effects of water temperature on Bluefin Tuna spawning biology in captivity. Bulletin of Fisheries Research Agency 4:157–171.

Masuma, S., N. Tezuka, H. Obana, N. Suzuki, K. Nohara, and S. Chow. 2003. Spawning ecology of captive bluefin tuna (Thunnus thynnus orientalis) inferred by mitochondrial DNA analysis. Bulletin of Fisheries Research Agency 6:9–14.

Miyashita, S. 2002. Studies on the seedling production of the Pacific Bluefin Tuna, Thunnus thynnus orientalis. Bulletin of Fisheries Laboratory Kinki University 8:1–171. (In Japanese).

Miyashita, S., Y. Sawada, T. Okada, O. Murata, and H. Kumai. 2001. Morphological development and growth of laboratory-reared lar-val and juvenile Thunnus thynnus (Pisces: Scombridae). Fisheries Bulletin 99:601–616.

Mylonas, C. C., C. Bridges, H. Gordin, A. B. Ríos, A. García, F. de La Gándara, C. Fauvel, M. Suquet, A. Medina, M. Papadaki, G. Heinisch, G. de Metrio, A. Corriero, R. Vassallo-Agius, J.-M. Guzmán, E. Mañanos, and Y. Zohar. 2007. Preparation and ad-ministration of gonadotropin-releasing hormone agonist (GnRHa) implants for the artificial control of reproductive maturation in captive-reared Atlantic Bluefin Tuna (Thunnus thynnus thynnus). Reviews in Fisheries Science 15:183–210.

Mylonas, C. C., A. Fostier, and S. Zanuy. 2010. Broodstock manage-ment and hormonal manipulations of fish reproduction. General and Comparative Endocrinology 156:516–534.

Nakadate, M., T. Kusano, H. Fushimi, H. Kondo, I. Hirono, and T. Aoki. 2011. Multiple spawning of captive Pacific Bluefin Tuna (Thunnus orientalis) as revealed by mitochondrial DNA analysis. Aquaculture 310:325–328.

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Niwa, Y., A. Nakazawa, D. Margulies, V. P. Scholey, J. B. Wexler, and S. Chow. 2003. Genetic monitoring for spawning ecology of cap-tive Yellowfin Tuna (Thunnus albacares) using mitochondrial DNA variation. Aquaculture 218:387–395.

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Reproduction of the Bluefin Tuna in Captivity. 2006. Final report of the FP5 project “Reproduction of the Bluefin Tuna in Captivity—fea-sibility study for the domestication of Thunnus thynnus.” Avail-able: http://www.uni-duesseldorf.de/MathNat/Zoophys/bridges/REPRODOTT%20Final%20Report.pdf. (August 2013).

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———. 2009. Project report “From capture based to SELF-sustained aquaculture and domestication of Bluefin Tuna, Thunnus thyn-nus.” Available: sites.google.com/site/selfdottpublic/reports2. (March 2012).

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Viñas, J., J. R. Alvarado Bremer, and C. Pla. 2004. Inter-oceanic ge-netic differentiation among Albacore (Thunnus alalunga) popula-tions. Marine Biology 145:225–232.

Wexler, J. B., V. P. Scholey, R. J. Olson, D. Margulier, A. Nakazawa, and J. M. Suter. 2003. Tank culture of Yellowfin Tuna, Thunnus albacares: developing a spawning population for research pur-poses. Aquaculture 220:327–353.


In Videos

MEMBER SPOTLIGHT

Milton Love and Alena PribylTwo of our members—Milton Love and Alena Pribyl—have cowritten

(along with John Hyde) the script for an edgy, fun, and informative video explaining how the long-lived rockfish (many live to be 100 years or more) get barotrauma. Fisheries professionals, as well as the public, can take home good knowledge from watching this and learning about the different devices used to recompress a rockfish by returning it to its depth of capture. Seven species of rockfish are considered overfished, and several states require these species to be discarded if captured. However, just throwing these fish overboard often leads to their demise because of pressure-related injuries from barotrauma. Is Barotrauma Keeping You Up? Try Getting Down with Recompression! begins with “Rockfish Barotrauma” rap (written and per-formed by none other than Ray Troll and Russell Wodehouse) and segues into a scene where Pribyl converses with a red rockfish puppet about baro-trauma. This is an impressive video—and almost as impressive is the fact that it already has circa 14,500 views on YouTube.

Here’s a peek at just a few of the lyrics for the rap song:

The genus is Sebastes and we like it a lot, It means magnificent in language of the GreeksSome cool fish effluvia for icthyo geeksYellow eye, Vermillion and Canary are a fewGet to know your rockfish and what you should doHave a rig and a plan how to get them to the bottomHave a heart, do your part, send them downto where you caught ’em

To watch the full video, visit: www.youtube.com/watch?v=EiZFghwVOyI

Photo credit: RockfishCompress/YouTube.

Photo credit: Paul Wellman.

Daniel M. Pauly You take the National Geographic … you will see that 99% of the film they

present shows beauty. What they don’t show you is they have to go further and further out to capture this beauty. In the dead zones, you actually can’t see a thing. It’s like a soup and you can’t film soup. To have a real presentation of the ocean that represents reality, people should be exposed to green or brown soup—really essentially a screen that is brown 80% of the time—that would then represent the situation we are in.

In celebration of the 20th anniversary of the International Cosmos Award—the objective of the prize being to develop the concept of “The Harmonious Coexistence between Nature and Mankind”—2005 winner and American Fisheries Society (AFS) member Daniel Pauly was featured in a video, which can be seen in its entirety here:

To watch the full video, visit: www.youtube.com/watch?v=nF6Vsl8D0rw

Photo credit: ubcpublicaffairs/YouTube.


In BlogsDerrick Ogle

R is a software environment for statistical analyses and graphics that has several characteristics that make it attractive for fisheries science work. Two good descriptions of these characteristics can be found on the Inside-R and R-project web pages. Because of these advantages the R environment is rapidly gaining popularity, both generally and within the fisheries science field. However, because R is a language that must be learned, many users may not adopt R as an analytical tool. This website has been developed to provide examples of fisheries-related analyses that can be performed in R. It is my hope that these examples will inspire or motivate you to use R or, if you are already an R user, provide examples on which you can build your R knowledge.

fishr.wordpress.com/2013/07/12/fsa-0-4-1Photo credit: Patrick Blanchard.

In NewsNew Fish Species Awaits AFS Name Approval

A new species of Black Bass was discovered by scientists with the Florida Fish and Wildlife Conservation Commission in the Escambia, Per-dido, Conecuh, Choctawhatchee, Yellow, and Blackwater Rivers, noting that the DNA profile was unique, belonging to no other recognized species. “We chose the name ‘Choctaw bass’ because the species’ range overlaps the historic range of the Choctaw Indians,” said Mike Tringali, who heads the genetics laboratory at the Florida Fish and Wildlife Conservation Com-mission’s Fish and Wildlife Research Institute. “As for our recommended

scientific name, Micropterus haiaka,’haiaka’ is a Choctaw word that means ‘revealed.’” The AFS must approve the suggested scientific name for it to take effect.

www.northescambia.com/2013/06/scientists-discover-new-fish- species-in-escambia-perdido-rivers

Photo credit: Florida Fish and Wildlife Conservation Commission.

www.sonotronics.com • (520) 746-3322

Celebrating 42 years“working together to make a difference in the world we share”

...Looking back over 42 years providing Equipment for Tracking Marine Animals.

Tortuguero, Costa Rica (early 70’s)

Tag used for Turtle Tracking

Acoustic Submersible

Receiver (early 80’s)

NEW AFS MEMBERS

Mike AllenLeah AlmeidaKristen AnsteadDavid ArcherChristine BaljkoJulia BeatyShane BushLeandro CastelloEric EvansKyle HerremanDavid HohlerTeresa JohnsonConor McManusDarcy McNichollJoe Miller

Reagan MillsReid NelsonBrett OldsTed ParkerJacob RasmussenJack RuggirelloMegan SabalMike SandlerMichael SchmidtkeBenjamin SchreiberWilliam StarkJoshua ThomasCaleb TomlinsonDaniel WieferichHolly Wilson


AFS ANNUAL MEETING 2014

First Call for Papers: Québec City 2014

Fisheries and Oceans Canada, the Atlan-tic International Chapter, and the North-east Division of the American Fisheries Society (AFS) are pleased to announce the first call for papers for the 144th Annual Meeting of the American Fish-eries Society in Québec City, Canada! The meeting’s theme, “From fisheries research to management: think and act locally and globally,” should foster pre-sentations and discussions that consider topics such as:

• Growing evidence for meaningful local adaptation despite the lack of neutral genetic differentiation;

• Incorporating metapopulation con-cepts into regional assessment and management actions;

• Research on and management of transboundary stocks;

• Shifting source–sink dynamics in metapopulations under climate change and their implications for conserva-tion;

• Eel and salmon biology, ecology, and management;

• Any other topic relevant to the theme.

AFS 2014 will be held on 17–21 August 2014 at the Québec City Convention Centre, next to the historic Old City. This fortified city on the banks of the Saint Lawrence River is a UNESCO World Heritage Treasure. Come experience the “Joie de Vivre” and hospitality of Québec City’s people and prepare yourself to be amazed!

GENERAL INFORMATIONThe scientific program consists of three types of sessions: Symposia (oral presen-tations organized by individuals or groups with a common interest), Contributed


Oral Presentations (grouped into sessions by topic), and Contributed Poster Presen-tations (organized to coincide with either symposia or contributed oral presentation topics). Fisheries professionals are in-vited to submit symposia proposals and abstracts for oral or poster presentations that address the meeting’s theme or that are relevant to fisheries. We encourage participation of fisheries professionals from academia (professors and especially students), from all levels of government, from First Nations, and from the private sector. We are hoping that topics related to marine systems and invertebrate re-sources will be well represented at the meeting.

SYMPOSIAThe Program Committee invites propos-als for symposia. Symposia related to the meeting theme will receive priority, and those not addressing the meeting theme should be of general interest to AFS members. The Program Committee also strongly encourages integrative symposia that span freshwater and marine systems (e.g., freshwater and marine phases of eel and Atlantic Salmon, stock assessment methods, etc.).

Symposium organizers are responsible for recruiting presenters, soliciting their abstracts, and directing them to submit their abstracts and presentations through the AFS online submission forms. The Program Committee will work with symposium organizers to incorporate ap-propriate presentations that were submit-ted as contributed papers. A symposium should include a minimum of 10 presen-tations. Time slots for oral presentations are limited to 20 minutes, but multiple time slots (i.e., 40 or 60 minutes) may be offered to keynote symposia speakers.

Symposium proposals must be submitted by 10 January 2014. All symposium proposal submissions must be made using the AFS online symposium proposal submission form available on the AFS website (www.fisheries.org). The Program Committee will review all symposium proposals and notify organizers of acceptance or refusal by 31 January 2014. Please note that once the core speakers of a symposium are

confirmed, organizers will use the AFS list server to contact additional potential speakers, especially students and young professionals with whom they may not be familiar, to broaden participation by the membership. If accepted, symposium organizers must submit a complete list of all confirmed presentations and titles by 7 March 2014. Symposium abstracts (in the same format as contributed oral and poster abstracts; see below) are due by 14 March 2014.

FORMAT FOR SYMPOSIUM PROPOSALS

(Submit using AFS online symposium submission form)

When submitting your abstract, include the following:

1) Symposium title: Brief but descrip-tive.

2) Organizer(s): Provide name, affilia-tion, telephone number, e-mail address of each organizer. The first name entered will be the main contact person.

3) Chairs: Supply name(s) of individual(s) who will chair the sympo-sium.

4) Description: In 300 words or less, describe the topic addressed by the pro-posed symposium, the objective of the symposium, and the value of the sympo-sium to AFS members and participants.

5) Format: Indicate whether the sympo-sium format is for oral presentations only or a mix of oral and poster presentations.

6) Presentation requirements: Speakers should use PowerPoint for presentations.

7) Audiovisual requirements: LCD projectors and laptops will be available in every room. Other audiovisual equip-ment needed for the symposium will be considered, but computer projection is strongly encouraged. Please list special audiovisual requirements.

8) Special seating requests: Standard rooms will be arranged theater style. Please indicate special seating requests

(for example, “After the break, a panel discussion with seating for 10 panel members will be needed”).

9) List of presentations: Please supply information on potential presenters, tenta-tive titles, and oral or poster designations.

10) Sponsors: If applicable, indicate sponsorship. Please note that a sponsor is not required.

CONTRIBUTED ORAL AND POSTER PRESENTATIONS

The Program Committee invites abstracts for sessions of contributed oral and poster presentations. Authors must indicate their preferred presentation format:

1. Contributed oral presentation only;

2. Contributed poster presentation only;

3. Contributed oral presentation preferred, but poster presentation acceptable.

Only one contributed oral presentation will be accepted for each senior author. Con-tributed oral presentations will be orga-nized by 20-minute time slots (14 minutes for presentation, 3 minutes for questions, and 3 minutes for room change or further questions). All oral presenters are expected to deliver PowerPoint presentations.

We encourage poster submissions be-cause of the limited time available for oral presentations. The program will include a dedicated poster session to encourage discussion between poster authors and at-tendees. Presenters are currently expected to have hard copies of their poster, but the Program Committee is exploring the pos-sibility of incorporating electronic post-ers. Further details will be provided in subsequent calls for papers.

STUDENT PRESENTERSStudent presenters must indicate whether they wish their contribution to be consid-ered for competition for a best presenta-tion (paper or poster, but not both) award. If the response is “no,” the presentation will be considered for inclusion in the An-nual Meeting by the Program Committee but will not receive further consideration


by the Student Judging Committee. If the response is “yes,” the student will be re-quired to submit an application to the Stu-dent Judging Committee. Components of the application will include an extended abstract and a check-off from the stu-dent’s mentor indicating that the study is at a stage appropriate for consideration for an award.

ABSTRACT SUBMISSIONAbstracts for contributed papers and poster papers must be received by 14 February 2014. All submissions must be made using the AFS online abstract sub-mission form, available at www.fisheries.org. When submitting your abstract:

• Provide a brief but descriptive title, avoiding acronyms or scientific names in the title unless the common name is not widely known;

• List all authors, their affiliations, ad-dresses, telephone numbers, and e-mail addresses;

• Provide a summary of your find-ings and restrict your abstract to 200 words;

• Provide two prioritized keywords.

All presenters will receive an e-mail con-firmation of their abstract submission and will be notified of acceptance and the des-

ignated time and place of their presenta-tion by 18 April 2014.The Program Committee will group con-tributed papers by topic based on the title and the two prioritized keywords.

Late submissions will not be accepted. The AFS does not waive registration fees for presenters at symposia, workshops, or contributed oral or poster presentation sessions. All presenters and meeting at-tendees must pay registration fees. Reg-istration forms will be available on the AFS website (www.fisheries.org) begin-ning May 2014. Register early for cost savings.

FORMAT FOR ABSTRACTSTitle: An Example Abstract for the AFS 2014 Annual Meeting

Format: Oral

Authors: Castonguay, Martin. Fisheries and Oceans Canada, Maurice Lamon-tagne Institute, 850 route de la Mer, C.P. 1000 Mont-Joli, QC G5H 3Z4; 418-775-0634; [email protected] Sainte-Marie, Bernard. Pêches et Océans Canada, Institut Maurice-Lamontagne, 850 route de la Mer, C.P. 1000 Mont-Joli, QC G5H 3Z4; 418-775-0617; [email protected]

Presenter: Martin Castonguay

Abstract: Abstracts are used by the Pro-gram Committee to evaluate and select papers for inclusion in the scientific and technical sessions of the 2014 AFS An-nual Meeting. An informative abstract contains a statement of the problem and its significance, study objectives, prin-cipal findings, and applications. The ab-stract conforms to the prescribed format and must be no more than 200 words in length.

Student presenter: No.

PROGRAM COMMITTEE CONTACTS

Program Co-Chairs:

Martin CastonguayPêches et Océans Canada / Fisheries and Oceans [email protected]

Bernard Sainte-MariePêches et Océans Canada / Fisheries and Oceans [email protected]

General information:Questions regarding the AFS 2014 meeting and Québec City, please contact [email protected] or visit www.afs2014.org


ANNOUNCEMENT

We are pleased to announce the re-lease of three new publications from the Fisheries and Aquaculture Department of the Food and Agriculture Organization of the United Nations (FAO):

Advances in Geographic Information Systems and Remote Sensing for Fisheries and Aquaculture

The essential guide to understanding the role of spatial analysis in the sustain-able development and management of fisheries and aquaculture is now avail-able in an easy-to-understand publica-tion that emphasizes the fundamental skills and processes associated with geo-graphic information systems (GIS) and remote sensing. The FAO Fisheries and Aquaculture Technical Paper “Advances in Geographic Information Systems and Remote Sensing for Fisheries and Aquaculture” outlines the re-quired spatial data and computer hardware and software as well as considerations necessary to implementing a GIS. It describes current issues, status, and applications of GIS and remote sensing to aquaculture, inland fisheries, and marine fisheries to illustrate the capabilities of these technologies. It addresses emerging thematic issues with a spatial context in fisheries and aquaculture in the near future and ways to overcome challenges in GIS work.

This publication is organized in two parts: the first is a summary version for administrators and managers, and the sec-ond contains the entire document intended for professionals in technical fields and academics. The full document is available on the CD-ROM that accompanies the summary version of the publication.

Meaden, G. J., and J. Aguilar-Manjarrez, editors. 2013. Advances in geographic information systems and remote sens-ing for fisheries and aquaculture. Summary version. FAO, FAO Fisheries and Aquaculture Technical Paper No. 552, Rome. 98 pp. Includes a CD-ROM containing the full document (425 pp.). www.fao.org/docrep/017/i3102e/i3102e00.htm

A Global Assessment of Potential for Offshore Mariculture Development from a Spatial Perspective

With the expected increase in human population and re-sulting competition for access to land and clean water, there is a growing need to transfer land-based and coastal aquaculture production systems farther offshore to increase the availability of fish and fishery products for human consumption. Maricul-

New FAO Fisheries Reports

ture, in particular offshore, offers significant opportunities for sustainable food production and development of many coastal communities, especially in regions where the availability of land, nearshore space, and freshwater are limited. A new FAO Fisheries and Aquaculture Technical Paper, “A Global Assess-ment of Potential for Offshore Mariculture Development from a Spatial Perspective,” provides, for the first time, measures of the status and potential for offshore mariculture development from a spatial perspective that are comprehensive of all mari-time countries and comparable among them. It also identifies countries that do not yet practice mariculture but have a high offshore potential.

The underlying purpose of this document is to stimulate interest in detailed assessments of offshore mariculture potential at the national level. An annex report examines remote sensing for the sustainable development of offshore mariculture.

Kapetsky, J. M., J. Aguilar-Manjarrez, and J. Jenness. 2013. A global assessment of potential for offshore mariculture development from a spatial perspective. FAO, FAO Fisheries and Aquaculture Technical Paper No. 549, Rome. 181 pp. www.fao.org/docrep/017/i3100e/i3100e00.htm

National Aquaculture Sector Overview Map Collection—User Manual

The National Aquaculture Sector Overview (NASO) map collection aims to assist FAO members in inventorying and monitoring aquaculture, using Google Earth and Google Maps technology. The collection has the potential to be used for a number of purposes, such as monitoring the status of and trends


in aquaculture development and addressing site selection and zoning issues. This user manual, available as a bilingual docu-ment in English/French, is meant to facilitate the completion of the Microsoft Excel form needed to create the NASO maps. The manual is intended for all FAO members who report aquaculture statistics to the FAO and for inventorying and monitoring aqua-culture in their respective countries and territories.

The NASO map collection is being developed by the Aqua-culture Branch in collaboration with the Fisheries and Aquacul-ture Statistics and Information Branch of the FAO Fisheries and Aquaculture Department.

Aguilar-Manjarrez, J. and V. Crespi. 2013. National aquaculture sector overview map collection. User manual [Vues générales du secteur aquacole national (NASO). Manuel de l’utilisateur]. FAO, Rome. 65 pp. www.fao.org/docrep/018/i3103b/i3103b00.htm

For feedback about these publications and/or for collaborative work please contact:

Dr. José Aguilar-ManjarrezAquaculture Officer, Aquaculture BranchFisheries and Aquaculture DepartmentFood and Agriculture Organization of the United NationsViale delle Terme di Caracalla, 00153 Rome, ItalyTel.: +39 06 570 55452 Fax: +39 06 570 53020E-mail: [email protected]


Temperature-Driven Decline of a Cisco Popu-lation in Mille Lacs Lake, Minnesota. Rajeev Kumar, Steven J. Martell, Tony J. Pitcher, and Divya A. Var-key. 33:669–681.

Influences of Ripar-ian Vegetation on Trout Stream Temperatures in Central Wisconsin. Ben-jamin K. Cross, Michael A. Bozek, and Matthew G. Mitro. 33:682–692.

Genetic Monitor-ing of Threatened Chi-

nook Salmon Populations: Estimating Introgression of Nonnative Hatchery Stocks and Temporal Genetic Changes. Donald M. Van Doornik, Debra L. Eddy, Robin S. Waples, Stephen J. Boe, Timothy L. Hoffnagle, Ewann A. Berntson, and Paul Moran. 33:693–706.

[Management Brief] Evaluating Salmon Spawning Habitat Capacity Using Redd Survey Data. Phillip A. Groves, James A. Chandler, Brad Alcorn, Tracy J. Richter, William P. Connor, Aaron P. Garcia, and Steven M. Bradbury. 33:707–716.

[Management Brief] Post-Release Survival and Behavior of Adult Shoal Bass in the Flint River, Georgia. Travis R. Ingram, Josh E. Tannehill, and Shawn P. Young. 33:717–722.

Self-Reporting Bias in Chinook Salmon Sport Fisheries in Idaho: Implications for Roving Creel Surveys. Joshua L. McCor-mick, Michael C. Quist, and Daniel J. Schill. 33:723–731.

[Management Brief] Precision Analysis of Three Aging Struc-tures for Amphidromous Dolly Varden from Alaskan Arctic Rivers. Jason T. Stolarski and Trent M. Sutton. 33:732–740.

Use of Stable Isotopes to Identify Redds of Putative Hatchery and Wild Atlantic Salmon and Evaluate Their Spawning Habitat and Egg Thiamine Status in a Lake Ontario Tributary. John D. Fitzsimons, Alex Dalton, Brydon MacVeigh, Mark Heaton, Chris Wil-son, and Dale C. Honeyfield. 33:741–753.

A Comparison of Methods to Estimate Shovelnose Sturgeon Mortality in the Mississippi River Adjacent to Missouri and Illi-nois. Quinton E. Phelps, Ivan Vining, David P. Herzog, Ross Dames, Vince H. Travnichek, Sara J. Tripp, and Mark Boone. 33:754–761.

Summer Habitat Use of Large Adult Striped Bass and Habi-tat Availability in Lake Martin, Alabama. Steven M. Sammons and David C. Glover. 33:762–772.

Insight into Age and Growth of Northern Snakehead in the Potomac River. John Odenkirk, Catherine Lim, Steve Owens, and Mike Isel. 33:773–776.

JOURNAL HIGHLIGHTSNorth American Journal of Fisheries ManagementVolume 33, Number 4, August 2013

[Management Brief] Homing of Sockeye Salmon within Hidden Lake, Alaska, Can Be Used to Achieve Hatchery Man-agement Goals. Christopher Habicht, Terri M. Tobias, Gary Fandrei, Nathan Webber, Bert Lewis, and W. Stewart Grant. 33:777–782.

Evaluation of Internal Tag Performance in Hatchery-Reared Juvenile Spotted Seatrout. Jonathan P. Wagner, Regi-nald B. Blaylock, and Mark S. Peterson. 33:783–789.

[Management Brief] Posthandling Survival and PIT Tag Retention by Alewives—A Comparison of Gastric and Surgical Implants. Theodore Castro-Santos and Volney Vono. 33:790–794.

[Management Brief] Juvenile Movement among Differ-ent Populations of Cutthroat Trout Introduced as Embryos to Vacant Habitat. Tessa M. Andrews, Bradley B. Shepard, Andrea R. Litt, Carter G. Kruse, Alexander V. Zale, and Steven T. Kalin-owski. 33:795–805.

[Management Brief] Mortality of Palmetto Bass Follow-ing Catch-and-Release Angling. Matthew J. Petersen and Phillip W. Bettoli. 33:806–810.

[Management Brief] Sexual Dimorphism in Alligator Gar. D. L. McDonald, J. D. Anderson, C. Hurley, B. W. Bumguardner, and C. R. Robertson. 33:811–816.

Spatiotemporal Predictive Models for Juvenile Southern Flounder in Texas Estuaries. Bridgette F. Froeschke, Philippe Tissot, Gregory W. Stunz, and John T. Froeschke. 33:817–828.

Field and Laboratory Evaluation of Dam Escapement of Muskellunge. Max H. Wolter, Corey S. DeBoom, and David H. Wahl. 33:829–838.

[Management Brief] Evaluation of Aging Structures for Silver Carp from Midwestern U.S. Rivers. Justin R. Seibert and Quinton E. Phelps. 33:839–844.

An Evaluation of Harvest Control Rules for Data-Poor Fisheries. John Wiedenmann, Michael J. Wilberg, and Thomas J. Miller. 33:845–860.

[Management Brief] A Standardized Technique to Back-Calculate Length at Age from Unsectioned Walleye Spines. Jonathan R. Meerbeek and Kimberly A. Hawkins. 33:861–868.

The Gulf of California, alsoknow as the Sea of Cortez,has 17 families and 67 speciesof chondrichthian fish, 115 families and 753 species ofbony Fish, plus two speciesof myxinids present in the of myxinids present in the Gulf of California. This work covers 105 families and over400 species.

Includes an illustrated glossary, 38 color plates,with over 200 individual fish images, and more.fish images, and more.

To Order:www.afsbooks.org


Swanson, C. 2013. Comments on USEPA’s 30 April 2013 draft as-sessment of potential mining impacts on Salmon ecosystems of Bristol Bay, Alaska. Western Division American Fisheries Soci-ety, Docket # EPA-HQ-ORD-2013-0189. Available: http://wdafs.org/meetings/archives/resolutions/. (August 2013).

U.S. Environmental Protection Agency. 2006. TENORM (technologi-cally enhanced naturally occurring radioactive materials) uranium location database. Office of Radiation & Indoor Air, Radiation Protection Division, Washington, D.C.

U.S. Geological Survey. 2009. Mineral resources program. Avail-able: http://tin.er.usgs.gov/metadata/mineplant.faq.html. (August 2013).

———. 2012. National coal resources data system ustratigraphic (US-TRAT) database. Available: http://energy.usgs.gov/Tools/Nation-alCoalResourcesDataSystem.aspx. (August 2013).

Whittier, T. R., R. M. Hughes, G. A. Lomnicky, and D. V. Peck. 2007. Fish and amphibian tolerance values and an assemblage tolerance index for streams and rivers in the western USA. Transactions of the American Fisheries Society 136:254–271.

Continued from page 391

Continued from page 399Those words apply to many fields, certainly including fish

habitat. With our academic skills and professional experiences, we are expected to perform feats of greatness. We can’t part the Red Sea or eradicate the most aggressive invasive species, but we can make heroic strides toward economic and ecologi-cal success. Just as other fields encounter significant paradigm shifts, some of us might need remedial training or peer educa-tion so we can embrace fish habitat in our daily routines. Those latest lessons will prove valuable as we strive to convince our leaders to engage on behalf of habitat. The fish will thank us!

This commitment to discovery and learning begins with people. Young professionals and trusted employees alike must be familiar with the full breadth of habitat topics, threats, man-dates, and opportunities. We all should find inspiration in ex-periencing habitat through educational fish camps organized across the nation, local wetland conservation opportunities such as plantings for erosion control, special events such as Salmon and Steelhead Days in the mountains, Coastweeks in early autumn, and ecology clubs in most schools. Those and other opportunities nourish the latent student in all of us while also garnering support for habitat fields ranging from obscure to vital.

This dual need for education and capacity also extends to organizations. At a National Fish Habitat Partnership (fishhabi-tat.org) board meeting in late June 2013, Wendy Wilson of the River Network (rivernetwork.org) implored the board and its 18 regional fish habitat partnerships to create an “interlocking national program” that is flexible and accountable and that is forever committed to matching goals with the capacity to suc-ceed. The River Network focuses on program success, which in turn hinges on individual commitments. It is akin to the circle of life, to borrow from the Disney film The Lion King (www.sub-zin.com/quotes/The+Lion+King/The+circle+of+life). Indeed, because “most fish biologists are not entrepreneurs” (Wendy Wilson, National Fish Habitat Partnership board meeting, June

25, 2013), organizations must strive for the best mix of funding streams, communications strategies, staffing, and partnerships with experts from other sectors as they seek to educate and in-fluence.

Fortunately, for individuals and organizations, there are outstanding models to help us meet increasing expectations for habitat science, management, and policy. Public perceptions seem to be shifting. The strong engagement spurred by the early Earth Days of the 1970s waned within a few decades. Now, with weather extremes, coastal wetland loss, and fish population col-lapses served up more and more as regular features on the eve-ning news, the public seems to have connected the dots. They want more information. And corporate business seems pleased to deliver. We can use that to our advantage, because each story is an opportunity for us to excel and for American Fisheries Society to provide an expert witness or fish story.

I offer two final thoughts about education and perceptions. Budgets seem to have shifted the traditional model of separate sector meetings and conferences for each society. Joint society conferences now offer less economic risk and greater oppor-tunity for interdisciplinary connections and awareness; that is, education. Another way to rally people to benefit habitat is to develop best management practices for topics of mutual inter-est. I’ve been involved in many efforts, each of which led to cost-efficient, effective, and consistent approaches to recurring challenges. Such efforts are natural opportunities to learn and advance our field.

As our careers course through lecture halls, cubicles, and the wilds of fish habitat, let us always remember the need to learn and share. I remain excitedly optimistic that as leaders and the public learn more about the importance of fish habitat, we’ll have increased success across our fields of interest. Healthier habitat will support harvested and protected species and the eco-system components that provide forage and structure.

From the Archives

The history of practical trout culture, by itself; hardly dates back ten years. Without meaning to detract from the mer-its of Mr. Ainsworth’s exertions, which were all the more meritorious because they were, in the practical sense in which he made them, original, I think we can safely say that the art of practical trout culture dates from the time that Seth Green bought the Caledonia stream, and demonstrated to the world what a great success could be made of practical trout breeding.

Livingston Stone (1872): Trout Culture, Transactions of the American Fisheries Society, 1:1, 46-56.


DATE EVENT LOCATION WEBSITE

September 23–25, 2013 2nd Annual World Congress of Mariculture and Fisheries-2013 (WCMF-2013)

Hangzhou, China bitconferences.com/wcmf2013/default.asp

September 23–26, 2013 OCEANS ‘13 MTS/IEEE - The Largest Ocean Conference in U.S. History

San Diego, CA oceans13mtsieeesandiego.org

September 28–October 4, 2013

2013 World Seafood Conference Newfoundland and Labrador, Canada

wsc2013.com

October 1–4, 2013 The Wild Trout Symposium XI Yellow Stone Na-tional Park, USA

wildtroutsymposium.com

October 7–11, 2013 40th Annual Meeting of the Alaska Chapter of AFS

Fairbanks, AK afs-alaska.org/annual-meetings/2011-2

October 21–27, 2013 3rd International Marine Protected Areas Congress

Marseille, France impac3.org

January 23–26, 2014 Southern Division Spring Meeting Charleston, SC sdafs.org/meeting2014

April 7–12, 2014 The Western Division Meeting’s 2nd International Mangroves as Fish Habitat Symposium

Mazatlan, Mexico fishconserve.org/email_messages/ Mangrove_Symposium.html

August 3–7, 2014 International Congress on the Biology of Fish Edinburgh, United Kingdom

icbf2014.sls.hw.ac.uk

August 17–21, 2014 AFS Annual Meeting 2014 Québec City, Canada

afs2014.org

August 31–September 4, 2014

International Symposium on Aquatic Animal Health (ISAAH)

Portland, OR afs-fhs.org/meetings/meetings.php

CALENDARFisheries Events

To submit upcoming events for inclusion on the AFS web site calendar, send event name, dates, city, state/ province, web address, and contact information to [email protected].

(If space is available, events will also be printed in Fisheries magazine.)

More events listed at www.fisheries.org


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In waterways around the world, smolt survival continues to be a problem despite efforts to reverse these trends. Predation has been identified as one of the causes of the decline. Acoustic telemetry is commonly used to track downstream migration of salmonids and has recently been used to identify predatory behavior in other species. Over the past decade, fine-scale fish tracks have illustrated migration behavior and survival in river systems throughout the world. New questions have emerged in recent years as more data becomes available from various species via fine-scale 2D/3D telemetry. A river system example is California’s Bay-Delta region. One of the principal questions of great importance in this region is: Can we determine whether or not an acoustically-tagged fish has been eaten by a predator? A critical assumption of survival estimation for tagged migrating species is that the detected tag signals are from distinctly unconsumed and freely migrating fish. With that, protocols for determining predatory-like movement have been objectively defined for use in analyzing telemetry data. Sound pulses from acoustic tags easily pass through the body wall of a fish, even if a smaller fish is consumed by a larger fish. To correctly interpret tag data it is important to recognize a predation event in order to correctly classify a tagged fish for survival studies. If the acoustic tags have short, precisely controlled transmission intervals, detection & ID ranges that are the

same, and are detected on multiple hydrophones at once, then accurate tracks of individual fish can be generated (Ehrenberg and Steig 2009). Two tagged smolts whose tracks overlap in space and time (appear to swim together) may indicate a predator has consumed two tagged smolts. Another possibil-ity is the tagged smolts are exhibiting schooling behavior. A likely predation event (shown left) shows two tags have continuously overlapping tracks for over three days and one of the tags became completely stationary (likely defecated) within the array. That tag remained stationary and slowly sunk into the substrate for several more days until the end of the tag battery life. Understanding predator behavior and distinguishing it from migrating smolt behavior is key to correctly interpreting acoustic tag results (Vogel 2010). If fine scale 2D or 3D track data is available, then sudden behavioral changes or characteristic, quantifiable behavioral patterns can be used to infer predation events. However, behavior results should be interpreted in context with concurrent environmental factors, including simple sinuosity and average speed over ground for known tagged predators, tagged smolts, high tide (low water velocity) and low tide (high water velocity). As acoustic telemetry advances, find out more about how scientists currently track fish behavior and determine predation events by contacting the fisheries scientists at [email protected].

Monitoring Survival: Steps Toward Evaluating Predation Using Acoustic Tags

Fish

Imag

es: F

ISH

BIO

(206) 633-3383HTIsonar.com

Measuring the impact of predator species on its prey is difficult, that isn’t new. What is new is how we track fish behavior & measure species interactions. Technological advances in acoustic transmitters continue to show promise in revealing complex ecological dynamics.Sam Johnston, Senior Scientist, HTI Hydroacoustic Technology, Inc.

‘Radar’ plots of movement of predatory fish and salmon smolts.The complete track of each fish was broken up into line segments of approximately 10 seconds in duration. The direction of travel of each segment was then calculated and summarized in the above plots (0 degrees is True North).

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Increasing rates of predation on juvenile salmonids is a challenge.