AAVP 2010 Annual Meeting Proceedings

American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Proceedings

1 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS AWARDS HISTORY

AAVP-Merial Distinguished Veterinary Parasitologists 1985 Jitender P. Dubey* 1986 Norman D. Levine 1987 E. J. Lawson Soulsby 1988 Jeffrey F. Williams 1989 K. Darwin Murrell 1990 William C. Campbell 1991 Jay Hal Drudge and Eugene T. Lyons 1992 Gilbert F. Otto 1993 Thomas R. Klei 1994 Peter M. Schantz 1995 James C. Williams 1996 T. Bonner Stewart 1997 J. Owen D. Slocombe 1998 J. Ralph Lichtenfels 1999 Roger K. Prichard 2000 Edward L. Roberson 2001 Byron L. Blagburn 2002 Sidney A. Ewing 2003 Louis C. Gasbarre 2004 David S. Lindsay 2005 Jorge Guerrero 2006 John W. McCall 2007 Ronald Fayer 2008 Dwight D. Bowman 2009 Ellis C. Greiner 2010** *National Academy of Sciences 2010 ** As of 2010, the award name has been changed to “AAVP-Merial Distinguished Veterinary Parasitologist Award”.

2010 AAVP-Merial Distinguished Veterinary Parasitologist Award Winner

George A. Conder, PhD

2 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Biographical Sketch

GEORGE A. CONDER After completing his M.S. in 1975, George Conder went to work with Ferron Anderson at BYU, Provo UT, where he completed his Ph.D. in 1979. He then did postdoctoral work with Jeff Williams at Michigan State University (1979-81), East Lansing, MI, before accepting a position (1981) as a Scientist I with Upjohn Company, Kalamazoo, MI. While at Upjohn, George progressed through the ranks to Scientist IV and became an Adjunct Professor at UTEP, where he was able to do collaborative work with Jack Bristol and Lil Mayberry. In 1994, George moved to Pfizer, Inc. to work as a Clinical Project Manager of their Animal Health Product Development group, in Groton, CT. George moved through the Pfizer ranks and in 2007, assumed the position of Director & Therapeutic Area Head of anti- parasitic drugs for Pfizer, but at their office in Kalamazoo, MI. Over the years he has continued a highly productive professional career as a parasitologist and has won many honors in the pharmaceutical industry, including: Fred Kagan Lead Finding Award, The Upjohn Company, 1988; W.E. Upjohn Award, The Upjohn Company, 1993; and the Central Research Achievement Award, Pfizer Inc., 1997. He also has served as Vice President (1992), President Elect (1993), President (1994), and Immediate Past President (1995) of the American Association of Veterinary Parasitologists.

3 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS AWARDS HISTORY

Hoechst-Roussel Agri-Vet Company Graduate Student Research Award 1987 Lora G. Rickard 1988 Debra A. Cross 1989 Stephen C. Barr 1990 Jim C. Parsons 1991 Carlos E. Lanusse 1992 David G. Baker 1993 Rebecca A. Cole 1994 Ray M. Kaplan 1995 Steve T. Storandt 1996 A. Lee Willingham III 1997 Carla C. Siefker 1998 Ryan M. O’Handley 1999 John S. Mathew 2000* Sheila Abner 2001 Andrew Cheadle 2002 No recipient 2003 Mary G. Rossano 2004 Andrea S. Varela 2005 Alexa C. Rosypal 2006 Sheila M. Mitchell 2007 Martin K. Nielsen 2008** Heather D. Stockdale 2009 Kelly E. Allen

*As of 2000, this award was renamed the “AAVP/Intervet Graduate Student Research Award”. **As of 2008 this award was renamed the “AAVP-Intervet/Schering Plough Outstanding Graduate Student Research Award”.

AAVP-Companion Animal Parasite Council (CAPC) Graduate Student Award in Zoonotic Disease 2008 David G. Goodman 2009 Stephanie R. Heise

Distinguished Service 1976 Rurel R. Bell 1983 Terrance J. Hayes 1987 Norman F. Baker 1988 Donald E. Cooperrider 1994 S. D. “Bud” Folz 1997 Honorico Rick Ciordia 2006 Raffaele “Raf” Roncalli 2008 Anne M. Zajac

4 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS Founded 1956

Officers 2009-2010

President Lora R. Ballweber Colorado State University Fort Collins, CO

President-Elect Karen Snowden Texas A&M University College Station, TX

Vice-President Patrick F. M. Meeus Pfizer Animal Health Kalamazoo, MI

Secretary/Treasurer Alan A. Marchiondo Pfizer Animal Health Kalamazoo, MI

Immediate Past-President Susan E. Little Oklahoma State University Stillwater, OK

2009-2010 AAVP Committee Chairs

Program - Patrick F. M. Meeus, Kalamazoo, MI Archives - Tom J. Nolan, Philadelphia, PA Historian - Raf Roncalli, Milltown, NJ Awards - Andrew S. Peregrine, Guelph, Onterio, Canada Constitution/Bylaws - Wendell L. Davis, Overland Park, KS Education - Gary Conboy, Charlottetown, PEI, Canada Finance - Ray M. Kaplan, Athens, GA Newsletter - Jenifer Edmonds, Parma, ID Nominations - Dwight D. Bowman, Ithaca, NY Outreach/Research - Anne M. Zajac, Blacksburg, VA Publications/Internet - Tom J. Nolan, Philadelphia, PA Student Representatives - Stephanie Heise, Stillwater, OK & Ashley Linton, Fort Collins, CO List Serve Manager - Bert Stromberg, St. Paul, MN

5 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

PAST PRESIDENTS OF THE AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS

1956-1958 L. E. Swanson 1958-1960 Fleetwood R. Koutz 1960-1962 Wendell H. Krull 1962-1964 Saeed M. Gaafar 1964-1966 E. D. Besch 1966-1968 George C. Shelton 1968-1970 John H. Greve 1970-1972 Harold J. Griffiths 1972-1973 Donald E. Cooperrider 1973-1975 Demetrice L. Lyles 1975-1977 Harold J. Smith 1977-1979 Norman F. Baker 1979-1981 Edward L. Roberson 1981-1983 Jeffrey F. Williams 1983-1985 John B. Malone 1985-1986 Robert M. Corwin 1986-1987 K. Darwin Murrell 1987-1988 Thomas R. Klei 1988-1989 Harold C. Gibbs 1989-1990 Bert E. Stromberg 1990-1991 Roger K. Prichard 1991-1992 J. Owen D. Slocombe 1992-1993 Ronald Fayer 1993-1994 George A. Conder 1994-1995 Charles H. Courtney 1995-1996 Byron L. Blagburn 1996-1997 Peter M. Schantz 1997-1998 James C. Williams 1998-1999 Louis C. Gasbarre 1999-2000 Robert S. Rew 2000-2001 Thomas J. Kennedy 2001-2002 Anne M. Zajac 2002-2003 Joseph F. Urban, Jr. 2003-2004 Craig R. Reinemeyer 2004-2005 Linda S. Mansfield 2005-2006 Ann R. Donoghue 2006-2007 Daniel E. Snyder 2007-2008 David S. Lindsay 2008-2009 Susan E. Little 2009-2010 Lora R. Ballweber

6 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

PAST SECRETARY-TREASURERS PAST PRESIDENTS OF THE OF THE AMERICAN ASSOCIATION OF AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS VETERINARY PARASITOLOGISTS Wendell H. Krull 1956-1959 1956-1958 L. E. Swanson Edward G. Batte 1960 1958-1960 Fleetwood R. Koutz Donald E. Cooperrider 1961-1969 1960-1962 Wendell H. Krull Rurel Roger Bell 1969-1977 1962-1964 Saeed M. Gaafar Terence J. Hayes 1978-1983 1964-1966 E. D. Besch Vassilios J. Theodorides 1983-1986 1966-1968 George C. Shelton S.D. “Bud” Folz 1987-1992 1968-1970 John H. Greve Thomas J. Kennedy 1993-1998 1970-1972 Harold J. Griffiths Daniel E. Snyder 1998-2004 1972-1973 Donald E. Cooperrider Alan A. Marchiondo 2004-2010 1973-1975 Demetrice L. Lyles 1975-1977 Harold J. Smith 1977-1979 Norman F. Baker PAST AAVP ANNUAL MEETINGS 1979-1981 Edward L. Roberson 1981-1983 Jeffrey F. Williams 1956 1st Annual Meeting – SAN ANTONIO, TX 16 OCT 1983-1985 John B. Malone 1957 2nd Annual Meeting – COLUMBUS, OH 17 AUG 1985-1986 Robert M. Corwin 1958 3rd Annual Meeting – PHILADELPHIA, PA 18 AUG 1986-1987 K. Darwin Murrell 1959 4th Annual Meeting – KANSAS CITY, MO 23 AUG 1987-1988 Thomas R. Klei 1960 5th Annual Meeting – DENVER, CO 14 AUG 1988-1989 Harold C. Gibbs 1961 6th Annual Meeting – WEST LAFAYETTE, IN 20 AUG 1989-1990 Bert E. Stromberg 1962 7th Annual Meeting – MIAMI BEACH, FL 12 AUG 1990-1991 Roger K. Prichard 1963 8th Annual Meeting – NEW YORK CITY, NY 28 JUL 1991-1992 J. Owen D. Slocombe 1964 9th Annual Meeting – CHICAGO, IL 19 JUL 1992-1993 Ronald Fayer 1965 10th Annual Meeting – PORTLAND, OR 11 JUL 1993-1994 George A. Conder 1966 11th Annual Meeting – LOUISVILLE, KY 13-14 JUL 1994-1995 Charles H. Courtney 1967 12th Annual Meeting – DALLAS, TX 9 JUL 1995-1996 Byron L. Blagburn 1968 13th Annual Meeting – BOSTON, MA 21 JUL 1996-1997 Peter M. Schantz 1969 14th Annual Meeting – MINNEAPOLIS, MN 13 JUL 1997-1998 James C. Williams 1970 15th Annual Meeting – LAS VEGAS, NV 22 JUN 1998-1999 Louis C. Gasbarre 1971 16th Annual Meeting – DETROIT, MI 18 JUL th 1999-2000 Robert S. Rew 1972 17 Annual Meeting – NEW ORLEANS, LA 17 JUL th 2000-2001 Thomas J. Kennedy 1973 18 Annual Meeting – PHILADELPHIA, PA 15 JUL th 2001-2002 Anne M. Zajac 1974 19 Annual Meeting – DENVER, CO 21 JUL th 2002-2003 Joseph F. Urban, Jr. 1975 20 Annual Meeting – ANAHEIM, CA 13 JUL st 2003-2004 Craig R. Reinemeyer 1976 21 Annual Meeting – CINCINNATI, OH 19 JUL nd 2004-2005 Linda S. Mansfield 1977 22 Annual Meeting – ATLANTA, GA 11 JUL 2005-2006 Ann R. Donoghue 1978 23rd Annual Meeting – DALLAS, TX 17 JUL 2006-2007 Daniel E. Snyder 1979 24th Annual Meeting – SEATTLE, WA 22-24 JUL 2007-2008 David S. Lindsay 1980 25th Annual Meeting – WASHINGTON, D.C. 20-22 JUL 2008-2009 Susan E. Little 1981 26th Annual Meeting – ST. LOUIS, MO 19-20 JUL 2009-2010 Lora R. Ballweber 1982 27th Annual Meeting – SALT LAKE CITY, UT 18-19 JUL 1983 28th Annual Meeting – NEW YORK, NY 17-18, JUL 1984 29th Annual Meeting – NEW ORLEANS, LA 15-17 JUL 1985 30th Annual Meeting – LAS VEGAS, NV 22-24 JUL

7 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

1986 31st Annual Meeting – ATLANTA, GA 20-22 JUL 1987 32nd Annual Meeting – CHICAGO, IL 19-21 JUL th 1988 33rd Annual Meeting – PORTLAND, OR 17-18 JUL AAVP 55 ANNUAL MEETING 1989 34th Annual Meeting – ORLANDO, FL 16-18 JUL Loews Atlanta Midtown Hotel 1990 35th Annual Meeting – SAN ANTONIO, TX 21-24 JUL 1991 36th Annual Meeting – SEATTLE, WA 28-30 JUL 1992 37th Annual Meeting – BOSTON, MA 2-4 AUG REGISTRATION 1993 38th Annual Meeting – MINNEAPOLIS, MN 17-20 JUL Ellington Ballroom PreFunction 1994 39th Annual Meeting – SAN FRANCISCO, CA 9-12 JUL th Saturday, 31 July 2010, 1:00 PM – 6:00 PM 1995 40 Annual Meeting – PITTSBURGH, PA 6-10 JUL Sunday, 1 August 2010, 8:00 AM – 3:00 PM (Joint meeting with the American Society of Parasitologists) 1996 41st Annual Meeting – LOUISVILLE, KY 20-23 JUL Monday, 2 August 2010, 8:00 AM – 12:00 Noon 1997 42nd Annual Meeting – RENO, NV 19-22 JUL 1998 43rd Annual Meeting – BALTIMORE, MD 25-28 JUL SPEAKER READY ROOM th 1999 44 Annual Meeting – NEW ORLEANS, LA 10-13 JUL OFFICE 2 2000 45th Annual Meeting – SALT LAKE CITY, UT 22-25 JUL 2001 46th Annual Meeting – BOSTON, MA 14-17 JUL Saturday, 31 July 2010, 12:00 PM – 5:00 PM 2002 47th Annual Meeting – NASHVILLE, TN 13-16 JUL Sunday, 1 August 2010, 8:00 AM – 5:00 PM 2003 48th Annual Meeting – DENVER, CO 19-23 JUL th Monday, 2 August 2010, 8:00 AM – 5:00 PM 2004 49 Annual Meeting – PHILADELPHIA, PA 24-28 JUL (Joint meeting with the American Society of Parasitologists) 2005 50th Annual Meeting – MINNEAPOLIS, MN 16-19 JUL SOCIAL PROGRAM 2006 51st Annual Meeting – HONOLULU, HI 15-18 JUL Welcoming Reception – Bayer Health Care, Animal Health 2007 52ndAnnual Meeting – WASHINGTON, DC 14-17 JUL The Terrace rd 2008 53 Annual Meeting – NEW ORLEANS, LA 19-22 JUL (Weather Backup: Salons D-F) 2009 54th Annual Meeting – CALGARY, CANADA 9-13 AUG (Joint meeting with the World Association for the Advancement of Saturday, 31 July 2010, 7:00 PM – 9:00 PM Veterinary Parasitology and the International Commission on Trichinellosis) Merial Ltd Social The Terrace (Weather Backup: Ellington Prefunction or Salons A-C) Sunday, 1 August 2010, 7:00 PM – 9:00 PM

No Social - On Your Own Monday, 2 August 2010, 7:00 PM – 9:00 PM

AAVP Graduate Student/Post Doc Luncheon/Mixer Dunwoody Sunday, 1 August 2010, Noon – 1:30 PM

AAVP 55Th ANNUAL MEETING Salon D

8 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

AAVP 55th ANNUAL MEETING Loews Atlanta Midtown Hotel

REGISTRATION Ellington Ballroom PreFunction Saturday, 31 July 2010, 1:00 PM – 6:00 PM Sunday, 1 August 2010, 8:00 AM – 3:00 PM Monday, 2 August 2010, 8:00 AM – 12:00 Noon

SPEAKER READY ROOM OFFICE 2 Saturday, 31 July 2010, 12:00 PM – 5:00 PM Sunday, 1 August 2010, 8:00 AM – 5:00 PM Monday, 2 August 2010, 8:00 AM – 5:00 PM

SOCIAL PROGRAM Welcoming Reception – Bayer Health Care, Animal Health The Terrace (Weather Backup: Salons D-F) Saturday, 31 July 2010, 7:00 PM – 9:00 PM

Merial Ltd Social The Terrace (Weather Backup: Ellington Prefunction or Salons A-C) Sunday, 1 August 2010, 7:00 PM – 9:00 PM

No Social - On Your Own Monday, 2 August 2010, 7:00 PM – 9:00 PM

AAVP Graduate Student/Post Doc Luncheon/Mixer Dunwoody Sunday, 1 August 2010, Noon – 1:30 PM

AAVP 55Th ANNUAL MEETING Salon D

9 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS 55th ANNUAL MEETING SPONSORS

EVENT SPONSORS

BAYER HEALTH CARE, ANIMAL HEALTH - Social MERIAL LTD – Social AWARDS SPONSORS BAYER HEALTH CARE, ANIMAL HEALTH1 COMPANION ANIMAL PARASITE COUNCIL2 INTERVET SCHERING PLOUGH ANIMAL HEALTH3 MERIAL, LTD.4

1AAVP Best Student Paper Presentation Awards 2Honorarium for AAVP-CAPC Graduate Student Award – Zoonotic Diseases 3Honorarium for AAVP-Intervet Schering Plough Graduate Student Award - Research 4Honorarium for AAVP-Merial Distinguished Veterinary Parasitologist Award

GENERAL MEETING SPONSORSHIP PLATINUM LEVEL CLINVET INTERNATIONAL (PTY) Ltd PFIZER ANIMAL HEALTH MERIAL, LTD

GOLD LEVEL CHARLES RIVER BIOLABS, BLE, IRELAND ELANCO ANIMAL HEALTH IDEXX LABORATORIES, INC.

SILVER LEVEL CENTRAL LIFE SCIENCES COMPANION ANIMAL PARASITE COUNCIL DIVERGENCE, INC. JOHNSON RESEARCH, LLC HMS VETERINARY DEVELOPMENT, INC. PIEDMONT ANIMAL HEALTH PROFESSIONAL LABORATORY & RESEARCH SERVICES, INC. STILLMEADOW, INC. YOUNG VETERINARY RESEARCH SERVICES

The American Association of Veterinary Parasitologists (AAVP) gratefully acknowledges the above companies for their support and sponsorship of the 2010 annual meeting. AAVP thanks Pfizer Animal Health for In-Kind Annual Meeting Support during 2009- 2010, for the mailing of the 2009 AAVP Proceedings, and sponsorship for the coffee breaks at the 2010 AAVP meeting.

10 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA NOTES

11 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA NOTES NOTES

12 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA NOTES

13 FOOD, FRIENDS AND FUN Don’t miss the reception for the 2010 meeting of the AAVP, sponsored by Bayer.

Saturday, July 31, 2010 | 7–9 pm The Terrace | Loews Atlanta Hotel

Join your colleagues for an evening of hors d’oeuvres, drinks and conversation at the Bayer reception. We hope to see you there.

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21 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Saturday July 31st Sunday August 1st Monday August 2nd Tuesday August 3rd

Centennial Salon D* Salon E (F) Salon D* Salon E (F) Salon D* Salon E (F)

8:00 8:00 8:00 8:00 Session 14 8:15 8:15 8:15 8:15 Session 13 Livestock Livestock helminth 8:30 8:30 Intervet/AAVP Outstanding 8:30 8:30 anthelmintic control, Graduate Student Talk 8:45 8:45 8:45 8:45 efficacy and diagnosis and resistance impact 9:00 9:00 9:00 Session 9 9:00 Education symposium 9:15 9:15 Session 1 Session 2 9:15 9:15 9:30 9:30 Students Students 9:30 9:30 Pfizer Coffee Break 9:45 9:45 9:45 9:45

10:00 10:00 10:00 10:00 AAVP Executive Pfizer Coffee Break Pfizer Coffee Break 10:15 Committee Meeting & 10:15 10:15 10:15 10:30 Luncheon 10:30 10:30 10:30 Session 15 President’s symposium 10:45 10:45 10:45 10:45 Livestock anthelmintic resistance 11:00 11:00 11:00 11:00 Session 3 Session 4 Session 10 11:15 11:15 Students Students 11:15 Session 11 11:15 Companion Protozoa 11:30 11:30 11:30 animal 11:30 helminths

11:45 11:45 11:45 11:45

12:00 12:00 12:00 12:00 * Default meeting room is Salon D ** Registration is in Ellington pre- 12:15 12:15 12:15 12:15 function - Sat : 13:00-18:00 12:30 12:30 12:30 12:30 - Sun : 8:00-15:00 Lunch Break - Mon : 8:00-12:00 12:45 12:45 12:45 12:45 Lunch Break *** Ready room is in Office 2 - Sat : 12:00-18:00 13:00 13:00 13:00 Student lunch in Dunwoody 13:00 (12:00-13:30) - Sun : 8:00-18:00 - Mon : 8:00-18:00 13:15 13:15 13:15 13:15

13:30 13:30 13:30 13:30 13:45 13:45 13:45 13:45 14:00 14:00 14:00 14:00

14:15 14:15 14:15 14:15

14:30 14:30 Session 5 Session 6 14:30 Session 12 14:30 Students / Companion Interesting clinical cases 14:45 Salon D 14:45 equine animal ecto 14:45 14:45 15:00 Opening Remarks 15:00 15:00 15:00

15:15 15:15 15:15 15:15

15:30 15:30 15:30 15:30 Pfizer Coffee Break 15:45 15:45 15:45 15:45 Pfizer Coffee Break 16:00 16:00 16:00 16:00

16:15 16:15 16:15 16:15 Plenary session 16:30 16:30 Session 7 16:30 16:30 16:45 16:45 Epidemiology 16:45 16:45 Session 8 17:00 17:00 Wildlife 17:00 Business meeting 17:00 17:15 17:15 17:15 17:15

17:30 17:30 17:30 17:30

17:45 17:45 17:45 17:45

18:00 18:00 18:00 18:00 Bayer Welcome 19:00 19:00 Merial Social 19:00 19:00 Social “The Terrace” 21:00 “The Terrace” 21:00 21:00 21:00

22 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Meeting floor plan

American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Program for 2010 AAVP Meeting

Saturday July 31, 2010 Sunday August 1st, 2010

8:00 to 13:00 Centennial 8:30 to 10:00 Session 1 Salon D AAVP Executive Committee Meeting & Luncheon. Students All AAVP officers and committee chairs please plan to attend! 8:30 4 Intervet/AAVP Outstanding Graduate Student Talk: On the Tracks of Erythema Migrans – Lone Star Ticks, 15:00 to 17:30 Plenary session Salon D Bacteria, and Disease. Stephanie Heise Opening Remarks President: Lora Balweber 9:00 5 Status of Cytauxzoon felis in wild felid populations in Vice President and Program Chair: Patrick Meeus the United States. Barbara C. Shock*; Staci M. Murphy; Laura L. Patton; Philip M. Shock; Colleen Olfenbuttel; Jeff Plenary session : “PARASITE EVOLUTION” - Veterinary Beringer; Suzanne Prange; Daniel M. Grove; Matthew Peek,; parasites and their strategies to survive human Mitchell Lockhart; Jay Butfiloski; Daymond W. Hughes; Sarah intervention Bevins; Victor F. Nettles; Holly M. Brown; David S. Peterson; M.J. Yabsley. 15:15 1 Shift in rules of engagement for eradication of cattle fever ticks in the United States. Adalberto A. Pérez de 9:15 6 An Ongoing Study of the Effects of Prescribed León*, J. Mathews Pound, Greta Schuster, Felix D. Guerrero, Burns on Tick and Tick-borne Pathogen Prevalence. Ronald B. Davey, Robert J. Miller, Kimberly H. Lohmeyer, Elizabeth R. Gleim*; M.J. Yabsley; Mike Conner; Michael David G. Hewitt, J. Alfonso Ortega, Tyler A. Campbell, Alex Levin E. Racelis, John A. Goolsby, Patricia Holman, Matthew Messenger, Dee Ellis, Roberta Duhaime, Liza Soliz, Andrew 9:30 7 Environmental contamination of public use areas Y. Li, Pamela L. Phillips, Kevin B. Temeyer, Pete Teel, with Toxocara spp. and Trichuris vulpis in Athens, GA. Stephen Wikel, Diane M. Kammlah, G. Gale Wagner, Kevin Jessica H. Murdock*; Joshua O. Cook; Adrienne B. Zercher; P. Varner, Danett K. Brake Dana Ambrose ; David Stallknecht; Ray M. Kaplan-; Andrew R. Moorhead 15:45 2 Clinically silent reemergence of Babesia equi infection in U. S. horses. Don Knowles 9:45 8 Effect Of Exposure To Household Laundering And Drying On Eggs Of Common Canine Helminths. Brian 16:15 3 The human filariasis elimination program - lessons Neumann*; Courtney Mosley; Anne M. Zajac learned for veterinary medicine. Charles MacKenzie 9:00 to 10:00 Session 2 Salon E-F 16:45 Behind the scenes of “Monsters inside me” – to be Students confirmed. 9:00 9 Determining Virulence Factors in Histomonas 19:00 - 21:00 Dinner The Terrace meleagridis. Elizabeth Carolyn Lynn*; Richard W. Gerhold; Larry R. McDougald; Robert Beckstead

Bayer Welcome Social 9:15 10 Molecular characterization of the Histomonas meleagridis complex. Lori Lollis

9:30 11 Altered Leukocyte Profiles in Turkey Poults Following a Primary Infection with Eimeria adenoeides. Ujvala Deepthi Gadde*; Hilary David Chapman; Thilakar Rathinam; Gisela F. Erf

9:45 12 Immunization of Northern Bobwhites With a Low Dose of Eimeria lettyae Provides Protection Against a High Dose Challenge. Richard W. Gerhold*; A. Lorraine Fuller; Larry R. McDougald

10:00 to 10:30 Pfizer coffee break Terras

25 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

10:30 to 12:00 Session 3 Salon D 11:30 23 Results of immunostimulation of dams with INF- Students gamma on the behavior of their pups congenitally infected with Toxoplasma gondii. David G. Goodwin*; Terry Hrubec; 10:30 13 Feasibility of Using Endoscopic Capsules in Dogs Anne M. Zajac; Jeannine Strobl; Bradley Klein; David S. to Detect Intestinal Helminths. Alice C.Y. Lee*; Norwood R. Lindsay Neumann; Michael A. Ulrich; Dawn E. Manthei; Dwight D. Bowman 11:45 24 Effect Of An Orange Oil Emulsion On Gastrointestinal Nematodes In Naturally Infected Sheep. 10:45 14 Optimization and validation of a real time PCR Casey Burke*; Joyce G. Foster; Anne M. Zajac assay for identification and quantification of Trichostrongyle nematodes in goats. Jennifer N. Towner*; Andrew R. 8:45 25 Dose titration effect of Sericea lespedeza feed Moorhea; Ray M. pellets on gastrointestinal nematode infection in lambs. James E. Miller*; Thomas H. Terrell; Joan M. Burke; Jorge A. 11:00 15 Replicate fecal egg counts and sensitive detection Mosjidis; Niki C. Whitley methods decrease variability and improve the accuracy of fecal egg count reduction tests in horses. Danielle E. Dimon; Ray M. Kaplan 12:00 to 14:00 Lunch break

11:15 16 Toward a Further Optimization of a Larva 14:00 to 15:30 Session 5 Salon D Migration Inhibition Assay for Detection of Resistance to Student / Equine Macrocyclic Lactone Drugs in Equine Cyathostomins. Daniel A. Zarate Rendon*; Ray M. Kaplan; Bob E. Storey 14:00 26 Toxicant-parasite interactions: the role of macroparasites in mercury dynamics within the 11:30 17 Development of an in vitro bioassay to detect gastrointestinal tract of mammalian hosts. Ashley Linton*; anthelmintic resistance in Dirofilaria immitis. Christopher Kimberlee B. Beckmen; Todd M. O'Hara; Mo D. Salman; Charles Evans*; Andy R Moorehead; Bob E. Storey; Mike T Lora R. Ballweber Dzimianski; Ray M. Kaplan 14:15..27 A Role for Collagenase in the Filariid-Wolbachia 11:45 18 Geospatial patterns of reported resistance of Mutualism: in situ Characterization of MMPs, W. pipientis, Dirofilaria immitis to macrocyclic lactone drugs in Louisiana. and Onchocerca volvulus. Michelle L. Gourley*; Rob Cassan Pulaski*; Jb Malone; TB Spencer; JA Fletcher; Jc Eversole; Charles Mackenzie McCarroll 14:30 28 A Survey of Parasites of Wild Caught Tokay 10:30 to 11:45 Session 4 Salon E-F Geckos (Gekko gecko) from Java, Indonesia. Roxanne A. Students Charles*; M.J. Yabsley; Angela E. Ellis; Ashley M. Rogers; Katherine F. Smith 10:30 19 Indirect fluorescent antibody (IFA) method for detection of antibodies against Hepatozoon americanum. 14:45 29 The Effect of Water Salinity on the Distribution and Kelly Allen*; Sanjay Kapil; Eileen M. Johnson; Susan E. Little Abundance of Snail Intermediate Host of Opisthorchis viverrini and Human Prevalence in Northeast Thailand. 10:45 20 Characterization of Diversity of Hepatozoon spp. Apiporn Suwannatrai*; Thitima Wongsaroj; John B. Malone; in Coyotes. Lindsay Starkey*; Roger Panciera; Kelsey Kulwadee Suwannatrai; Jutharat Kulsantiwong; Supawadee Paras; Misti West; Kelly Allen; Stephanie Heise; Michael Piratae; Chalida Thammasiri; Sattrachai Prasopdee; Panita Reiskind; Mason Reichard; Susan Little Khampoosa; Rasamee Suwanwerakamtorn; Pairat Tarbsripair; Thidarat Boonmars; Somsak Sukchan; Jennifer 11:00 21 Prevalence of antibodies to Trypanosoma cruzi, C. McCarroll; Smarn Tesana Toxoplasma gondii, Encephalitozoon cuniculi, Sarcocystis neurona, Besnoitia darlingi, and Neospora caninum in North 15:00 30 Effect of two different chemical formulations of a American opossums. Alice E. Houk*; David G. Goodwin; commercial disinfectant on the development of Parascaris Anne M. Zajac; Stephen C. Barr; J. P. Dubey; David S. equorum eggs. Jessica C. Gould*; Mary G. Rossano* Lindsay 15:15 31 Larvicidal Efficacy of Fenbendazole Against a 11:15 22 Cysticercus ovis infection and the Canadian sheep Macrocyclic Lactone-Resistant Isolate of Parascaris industry: an emerging problem. Bradley D. De Wolf*; Paula I. equorum. Craig R. Reinemeyer*; Julio C. Prado; Wendy E. Menzies; Andria Q. Jones; Andrew S. Peregrine; Jocelyn T. Vaala Jansen; Jennifer MacTavish

26 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

14:00 to 15:30 Session 6 Salon E-F 17:00 42 Seroprevalence of Anoplocephala perfoliata Companion animal ecto Infection Among Horses in West Coast States of the US. Patrick Meeus*; Cari Lagrow; Steven Kania; Craig 14:00 32 Comparative Efficacy of Advantage® Topical Reinemeyer; Vickie King; Sallie Cosgrove Solution (Imidacloprid) and Comfortis® Chewable Tablets (Spinosad) for the Control of Fleas on Dogs. Byron L. 17:15 43 Trypanosoma cruzi infection in dogs in south Blagburn*; Larry Cruthers; Jennifer Ketzis; Robert Arther; central Louisiana. Prixia del Mar Nieto; John Malone*

Wendell Davis; Iris Schroeder; Terry Settje; Jamie M. Butler 16:00 to 17:30 Session 8 Salon E-F 14:15 33 Inhibition of flea (Ctenocephalides felis) feeding on Wildlife dogs treated with dinotefuran, pyriproxfen, and permenthrin (Vectra 3D) based on quantitative polymerase chain reaction 16:00 44 Baylisascaris procyonis infection in raccoons and (qPCR) detection of a canine HMBS gene sequence. dogs on Prince Edward Island, Canada. Gary Conboy*; Chengming Wang; Bernhard Kaltenboeck; Jane D. Mount; Tonya A Stewart; Amanda Taylor Joy V. Bowles; Jamie M. Butler; Sherry Wilson; Sheila Gross; Cathy Ball; Byron L. Blagburn* 16:15 45 Distribution, prevalence and genetic characterization of Baylisascaris procyonis from selected 14:30 34 Efficacy of topically applied dinotefuran regions of Georgia and Florida. Emily L. Blizzard; Cheryl D. formulations and orally administered spinosad tablets against Davis; Scott Henke; David B. Long; Margaret Beck; Michael the KS1 flea strain infesting dogs. M. Dryden*; P Payne; V J. Yabsley* Smith; A McBride; D Ritchie 16:30 46 Prevalence of Baylisascaris procyonis in 14:45 35 Efficacy of flavored spinosad tablets administered Raccoons (Procyon lotor) from eastern Colorado. Deanna J. orally to dogs in a Simulated Home Environment (SHE), for Chavez; Ivy LeVan; Michael W. Miller; Lora R. Ballweber* the control of existing flea (Ctenocephalides felis) infestations. Daniel Snyder 16:45 47 What's Killing Our Deer? Investigation of Biting Fly Vectors of Epizootic Hemorrhagic Disease in Texas. Tracy 15:00 36 Rapid onset of action of spinosad against adult cat Cyr flea (Ctenocephalides felis) infestations on dogs. Daniel Snyder 17:00 48 White-Tailed Deer - Alternate Host for Babesia bovis? Patricia Holman 15:15 37` Insecticide Resistance Profiles of Field-Collected Isolates of Cat Fleas (Ctenocephalides felis). Michael K. 17:15 49 Identification, distribution and hosts of ticks in Rust*; Byron L. Blagburn; Iris Schroeder; Sarah Weston Kansas 2000 - 2007. M. Dryden*; P Payne.; V Smith.; A McBride.; M Hobson 15:30 to 16:00 Pfizer coffee break Terras 17:30 50 Effects of Different Burning Regimes on Tick 16:00 to 17:30 Session 7 Salon D Populations in Rangelands. Mason V. Reichard*; Kristen A. Parasite prevalence Baum

16:00 38 Geospatial Analysis and Ecological Niche 17:45 51 Genetic characterization of Toxoplasma gondii Modeling of Chagas Disease in Bolivia. Paula Mischler from Sand cats (Felis margarita). C. Rajendrana*; An Pasb; J. P. Dubeya; C. Suc 16:15 39 Prevalence of canine vector-borne diseases in heartworm-tolerant Jindo dogs. SungShik Shin*; Seok-Il Oh; 19:00 - 21:00 Dinner The Terrace DaeSung Oh; KyuSung Ahn; Kyung-Oh Cho

16:30 40 Seroprevalence of Borrelia burgdorferi, Merial Social Anaplasma phagocytophilum, Ehrlichia canis and Dirofilaria immitis among dogs in Canada. Alain Villeneuve*; Jonas K. Goring; Lynne Marcotte; Sebastien Overvelde

16:45 41 Seroprevalence of Heartworm, Toxoplasma gondii, FIV and FeLV Infections in Pet Cats in Bangkok and Suburban Area, Thailand. Woraporn Sukhumavasi*; Mary L. Bellosa; Araceli Lucio-Forster; Janice L. Liotta; Alice C.Y. Lee; Pitcha Pornmingmas; Sudchit Chungpivat; Leif Lorentzen; Dwight D. Bowman; J. P. Dubey

27 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

10:30 to 12:15 Session 11 Salon E-F Monday August 2nd, 2010 Protozoa

8:30 to 10:00 Session 9 Salon D 10:30 63 Isospora suis and its association with post- Education symposium weaning performance on three Ontario swine farms. Andrea Aliaga-Leyton; Robert M. Friendship; Cate Dewey; Cory 8:30 52 2009 AAVP Veterinary Parasitology Education Todd; Andrew S. Peregrine* Symposium. Gary Conboy; Susan Little; Karen F. Snowden; Tammi Krecek 10:45 64 Adaptive Evolution in Toxoplasma gondii ROP16 and ROP18 Genes Involved in Parasite Invasion. Hany M. 9:00 53 What is NAVMEC? Karen Snowden Elsheikha*; Miao-Miao Liu; Zi-Guo Yuan; Rui-Qing Lin; Xing- Quan Zhu 9:30 54 Survey of parasitology teaching at North American veterinary colleges: initial results from the 2009 AAVP/CAPC 11:00 65 Extracellular pH Governs the Entry and Education Symposium. Susan Little; Amy Edwards; Gary Phenotypic Plasticity of Neospora caninum. Hany Elsheikha Conboy; Karen F. Snowden 11:15 66 Characterization of giardin protein expression 10:00 to 10:30 Pfizer coffee break Terras during encystation of Giardia duodenalis. Mark C. Jenkins*; Celia O'Brien; Dumitru Macarasin; Jeffrey Karns; Monica 10:30 to 12:15 Session 10 Salon D Santin-Duran; Ronald Fayer Companion animal endo 11:30 67 Survival of Trypanosoma cruzi in Aca’ juice: 10:30 55 Evaluation of testing and treatment procedures for Implications for food borne Chagas disease outbreaks. heartworm (Dirofilaria immitis) in animal shelters in Georgia, David Lindsay South Carolina, and North Carolina. Andrew R. Moorhead*; David S. Boardman; Ruth D. Usher; Natalie D. Duncan; 11:45 68 Morphometrics of Assemblages of Giardia Maria T. Correa duodenalis Cysts from the Feces of Dogs and Cats. Dwight D. Bowman*; Stephanie B. Yager; Britta A. Okyere; Bo Li; 10:45 56 Genetic changes in Dirofilaria immitis populations Kyuhyung D. Kang; Heejeong Youn; Marissa Karpoff; Hussni possibly associated with exposure to macrocyclic lactones. O. Mohammed; Araceli Lucio-Forster; Janice L. Liotta Timothy G. Geary*; Catherine Bourguinat; Byron L. Blagburn; Kathy Keller; Rudolph Schenker; Roger K. Prichard 12:00 69 A new Besnoitia sp. f rom the southern plains woodrat (Neotoma micropus). J. P. Dubey*; M.J. Yabsley 11:00 57 Performance Comparison of a New, In-Clinic Method for the Detection of Canine Heartworm Antigen. 12:15 to 13:30 Lunch Alice C.Y. Lee*; Dwight D. Bowman; Araceli Lucio-Forster; Melissa J. Beal; Janice L. Liotta; Ray Dillon 13:30 to 15:45 Session 12 Salon D Interesting clinical cases 11:15 58 Efficacy of a Single Oral Administration of Milbemycin Oxime (Interceptor® Flavor Tabs® and Sentinel® 13:30 70 Equine abortion caused by Encephalitozoon Flavor Tabs®) Against Natural Infections of Ancylostoma cuniculi. Karen F. Snowden*; Travis Heskitt; Barbara braziliense in Dogs. Stephen Bienhoff Sheppard

11:30 59 The Efficacy of Milbemycin Oxime Against 13:45 71 Unusual Cause of Ectoparasitic Pruritus in a Migrating Pre-Adult Spirocerca lupi in Experimentally Infected Horse. Araceli Lucio-Forster*; Mary C. Smith; Ann Georgi Dogs. Dawid J. Kok*; Rudolph Schenker Leonard, Ithaca Artisan; Dwight D. Bowman

11:45 60 The Efficacy of Milbemycin Oxime to Protect Dogs 14:00 72 Aberrant migration of Dirofilaria immitis in three Against Infection with Spirocerca lupi (Nematoda: Spirurida) dogs - case report. Rhonda Pinckney*; Tara E Paterson in an Endemic Area. Dawid J. Kok; Rudolph Schenker 14:15 73 Autochtonal Dirofilaria repens in a dog in The 12:00 61 Attributes, Knowledge, Beliefs, and Behaviors Netherlands. Paul A.M. Overgaauw*; Evert P. Van Dijk Relating to Prevention of Heartworm Infection among Members of a National Hunting Dog Club. Sharron Patton*; 14:30 74 Uncommon Complications in a Dog with Dirofilaria Amanda Lutzy; Barton Rohrbach immitis. Jennifer E. Carter*; Guillaume Chanoit; Cheryl Kata

12:15 62 Obtaining a New Iisolate of Ancylostoma 14:45 75 Babesia canis rossi Infection in a Texas Dog braziliense without the Need of Necropsy. Janice L. Liotta*, Mason V. Reichard*; Robin W. Allison; Todd J. Yeagley Alice C.Y. Lee, Sarp Aksel, Ibrahim Alkhalife, Alejandro Cruz- Reyes, Heejeong Youn, Dwight D. Bowman

28 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

15:00 76 Hepatic alveolar echinococcosis in a dog in British 9:15 83 Multiple Anthelmintic Resistance on a Llama Farm Columbia, Canada. Andrew S. Peregrine*; Emily Jenkins; in the Southeastern United States. Bob E. Storey*; Sue B. Brian Barnes; Shannon Johnson; Janet Hill; Lydden Polley; Howell; Anand N. Vidyashankar; Lisa H. Williamson; Ray M. Ian K. Barker; Bruno Gottstein Kaplan

15:15 77 The use of ponazuril to treat a Toxoplasma gondii 8:00 to 9:30 Session 14 Salon E-F outbreak in a zoo setting. Jennifer A. Spencer*; Leah A. Helminth efficacy / diagnosis Kuhnt; John F. Roberts; Byron L. Blagburn; Bishop / impact Thompson 8:00 84 A Pilot Study on the Effect of an Integrated Control 15:30 78 Parasitologic Pet Peeves. Craig R. Reinemeyer*; Program of Fasciola hepatica in Cajamarca, Peru. Francisco Wendell L. Davis Raunelli*; Sergio Gonzalez; Jorge Guerrero

15:45 to 16:15 Pfizer coffee break Terras 8:15 85 Coprological Evaluation of Pour-on and Injectable Formulation of Moxidectin in Beef Cattle. J.G. Powell*; S.A. 16:15 to 18:00 AAVP Business Salon D Gunter; C.A. Tucker; J.L. Reynolds; Z.B. Johnson Meeting 8:30 86 The Effects of Cooperia punctata on cattle 16:15 PRESIDENTIAL ADDRESS. Lora Ballweber productivity. Bert E. Stromberg*; Louis C. Gasbarre; Audie Waite; David T. Bechtol; Michael S. Brown; Nicholas 16:45 AAVP BUSINESS MEETING Robinson; Erik Olson; Harold Newcomb

17:45 AAVP AWARDS / PICTURES 8:45 87 Modification and further evaluation of a fluorescein- All award recipients, including student travel grant labeled peanut agglutinin test for identification of awardees, and all students in the paper competition, Haemonchus contortus ova in fecal samples and findings of please plan to attend! note since the inception of offering the diagnostic test. Janell K. Bishop-Stewart*; ME Jurasek; JA Hall; JM O'Hara, O; J. Dinner on your own and school reunions at AVMA. Snyder; ML Kent

9:00 88 Comparison of a McMaster’s Chamber with increased detection sensitivity to the Stoll and Modified rd Wisconsin Fecal Egg Count Methods. B. Howell*; Bob E. Tuesday August 3 , 2010 Storey; Ray M. Kaplan

8:00 to 9:30 Session 13 Salon D 9:30 to 10:00 Pfizer coffee break Terras Anthelmintic efficacy / resistance 10:00 to 11:30 Session 15 Salon D President’s symposium 8:00 79 Pharmacological Characterization of New Cholinergic Anthelmintics in C. elegans. Timothy G. Geary*; 10:00 89 Molecular Mechanisms of Resistance : Nicotinic Charles Viau; Elizabeth Ruiz Lancheros; Abdel Francis; Tita and Macrocyclic Lactone Anthelmintics. Richard J. Martin*; N. Walter Alan P. Robertson

8:15 80 Anti-Parasitic Efficacy of Herb Extracts on Ovine, 10:30 90 Molecular mechanisms of resistance to Equine, and Canine Strongylid Eggs and Larvae. Heejeong benzimidazole and macrocyclic lactone anthelmintics. Roger Youn*; Kyonghee Kim; Yeongsuk Lim; Kyongeun Lee; Prichard Bongkyun Park; Janice L. Liotta; Clement Alawa; Araceli 10:30 91 Cattle Internal Parasitism and Deworming Lucio-Forster; Dwight D. Bowman Effectiveness from the 2008 USDA NAHMS Beef Cow/Calf Study. Bert E. Stromberg*; Louis C. Gasbarre; Lora R. 8:30 81 On the Mode of Action of Tribendimidine. Alan P. Ballweber; David A. Dargatz; Judy M. Rodriguez; Dante S. Robertson*; John A Carr; Sreekanth Puttachary; Santosh Zarlenga Pandey; Richard J. Martin 11:00 AM – MEETING ADJOURNS 9:00 82 Investigating Candidate Resistance Genes in an - SAFE TRAVELS Ivermectin-resistant Isolate of Haemonchus contortus. Sally M. Williamson*; Gerald C. Coles; Samantha McCavera; Adrian J. Wolstenholme

ABSTRACTS 2010

American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

ABSTRACTS Plenary Session 1 Shift in rules of engagement for eradication of cattle fever ticks in the United States Beto Perez de Leon1*, J. Mathews Pound1, Greta Schuster1, Felix D. Guerrero1, Ronald B. Davey1, Robert J. Miller1, Kimberly H. Lohmeyer1, David G. Hewitt2, J. Alfonso Ortega2, Tyler A. Campbell3, Alex E. Race- lis4, John A. Goolsby4, Patricia Holman5, Matthew Messenger6, Dee Ellis7, Roberta Duhaime8, Liza Soliz1, Andrew Y. Li1, Pamela L. Phillips1, Kevin B. Temeyer1, Pete Teel9, Stephen Wikel10, Diane M. Kammlah1, G. Gale Wagner5, Kevin P. Varner11, Danett K. Brake1 1Knipling-Bushland U.S. Livestock Insects Research Laboratory, USDA-ARS, Kerville, TX, 2Agronomy & Resource Sciences Department, Texas A&M, Kingsville, TX, 3National Wildlife Research Center, USDA- APHIS-WS, Kingsville, TX, 4Kika de la Garza Subtropical Agriculture Research Center, USDA-ARS, Wesla- co, TX, 5College of Veterinary Medicine and Biomedical Sciences, Texas A&M, College Station, TX, 6Cattle Fever Tick Eradication Program, USDA-APHIS, Riverdale, MD, 7Texas Animal Health Commission, Austin, TX, 8Cattle Fever Tick Eradication Program, USDA-APHIS, San Juan, TX, 9Entomology Department, Texas A&M, College Station, TX, 10Center for Biodefense and Emerging Infectious Diseases, University of Texas Medical Branch, Galveston, TX, 11USDA-APHIS, Austin, TX

With the exception of a systematic quarantine zone in south Texas along the border with Mexico, cattle fever ticks (CFT), i.e. Rhipicephalus (Boophilus) microplus and R. (B.) annulatus, were officially eradicated from the U.S. in 1943. Because of their roles as vectors of bovine babesiosis, the re-emergence of CFT is a real and imminent threat to the livestock industry. It is estimated that the livestock industry realizes annual savings totaling over 3 billion dollars at today’s currency rate since the U.S. was declared free of CFT and bovine babesiosis. The increased risk for bovine babesiosis in U.S. cattle associated with the apparent surge in CFT outbreaks observed since 2004 prompted a revision of the research agenda through a public workshop organized in 2009. Revised initiatives are underway to address knowledge gaps related to epidemiology and surveillance; ecology and biology of tick vectors and wildlife; diagnosis, treatment, and prevention; integrated approaches for CFT eradication; and anti-tick vaccines. Changes in agricultural practices as well as environmental and ecological conditions promoting the abundance of white-tailed deer (WTD) and free-ranging non-native ungulates help sustain CFT populations in south Texas even in the absence of cattle. Evidence based on serologic and molecular findings revealed the presence ofB . bovis-like organisms in WTD. These events represent serious complicating factors for eradication efforts. A critical assessment of traditional approaches is required to enable strategies for sustainable CFT eradication. For example, extensive surveillance on WTD and non-native ungulates throughout the year would enhance our understanding of CFT epidemiology along the U.S. – Mexico border. Similarly, partnerships with wildlife experts will facilitate the development of management practices to mitigate the risk for eradication failure associated with WTD as hosts for CFT. Current research efforts are expected to deliver the tools veterinary regulatory agencies need to ensure continued success with the mission to keep the U.S. free of CFT. Re- search supported in part by USDA-AFRI grant no. 10400992.

33 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

2 Clinically silent reemergence of Babesia equi infection in U. S. horses Donald Knowles Animal Disease Research Unit, United States Department of Agriculture, Agricultural Research Service, Pullman, WA

Equine babesiosis, also referred to as piroplasmosis is caused by tick-borne infection of equids by either Babesia equi or Babesia caballi. Infection with either parasite involves erythrocyte invasion, lysis and anemia. Babesia equi has also been referred to as Theileria equi, however this designation remains controver- sial. Subsequent to resolution of acute infection, both parasites persist in the equid host, possibly for life. Therefore persistently infected equid hosts represent a clinically silent transmission reservoir. A significant component of the B. caballi life cycle is that Dermacentor nitens, a competent vector for B. caballi has been shown to maintain infection across tick generations. The recent reemergence of B. equi infection in U. S. horses has increased interest and concern regarding equine babesiosis. Three questions receiving increased consideration due to the recent re-emergence of B. equi infection are: (1) why was clinical disease not detected in the acute phase in the majority of infected horses; (2) how did infection enter the U. S., and (3) is it possible to treat and remove transmission risk from infected horses?

3 The human filariasis elimination program - lessons learned for veterinary medicine Charles Mackenzie Dept. of Pathobiology and Diagnostic Investigation, Michigan State, East Lansing, MI

Abstract text not available.

Session 1 - Students 4 On the Tracks of Erythema Migrans – Lone Star Ticks, Bacteria, and Disease Stephanie R. Heise Veterinary Pathobiology, Oklahoma State University, Stillwater, OK

Amblyomma americanum, the lone star tick (LST) is the most commonly reported tick from humans in the southern United States and is associated with transmission of diseases, including STARI (southern tick-associated rash illness), a Lyme disease-like illness of people of unknown etiology, and ehrlichioses caused by Ehrlichia chaffeensis and Ehrlichia ewingii. To better define the microbial communities within A. americanum, colony-raised and wild-caught ticks, including those associated with a STARI case, were analyzed before and after feeding using a 16S rDNA-wide PCR approach followed by phylogenetic analysis of sequences; ticks were further evaluated by specific PCR for Borrelia spp., Rickettsia spp., and Ehrlichia spp. Work towards an animal model for STARI was also initiated. The data gained in these studies provides a thorough microbial characterization of the tick of interest, A. americanum. It provides support for previous work documenting frequent infection of LST with Rickettsia spp., all members of the spotted fever group, and confirms the preponderance of R. amblyommii in all life stages as well as the presence of Borrelia lonestari, another putative agent of STARI, in wild LST populations. Upon feeding, the bacterial community present in these ticks shifts significantly, with a dominant rise of the proportion of sequences associated with the genus Rickettsia. The development of an animal model provides a new prospective for the study of STARI. Taken together these studies work towards greater understanding of the etiology of STARI and provide valuable tools to be used in future research.

34 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

5 Status of Cytauxzoon felis in wild felid populations in the United States Barbara C. Shock1*, Staci M. Murphy2, Laura L. Patton3, Philip M. Shock4, Colleen Olfenbuttel5, Jeff Bering- er6, Suzanne Prange7, Daniel M. Grove8, Matthew Peek9, Mitchell Lockhart10, Jay Butfiloski11, Daymond W. Hughes12, Sarah Bevins13, Victor F. Nettles14, Holly M. Brown15, David S. Peterson16, M.J. Yabsley2 1Department of Infectious Diseases and the Southeastern Cooperative Wildlife Disease Study, Department of Population Health, College of Veterinary Medicine, University of Georgia, Athens, GA, 2University of Georgia, Athens, GA, 3Kentucky Department of Fish and Wildlife Resources, Frankfort, KY, 4West Virginia Division of Natural Resources, Charleston, WV, 5North Carolina Wildlife Resources Commission, Apex, NC, 6Missouri Department of Conservation, Columbia, MO, 7Ohio Department of Natural Resources, Athens, OH, 8North Dakota Game and Fish Department, Bismarck, ND, 9Kansas Department of Wildlife and Parks, Pratt, KS, 10Valdosta State University, Valdosta, GA, 11South Carolina Department of Natural Resources, Columbia, SC, 12USDA Wildlife Services, Athens, GA, 13Colorado State University, Fort Collins, CO, 14Uni- versity of Georgia, Athens, GA, 15Department of Pathology, University of Georgia, Athens, GA, 16Depart- ment of Infectious Diseases, University of Georgia, Athens, GA

Cytauxzoon felis, a protozoan parasite of wild and domestic felids, is the causative agent of cytauxzo- onosis in domestic and some exotic felids. C. felis is known to be transmitted by two ticks, Dermacentor variabilis and Amblyomma americanum, which have overlapping distributions throughout the Southern US; however, D. variabilis ranges further into northern states. Our objective was to determine the distribution and prevalence of C. felis in wild felid populations and to characterize the intraspecific variability. Fourteen states were included in the study (California, Colorado, Florida, Georgia, Kansas, Kentucky, Louisiana, Mis- souri, North Carolina, North Dakota, Ohio, Oklahoma, South Carolina, and West Virginia). Blood or spleen samples from hunter/trapper-killed felids (n=706) were tested for C. felis by PCR, targeting the ribosomal internal transcribed spacer regions 1 and 2 (ITS-1; ITS-2). We detected prevalence rates of 79% in Mis- souri (39 bobcats [Lynx rufus]), 65% in Oklahoma (20 bobcats), 63% in North Carolina (8 bobcats), 57% in South Carolina (7 bobcats), 55% in Kentucky (74 bobcats), 36% in Florida (45 bobcats), 33% in Louisi- ana (1 bobcat, 1 cougar [Puma concolor], 1 serval [Leptailurus serval]), and 31% in Kansas (39 bobcats). The prevalences were lower in Georgia (9%, 143 bobcats), North Dakota (2%, 172 bobcats, 5 cougars), California (0%, 26 bobcats), Colorado (0%, 67 bobcats) Ohio (0%, 19 bobcats), and West Virginia (0%, 37 bobcats). We also characterized the ITS-1 and ITS-2 genes and found greater intraspecific variability in wild felids compared to previous reports in domestic felids. These data indicate that C. felis is widespread and quite diverse in wild felid populations.

6 An Ongoing Study of the Effects of Prescribed Burns on Tick and Tick-borne Pathogen Prevalence Elizabeth R. Gleim1*, M.J. Yabsley2, Mike Conner3, Michael Levin4 1Wildlife, University of Georgia, Warnell School of Forestry & Natural Resources, Athens, GA 2University of Georgia, Athens, GA, 3Wildlife, Joseph W. Jones Ecological Research Center at Ichauway, Newton, GA, 4Vector-borne Diseases, Centers for Disease Control and Prevention, Atlanta, GA

Prescribed fire has become a common forest management tool, particularly in the southeastern United States. As land-use changes are being implicated in the spread and emergence of disease and vectors around the world, it is critical that we understand the impact that land management practices have on wildlife, domestic animal and human diseases and identify practices which may prevent the spread of disease. To better understand the effects of long-term prescribed fire on tick and tick-borne pathogen dynamics, twenty-one sites in southwestern Georgia with variable prescribed fire regimes were sampled for ticks which were subsequently tested for tick-borne pathogens. During initial field work in summer 2009 and winter and spring 2010, 212 Amblyomma americanum, 56 adult A. maculatum, 4,433 larval Amblyomma

35 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA sp., 21 adult Dermacentor variabilis, and 39 adult Ixodes sp. were collected. Fewer ticks were collected in burned sites compared to unburned sites. Furthermore, distinct differences in tick species composition were observed, with Amblyomma maculatum dominating burned sites, while A. americanum dominated unburned sites. These findings have highlighted a new public health threat in southwestern Georgia, asA. maculatum transmits the causative agent of Rickettsia parkeri rickettsiosis (Rickettsia parkeri). Molecular analysis of ticks for tick-borne pathogens (Ehrlichia, Borrelia, Hepatozoon and Rickettsia spp.) is underway.

7 Environmental contamination of public use areas with Toxocara spp. and Trichuris vulpis in Athens, GA Jessica H. Murdock1*, Joshua O. Cook1, Adrienne B. Zercher1, Dana Ambrose1, David Stallknecht2, Ray M. Kaplan1, Andrew R. Moorhead1 1Infectious Diseases, College of Veterinary Medicine, Athens, GA, 2Southeastern Wildlife Cooperative Dis- ease Study, College of Veterinary Medicine, Athens, GA

Intestinal parasites of cats and dogs (e.g. Toxocara spp.) in the Athens, Georgia area could represent a zoonotic risk to humans through contamination of the environment. The eggs of Toxocara spp. remain infective in soil for extended periods of time following fecal deposition. Incidental ingestion of Toxocara eggs by humans, in particular children, can result in ocular and visceral larva migrans. To determine the potential risk of human exposure in the greater Athens area, the level of environmental contamination must be established. We hypothesize that the prevalence of Toxocara spp. eggs in soil in Athens, GA will be similar to or higher than that documented in other regions. To answer this question, we examined soil samples using a novel technique for the presence of helminth eggs, specifically Toxocara spp., focusing on heavily-used public facilities (i.e. playgrounds, public parks, dog parks). Three soil samples will be collected from each of forty sites. Samples will be soaked for 12 hours in deionized water, then sieved (38µm screen size); the contents will be rinsed into a test tube, floated using centrifugation and supersaturated Sheather’s sugar solution (SG=1.27), and then examined using light microscopy. Soil particle size, substrate type (e.g. gravel, sand, tire shreds, etc.) relative exposure to sunlight, and intended use of each of the collection sites (e.g. dog park, playground) will be incorporated into the study as potential variables affecting distribution. Preliminary results indicate that Toxocara cati and Trichuris vulpis eggs are present at least one local playground and dog park, respectively. The data collected from this study will provide a baseline of the level of environmental contamination with Toxocara spp., and may elucidate factors important in the management and prevention of infection and disease in human populations.

8 Effect Of Exposure To Household Laundering And Drying On Eggs Of Common Canine Helminths Brian Neumann*, Courtney Mosley, Anne M. Zajac Biomedical Sciences and Pathobiology, Virginia Tech, Blacksburg, VA

A growing portion of the pet-owning public has become aware of the risks of infection with the common canine helminths Toxocara and Ancylostoma. Discussions with pet owners often lead to questions about the ability of environmental stages of these parasites to survive normal household conditions. This experiment was designed to investigate the effects on parasite survival of exposure to conditions in a household washing machine and dryer. Eggs of Toxocara canis were isolated by passing feces from a naturally infected puppy through a series of sieves. Eggs were then added to nylon mesh bags with a 10µm mesh opening. The bags were sealed with epoxy glue. Six bags of eggs were added to a full load of laundry and exposed to a standard hot/cold regular wash cycle with detergent in a household washing machine. Following the wash cycle, the clothes and 3 of the bags of eggs were transferred to a household dryer, which was run at a high setting until the clothing was dry (approximately 70 minutes). A third set of 3 bags served as an untreated

36 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA control. Following the experimental treatments, bags were opened and parasite eggs were recovered and placed in Petri dishes in a shallow layer of water. The eggs were kept at room temperature and examined daily for evidence of development. After 21 days an average of 97% of eggs in the untreated control group showed evidence of development. At the end of the same period, a mean of 92% of eggs added to the washer only showed comparable development to the control. In contrast, only 9% of eggs from the 3 bags exposed to the washer and dryer showed similar development. These preliminary results indicate that Toxo- cara eggs exposed to at least some normal dryer conditions will not survive.

Session 2 – Students 9 Determining Virulence Factors in Histomonas meleagridis Elizabeth Carolyn Lynn1*, Richard W. Gerhold2, Larry R. McDougald2, Robert Beckstead2 1Poultry Science, University of Georgia, Athens, GA, 2The University of Georgia, Athens, GA

Histomoniasis (blackhead disease) often causes high mortality in turkeys (50-100%) and morbidity in broiler breeder pullets. There is considerable variation in the severity of outbreaks. A national study of H. meleagridis strains is needed to find associations of variants with expression of virulence factors. By understanding the molecular basis of strain variation, virulence factors can be found and new targets for immunization and/ or treatment identified. Using sequence information for virulence genes found inEntamoeba histolytica and Trichomonas vaginalis we have generated degenerate PCR primers and amplified and cloned two cysteine proteases genes from H. meleagridis. Additionally, we are using RNA subtractive hybridization screens to identify differentially expressed genes found in virulent strains but not expressed in attenuated strains of Histomonas. Real-time PCR will be used to determine the expression level of these genes in virulent and attenuated strains. Putative virulence genes will also be expressed alone or in combination in attenuated strains of Histomonas and tested for their ability to confer virulence. To accomplish this, we are identifying the promoters of housekeeping genes in Histomonas through Splinkerette PCR technology and testing their ability to drive GFP expression in transfected Histomonads.

10 Molecular characterization of the Histomonas meleagridis complex Lori A. Lollis Poultry Science, University of Georgia, Athens, GA

Histomoniasis, caused by Histomonas meleagridis, is a major disease of gallinaceous birds and causes significant economic losses to the poultry industry. Although it is known that there are differences in virulence and tropism of H. meleagridis, genetic analysis of the parasite has not been conducted to determine if there are genotypic variations responsible for these differences. Histomonas DNA was extracted from paraffin embedded tissues of domestic poultry cases previously diagnosed as histomoniasis. The ITS1, 5.8S, and ITS2 rRNA regions were amplified by PCR, ligated into vector and cloned into competent E.coli. Resulting plasmids were sequenced using the forward and reverse plasmid primers. Nucleotide sequences of approximately thirty amplicons were compared to each other as well as a single H. meleagridis sequence, and other closely related protozoan sequences available from GenBank. The results of the sequence analysis suggest that there are at least two different genotypes within the H. meleagridis morphologic complex. One group is closely related to the H. meleagridis sequence available from GenBank, while the other group of our sequences has a higher nucleotide identity to Dientamoeba fragilis (70%) than to H. meleagridis (65%) and had at least thirty-eight nucleotide polymorphisms compared to the H. meleagridis sequence from GenBank.

37 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

11 Altered Leukocyte Profiles in Turkey Poults Following a Primary Infection withEimeria adenoeides Ujvala Deepthi Gadde*, Hilary David Chapman, Thilakar Rathinam, Gisela F. Erf Department of Poultry Science, University of Arkansas, Fayetteville, AR

Coccidiosis is a common disease of the intestinal tract of turkeys caused by Protozoan parasites of the genus Eimeria. Seven species have been described that infect the turkey and of these, E.adenoeides is one of the most pathogenic.Turkey poults at 20 d of age, given a primary inoculum of 12.5 × 103 oocysts, showed reduced weight gain, and the production of large numbers of oocysts in the feces, compared to uninfected poults. Poults were raised under conditions to prevent re-infection to determine the ability of the primary infection to confer protective immunity against a challenge infection of 5 × 104 oocysts given at 34 d of age. Using weight gain and oocyst production following challenge as criteria for protection, the results indicated that immunity had developed. The concentration and proportions among white blood cell (WBC) in periph- eral blood were determined at different times following the primary infection. The WBC concentration of infected poults was elevated on d 7 and 11, primarily due to elevated levels of lymphocytes and monocytes on d 7, and eosinophils on day 11. There were no differences in heterophil and basophil concentrations. With the exception of increased percentages of eosinophils on d 11, infection was not associated with alterations in the proportions among WBC populations. Comparison of CD4 and CD8 defined lymphocyte subpopula- tions in the blood revealed higher concentrations of CD4+ lymphocytes on d 11, lower concentrations of CD8+ cells on day 4, and higher concentrations of CD8+ cells on d 11, as well as elevated ratios of CD4+ to CD8+ lymphocytes on d 4 and 11 in infected compared to non-infected poults, respectively. These alterations in WBC profiles suggest initiation of both innate and adaptive cellular immune activities designed to effectively cope with a parasitic, intracellular pathogen.

12 Immunization of Northern Bobwhites With a Low Dose of Eimeria lettyae Provides Protec- tion Against a High Dose Challenge Richard W. Gerhold*, A. Lorraine Fuller, Larry R. McDougald The University of Georgia, Athens, GA

To determine if Northern Bobwhites (Colinus virginianus) can be immunized against Eimeria lettyae by a low dose inoculation of oocysts, we inoculated thirty birds each with either 100 or 1,000 oocysts within the first week of life. Four weeks following the immunization, the immunized birds were challenged with 1 X 106 oocysts of E. lettyae. Eight days following the challenge, birds were killed, weighed, and intestines examined for gross lesions. Parameters used to determine the effectiveness of the immunization during the challenge period included weight gain, severity of gross intestinal lesions, severity of diarrhea, feed to gain conversion ratio, and oocysts production compared to the unimmunized unchallenged as well as unimmunized challenged quail of the same age. Immunized birds gained an average of 33.3 gm (immunized with 100 oocysts) or 28.9 gm (immunized with 1,000 oocysts); whereas unimmunized challenged birds gained 11.5 gm. Immunized quail produced 99.7% fewer oocysts, contained minimal gross intestinal lesions, had minimal diarrhea, and had a 50% lower feed to gain conversion ratio compared to unimmunized challenged controls. Our findings indicate that immunization not only aids in suppression of weight loss, but also leads to a significant decrease in oocyst production. These findings indicate that vaccination is a viable option for controlling coccidiosis in quail and that research aimed at creating vaccine strains is warranted.

38 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Session 3 – Students 13 Feasibility of Using Endoscopic Capsules in Dogs to Detect Intestinal Helminths Alice C.Y. Lee1*, Norwood R. Neumann2, Michael A. Ulrich2, Dawn E. Manthei2, Dwight D. Bowman1 1Microbiology & Immunology, Cornell University, Ithaca, NY, 2Cheri-Hill Kennel & Supply Inc., Stanwood, MI

Modern technology has introduced more efficient and less invasive ways of conducting animal research. In the parasiticide industry, however, the gold standard for determining anthelmintic efficacy is still necropsy. A previous study showed that conventional gastrointestinal (GI) endoscopy did not provide accurate counts of intestinal helminths, largely because the middle small intestine could not be viewed. Endoscopic capsules (wireless, ingestible cameras) are used in people to provide more comprehensive intestinal examination. A preliminary study was undertaken to investigate the safety and utility of capsule endoscopy in dogs.

Five clinically healthy dogs of varying age (mean: 1.5 years, range: 0.4-3), weight (mean: 27.1 kg, range: 9.5-57.7), breed (Great Dane, Lab, pitbull, hound, and beagle), and gender (3 male, 2 female) were recruit- ed. After a 12-hour fast, a data recorder and sensors were secured to the dog with a jacket. The endoscopic capsule was administered orally. Equipment was removed 5-8 hours later, and images were reviewed.

Dogs exhibited no pain or distress during the procedure. Images were captured along the entire small bowel as well as portions of esophagus, stomach, and colon. Findings included hair, foreign bodies, erosions, and luminal parasites. Mean gastric and small bowel transit times (TT) were 1.8 hours (range: 0.2-3.6) and 1.8 hours (range: 0.2-2.5), respectively. The capsule passed in the stool within 24 hours.

Capsule endoscopy was well tolerated by the 5 dogs. Captured images were of sufficient quality to detect roundworms, hookworms, and tapeworms, as well as associated mucosal lesions. Given the small study population, it is not possible to draw definite conclusions about the effects of age, size, gender, or breed on TT. Preliminary results show that capsule endoscopy is a safe and minimally invasive imaging method that allows for the detection and identification of intestinal helminths in dogs.

14 Optimization and validation of a real time PCR assay for identification and quantification of Trichostrongyle nematodes in goats Jennifer N. Towner*, Andrew R. Moorhead, Ray M. Kaplan Infectious Diseases, University of Georgia College of Veterinary Medicine, Athens, GA

Gastrointestinal nematodes of the family Trichostrongylidae are important pathogens of ruminants with significant economic impact worldwide. Improved diagnostic techniques are needed to rapidly differentiate among common trichostrongyle species. Due to morphological similarities among trichostrongyle eggs, species differentiation is only possible through identification of larvae isolated from coprocultures. This technique is time consuming, taking up to 14 days to process, and requires experienced technicians for accuracy. Recent studies demonstrate that molecular techniques are more reliable and focus on using Real Time PCR as a means for rapidly differentiating and semi-quantifying trichostrongyle species. The aim of the present study is to optimize a Real Time PCR assay in order to identify and quantify the eggs of the three major trichostrongyle species of goats within a fecal sample. DNA from eggs as well as plasmid DNA containing the ribosomal ITS-2 (Second Internal Transcribed Spacer) region of Haemonchus contortus, Trichostrongylus colubriformis, and Teladorsagia circumcincta were used to evaluate the specificity and detection range of this assay. Samples of pure and mixed populations were used to access the detection of various percentages of each species present. Threshold Cycle (Ct) values obtained from amplification were

39 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA compared among the different percentages tested. Using this assay, we were able to estimate the relative percentage of each species present within a mixed fecal sample. Using a total of 800 eggs, we were able to demonstrate linear quantification (R²=0.93) ofHaemonchus contortus eggs from a range of 1-800. We also demonstrated linear quantification (R²=0.88) of Trichostrongylus colubriformis eggs from a range of 1-800. Within a mixed sample, as little as 0.1% of each species could be detected. This study demonstrates that Real Time PCR assay can be used to rapidly identify and quantify DNA of common trichostrongyle species at a wide detectable range.

15 Replicate fecal egg counts and sensitive detection methods decrease variability and improve the accuracy of fecal egg count reduction tests in horses Danielle E. Dimon*, Ray M. Kaplan Infectious Diseases, University of Georgia, Athens, GA

Fecal egg count reduction test (FECRT) is the current gold standard for diagnosing anthelmintic resistance on horse farms. However, variability in the measurement of fecal egg count (FEC) yields a wide spectrum of results within individual horses, which ultimately impacts the accuracy of the inference made regarding resistance. This problem is magnified when small numbers of horses are tested; a common situation on horse farms. The primary objective of this study was to investigate the impact of using 3 replicate FEC both before and after treatment on the consistency of the results. In addition we wanted to compare a 2-chamber McMaster (0.3 ml volume) with a large volume 3-chamber McMaster (0.9 ml), which have detection levels of 25 eggs per gram (epg) and 8.3 epg, respectively. FECRT were performed using ten adult horses on 2 separate farms. All horses were treated with pyrantel pamoate, and feces were collected on the day of, and 14 days after treatment. Three replicate FEC were performed using each method. Results demonstrate that variability in single FEC is large and can lead to erroneous conclusions when comparing pre and post-treatment FEC. Performing replicate FEC greatly reduced level of variability and improved accuracy. In addition, analysis using a Poisson regression with random effects model demonstrated a significant difference (p<0.0025) between mean FEC of the 2- and 3-chambered slides, indicating improved accuracy with the more sensitive method. Differences between the 2 slide types were magnified when FEC were low, which is almost always the case post-treatment. In several horses the 2-chamber McMaster yielded 100% reduction, but with the 3-chambered slide no horses ever demonstrated 100% reduction. Measurement replication is a standard practice in biological assays, and the results of this study clearly indicate that when conducting a FECRT, repeating FEC should be performed as a standard practice.

16 Toward a Further Optimization of a Larva Migration Inhibition Assay for Detection of Re- sistance to Macrocyclic Lactone Drugs in Equine Cyathostomins Daniel A. Zarate Rendon*, Ray M. Kaplan, Bob E. Storey Infectious Diseases, University of Georgia, Athens, GA

Currently there are no accurate and validated in vitro tests for detection of resistance to macrocyclic lactone anthelmintics in cyathostomin parasites of horses. Among the in vitro assays developed, previous studies suggest that the larval migration inhibition assay (LMIA) may be a promising option. However, more research is needed before the assay is validated and ready for use at the farm level. A major impediment to the successful optimization of this assay in cyathostomins is inconsistency in the level of motility of the L3, which causes variability in the rate of migration during the assay. This in turn can strongly influence the dose-response measured to the drug. Thus, it is important to investigate means of improving the consistency of the migration in the untreated control L3. Working with a pool of cyathostomin L3 we tested the effect of three different incubation media (PBS, RPMI, Earle’s solution) on the motility of L3 using a subjective scale (1 -3). No significant differences were observed between the different media (p>0.05)

40 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA and, in general, a very low level of motility was observed after 2 hrs. of incubation. To further investigate other possible parameters that might play a role in the level of motility displayed, we also examined type of incubation plate, level of media in well and length of incubation. In all cases, no significant differences were found. Another important assay parameter is the size of the mesh used, therefore we tested several different mesh sizes (20, 25 and 30 um) during a two hour migration. There were no significant differences on migration (p>0.05). Despite the problems encountered with the LMIA, the experience we have gained in the LMIA provide us with optimism that this assay has promise for detecting resistance in cyathostomins.

17 Development of an in vitro bioassay to detect anthelmintic resistance in Dirofilaria immitis Christopher Charles Evans*, Andy R Moorehead, Bob E. Storey, Mike T Dzimianski, Ray M. Kaplan Department of Infectious Diseases, University of Georgia, Athens, GA

The canine heartworm (Dirofilaria immitis) is a parasite of significant veterinary importance, causing disease in dogs, cats, and other animals. Since being introduced more than 25 years ago, macrocyclic lactones have been widely used in prophylactic therapy to prevent heartworm disease. However, increasing reports of adult heartworm infections in dogs, where the administration of monthly prophylaxis is documented, indicate increasing failure of prophylaxis. This suggests that there is either an increase in the frequency with which owners fail to properly administer the drugs, or resistance to the drugs has developed in the worms. Objectively verifying the former situation is not possible, and no assays are currently validated for assess- ing the latter. To address this important issue, we focused on developing an assay that uses the third-stage infective larvae (L3) as a target, since this is a stage targeted by the prophylactic treatments. Macrocyclic lactones are believed to act on worms by a mechanism that induces paralysis, therefore, we developed a larval migration inhibition assay (LMIA) that directly measures this phenotype. The LMIA involves incu- bating L3 in vitro with a range of drug concentrations, after which the ability of L3 to migrate through a fine mesh is determined microscopically and quantified. Data is then analyzed to measure the dose-response. Significant shifts in the dose-response in clinical isolates suspected of being resistant as compared to the known susceptible isolates are indicative of anthelmintic resistance. Performing this assay with a laboratory strain of D. immitis using eprinomectin as a representative macrocyclic lactone has yielded a reproducible sigmoidal dose-response with approximate EC50 and EC95 of 11µM and 27µM, respectively. The goal of this study is to compare parasite isolates from cases of suspected prophylactic failure using the LMIA as a parameter for drug susceptibility, allowing insight into the nature of drug resistance in heartworms.

18 Geospatial patterns of reported resistance of Dirofilaria immitis to macrocyclic lactone drugs in Louisiana Cassan Pulaski1*, Jb Malone2, TB Spencer3, JA Fletcher4, Jc McCarroll5 1LSU School of Veterinary Medicine, Class of 2012, Baton Rouge, LA, 2Pathobiological Sciences, School of Veterinary Medicine, Baton Rouge, LA, 3Avoyelles Animal Clinic, Mansura, LA, 4Veterinary Clinic of Avoy- elles, Inc, Marksville, LA, 5Pathobiological Sciences, Louisiana State University, Baton Rouge, LA

Strains of Dirofilaria immitis resistant to currently used preventive drugs are reported to be emerging in spotty locations in dogs by practicing veterinarians in Louisiana. If not controlled in the early stages, resistance threatens to become a widespread problem in the US that may limit the effectiveness of current preventive drug treatment methods. To validate these practice reports, a statewide survey of veterinarians was done to define the extent of the problem and identify focal ‘hotspots’ of reported macrocyclic lactone (ML) lack of efficacy (LOE). Geographic information systems (GIS) methods were used to develop statewide and community-level risk maps for one parish (Avoyelles) based on practitioner-reported cases of preventive drug LOE and to define clustering, time-space patterns or other geospatial evidence of focal resistance of D. immitis to ML drugs. Statewide survey results (25.8% response rate; 221/855) indicated there are focal

41 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA locations with high rates of reported LOE of ML drugs against heartworms and that numbers increased from 2005-2008. Geospatial analysis using kernel density and space-time permutation modeling software revealed focal ‘hotspots’ of population-adjusted cases of preventive drug failure in Avoyelles Parish in 2007, 2008 and 2009. Buffers zones of 0.5 km radius were created centered on case locations (to simulate flight range of vector mosquitos). Between 2007-2008, 40% of buffer zones intersected and 53% of buffers intersected between 2008-2009, suggesting potential dog-to-dog transmission of resistant heartworms between neighboring dogs. Further study is needed on focal geospatial dynamics and isolation of resistant strains by 1) in vitro drug assays on microfilariae and infective larvae and 2)in vivo experimental infection of dogs with D. immitis isolates from focal areas with high lack of efficacy rates.

Session 4 – Students 19 Indirect fluorescent antibody (IFA) method for detection of antibodies againstHepato - zoon americanum Kelly Allen1*, Sanjay Kapil2, Eileen M. Johnson1, Susan E. Little1 1Veterinary Pathobiology, Oklahoma State University, Stillwater, OK, 2Animal Disease Diagnostic Lab, Okla- homa State University, Stillwater, OK

Hepatozoon americanum is a tick-borne apicomplexan parasite that causes a debilitating, sometimes fatal, disease in dogs referred to as American canine hepatozoonosis (ACH). Diagnosis of ACH can be difficult, as the parasite is rarely observed in blood smears, detection by PCR is not always possible due to fluctuat- ing levels of organism in circulation, and muscle biopsy to look for parasite stages or associated lesions, although reliable and sensitive, is invasive. To avoid these complications, we developed an indirect fluorescent antibody (IFA) assay using sporozoites isolated from experimentally infected Amblyomma maculatum ticks. Serum from experimentally infected dogs in which infection was confirmed by muscle biopsy and PCR was used as positive control; serum samples from uninfected dogs and from dogs with antibodies reactive to Toxoplasma gondii, Neospora caninum, Babesia gibsoni, and B. canis were included as negative controls. Initial screening of dogs from endemic areas (n=48) and coyotes with documented H. americanum infections (n=12) revealed antibodies reactive to sporozoites in 2.1% and 33%, respectively. Several (n=9) coyotes naturally infected with H. americanum as confirmed by muscle biopsy and/or PCR did not have antibodies reactive to sporozoites in this assay. The route of infection or strain of parasite may influence development of antibodies to H. americanum, issues which should be taken into account in developing serologic tests for this parasite.

20 Characterization of Diversity of Hepatozoon spp. in Coyotes Lindsay Starkey*, Roger Panciera, Kelsey Paras, Misti West, Kelly Allen, Stephanie Heise, Michael Reis- kind, Mason Reichard, Susan Little Oklahoma State University, Stillwater, OK

In the southern United States, infection of domestic and wild canids with Hepatozoon americanum can lead to severe disease characterized by fever, myalgia, lethargy, and periosteal proliferation; Hepatozoon canis has also been described in canids from this area. To better define the species and strains of Hepatozoon spp. infecting coyotes in the southern U.S., whole blood and muscle samples collected from 44 coyotes at 8 different locations were evaluated by a nested PCR using primers amplifying a variable region of the apicomplexan 18S rRNA gene as well as histopathology (muscle only) for presence of tissue cysts. Hepa- tozoon spp. infection was identified in 79.5% (35/44) of coyotes tested, including 28 of 44 (63.6%) whole blood samples and 28 of 44 (63.6%) muscle samples tested by PCR, and 23 of 44 (52.3%) muscle samples

42 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA evaluated by histological examination. Positive coyotes were more likely to be identified in areas with A. maculatum ticks (35/39; 89.7%) than areas without (0/5; 0%). Preliminary sequence analysis revealed 14 different sequence types comprising two major clusters of Hepatozoon spp.: one most closely related to H. americanum and a second intermediate between H. americanum and H. canis. Characterization of the Hepatozoon spp. infecting coyotes will help us better understand this parasite in wild canid populations and serves as a model of infection dynamics in domestic dogs that are not managed with acaricides or prevented from scavenging as a part of their diet.

21 Prevalence of antibodies to Trypanosoma cruzi, Toxoplasma gondii, Encephalitozoon cuniculi, Sarcocystis neurona, Besnoitia darlingi, and Neospora caninum in North American opossums Alice E. Houk1*, David G. Goodwin1, Anne M. Zajac1, Stephen C. Barr2, J. P. Dubey3, David S. Lindsay1 1Biomedical Sciences and Pathobiology, Virginia Tech, Blacksburg, VA, 2Clinical Sciences, Cornell Univer- sity, Ithaca, NY, 3Animal Parasitic Disease Laboratory, United States Department of Agriculture, Beltsville, USA, Beltsville, MD

We examined the prevalence of antibodies to zoonotic protozoan parasites (Trypanosoma cruzi, Toxoplas- ma gondii, and Encephalitozoon cuniculi) and protozoan+s of veterinary importance (Neospora caninum, Sarcocystis neurona and Besnoitia darlingi) in a population of North American opossums (Didelphis virgin- iana) from Louisiana. Samples from 30 opossums were collected as part of a survey for T. cruzi in Louisi- ana opossums. Frozen sera from these 30 opossums were examined using an indirect immunofluorescent antibody test (IFAT) against in vitro produced antigenic stages of these protozoans. Additionally, 24 of the 30 samples were examined using haemoculture and all 30 were examined in the modified direct agglutination test (MAT) for antibodies to T. gondii. The prevalence of reactive IFAT samples were as follows: 60% for T. cruzi, 27% for To. gondii, 23% for E. cuniculi, 17% for S. neurona, 47% for B. darlingi, and 0% for N. caninum. Hemaculture revealed that 16 (67%) of 24 samples were positive for T. cruzi compared to 18 of 30 (60%) by IFAT. The sensitivity and specificity for the IFAT compared to hemaculture was 100% for each. The modified direct agglutination test revealed that 9 (30%) the 30 samples from opossums had antibodies to T. gondii compared to 8 (27%) using the IFAT. The sensitivity and specificity of the IFAT compared to the MAT was 100% and 72%, respectively.

22 Cysticercus ovis infection and the Canadian sheep industry: an emerging problem Bradley D. De Wolf1*, Paula I. Menzies2, Andria Q. Jones2, Andrew S. Peregrine2, Jocelyn T. Jansen2, Jen- nifer MacTavish3 1Population Medicine, Ontario Veterinary College, University of Guelph, Guelph, ON, 2Ontario Veterinary College, University of Guelph, Guelph, ON, 3Canadian Sheep Federation, Guelph, ON

Cysticercus ovis is the intermediate stage of a canine tapeworm, Taenia ovis, which produces cystic lesions in the skeletal and cardiac muscle of sheep. If numerous, these lesions can result in condemnation of the entire carcass. In 2008, an outbreak of C. ovis occurred in animals that originated from a feedlot in southwestern Ontario. The feedlot handles 25 000 animals per year that originate from multiple farms in multiple Canadian provinces and the United States. Between February and May 2008, 12% of all animals marketed from this feedlot were condemned due to C. ovis. Confirmation of C. ovis was determined based on microscopic examination of the muscle lesions and dimensions of the parasite’s rostellar hooks. Historically, while the number of lambs slaughtered in Ontario abattoirs remained constant between 2003 and 2008, the proportion of all carcasses condemned due to C. ovis increased from 0% to 50%, respectively. The rise in carcass condemnations due to C. ovis suggests that the prevalence of this infection on Canadian sheep farms is increasing. Until recently, it was not possible to trace a condemned lamb carcass in Ontario back

43 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA to the animal’s farm of origin. Through development of the Canadian Sheep Identification Program (CSIP), and direct intervention from the Canadian Sheep Federation, it has been possible to address this problem. Research has shown that in 2009, lambs condemned in Ontario due to C. ovis originated from the Cana- dian provinces of Manitoba, Alberta, Ontario, Quebec and Saskatchewan; 85% of the total Canadian sheep flock resides within the latter four provinces. The data suggest that, in recent years, C. ovis has spread across Canada and that endemic transmission through domestic dogs and/or wild canids is now occurring in multiple provinces. Within Ontario, the data indicate that endemic transmission occurred for the first time in 2009.

23 Results of immunostimulation of dams with INF-gamma on the behavior of their pups congenitally infected with Toxoplasma gondii. David G. Goodwin1*, Terry Hrubec2, Anne M. Zajac1, Jeannine Strobl2, Bradley Klein3, David S. Lindsay1 1Biomedical Sciences and Pathobiology, Virginia Tech, Blacksburg, VA, 2Edward Via Virginia College of Osteopathic Medicine, Blacksburg, VA, 3Virginia Tech, Blacksburg, VA

Toxoplasma gondii can cause congenital infection in humans. Approximately 85% percent of women in the U.S population are at risk of delivering offspring congenitally infected with T. gondii. Pregnancy brings about a state of immunosuppression making women more susceptible to infectious agents. It is hypothesized that administering immune stimulation to pregnant individuals will decrease the deleterious behavioral effects caused by T. gondii. Chronic infection with T. gondii has been shown to result in decreased motor function, increased open field activity and decreased memory in mice. Our experiments used immune stimulation of the pregnant dam with interferon-gamma (INF-g) prior to inoculation with T. gondii. The Barnes maze test is used to look at spatial memory, rate of learning, memory acquisition, and open field activity. We used 4 treatment groups; 1) uninfected not given INF-g or T. gondii, 2) immune stimulated with INF-g not given T. gondii, 3) immune stimulated with INF-g and T. gondii infected, and 4) T. gondii infected not given INF-g. One week after weaning (4 weeks of age), the pups were tested using the Barnes maze test. Mice of the same sex and litter were tested again at 8 weeks of age using the Barnes maze test. At 4 weeks of age T. gondii infected mice and control mice, males and females showed no difference (P >0.1) in 7 of 8 maze parameters. There was a difference in memory retention with the T. gondii infected male mice showing a decreased in memory retention. At 8 weeks of age male and female mice had differences (P < 0.1) in 4 of 8 parameters measured. Immune stimulation prior to infection appears to have little influence in correcting altered behavior in congenitally infected mice.

24 Effect Of An Orange Oil Emulsion On Gastrointestinal Nematodes In Naturally Infected Sheep Casey Burke1*, Joyce G. Foster2, Anne M. Zajac1 1Biomedical Sciences and Pathobiology, Virginia Tech, Blacksburg, VA, 2Appalachian Farming Systmes Research Center, USDA, ARS, Beaver, WV

Increasing levels of anthelmintic resistance in ovine gastrointestinal strongylids, especially Haemonchus contortus have led many investigators worldwide to examine potential anthelmintic effects of naturally occurring plant products. In previous work, we have shown that 1200 mg/kg of an orange oil emulsion given daily for 5 days showed 87.8% efficacy against H. contortus in a gerbil model. In sheep experimentally

44 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA infected with H. contortus a single treatment of a modified orange oil emulsion reduced fecal egg counts by 97%. Our objective in this study was to examine the effects of the orange oil emulsion on natural strongylid infection in sheep grazing contaminated pasture.

Twelve mixed breed weaned lambs were exposed to naturally infected pasture at the Virginia Tech Sheep Center. Five days before treatment, the lambs were housed and strongylid fecal egg counts determined (Modified McMaster’s test). Sheep were allocated to 2 groups balanced for fecal egg count. On Day 0, 600 mg/kg of orange oil emulsion was administered orally to sheep in one of the groups. Rectal fecal samples were collected on Days 1, 2 and 4 post treatment. On Day 6, sheep were euthanized and abomasal and intestinal samples collected for determination of parasite numbers and species. Four days after treatment the mean fecal egg counts were 292 and 3425 in the treated and untreated lambs, respectively. Mean total worm burden in the treated animals was 7429 compared to 15,588 in untreated animals. The difference between worm burdens of treated and untreated animals was greater in abomasal worm counts than in intestinal counts. These results suggest that citrus oils should be further explored for activity against ovine gastrointestinal nematodes.

25 Dose titration effect of Sericea lespedeza feed pellets on gastrointestinal nematode infection in lambs James E. Miller1*, Thomas H. Terrell2, Joan M. Burke3, Jorge A. Mosjidis4, Niki C. Whitley5 1Pathobiological Sciences, Louisiana State University, Baton Rouge, LA, 2Agricultural Research Station, Fort Valley State University, Fort Valley, GA, 3Dale Bumpers Small Farms Research Center, USDA, ARS, Booneville, AR, 4Agronomy and Soils, Auburn University, Auburn, AL, 5Agricultural and Environmental Sci- ences, North Carolina A&T University, Greensboro, NC

Twenty-two 7-8 month old naturally infected lambs were removed from pasture and randomly allocated, based on FEC and breed, to 4 treatment groups. Lambs were confined indoors in individual pens. Group 1 (n = 6) was fed a commercial pelleted feed. Groups 2 (n = 6), 3 (n = 5) and 4 (n = 5) were fed a 25%, 50% and 75% sericea lespedeza leaf meal (SLLM) pelleted feed, respectively, at 3.5% of body weight for 6 weeks. All rations were balanced. Feces and blood were collected weekly to monitor infection level based on FEC and PCV. Weights were taken at the beginning and at the end of the study. During the last week of SLLM feeding, feces from each group were cultured and recovered L3 were used to infect worm free lambs to evaluate viability after exposure to SLLM condensed tannin in the feces. At the start of the study, FEC for all groups was similar. Group 1 FEC remained high throughout the study. By week 2, FEC was reduced by 85.0%, 61.1% and 76.6% for Groups 2, 3 and 4, respectively. Group 2 and 4 FEC remained low for the duration of the study, but Group 3 FEC steadily increased and was similar to Group 1 at the end of the study. Weight gain for the 4 groups was 7.0, 6.1, 5.4 and 4.4 kg, respectively. Results indicate that all 3 dose levels of SLLM pellets reduced FEC, but the 50% dose level reduction was not maintained. As SLLM dose increased, weight gain decreased. Infection of worm free lambs was highest to lowest with L3 from Groups 1 to 3 (not enough L3 were recovered from Group 4), respectively. Supplement feeding SLLM at 25% of total intake may be useful to reduce FEC and reinfection in grazing lambs.

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Session 5 - Student / Equine 26 Toxicant-parasite interactions: the role of macroparasites in mercury dynamics within the gastrointestinal tract of mammalian hosts Ashley Linton1*, Kimberlee B. Beckmen2, Todd M. O’Hara3, Mo D. Salman4, Lora R. Ballweber4 1Colorado State University, Fort Collins, CO, 2Alaska Department of Fish & Game, Fairbanks, AK, 3Univer- sity of Alaska-Fairbanks, Fairbanks, AK, 4Colorado State University, Fort Collins, CO

Certain intestinal macroparasites have been shown to bioaccumulate heavy metals (e.g. Pb, Cd) at significantly higher concentrations than that of their fish hosts. We, therefore, hypothesize that toxicant-parasite interactions within the host intestine may have a positive effect on overall host-health. Thus, the objective of this ongoing study is to assess the ability of gastrointestinal macroparasites to bioaccumulate mercury, and to determine their role in mercury distribution and biotransformation within the host. Intestinal tracts were processed from Alaskan Gray wolves (Canis lupus). Macroparasites were removed and weighed, and nematodes were enumerated; additionally, host luminal contents and various tissue samples were collected for total mercury (THg) analysis. Prevalence of cestodes and ascarids in the 89 intestinal tracts examined was 61.8% (55/89) and 19.1% (17/89), respectively. Nine wolves contained both cestodes and ascarids, out of 63 parasitized animals (14.3%). Ascarids from 15 of the 17 animals were identified morphologically, and prevalence of Toxocara canis and Toxascaris leonina was found to be 33.3% (5/15) and 80.0% (12/15), respectively. Two individuals were co-infected with both Toxocara canis and Toxascaris leonina. All cestodes were of the genus Taenia. Preliminary THg results showed concentrations in pooled, homogenized cestodes ranging from 2.95 to 75.03 ppb (ww), with a median of 12.98 ppb (ww). Nematode THg concentrations ranged from 3.34 to 6.32 ppb ww, with a median of 4.72 ppb (ww), possibly suggesting a greater potential for uptake by the cestodes. Initial results confirm that these parasites are capable of mercury uptake, and that THg concentrations in these parasites lie within a detectable range. These data will be of critical importance as we move forward in addressing the role of macroparasites in mercury distribution and biotransformation within the host.

27 A Role for Collagenase in the Filariid-Wolbachia Mutualism: in situ Characterization of MMPs, W. pipientis, and Onchocerca volvulus. Michelle L. Gourley1*, Rob Eversole1, Charles Mackenzie2 1Dept. of Biological Sciences, Western Michigan University, Kalamazoo, MI, 2Dept. of Pathobiology and Diagnostic Investigation, Michigan State, East Lansing, MI

Parasitic nematodes that cause canine and feline heartworm (Dirofilaria immitis),as well as bovine and human filariasis Onchocerca( spp.) harbor the bacterial mutualist Wolbachia pipientis. Antibiotic elimination of the bacteria arrests worm reproduction and molting. Wolbachia have therefore been slated as novel drug targets for the treatment of veterinary and human filariasis. However, the cellular mechanisms that facilitate the worm-bacteria mutualism have not been sufficiently characterized. Specifically, no proteins have been identified which could account forWolbachia ’s ability to penitrate multiple host tissue layers or contribute to worm embryogenesis and molting. To address this issue, we conducted an in situ study of W. pipientis and collagenase. Collagenases are extracellular matrix degrading enzymes common to bacterial pathogens and required for eukaryotic tissue remodeling. We used immunocytochemistry to examine W. pipientis and Onchocerca volvulus within nodules collected from 40 Ecuadorian patients. Collagenases MMP-2 and MMP-9 were present in worm intestinal wall, mature spermatocytes, and tissues commonly harboring W. pipientis (body wall, oocytes, embryos throughout development, and microfilaria). Collagenase staining was stronger in body wall regions where Wsp signal was localized, and was only punctate when coinciding

46 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA with Wsp signal in consecutive tissue sections. Wsp staining was diffuse in unfertilized oocytes, but punctate and localized to the posterior pole of many-cell stage embryos. Wsp signal was frequently constrained to single embryonic germ cells, suggesting control over bacterial distribution during germ cell division. The colocalization of W. pipientis with collagenase in O. volvulus tissue is noteworthy because it places the bacteria within cellular proximity to an extracellular matrix degrading enzyme. This enzymatic association could facilitate bacterial movement through nematode tissues and contribute to tissue remodeling during oogenesis and embryogenesis. These results offer one of the first molecular mechanisms for bacterial activity in the Wolbachia-filariid mutualism.

28 A Survey of Parasites of Wild Caught Tokay Geckos (Gekko gecko) from Java, Indonesia Roxanne A. Charles1*, M.J. Yabsley2, Angela E. Ellis3, Ashley M. Rogers4, Katherine F. Smith5 1Department of Population Health, Southeastern Cooperative Wildlife Disease Study, College of Veterinary Medicine, University of Georgia, Athens, GA, 2Southeastern Cooperative Wildlife Disease Study, College of Veterinary Medicine, Warnell School of Forestry and Natural Resources, University of Georgia, Athens, GA, 3Veterinary Diagnostic Lab, College of Veterinary Medicine, University of Georgia, Athens, GA, 4Southeast- ern Cooperative Wildlife Disease Study, Department of Population Health, College of Veterinary Medicine, University of Georgia, Athens, GA, 5Brown University, Providence, RI

In the current study we surveyed 81 wild-caught Tokay Geckos (Gekko gecko) from the island of Java, Indo- nesia for endo- and ecto-parasites. Based on necropsy and fecal examination, 94% of geckos were infected with at least one parasite, including at least six helminth species, one pentastomid species, and two species of coccidia. No ectoparasites were detected on any gecko. At necropsy, we identified Paradistomum geckonum in the bile duct (prevalence (P)=1.2%, mean infection intensity (MII) = 7.0), Oochoristica sp. in the small intestine (1.2%, 1.0), larval acanthocephalans and nematodes in the coelomic cavity (13.5%, 2.0 and 1.2%, 1 respectively), Physalopteroides sp. in the stomach (11.1%, 2.3), several species of pinworms in the family Pharyngodonidae in the large intestine (54.3%, 29.0), and Raillietiella affinis (40.7%, 8.7) in the lungs. Histological examination of stomachs indicated that at least 4.9% of geckos were infected with Cryptosporidium spp. In addition, an unidentified trematode was observed in the pancreas of two geckos and endogenous stages of Eimeria tokayae were detected in the small intestine of numerous animals. At necropsy, a fecal sample was examined for ova and oocysts. We found eggs morphologically consistent with Paradistomum geckonum (P=2.6%), Oochoristica sp. (P=1.3%), Physalopteroides sp. (P=9.2%), pinworms (P=50%), and an unidentified trematode (P=5.3%). Oocysts of E. tokayae were found in 67.1% of geckos. These data indicate that wild-caught geckos from Indonesia are infected with several species of parasites, some of which might be zoonotic (e.g., Cryptosporidium).

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29 The Effect of Water Salinity on the Distribution and Abundance of Snail Intermediate Host of Opisthorchis viverrini and Human Prevalence in Northeast Thailand Apiporn Suwannatrai1*, Thitima Wongsaroj2, John B. Malone1, Kulwadee Suwannatrai3, Jutharat Kulsanti- wong3, Supawadee Piratae3, Chalida Thammasiri3, Sattrachai Prasopdee3, Panita Khampoosa3, Rasamee Suwanwerakamtorn4, Pairat Tarbsripair5, Thidarat Boonmars6, Somsak Sukchan7, Jennifer C. McCarroll1, Smarn Tesana3 1Pathobiological Sciences, LSU, Baton Rouge, LA, 2Department of Disease control, Ministry of Public Health, Bangkok, Thailand, 3Food-Borne Parasite Research Group, Department of Parasitology, Faculty of Medicine, KKU, Khon Kaen, Thailand, 4Regional Centre for Geo-Informatic and Space Technology, North- east Thailand, Department of Computer Science, Faculty of Sciences, KKU, Khon Kaen, Thailand, 5Depart- ment of Biology, Faculty of Sciences, KKU, Khon Kaen, Thailand, 6Department of Parasitology, Faculty of Medicine, KKU, Khon Kaen, Thailand, 7Office of Soil Survey and Land Use Planning, Land Development Department, Bangkok, Thailand

Liver fluke infection caused by Opisthorchis viverrini remains a major public health problem in Southeast Asia including Thailand, Lao PDR, Cambodia and some villages in South Vietnam. The incidence of cholan- giocarcinoma in northeast Thailand is remarkable, the highest in the world, and is coincident with areas with high prevalence rates of O. viverrini. Understanding how changes in climate and environment impact the distribution and emergence of opisthorchiasis in northeast Thailand is important to prevention and control. We used maximum entropy algorithm software (Maxent) and ArcGIS 9.3 to develop an ecological niche model based on the relationships between village prevalence site records of O. viverrini in northeast Thai- land with bioclimatic data, altitude, and other relevant environmental factors, including soil surface salt data and enhanced vegetation index and day-night land surface temperature data from the MODIS earth observ- ing satellite. Soil surface salt was the most important variable in the prediction model (27.5 % contribution). Precipitation in the wettest month and altitude were also important model variables (14.5% and 13.1%, respectively). A separate malacological survey was carried out on 56 water bodies in the Khorat basin at locations representative of six classes of soil surface salt. The preferred habitat of B. siamensis goniomphalos was found to be clear, shallow shoreline waters. Snails were found in water with salinity levels ranging from 0.05-22.11ppk, an observation that provides new evidence that B. siamensis goniomphalos has a preference for waters with some saline content rather than fresh water. The highest density of the snail was found in water within a salinity range 2.51-5.00 ppk in rice paddies, ponds, ditches and canals. Snail vector density and distribution in the Khorat basin was negatively correlated with water salinity >5ppk (r = -0.361, p<0.05). This result supports findings of the relationship of salinity to human prevalence surveillance.

30 Effect of two different chemical formulations of a commercial disinfectant on the development of Parascaris equorum eggs Mary G. Rossano*, Jessica C. Gould Animal & Food Sciences, University of Kentucky, Lexington, KY

The purpose of this study was to determine the effects of two different chemical formulations of commercial brand Professional Amphyl disinfectant on the development of Parascaris equorum eggs. The different commercial formulations tested were Professional Amphyl II Deodorant Spray (an aerosol spray containing 0.106% dimethyl benzyl ammonium saccharintate and 79.646% Ethanol) and Professional Amphyl Disin- fectant Cleaner (a liquid concentrate containing 10.5% o-phenylphenol and 5.0% 0-benzyl-p-chlorophenol). Deionized water was used as the control. Eggs were incubated at 25oC and evaluated by light microscopy on days 10 (d10) and 58 (d58) post treatment, when they were categorized as larvated or non-larvated. Overall treatment efficacy was assessed by the Kruskal-Wallis ANOVA and treatment groups were compared to each other by the Wilcoxon Rank Sum test. A p-value of < 0.05 was deemed to be significant. On

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D10, the average % larvated eggs in the aerosol spray, the liquid product and the deionized water treatment groups were 6.71, 81.30, and 89.24 respectively. On D58, the average % larvated eggs in the aerosol spray, the liquid product and the deionized water treatment groups were 21.95, 63.71, and 78.43 respectively. There was a significant difference (P<0.0001) between the three treatments on both day 10 and day 58. At each time point, both forms of the disinfectant produced significantly lower percentages of larvated eggs compared to the control, and the spray product was significantly more effective than the liquid product. Professional Amphyl II Deodorant Spray showed the most promise as a chemical disinfectant against P. equorum eggs. Used in combination with other parasite management practices, Professional Amphyl II Deodorant Spray may prove effective in reducing the incidence of P. equorum infection via stalls or fomites.

31 Larvicidal Efficacy of Fenbendazole Against a Macrocyclic Lactone-Resistant Isolate of Parascaris equorum Craig R. Reinemeyer1*, Julio C. Prado1, Wendy E. Vaala2 1East Tennessee Clinical Research, Inc., Rockwood, TN, 2Intervet/Schering Plough Animal Health, Alma, WI

During 2008, 16 suckling foals were each infected orally with ~600 larvated eggs of a macrocyclic lactone- resistant (M/L-R) isolate of P. equorum. Foals were assigned randomly to two treatment groups: Group 1 - fenbendazole (FBZ) paste, 10 mg/kg PO daily for 5 consecutive days (Days 11 to 15 post-infection), or Group 2 - ivermectin (IVM) paste, 0.2 mg/kg PO on Day 15 PI. Inoculated foals were confined with their dams PI. Approximately 80 days PI, foals were weaned, placed in individual stalls, and sampled regularly to detect the onset of patency. When six foals achieved egg counts >150 Parascaris EPG, the first cohort of 12 foals was scheduled for necropsy 10 days later. Four additional foals were processed identically at a later date. Individual fecal samples were collected on the day of necropsy. Geometric mean egg counts of Group 1 foals (1.35 EPG) were significantly lower (P<0.0001) than those of Group 2 foals (GM = 281.03 EPG), representing a FECR of 99.5%. At necropsy, the contents of the stomach, small intestine, and ce- cum were collected and sieved. P. equorum specimens were recovered, counted, and identified to sex and stage. Mean numbers of adult P. equorum were significantly lower (P<0.0018) in foals treated with FBZ for five consecutive days (GM = 4.28; range 0 - 234) compared to those treated once with IVM (GM = 115.72; range 31 - 593). All eight foals treated with IVM met adequacy of infection criteria for negative controls. This study confirmed that the isolate tested was M/L-R, and that FBZ treatment (10 mg/kg) for five consecutive days was 96.3% effective as a larvicide. This regimen has practical applications for treating foals that may be carrying occult, M/L-R infections, and for synchronizing ascarid control programs in juvenile horses.

Session 6 - Companion animal ecto 32 Comparative Efficacy of Advantage® Topical Solution (Imidacloprid) and Comfortis® Chewable Tablets (Spinosad) for the Control of Fleas on Dogs Byron L. Blagburn1*, Larry Cruthers2, Jennifer Ketzis3, Robert Arther4, Wendell Davis4, Iris Schroeder5, Terry Settje4, Jamie M. Butler1 1Dept. Pathobiology, CVM, Auburn University, Auburn, AL, 2Professional Laboratory and Research Services Inc., Corapeake, NC, 3Charles River Biolab, Co. Mayo, Ireland, 4Bayer Health Care, Shawnee, KS, 5Bayer Animal Health GmbH, Leverkusen, Germany,

Three controlled laboratory cohort studies were conducted to evaluate the initial and residual flea control efficacy of Advantage® Topical Solution ([ATS] 9.1 % w/w imidacloprid) vs oral treatment with Comfortis® Chewable Tablets ([CCT] spinosad), when administered per label directions. A total of 69 beagles, 14-16

49 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA weeks of age, were acclimatized to the test facilities. During acclimation, each dog was infested with 100 colony-reared adult fleas (C. felis). Dogs were randomized to 3 groups of 7-8 dogs/group based on acclima- tization flea counts performed 24 hours after infestation. Each dog was infested with 100 fleas on study day (SD) -1. On SD 0, group 1 dogs were treated with ATS, group 2 dogs were treated with CCT, while group 3 dogs remained non-treated. All dogs were offered food following treatment. Each dog was reinfested with 100 fleas on SD 6, 13, 20, and 27. Fleas were counted and removed on SD 2, 8, 15, 22, and 29. Flea efficacy was calculated by comparing the geometric mean number of live fleas on the treated dogs to fleas on the control dogs. Combined flea efficacy for all study sites on SD 2, 8, 15, 22, and 29 for ATS was 100, 100, 100, 98, and 90%, respectively, and 100, 100, 85, 63, and 2% for CCT, respectively. Geometric mean flea counts for the control dogs ranged from 52.8-98.2 fleas per dog. Significantly fewer fleas (p< 0.05)- re mained on the dogs treated with ATS, compared to dogs treated with CCT on SD 15, 22, and 29 at all three study sites. There were no statistical differences in the number of live fleas recovered from the dogs treated with CCT vs the control dogs on SD 22 and 29 for two study sites, and on SD 29 for the third site (p>0.05).

33 Inhibition of flea Ctenocephalides( felis) feeding on dogs treated with dinotefuran, pyriproxfen, and permenthrin (Vectra 3D®) based on quantitative polymerase chain reaction (qPCR) detection of a canine HMBS gene sequence Byron L. Blagburn1*, Bernhard Kaltenboeck1, Jane D. Mount1, Joy V. Bowles1, Jamie M. Butler1, Sherry Wilson2, Sheila Gross3, Cathy Ball2, Chengming Wang1 1Pathobiology, Auburn University, Auburn, AL, 2Summit Vetpharm, Rutherford, NJ, 3Contract Statistician, Piscataway, NJ

Fleas (C. felis) infestations on dogs and cats can result in irritation, anemia, and flea allergy dermatitis. Pre- vention of flea feeding by flea control products can help prevent primary diseases in pets and transmission of disease agents to pets and people. We compared flea feeding on control dogs and dogs treated with dinotefuran, pyriproxyfen, and permethrin (Vectra 3D®) using qPCR for detection of a canine hydroxymeth- ylbilane synthase (HMBS) housekeeping gene sequence. The canine HMBS gene codes for an enzyme necessary for heme formation and is a component of the genome of every canine blood leukocyte. Its presence can be quantified by qPCR and used as a unit of measurement for blood volume consumed by fleas. Sixteen Beagle dogs were placed in two groups of 8 dogs each. Eight dogs were treated topically with Vectra 3D® per label recommendations and 8 dogs were treated with excipients without active ingredients. All dogs were infested with 150 insectary-reared adult fleas (50% male/50% female) at time 0. Fleas were removed from dogs at 5 min, 10 min, 15 min, 20 min, 30 min, 45 min and 60 min after infestation. Canine HMBS DNA was quantified by comparison to PicoGreen® assayed copies of the standard during FRET- PCR performed on a LightCycler® 1.5 real-time PCR platform with Software version 3.53. Blood consump- tion by fleas feeding on treated dogs vs. control dogs, as determined by mean number of HMBS copies per 10 fleas, was significantly different at all time points except 15 min. post treatment (p≤ 0.05). Results of this study indicate that Vectra 3D® significantly reduces flea feeding on treated dogs based on qPCR detection of an HMBS gene sequence.

This study was supported in part by a research grant to B. L. Blagburn from Summit VetPharm, Rutherford, NJ.

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34 Efficacy of topically applied dinotefuran formulations and orally administered spinosad tablets against the KS1 flea strain infesting dogs M. Dryden*, P Payne, V Smith, A McBride, D Ritchie Department of Diagnostic Medicine/Pathobiology, College of Veterinary Medicine, Kansas State University, Manhattan, KS

Two studies were conducted to evaluate and compare the efficacy of two different topical dinotefuran formulations and orally administered spinosad against the KS1 flea strain infesting dogs. Both studies had almost identical study designs. Dogs (15m:15f) were weighed and distributed into 3 treatment groups (10 dogs each) and treated on day 0. In study 1 treatment groups were untreated controls, dinotefuran - pyriproxyfen - permethrin topical spot-on (DPP) and spinosad chewable tablet. In study 2 treatment groups were untreated controls, dinotefuran - pyriproxyfen (DP) topical spot-on and spinosad chewable tablet. Products were administered according to label directions. Spinosad treated dogs were fed prior to dosing and observed to ensure dogs did not vomit. Dogs were infested with 100 fleas on Day –2 and efficacy determined by removing fleas with a flea comb at 6 hrs and at 24 hrs post-treatment. Dogs were reinfested with 100 fleas on days 7, 14, 21 & 28 post treatment and then combed at 6 hrs and 24 hrs post-reinfestation. In both studies spinosad and DPP had 100% and > 97.2% efficacy, respectively at 6 hours post-treatment. In Study #1 both formulations provided > 93.94% efficacy within 6 hr post-reinfestation through day 14. At day 21 the 6 hr efficacy of the spinosad formulation was 54.9%. The DPP formulation provided 92.3% efficacy 6 hr post infestation on day 28. In study # 2 the DP formulation provided 99.47% efficacy through day 28 at 24 hours post-infestation. Spinosad provided > 96.6% through day 14 at 24 hrs post-re-infestation, but efficacy then declined to 64.8% at day 21 The dinotefuran topical formulations were highly effective against the KS1 flea strain, whereas the spinosad oral tablets did not provide a high level of residual efficacy on days 21 and 28 when dogs were combed at 24 hrs post-infestation.

35 Efficacy of flavored spinosad tablets administered orally to dogs in a Simulated Home Environment (SHE), for the control of existing flea Ctenocephalides( felis) infestations Daniel E. Snyder Elanco Animal Health, Greenfield, IN

The Simulated Home Environment (SHE) study has been used to simulate real world conditions to assess flea efficacy in the face of an ongoing flea challenge. Twenty dogs were equally and randomly allocated to two groups. Dogs were dosed on days 0, 28 and 56 with either Spinosad tablets (30-60 mg/kg) or 0 mg/kg (control). The flea life cycle was established in each dog’s individual carpeted pen prior to the st1 treatment, and was maintained at high levels over the entire study period so reinfestation would occur from the dog’s ‘home’ environment. Throughout the study to Day 84, individual dog flea comb counts were performed weekly and up to 50 fleas combed off were placed back on the dog, further augmenting the flea challenge. The increasing calculated geometric mean (GM) percent reduction in flea counts in the Spinosad group compared to the control group demonstrated a progressive decline in flea challenge as a result of the repeated monthly treatments with Spinosad. The control group GM flea counts on Days 56 and 84 were 441 and 349 fleas per dog, and the reduction in flea counts in the Spinosad group at these times were 98.3% and 99.4%, respectively. The results from this SHE study demonstrate sustained month-long effectiveness of each Spinosad treatment. The Spinosad flea treatment program as assessed in this SHE study successfully controlled the simulated flea problem under challenge conditions of far greater severity than normally encountered in the ‘real world’. The results also confirm the results from the clinical field study using client owned dogs where flea counts at 15 days after enrollment were reduced by 98.6% and at 90 days were

51 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA reduced by 99.9% further confirming that monthly Spinosad treatments are highly effective for the prevention and control of flea problems on dogs, causing a cumulative reduction in flea challenge from their local environment.

36 Rapid onset of action of spinosad against adult cat flea Ctenocephalides( felis) infestations on dogs Daniel E. Snyder Elanco Animal Health, Greenfield, IN

Spinosad is a novel flea adulticide with a unique mode of action among insecticides. In the treatment of flea infestations, the speed of onset of flea killing activity may be important in alleviating canine flea allergy dermatitis. Objective. To assess the speed of kill efficacy of spinosad tablets, administered at 30 to 60 mg/ kg to flea infested dogs, compared to untreated controls. Trial Design. On the basis of flea counts from an infestation of 100 applied 9 or 10 days pre-study, dogs were ranked and blocked into groups of 4, and allocated to receive either no treatment (control group), or to be treated with spinosad (7 groups). On Day -1, all dogs were infested with approximately 100 unfed adult fleas. Spinosad was administered at time 0, and flea counts were completed at that time on the control group, and in the spinosad groups at 0.5, 1, 2, 4, 8, 24 and 48 hours after treatment. Some dogs treated outside the targeted spinosad dose range were excluded from the analysis, leading to elimination of 1 dog from each of 5 spinosad groups. Results. Geo- metric mean flea counts in the control group were 97.5, validating the flea infestation source and methodology. At 0.5 hours post treatment, there was a significant mean flea count reduction (p<0.001) of 82.9% in the spinosad group, relative to the untreated control group. Geometric mean flea counts in spinosad treated dogs were significantly less (p≤0.002) than in untreated controls at all subsequent time points. Spinosad was 100% effective at all flea counts from 4 hours post treatment. Conclusion. These results indicate a rapid onset of action of spinosad administered orally at a dose rate of 30 to 60 mg/kg, with significant difference (p<0.001) from untreated controls as soon as 30 minutes post treatment.

37 Insecticide Resistance Profiles of Field-Collected Isolates of Cat Fleas (Ctenocephalides felis) Michael K. Rust1*, Byron L. Blagburn2, Iris Schroeder3, Sarah Weston3 1Entomology, University of California Riverside, Riverside, CA, 2Dept. Pathobiology, CVM, Auburn Univer- sity, Auburn, AL, 3Bayer Animal Health GmbH, Leverkusen, Germany

It has been 12 years since the last comprehensive review of insecticide resistance profiles for field-collected isolates of cat fleas,Ctenocephalides felis (Bouche), has been published. Very limited numbers of field-collected isolates have been tested for insecticide resistance to commonly used insecticides. We report on the susceptibility of 12 field isolates to deltamethrin, permethrin, sumithrin, tetrachlorvinphos, fipronil, and imidacloprid. In addition, four laboratory strains that had no history of insecticide exposure and data published by Moyses and Gefeller served as possible baselines for susceptibility to each of the toxicants. Insecticidal activity was determined by applying 0.2 µl droplets of each toxicant to adult fleas. Topical applications of dieldrin failed to kill any of the isolates and provide data for probit analysis. Resistance Ratios (LD50 field strain/LD50 lab strain) for permethrin ranged from 2.4 to 9.6. RR50 for sumithrin and deltamethrin ranged from 2.1 to 12.3 and 2.1 to 10.9, respectively. The RR50 for fipronil and imidacloprid ranged from 1.2 to 3.2 and <1 to 3.6, respectfully. Larval bioassays were conducted for imidacloprid by treating larval rearing media with a serial dilution of the insecticide. The RR50 of the larval bioassays provided a similar response as the adult topical bioassay validating the use of larval bioassays to determine susceptibility of cat fleas to insecticides. This work was in supported in part by a research grant from MGK Co. and Bayer Animal Health, Monheim, Germany.

52 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Session 7 - Parasite prevalence 38 Geospatial Analysis and Ecological Niche Modeling of Chagas Disease in Bolivia Paula D. Mischler PBS, LSU Vetmed, Baton Rouge, LA

An estimated 10-15 million people in the world are infected with Chagas disease. In Bolivia, it is estimated that 10 percent of the population is infected with the disease with another 3.5 million people at risk. A myriad of factors affect the high prevalence of the disease in Bolivia including poverty, the presence of domestic and sylvatic populations of Triatoma infestans, and societal habits. This study focused on the use of two modeling techniques to predict the risk of Chagas disease in Bolivia. A comprehensive literature search yielded seroprevalence survey results for 14 municipalities in 5 Bolivian departments. Records of Triatoma infestans positive for chagas infection was obtained for 17 municipalities in 5 Bolivian departments. The seroprevalence and vector occurrence data was used in conjunction with health data (human development index data at the municipal level) and environmental data (ecological region, elevation, Bioclim variables) to create both an ecological niche model in Maxent on vector records data and a geospatial query model based on ranges of variables for each seroprevalence study site using ArcGIS 9.3. Both models used the same environmental and socioeconomic variables. Both models predicted similar Chagas distributions, but there were differences with the geospatial model predicting a more easterly distribution when compared to the ecological niche model. The aim is to develop geospatial risk assessment models that are useful in the implementation and planning of Chagas disease surveillance and resource allocation to target the portion of the population most at risk for the disease.

39 Prevalence of canine vector-borne diseases in heartworm-tolerant Jindo dogs SungShik Shin1*, Seok-Il Oh2, DaeSung Oh1, KyuSung Ahn1, Kyung-Oh Cho3 1Parasitology, College of Veterinary Medicine, Chonnam National University, Gwangju, South Korea, 2Ko- rean Jindo Dog Center, Jindo 539-823, South Korea, 3Biotherapy Human Resources Center, College of Veterinary Medicine, Chonnam National University, Gwangju, South Korea

Previous studies indicate that Jindo, a Korean-native dog designated by the government as a natural treasure of Korea, is quite tolerant to heavy infection with canine heartworms. In 2004, we performed a serological survey on heartworm infection in Jindo dogs in which 69.6% of 365 dogs raised in the Jindo Island were infected with Dirofilaria immitis. In 2009, we performed another serological survey in the same area for D. immitis, Anaplasma phagocytophilum, Ehrlichia canis, and Borrelia burgdorferi infections using a commercially-available fast diagnostic test. From a total of 308 Jindo dogs, the number of serologically positive dogs for any of the four pathogens was 113 (36.7%). The highest prevalence observed was D. immitis (33.8%), followed by A. phagocytophilum (4.2%) and E. canis (1.0%). Seropositivity to B. burgdorferi was not observed in the survey. Majority of tick species collected from dogs in the study area was Hae- maphysalis longicornis. This study indicates that while tick-borne diseases are relatively limited in number, canine heartworm infection, a mosquito-borne disease, is still quite high among Jindo dogs despite the continuous effort of prophylactic measure by the local government.

53 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

40 Seroprevalence of Borrelia burgdorferi, Anaplasma phagocytophilum, Ehrlichia canis and Dirofilaria immitis among dogs in Canada Alain Villeneuve1*, Jonas K. Goring2, Lynne Marcotte2, Sebastien Overvelde2 1Pathology and Microbiology, Universite de Montreal, St. Hyacinthe, QC, 2IDEXX Reference Lab, IDEXX Labs, Montreal, QC

In Canada, the environmental suitability for Borrelia burgdorferi and Anaplasma phagocytophilum vectors, Ixodes scapularis and Ixodes pacificus, appears to be growing. In 1991, there was one known endemic location for I. scapularis while currently there are at least eleven. An estimated 8-12% of the 50-175 million adventitious I. scapularis ticks that enter Canada on birds are positive for B. burgdorferi. Heartworm prevalence has been relatively stable over the past 20 years.

The study assessed the seroprevalence of Heartworm (Dirofilaria immitis) and three tick-borne diseases (B. burgdorferi, A. phagocytophilum, Ehrlichia canis) in Canadian dogs. A total of 238 veterinary practices from nine Canadian provinces submitted 86,251 canine test results from an ELISA immunoassay, (SNAP ® 4Dx®, IDEXX Laboratories, Westbrook, Maine, USA) performed either in-house or at one of five Canadian reference labs.

The agent with the highest seroprevalence was B. burgdorferi (0.72%) followed by for D. immitis (0.22%), A. phagocytophilum (0.19%) and E. canis (0.05%). The majority of testing and positive cases were found in Central and Eastern Canada. In addition to the seroassessment, clinical and signalment data was provided for 913 of the patients testing positive for one or more vector borne disease. 79% of the patients testing positive did not have a travel history outside of their native province in the past 6 months. Roughly an equal number of patients testing positive had a history of tick contact as did not. Dogs co-infected with B. burgdorferi/D. immitis and B. burgdorferi/A. phagocytophilum had the highest likelihood to be ill, 20% and 18.9%, respectively. Of the dogs infected with a single agent, B. burgdorferi seropositivity had the highest percentage of ill patients (14.3%) followed by Heartworm (12.2%). The risk for vector borne infectious agents in the Canadian canine population is low but widespread with foci of higher prevalence.

41 Seroprevalence of Heartworm, Toxoplasma gondii, FIV and FeLV Infections in Pet Cats in Bangkok and Suburban Area, Thailand Woraporn Sukhumavasi1*, Mary L. Bellosa2, Araceli Lucio-Forster3, Janice L. Liotta4, Alice C.Y. Lee3, Pitcha Pornmingmas5, Sudchit Chungpivat1, Leif Lorentzen6, Dwight D. Bowman3, J. P. Dubey7 1Parasitology Unit, Department of Pathology, Faculty of Veterinary Science, Chulalongkorn University, Bangkok, Thailand, 2College of Veterinary Medicine, Cornell University, Ithaca, NY, 3Microbiology and Im- munoloy, Cornell University, Ithaca, NY, 4Department of Microbiology and Immunology, Cornell University, Ithaca, 5Suvarnachad Animal Hospital, Bangkok, Thailand, 6IDEXX Laboratories, Westbrook, ME, 7Animal Parasitic Disease Laboratory, United States Department of Agriculture, Beltsville, USA, Beltsville, MD

Seroprevalence of heartworm (Dirofilaria immitis), Toxoplasma gondii, feline immunodeficiency virus (FIV) and feline leukemia virus (FeLV) infections was examined in serum samples from seven hundred and forty- six pet cats collected from the private clinics or hospitals located in Bangkok and suburban areas between May and July 2009. Of these 746 samples, 4.56% (34/746) were positive for heartworm antigen, 20.11% (150/746) had antibodies against FIV and 24.53% (183/746) had circulating FeLV antigen. In addition a random subsample of sera was tested for T. gondii antibodies using a modified agglutination test. Of 348 serum samples, 10.06% (35/348) were seropositive against T. gondii. The percentage of FIV/FeLV, FIV/T. gondii and FeLV/T.gondii co-infections were 5.36% (40/746), 2.87% (10/348) and 2.87% (10/348), respectively. Interestingly, of 35 T. gondii-seropositive cats, 40% (14/35) were co-infected with aforementioned

54 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA pathogens. Currently, the association of these infections with gender, age and cat lifestyle is being investigated. This data is important since it is the first to reveal the prevalence of four important feline infectious disease agents simultaneously in Thailand, hence providing evidence for the need of disease awareness for veterinarians and pet owners.

42 Seroprevalence of Anoplocephala perfoliata Infection Among Horses in West Coast States of the US Patrick Meeus1*, Cari Lagrow2, Steven Kania3, Craig Reinemeyer4, Vickie King5, Sallie Cosgrove5, 1Parasi- tology Discovery, Pfizer Animal Health, Kalamazoo, MI 2Pfizer Animal Health, Kalamazoo, MI, 3University of Tennessee, Knoxville, TN, 4East Tennessee Clinical Research, Inc, Knoxville, TN, 5Pfizer Animal Health, Kalamazoo, MI

A survey was conducted in 634 randomly-selected horses residing in the Western Coastal region of the US to determine the seroprevalence of Anoplocephala perfoliata. Equine practitioners in the region (n = 174) enrolled a maximum of four horses each, and no more than two horses per farm. Enrolled horses were two years of age or older, in apparent good health, and had not been treated with anthelmintics within the prior 30 days. For each enrolled horse, a questionnaire was filled out to capture demographic characteristics, details of management and housing Subsequently, blood samples were collected and serum was analyzed with an ELISA for the presence or absence of A. perfoliata antibodies (sensitivity 81%, specificity 95%). Prevalences were calculated for location, animal characteristics, management features, and housing type.

The horse seroprevalences of A. perfoliata were 8.5% in California, 23.1% in Oregon, and 15.0% in Wash- ington. The farm seroprevalences, i.e. proportion of farms with at least one positive horse, were 17.3%, 36.5% and 25.3%, respectively. Higher temperatures and drier weather appear to adversely affect tapeworm transmission, and are accompanied by lower seroprevalence levels. No impact on the seroprevalence was observed from age, gender or breed. The use of praziquantel dewormers resulted in a numerical reduction in prevalence from 19 to 13.9 %, but this difference was not statistically significant. Access to healthy pastures, where orbatid mites presumably can thrive, resulted in a numerically higher prevalence.

43 Trypanosoma cruzi infection in dogs in south central Louisiana Prixia del Mar Nieto*, John B. Malone Pathobiological Sciences, Louisiana State University, Baton Rouge, LA

Dogs considered to be at high risk of infection with Trypanosoma cruzi in southern Louisiana were tested serologically using the indirect fluorescent antibody test (IFAT). Serum samples obtained from a total of 122 dogs from three kennels, and from client dogs from local veterinary practices tested by IFAT revealed a prevalence rate of 22.1%. Fifty randomly selected samples from this group were also tested using two rapid experimental immunochromatographic assays designed as alternative or complementary diagnostic tests for T. cruzi infection. Of the fifty samples tested thirteen animals tested positive using rapid assay A and eleven animals tested positive using rapid assay B. In the same group, 11 animals tested positive by IFAT. The sensitivity of rapid assay A and B were 100%; the specificity of rapid assay A was 95%, and rapid assay B was 100% as compared to the IFAT, the test standard. Clinico-pathological reports revealed that cardiac signs are the main indicators of Chagas disease. Rapid, easy and accurate screening assays, used in conjunction with confirmatory tests, would be an excellent tool for veterinarians to diagnoseT. cruzi infection. Greater awareness by veterinarians of the risk as well as the description of clinical and pathological findings will contribute greatly to an understanding of the true prevalence of Chagas disease in dogs in the United States.

55 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Session 8 – Wildlife 44 Baylisascaris procyonis infection in raccoons and dogs on Prince Edward Island, Canada Gary Conboy1*, Tonya A Stewart2, Amanda Taylor2 1Pathology and Microbiology, Atlantic Veterinary College-UPEI, Charlottetown, PE, 2Companion Animal, Atlantic Veterinary College-UPEI, Charlottetown, PE

Baylisascaris procyonis, the raccoon roundworm, is the cause of cerebrospinal nematodosis in rabbits, rodents, birds and various other animals including humans. Human exposure is through the ingestion of larvated eggs and infection can lead to ocular disease or severe central nervous system disease which can result in death. In addition to raccoons, patent B. procyonis infections also occur in dogs. At present, B. procyonis infection in dogs is considered rare, however, diagnoses may be missed due to the similarity in morphology between B. procyonis and Toxocara canis eggs. Since April 2005, patent B.procyonis infections were diagnosed by fecal flotation in 5 dogs on PEI. Each case was confirmed by recovery of adult worms passed in the feces. Dogs ranged in age from 9 months to 4 yrs (mean = 2.25 yrs). All dogs were treated with fenbendazole (50 mg/kg for 5-7 days). One dog was hospitalized and shed 17 adult worms in the feces during the treatment period. A necropsy survey found B. procyonis infection in 17/242 (7%) of the raccoons on PEI. A fecal flotation survey of dogs at the PEI Humane Society (2005-2007) detectedB. procyonis eggs in 2/555 (0.4%) samples. Infections were confirmed by the recovery of adult worms passed in the feces in both of these dogs. Prevalence of B. procyonis infection in raccoon populations in the midwestern USA and elsewhere may be as much as 10 times that of PEI. Presumably, exposure risk of infection and percentage of dogs shedding eggs would be greater than that observed on PEI in regions with a higher prevalence of B. procyonis infection in the raccoon population. Veterinarians need to be aware of the possibility of B. procyonis infection for the diagnosis, treatment and prevention of environmental contamination by dogs with the eggs of this dangerous zoonotic pathogen.

45 Distribution, prevalence and genetic characterization of Baylisascaris procyonis from selected regions of Georgia and Florida Michael J. Yabsley1*, Cheryl D. Davis2, Scott Henke3, David B. Long4, Margaret Beck5, Emily L. Blizzard1 1University of Georgia, Athens, GA, 2Western Kentucky University, Bowling Green, KY, 3Caesar Kleberg Wildlife Research Institute, Kingsville, TX, 4USDA-APHIS-Wildlife Services, Kingsville, TX, 5Goose Creek Wildlife Sanctuary, Tallahassee, FL

Baylisascaris procyonis, a parasitic intestinal nematode commonly found in raccoons (Procyon lotor), has historically been absent from the southeastern United States. In 2002, the parasite was first documented in Atlanta, Georgia. The goal of this study was to investigate distribution in Georgia and northern Florida. Intestinal tracts of 312 raccoons from 25 Georgia counties and 53 raccoons from three northwestern Florida counties were examined for B. procyonis. The only county where B. procyonis was detected was Clarke County where 12 of 116 (10.3%) raccoons were infected. In Clarke County, significantly more juveniles (p=0.049) were infected compared with adults, and no differences in prevalence were noted by sex, season of capture, or land-use (rural vs. urbanized); however, significantly (P=0.0370) higher worm burdens were found in infected raccoons from urban/suburban locations compared with rural areas. A single immature worm from a Florida raccoon was confirmed to be B. procyonis by PCR. No other raccoons were infected. To try and identify a source population for the parasites, we amplified and sequenced regions of the rRNA genes from worms from various locations. To date, ITS-1 sequences have been successfully obtained from 18 worms from Georgia (n=6), Kansas (1), Florida (1), Kentucky (4), and Texas (6). Although numerous polymorphic bases were observed among the samples, none were associated with a particular geographic

56 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA location. In addition, sequences from the 18S, 5.8S, and ITS-2 regions from six samples from Georgia, Kentucky, and Texas were 100% identical. These data indicate that the distribution of B. procyonis within Georgia is increasing and that limited genetic variation in the rRNA and ITS gene regions is present among widely distributed populations of B. procyonis. In addition, this is the first report of B. procyonis in Florida and increases the distribution of this important zoonotic parasite.

46 Prevalence of Baylisascaris procyonis in Raccoons (Procyon lotor) from eastern Colo- rado Lora R. Ballweber1*, Ivy LeVan2, Michael W. Miller2, Deanna J. Chavez1 1Colorado State University, Fort Collins, CO, 2Colorado Division of Wildlife, Fort Collins, CO

Baylisascaris procyonis is a zoonotic raccoon ascarid that is common in many parts of North America. To date, however, the prevalence of B. procyonis in Colorado has not been determined. The purpose of this study was to ascertain the prevalence of B. procyonis in raccoons in eastern Colorado. A total of 54 raccoons have, thus far, been collected by the Colorado Division of Wildlife or submitted to the Colorado State University Veterinary Diagnostic Laboratory as part of ongoing disease investigations during 2007-2010. Forty-seven were examined by necropsy and seven by a combination of fecal flotation and anthelmintic treatment. At necropsy, the gastrointestinal tracts were collected, opened longitudinally, and the ascarids removed and counted. Age (n = 36; juvenile, adult) and gender (n = 35) of the raccoons were recorded when intact carcasses were available. Prevalence and 95% confidence interval (CI), range, and mean intensity were determined. Mann Whitney U or Chi square statistics were used to examine correlations between age or gender, and the presence of ascarids or the number of ascarids. Baylisascaris procyonis was found in 32 of 54 raccoons (59.3 %; 95% CI = 45.0%, 72.4%). Mean intensity was 10.7 with a range of 1 to 49 per infected individual. Geographically, the parasite did not appear to be clustered in any particular region. There was no significant difference between age or gender and presence of ascarids or number of ascarids. Based on results to date, B. procyonis appears to be prevalent in raccoons of eastern Colorado. There does not appear to be any difference between age or gender and number of ascarids or the presence of ascarids as has been reported in some previous studies. However, this may be the result of small sample sizes.

47 What’s Killing Our Deer? Investigation of Biting Fly Vectors of Epizootic Hemorrhagic Disease in Texas Tracy L. Cyr Veterinary Pathobiology, College of Vet. Med. and Biomed. Sciences, Texas A&M University, College Sta- tion, TX

Epizootic hemorrhagic disease (EHD) is the most important and common viral disease of white-tailed deer in the United States, and outbreaks occur every year in parts of the southeast. EHDV was first reported in Texas in 1966 in a captive white-tailed deer and a bighorn sheep, but both wild and domestic ruminants can be infected by this arthropod-borne pathogen. The disease effecting white-tailed deer in the US is caused primarily by two of the at least 8 serogroups of epizootic hemorrhagic disease virus (EHDV-1 and EHDV-2), genus Orbivirus, in the family Reoviridae. Recent reports suggest that a third serotype, EHDV-6, associated with mortality in white-tailed deer may become endemic in several states, including Texas.

In order to collect and identify biting fly vector(s) of epizootic hemorrhagic disease virus (EHDV) suspected of causing debilitation and death in white-tailed deer in Texas, we set out CDC miniature light traps equipped with CO2 source around likely Culicoides breeding areas on Texas ranches near the San Angelo Extension Center where deer losses due to EHD have been reported. Traps were set out in the evening and

57 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA collected the next morning over a 5 month period, beginning in Spring, 2008. Counts and species identifica- tions have been made for over 3,000 adult female Culicoides collected from 2 of the larger collection sites in the San Angelo, Texas, area. At least 12 different species are represented. The majority of the specimens were collected during the late spring and early fall months (37% and 36%, respectively). Only 28% of the total flies were collected during the months of June and July, most likely due to the very dry environmental conditions during that time.

48 White-Tailed Deer - Alternate Host for Babesia bovis? Patricia J. Holman Veterinary Pathobiology, Texas A&M University, College Station, TX

In the western hemisphere bovine babesiosis is caused by Babesia bovis and Babesia bigemina, with B. bovis the more pathogenic agent. The disease, also known as Texas Cattle Fever, was officially eradicated from the United States in 1943 by eliminating the tick that vectors bovine Babesia, Rhipicephalus (Boophi- lus) spp. Since then, a permanent Boophilus tick quarantine buffer zone along the Texas-Mexico border has been maintained to prevent reestablishment of the ticks in the U.S. Fever tick outbreaks in the buffer zone proper are not uncommon, but 60% of new outbreaks in Texas from October 2008 through July 2009 were in “tick-free” areas outside of the quarantine zone. Boophilus ticks preferentially feed on cattle, but may infest other ungulates. Thus, this study was undertaken to determine if white-tailed deer in Texas harbor B. bovis.

Blood samples from free-ranging white-tailed deer (Odocoileus virginianus) in several Texas counties were screened for B. bovis and other hemoparasites by the polymerase chain reaction (PCR). Sera or plasma were assayed by the immunofluorescent antibody test to detect antibody activity to B. bovis. Overall, approximately 10% of the deer tested were positive for B. bovis by PCR. Molecular characterization of the resulting B. bovis rDNA amplicons showed 99% identity to B. bovis 18S rRNA gene sequences derived from cattle isolates. Weak seroreactivity to B. bovis was shown by the IFAT. Screening for additional hemoparasites of deer (Theileria cervi, Babesia odocoilei and other Babesia spp.) revealed the presence of T. cervi, B. odocoilei, and a genotypically unique Theileria sp. The finding of putative B. bovis in white-tailed deer necessitates further study to determine if deer may act as a transient host or even a reservoir of infection for B. bovis pathogenic to cattle.

49 Identification, distribution and hosts of ticks in Kansas 2000 – 2007 M. Dryden*, P Payne, V Smith, A McBride, M Hobson Department Of Diagnostic Medicine/Pathobiology, College Of Veterinary Medicine, Kansas State Univer- sity, Manhattan, KS

The Insect Diagnostic Laboratory Department of Entomology at K-State, the Veterinary Parasitology Diag- nostic Laboratory at K-State and ticks collected in the Veterinary Teaching Hospital, College of Veterinary Medicine at K-State were identified as to species. Additional information collected included host, life stage, date and county of collection. From the 3530 ticks submitted 7 different tick species were identified; A. americanum, A. maculatum, D. variabilis, D. albopictus, I. scapularis, R. sanguineus and O. megnini. These ticks were collected from 11 different mammalian hosts (cattle, dog, white-tailed deer, horse, cat, fox, human, llama, rabbit, raccoon and skunk. There were 262 ticks submitted where the host was not recorded. The most common tick species submitted were A. americanum (43.9%) and D. variabilis (42.6%), with A. maculatum (7.7%) the third most common. Of the total ticks collected 60.5%, 10.0% and 9.9% were from dogs, humans and cattle respectively. A total of 354 ticks were submitted from humans with A. americanum (57.6%) and D. variabilis (41.5%) being the most common. Of the 2135 ticks submitted from dogs D. varia-

58 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA bilis (53.8%) and A. americanum (42.6%) were the most common. From cattle 348 ticks were submitted with 65.8% being A. maculatum. From cats 158 ticks were submitted with the most common being A. americanum (63.9%) and D. variabilis (22.2%). There were 199 ticks submitted from horses with D. albipictus (48.2%) and A. americanum (42.2%) being most common. The only O. megnini (21) submitted were from cats in two western KS counties. I. scapularis (18) were submitted from 8 counties in eastern Kansas and host range included dog, horse, fox, human and rabbit.

50 Effects of Different Burning Regimes on Tick Populations in Rangelands Mason V. Reichard1*, Kristen A. Baum2 1Department of Veterinary Pathobiology, Oklahoma State University, Stillwater, OK, 2Zoology, Oklahoma State University, Stillwater, OK

Traditional rangeland management strategies, such as intense grazing by ruminants, frequent burning of grasses, and herbicide application, have focused on maintaining dominant forage species and reducing biological diversity across the landscape. Current efforts to return natural fire and grazing regimes to grasslands have focused on utilizing interactions between fire and grazing to create a shifting mosaic of grass- land patches with different fire return intervals and grazing pressures. These interactions are spatially and temporally controlled and the resulting heterogeneous pattern has been proposed as a critical feature for maintaining the structure, function, and diversity in grasslands. These changes in plant community structure also may have important implications for controlling ticks in grasslands. We are investigating the effects of heterogeneous versus homogeneous burning on the abundance, diversity, and survivability of ticks in rangelands. We are also comparing levels of tick burdens on infested cattle in pastures with heterogeneous versus homogeneous burning. The results of a three year data set will be presented and support the use of heterogeneous burning regimes as a potential method for controlling tick populations in grasslands.

51 Genetic characterization of Toxoplasma gondii from Sand cats (Felis margarita) C. Rajendran1*, An Pas2, J. P. Dubey1, C. Su3 1Animal Parasitic Disease Laboratory, U.S. Department of Agriculture, Beltsville, MD, 2Breeding Centre for Endangered Arabian Wildlife, Sharjah,, 3Department of Microbiology, The University of Tennessee, Knox- ville, TN

Until recently, Toxoplasma gondii was considered clonal with little genetic diversity. Recent studies indicate that there are geographic differences among isolates of T. gondii. However, little is known of genetic diversity of the parasite in the Arabian Peninsula. Here, we describe genetic characteristics of T. gondii from the Sand cat (Felis margarita) at the Breeding Centre for Endangered Arabian Wildlife (BCEAW), Shar- jah, United Arab Emirates. BCEAW has experienced high newborn mortality associated with congenital toxoplasmosis. In the present study, T. gondii DNA was isolated from three Sand cats; directly from livers and lungs of two cats, and from the viable isolate of T. gondii obtained by bioassay of tissues of one cat. PCR-RFLP genotyping at 10 genetic loci revealed that T. gondii isolates from these cats have an atypical genotype, which is different from the clonal type I, II and III strains. This genotype was previously reported in T. gondii isolates of dogs from Sri Lanka, indicating it is widely spread in Asia and the Arabian Peninsula.

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Session 9 - Education symposium 52 2009 AAVP Veterinary Parasitology Education Symposium Gary Conboy1*, Susan Little2, Karen F. Snowden3, Tammi Krecek4 1Pathology and Microbiology, Atlantic Veterinary College-UPEI, Charlottetown, PE, 2Oklahoma State Uni- versity, Stillwater, OK, 3Veterinary Pathobiology, Texas A&M University College of Veterinary Medicine, Col- lege Station, TX, 4Pathobiology, Ross University School of Veterinary Medicine, Basseterre

The 2009 AAVP Education Symposium funded by a grant from the Companion Animal Parasite Council and AAVP hosted by the National Center for Veterinary Parasitology was held December 11-12, 2009 at the Center for Veterinary Health Sciences, Oklahoma State University, Stillwater, Oklahoma. Thirty-four educators from 19 veterinary colleges in North America and the Carribean attended. The program included presentations by Dr. Sidney Ewing (Professor Emeritus, Oklahoma State University) - History of Veterinary Parasitology in North America, Amy Edwards and Susan Little (3rd year veterinary student/Professor respectively, Oklahoma State University) - Results from the Survey - How We Teach Parasitology Now and our keynote speaker, Dr. Elizabeth Hardie (Professor and Head, Clinical Sciences, North Carolina State University) - Outcome Assessment and Clinical Competencies in Veterinary Medicine. After the presentations, 3 breakout focus groups each discussed such topics as: 1. Clinical competencies in veterinary parasitology (ability to effectively manage clinical medical/surgical cases of parasitic infection by having the knowledge base to recognize clinical signs of parasitism and choose proper diagnostic methods, op- tions for treatment, control and prevention as well as deal with any relevant zoonotic issues); 2. Outcome assessment for veterinary parasitology (traditional testing formats of multiple choice/matching augmented by the use of clickers/on-line quizzes employed in the pre-clinical years due to restraints in manpower and the student level of clinical knowledge base and in the clinical year adopting more practical skill assessment through case presentations and direct evaluation of performance on diagnostic exercises with grading by rubric) and 3. Needs in veterinary parasitology education (recruitment of graduate students; improved communication/greater interaction with clinical faculty; continual review and renewal of course materials; greater interaction with and sharing of teaching resources between colleagues for mutual support and ben- efit; develop clinical veterinary parasitologists through NCVP/Board specialization/AAVP student chapters).

53 What is NAVMEC? Karen F. Snowden Veterinary Pathobiology, Texas A&M University College of Veterinary Medicine, College Station, TX

The North American Veterinary Medical Education Consortium (NAVMEC) was launched by the American Association of Veterinary Medical Colleges (AAVMC) in 2009 to ensure that veterinary medical education meets the needs of our changing society in the future. More than 200 stakeholders including representatives from North American colleges of veterinary medicine, umbrella professional organizations (AAVMC, AVMA Council on Education), specialty professional organizations (AAVP etc.), the animal health industry, business/marketing companies, state and national licensing agencies and private practitioners are participating in three multi-day meetings (Feb, Apr, July, 2010). Topics addressed in these meetings include 1) societal needs and clinical competencies, 2) veterinary educational models and 3) synthesis including accreditation and licensure issues. The programs include stimulation presentations by invited speakers as well as structured breakout groups, followed by summary discussions. The anticipated outcome of this series of meetings is a document that defines future needs and directions concerning “the relationship between education, accreditation, and licensure to ensure that the veterinary colleges/schools can be cre-

60 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA ative in planning their future educational programs while meeting accreditation standards, and that new graduates will be prepared to meet licensing requirements.” Additional information about NAVMEC is on the website: http://www.aavmc.org/navmec.htm

54 Survey of parasitology teaching at North American veterinary colleges: initial results from the 2009 AAVP/CAPC Education Symposium Susan Little1*, Amy Edwards2, Gary Conboy3, Karen F. Snowden4 1Oklahoma State University, Stillwater, OK, 2Center for Veterinary Health Sciences, Oklahoma State Uni- versity, Stillwater, OK, 3Pathology and Microbiology, Atlantic Veterinary College-UPEI, Charlottetown, PE, 4Veterinary Pathobiology, Texas A&M University College of Veterinary Medicine, College Station, TX

Instruction in veterinary parasitology varies among the veterinary medical education programs in North America. To begin to assess how veterinary parasitology is taught at different institutions, an initial survey was conducted of faculty responsible for teaching veterinary parasitology in conjunction with the 2009 AAVP/CAPC Education Symposium. Of the survey responses received (n=18), 50% were from respondents who had taught parasitology to veterinary students for more than 10 years, and 55.5% had taught at their current institution for 6 years or more. Most respondents (81.3%) reported 2 or 3 faculty members were involved in teaching veterinary parasitology at their school, and the most common delivery method reported was traditional lecture/laboratory (68.8%). Respondents reported an average of 37.4 contact hours of lecture and 28.2 contact hours of laboratory in the preclinical years, and all respondents reported clinical / 4th year rotations that involved parasitology. A formal process of evaluating outcomes assessment (the focus of the 2009 education symposium) was reported by a minority (18.8%) of respondents. This survey provided an initial view of how parasitology content is currently delivered to veterinary students and how the impact (learning) is assessed. Future plans include expansion of the ability of the survey instrument to assess content and depth of veterinary parasitology teaching as well as additional efforts to solicit responses from all institutions.

Session 10 - Companion animal endo 55 Evaluation of testing and treatment procedures for heartworm (Dirofilaria immitis) in animal shelters in Georgia, South Carolina, and North Carolina Andrew R. Moorhead1*, David S. Boardman1, Ruth D. Usher1, Natalie D. Duncan1, Maria T. Correa2 1Infectious Diseases, College of Veterinary Medicine, Athens, GA, 2Population Health and Pathobiology, North Carolina State University-College of Veterinary Medicine, Raleigh, NC

Heartworm (Dirofilaria immitis) is a major threat to both canine and feline health. The incidence of heartworm cases is increasing in both Georgia and the Carolinas, as determined by previous surveys of both clinics and animal shelters. We hypothesize that a large number of animals surrendered to animal shelters are currently not being administered anthelmintic prophylaxis and thus could be contributing to this increase. Furthermore, testing and treatment procedures for heartworm in animal shelters are not optimized, and could also be contributing to the increase in heartworm incidence. In an effort to understand the spread of the disease, as well as possibly institute potential control measures in animal shelters, we wished to determine the current testing and treatment procedures for heartworm in animal shelters in Georgia, South Carolina, and North Carolina. We performed a brief email survey of animal shelters regarding their heartworm testing and treatment procedures. Of the over 500 shelters surveyed, we received 122 responses. Of these 77.4% were non-profit organizations that were deemed no-kill/limited admission. 84.4% and 12.3% of the respondent shelters tested for heartworm in dogs and cats, respectively. Furthermore, over 90% of

61 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA non-profit shelters versus 60% of government-administered shelters tested for heartworm. These results seem to indicate that both non-profit and government facilities are aware of heartworm and the need to determine the heartworm status of animals. Also, due to the fact that only 34.2% of the respondent shelters were associated with a veterinarian, veterinary care does not appear to be a prerequisite for testing. We will perform a follow-up survey to gain detailed information about the type of test administered, as well as the decision-making process used regarding treatment.

56 Genetic changes in Dirofilaria immitis populations possibly associated with exposure to macrocyclic lactones Timothy G. Geary1*, Catherine Bourguinat1, Byron L. Blagburn2, Kathy Keller1, Rudolph Schenker3 Roger K. Prichard1 1Institute of Parasitology, McGill University, Ste-Anne-de-Bellevue, QC, 2Dept. Pathobiology, CVM, Auburn University, Auburn AL, 3Novartis Animal Health, Basel

Macrocyclic lactone (ML) endectocides are used, mostly at monthly intervals, during the mosquito transmission season for heartworm chemoprophylaxis in dogs and cats. Unconfirmed reports of loss of efficacy of ML heartworm preventatives have appeared in some locations in recent years. ML resistance is an increasing problem in nematode parasites of ruminants and horses and in the human filarial nematode Onchocerca volvulus. Heritable loss of efficacy of MLs would have a genetic basis; detection of relevant genetic changes could indicate a developing drug resistance situation in Dirofilaria immitis. A baseline genetic database was built from ML-naïve D. immitis from dogs. Genetic polymorphisms were been investigated in D. immitis populations obtained from dogs with different treatment histories or in which a loss of ML efficacy is suspected. We found evidence for significant differences in genetic polymorphism between parasite populations with different treatment histories, most notably from parasites obtained from putative loss-of-efficacy cases. This forms part of an ongoing project to evaluate evidence of genetic selection for ML resistance in D. immitis.

57 Performance Comparison of a New, In-Clinic Method for the Detection of Canine Heart- worm Antigen Alice C.Y. Lee1*, Dwight D. Bowman1, Araceli Lucio-Forster1, Melissa J. Beall2, Janice L. Liotta1, Ray Dillon3 1Microbiology and Immunology, Cornell University, Ithaca, NY, 2IDEXX Laboratories, Inc., Westbrook, ME, 3Clinical Sciences, Auburn University, Auburn, AL

Canine heartworm is endemic in many parts of the world, and veterinarians rely on rapid in-clinic antigen tests to screen for this infection. Recently, an in-clinic, instrument-based rotor employing an agglutination immunoassay was launched in the marketplace (VetScan VS2® Canine Heartworm Antigen Test; Abaxis). Because of the widespread use of heartworm prevention and possible false negative test results in dogs with low heartworm burdens, the performance of VetScan VS2® Canine Heartworm Antigen Test and a commercially available ELISA-based test (SNAP® Heartworm RT Test; IDEXX) was compared using samples from dogs with low heartworm burdens and/or low levels of circulating antigen.

Ninety serum samples were evaluated using the two methods. Testing was performed according to the manufacturer’s product insert by personnel blinded to sample status. The samples were derived from two sources: dogs with necropsy-confirmed heartworm status (40 with 1-4 female worms, 30 with no worms),

62 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA and field dogs (20) confirmed positive for antigen by microtiter plate ELISA (PetChek® Heartworm PF Anti- gen Test; IDEXX). All 40 dogs with heartworms on necropsy were also confirmed to have circulating antigen by PetChek.

In necropsy-negative dogs (n=30), neither VetScan nor SNAP RT detected heartworm antigen. Of the samples testing positive for antigen by PetChek (n=60), VetScan and SNAP RT detected antigen in 15 and 56 samples, respectively. Percent agreement (plus 95% confidence interval) for each test relative to the PetChek qualitative result was 50% (40-60%) for VetScan and 96% (89-98%) for SNAP RT. Relative to the presence or absence of female worms at necropsy, agreement was 61% (50-72%) for VetScan and 99% (92-99.6%) for SNAP RT.

It is clinically important that dogs with low heartworm burdens and/or low levels of circulating heartworm antigen are correctly identified by veterinarians, and the VetScan® rotor may not be an acceptable diagnostic methodology for these patients.

58 Efficacy of a Single Oral Administration of Milbemycin Oxime (Interceptor® Flavor Tabs® and Sentinel® Flavor Tabs®) Against Natural Infections of Ancylostoma braziliense in Dogs Stephen E. Bienhoff New Product Development, Novartis Animal Health US, Inc., Greensboro, NC

The objective of this randomized, blinded, placebo controlled laboratory study was to confirm the efficacy of a single oral administration of the marketed formulations of milbemycin oxime (Interceptor® Flavor Tabs® and Sentinel® Flavor Tabs®) at a dose of 0.23 mg/lb (0.5 mg/kg) against natural infections of Ancylostoma braziliense in dogs. Thirty-six hookworm infected dogs, a minimum of 10 weeks of age and of various breeds and genders were used. Fecal egg counts were done on 3 separate days prior to treatment for randomization purposes. Dogs were ranked by descending order of the fecal egg count arithmetic means and randomly assigned to treatment in blocks of 3 dogs each, 12 dogs per group (1 Interceptor, 1 Sentinel and 1 negative control group). Dogs were dosed according to the product label with blinding maintained by separation of function. Worm counts were done at necropsy 7 days after treatment in the same order as treatment group assignments. Reduction in A. braziliense in the treated groups were compared to the negative control group using analysis of varience of the A. braziliense geometric mean counts. Efficacy was defined as the ability of the test products to significantly (p≤ 0.05) reduce parasite load by 90% or greater in treated dogs when compared to adequately infected negative control dogs. The placebo control group had a geometric mean worm count of 19.2. The Interceptor treated group had a geometric mean worm count of 0.38 (p<0.0001) representing a 98% reduction in parasite load and the Sentinel treated group had a geometric mean worm count of 0.98 (p<0.0001) representing a 95% reduction in parasite load. In this study, Interceptor® Flavor Tabs® and Sentinel® Flavor Tabs®, administered at a minimum milbemycin oxime dose of 0.23 mg/lb (0.5 mg/kg), were efficacious for removing adult A. braziliense in dogs.

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59 The Efficacy of Milbemycin Oxime Against Migrating Pre-AdultSpirocerca lupi in Experi- mentally Infected Dogs Dawid J. Kok1*, Rudolph Schenker2 1ClinVet International (Pty) Ltd, Bloemfontein, South Africa, 2Novartis Animal Health, Basel

Introduction: Milbemycin oxime (MO) is known to be effective against a range of gastrointestinal roundworms, but its efficacy against Spirocerca lupi (S. lupi) has not been extensively investigated. This study evaluated the efficacy of MO against pre-adult stages ofS. lupi following the experimental infection of dogs.

Methods: Dogs, experimentally infected with L3 larvae recovered from scarabaeid beetles, were randomly assigned to treatment or control groups. MO, in a tablet formulation containing praziquantel (Milbemax®; Novartis Animal Health Inc), was administered to the treatment group dogs at a minimum dose of 0.5mg/kg/ bodyweight. MO was given once (day 30 post-infection), or repeatedly at 28-day or 14-day intervals (starting day 28 post-infection). Necropsy of all dogs took place 168 days after infection. Worms were recovered and counted, and any lesions attributable to S. lupi in the thoracic aorta and oesophagus, were described and quantified.

Results and Conclusions: All dogs were found to be adequately infected as confirmed by the extensive lesions within the thoracic aorta of both the control and treated animals. The treatment protocol was not expected to eliminate aortic damage as the first anthelmintic dose was given after parasite migration to the aorta would have occurred. The single treatment with MO demonstrated an efficacy of 79.8% in reducing oesophageal worm burden and strongly indicated that the anthelmintic is efficacious against pre-adult stages. Repeat treatments with MO, at either 14- or 28-day intervals, completely prevented establishment of S. lupi within the oesophagus and confirmed the efficacy of MO against migrating pre-adult larvae.

60 The Efficacy of Milbemycin Oxime to Protect Dogs Against Infection withSpirocerca lupi (Nematoda: Spirurida) in an Endemic Area Dawid J. Kok1*, Rudolph Schenker2 1ClinVet International (Pty) Ltd, Bloemfontein, South Africa, 2Novartis Animal Health, Basel

Introduction: Milbemycin oxime (MO) is known to be effective against a range of gastrointestinal roundworms, but its efficacy against Spirocerca lupi (S. lupi) has not been extensively investigated. This study evaluated the efficacy of a prophylactic treatment regimen with MO, to protect dogs, raised in an endemi- cally infected region of South Africa, against canine spirocercosis.

Methods: The pups were raised at their original homes in accordance with the normal husbandry procedures of their owners. They were alternately allocated to control and treatment groups. MO, in a tablet form containing praziquantel (Milbemax®; Novartis Animal Health Inc), was administered at a minimum dose of 0.5mg/kg bodyweight to all pups in the treatment group, when they were between 2 and 6 weeks old. Repeat doses were given on 5 occasions at intervals of approximately 28 days. At the end of the 6 month field phase, necropsies of the 31 treated and 27 untreated pups were conducted. Worms were recovered and counted, and any lesions attributable to S. lupi in the thoracic aorta and oesophagus, were described and quantified.

Results and Conclusions: The prophylactic treatment regimen prevented 86.5% of worms from becoming established in the thoracic aorta and 89.4% of worms from becoming established in the oesophagus. The

64 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA overall severity of infection was lower in the treatment group, with these animals showing a reduction in the grade of aortic lesions and a marked decrease in the number of aortic and oesophageal nodules. MO therefore deserves further consideration as a prophylactic treatment against S. lupi.

61 Attributes, Knowledge, Beliefs, and Behaviors Relating to Prevention of Heartworm Infec- tion among Members of a National Hunting Dog Club Sharron Patton*, Amanda Lutzy, Barton Rohrbach Department of Comparative medicine, University of Tennessee College of Veterinary Medicine, Knoxville, TN

A better understanding of the deficiencies in knowledge, beliefs, and behaviors of dog owners may help design better educational materials for heartworm prevention. Recently we completed a study of heartworm prevention practices used by commercial kennel owners, dog trainers, and dog owners that were members of a national hunting dog club (JAVMA 2010, In Press). This study raised additional questions about the knowledge, beliefs, and behaviors of dog owners concerning heartworm prevention. Respondents were asked to participate in a follow-up study to address these additional questions. The results suggest that the dog owners lack confidence in the accuracy of the heartworm test to identify infection, the efficacy of products sold for prevention of heartworm, and the effectiveness of treatment to eliminate heartworms from infected dogs. Knowledge about when to begin heartworm preventive medication in a new puppy, and the timing of heartworm tests was also lacking among a substantial number of respondents. To increase ac- ceptance of prophylaxis and reduce the likelihood of a false conclusion that prophylaxis failed, education of dog owners’ should focus on the importance of administering year-round heartworm prophylaxis and/or administering the last dose of monthly heartworm preventive after the last possible day transmission can occur, and the importance and timing of an annual heartworm test.

62 Obtaining a New isolate of Ancylostoma braziliense without the Need of Necropsy Janice L. Liotta1*, Alice C.Y. Lee1, Sharp Aksel1, Ibrahim Alkhalife2, Alejandro Cruz-Reyes3, Heejeong Youn4, Dwight Douglas Bowman4 1Microbiology & Immunology, Cornell University, Ithaca, NY, 2Department of Pathology, Parasitology, King Saud University, Riyadh,, 3Instituto de Biologfa, Universidad Nacional Aut=noma de MTxico, Mexico, 4Col- lege of Veterinary Medicine, Seoul National University, Seoul, South Korea

Isolation of a specific Ancylostoma species typically requires euthanasia of the source animal or holding an animal long enough to collect its feces after treatment for worm recovery and identification. The reason is that the eggs cannot be morphologically distinguished. In keeping with the 3 R’s of laboratory animal research (reduction, refinement, replacement), the objective of this study was to obtain an isolate of A. braziliense from one-time field-collected samples of canine feces for species identification and to isolate the strain without the need for the euthanasia of any animals. Fecal samples (n=152) were collected and identified as containing Ancylostoma eggs (n=66) while visiting the parasitology laboratory of the University of Florida, Gainesville: centrifugal sugar flotation, eggs from hookworm positive slide washed into tubes, DNA extracted, followed by PCR amplification and identification using RFLP with Hinf1 of samples containingA. braziliense (n=2). Larval cultures were initiated in Florida and driven to NY while split samples were shipped to Cornell. Larvae from the cultures initiated in FL were used to infect a laboratory reared kitten purchased and housed at Cornell for the purpose of inhibiting the growth of any contaminating A. caninum. The infection was patent at 15 days, and eggs were identified as A. braziliense by PCR/RFLP/DNA-sequencing. Using forceps and endoscopy, two adult worms (1 male, 1 female) were recovered from the cat and identified morphologically as A. braziliense. Larvae were cultured from the feces of this cat and used to infect a laboratory reared Beagle dog. Additionally, subsequent to treatment of the cat, worms recovered from the

65 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA sieved feces were A. braziliense but included one female A. caninum containing infertile eggs. The dog (patent 14 days post-infection) is being maintained as a source for further studies, and the cat has been placed in an adopted home.

Session 11 – Protozoa 63 Isospora suis and its association with post-weaning performance on three Ontario swine farms Andrew S. Peregrine*, Robert M. Friendship, Cate Dewey, Cory Todd, Andrea Aliaga-Leyton Population Medicine, Ontario Veterinary College, University of Guelph, Guelph, ON

In recent work, oocysts of Isospora suis were detected in the feces of piglets at 7-21 days of age on 35 of 50 (70%) swine farms in southwestern Ontario. Furthermore, litters of pigs infected with I. suis were significantly more likely to exhibit diarrhea than negative litters. However, while I. suis-positive farms had a mean standardized weaning weight that tended to be lighter than the weaning weight on non-infected farms, this was not significantly different. In order to determine whether piglets derived from litters infected withI. suis grow more slowly than piglets from non-infected litters in the post-weaned period, a prospective cohort study was conducted on a convenience sample of 3 swine farms in southwestern Ontario. Fecal samples were collected from 3 randomly selected piglets in each of 72 litters at 2, 3 and 5 weeks of age, and examined for oocysts using the Cornell-Wisconsin centrifugal floatation method. For all piglets in all 72 litters, weight was recorded at 1, 2, 3, 4, 5 and 8 weeks of age. If one or more piglets in a litter were found shedding I. suis oocysts at 2 or 3 weeks of age the litter was classified as infected. The percentage of litters infected with I. suis was 40% (4/10), 60% (18/30) and 84.4% (27/32) on the 3 farms. For the 678 piglets that were weighed, there were an average of 5.7 weight observations per animal. A linear mixed model with random intercepts was used to examine the effect of infection on weight gain. On average, pigs from infected litters were 1.4 kg (95% CI = 1.1–1.8 kg, P<0.001) lighter than pigs from non-infected litters at 62 days of age. Thus, infection with I. suis during the nursing stage was associated with significantly lower weights of pigs at 62 days of age.

64 Adaptive Evolution in Toxoplasma gondii ROP16 and ROP18 Genes Involved in Parasite Invasion Hany M. Elsheikha1*, Miao-Miao Liu2, Zi-Guo Yuan2, Rui-Qing Lin2, Xing-Quan Zhu2 1School of Veterinary Medicine and Science, University of Nottingham, Loughborough, 2Department of Parasitology, South China Agricultural University, Guangzhou

Toxoplasma gondii is a pathogenic protozoan parasite capable of infecting a wide range of warm-blooded vertebrate hosts including humans. DNA sequences of genes encoding rhoptry protein 16 (ROP16) and rhoptry protein 18 (ROP18) were obtained from T. gondii strains from different hosts and geographical locations. Nucleotide sequences were aligned using the ClustalX 1.81, and MP, NJ and ME trees were constructed using the software MEGA version 4.0. Also, sequences were used to identify sites and lineages subject to adaptive evolution. No statistically significant positive selection could be detected in the rhoptry 16 genes. However within the Rhoptry 18 genes the branch-sites methods identified a single branch of the tree with significant evidence (p < 0.05 after Bonferroni correction) of adaptive evolutiond N/dS > 1. Eighteen codons are detected as belonging to the dN/dS > 1 class of codons. These codons are dispersed throughout the gene with ten (297L 304S 363A 380K 383F 397K 441R 457A 512S 517D) residing in the Protein tyrosine kinase protein domain (positions 255-528, Pfam Pkinase_Tyr, PF07714) which give the rhoptry18 genes their protein kinase function. The identification of codons within the rhoptry gene with evidence of

66 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA positive selection suggests further evidence for this gene as important in the virulence capacity of this pathogen. It is perhaps unsurprising that some of the rhoptry genes are undergoing rapid evolution as these are at the forefront of competition between host and pathogen a classic arena for Darwinian evolution.

65 Extracellular pH Governs the Entry and Phenotypic Plasticity of Neospora caninum Hany M. Elsheikha School of Veterinary Medicine and Science, University of Nottingham, Loughborough

Neospora caninum, a strictly intracellular protozoan, is a major leading cause of parasitic-induced abortion in cattle. Despite this prevalence, very little is known about the pathogenesis of N. caninum infection. A widely held view of N. caninum infection is that both cellular proliferation and stage interconversion (tachyzoite-bradyzoite switching) are triggered, perhaps even modulated by, changes in culture conditions. The present study aimed to examine the effects of modification of the growth medium pH on N. caninum entry, proliferation and transformation in cultured K-652 and Vero cell lines. The entry pathway of N. caninum in K-562 cells was studied by using a cell membrane potential-sensitive fluorescent probe. Parasite invasion and proliferation of Vero cells was assessed by plaque formation assay. Stage transformation was studied using indirect double immunofluorescence and electron microscopy approaches. Findings indicate that a major pathway for N. caninum entry into the K-562 cell line is dependent on endocytosis and exposure to a low pH. Also, N. caninum proliferation in Vero cells was highest when pH level was optimum and parasite encystation increased when the pH level was alkaline or acidic. These data suggest that external pH has a determinable effect on the host cells and free N. caninum parasites and support the hypothesis that extracellular pH regulates the entry and phenotypic plasticity of N. caninum in mammalian cells.

66 Characterization of giardin protein expression during encystation of Giardia duodenalis Mark C. Jenkins*, Celia O’Brien, Dumitru Macarasin, Jeffrey Karns, Monica Santin-Duran Ronald Fayer, Animal Parasitic Diseases Laboratory, USDA/ARS, Beltsville, MD

Giardia duodenalis trophozoites attach to the gut surface by means of a ventral disk that contains various giardin proteins that appear to be important to VD structural integrity. One approach to preventing giardiasis is to stimulate giardin-specific antibodies and thereby block trophozoite attachment to the gut epithelium. Understanding giardin expression during encystation (cyst formation) or excystation (trophozoite release) might provide clues to the role of giardins in the Giardia life-cycle. In the present study expression of major giardin proteins during in vitro conversion of trophozoites to cysts was characterized. Quantitative RT-PCR using giardin-specific primers provided data on the expression of each respective gene sequence relative to house-keeping genes over time. Immunoblotting using giardin-specific antisera revealed the presence of beta- and delta-giardin at all timepoints during encystation. Moreover, immunofluorescence assay showed that both giardins were localized to a well- defined ventral disk early in encystation, and then was increas- ingly localized to an amorphous structure inside the cysts as early as 48 hr during encystation. Dual color labeling of G. duodenalis trophozoites with antisera to delta and beta giardin followed by confocal epiflu- orscence microscopy revealed the spatial organization of the two major giardin proteins in the trophozoite stage. These findings confirm earlier observations that the VD is disassembled during encystation, and provide interesting insight on the unique metabolic activities of G. duodenalis during encystation.

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67 Survival of Trypanosoma cruzi in Acai juice: Implications for food borne Chagas disease outbreaks David S. Lindsay* Biomedical Sciences and Pathobiology, Virginia Tech, Blacksburg, VA

American trypanosomiasis (Chagas disease) is a neglected tropical disease and has a potential to emerge in the US. Approximately 10 million people are infected with Trypanosoma cruzi and an additional 90 million are at risk of developing this often debilitating disease. The insect vector normally transmits the infection after feeding on the host and defecating near the feeding site. Infective stages then enter the wound and es- tablish infection. Outbreaks of T. cruzi associated with ingestion of Aca’ juice have been reported in Brazil. These outbreaks are characterized by having a large number of acute infections from a common source. It is assumed that the insect vectors are accidentally ground up in the preparation of the juice products or that the food is contaminated with anal gland secretions from South American opossums. The present study examined the survival of cell culture derived trypomastigotes of the Brazil strain T. cruzi in commercially purchased Aca’ juice (2 types), distilled water, tap water, and artificial gastric juice (acid-pepsin solution). Viability of treated trypomastigotes was assayed by inoculation of treated trypomastigotes on to bovine monocyte (BM) cell cultures. Trypomastigotes treated for 30 minutes at room temperature (RT) in tap or distilled water, and artificial gastric juice were not infectious for BM cells. Amastigotes were observed in BM cells inoculated with trypomastigotes treated for 30 minutes at RT in Aca’ juice indicating that treated trypomastigotes were infectious supporting field observations on outbreaks of acute Chagas disease caused by ingestion of contaminated Aca’ juice. Supported by grants from the Office of Research and Graduate Studies, and the Department of Biomedical Sciences and Pathobiology, VA-MDRCVM, Virginia Tech.

68 Morphometrics of Assemblages of Giardia duodenalis Cysts from the Feces of Dogs and Cats Dwight D. Bowman1, Stephanie B. Yager1, Britta A. Okyere1, Bo Li1, Kyuhyung D. Kang1, Heejeong Youn2, Marissa Karpoff1, Hussni O. Mohammed1, Araceli Lucio-Forster1, Janice L. Liotta1 1Microbiology and Immunology, Cornell University, Ithaca, NY, 2College of Veterinary Medicine, Seoul Na- tional University, Seoul, South Korea

Initially, area, length, width, and eccentricity of cysts from 7 dogs and 9 cats, isolated using MgSO4 flotation, were compared. Images (1000x) of 100 cysts from each animal were measured. DNA sequences from cysts from each animal was characterized as to Assemblage at the Triosephosphate Isomerase (TPI) and β-giardin (BG) gene loci. In the TPI locus: for canine cysts, one isolate was identified as Assemblage B, while the rest were identified as Assemblage C; for feline cysts, two isolates were Assemblage A, three Assemblage B, one Assemblage C, and three Assemblage F. Using the BG locus, the canine Assemblage C isolates were assigned to Assemblage D, and one feline Assemblage B isolate was assigned to As- semblage F; all others agreed with the TPI locus assignment. Using general linear models for both the TPI and BG metrics, the means of length, width, and area were significantly different between the different Assemblages, with cysts of Assemblage B being smaller than those of Assemblages A, F, C and D. As an additional test of the potential morphological distinctness of recovered cysts, 10 cysts from 6 cats (BG As- semblage determination: 4 F, 1 A, 2B) and 4 dogs (BG Assemblage: D) collected by ZnSO4 flotation and measured in a similar fashion before Assemblage determination. Assemblage B cysts were significantly smaller than those of Assemblage D, while Assemblage F cysts were significantly smaller than those of As- semblage D cysts and larger (although not significantly) than those of Assemblage B. The 10 Assemblage A cysts from one cat were highly variable in size and overlapped the distributions of the other Assemblage types. Overall, there are morphometric differences in the Assemblages of cysts, but it remains unknown to what extent these Assemblage traits are influenced by host factors.

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69 A new Besnoitia from the southern plains woodrat Neotoma micropus) J. P. Dubey1*, M.J. Yabsley2 1Animal Parasitic Disease Laboratory, United States Department of Agriculture, Beltsville, USA, Beltsville, MD, 2University of Georgia, Athens, GA

Certain species of the protozoan genus Besnoitia cause clinical disease in livestock and wildlife. A new species of Besnoitia was found from the southern planes woodrat (Neotoma micropus). The parasite was detected by bioassay of woodrat tissues in out-bred Swiss Webster mice in an attempt to isolate Toxoplas- ma gondii. Initially, the organism was misdiagnosed as T. gondii because it was highly pathogenic for mice and its tachyzoites resembled T. gondii tachyzoites. Further studies revealed that it differed structurally and biologically from T. gondii. Tachyzoites were successfully cultivated and maintained in vitro in bovine monocytes and African green monkey kidney cells, and in vivo in mice. Non-dividing, uninucleate tachyzoites were approximately 1 x 5 μm in size. Longitudinally-cut bradyzoites in tissue sections measured 1.5-1.6 x 7.7-9.3 μm. Tissue cysts were microscopic, up to 210 μm long, and were infective orally to mice. Cats fed tissue cysts shed unsporulated 13 x 14 μm sized oocysts. All mice inoculated with the wood rat Besnoitia died of acute besnoitiosis, irrespective of the dose, and Norwegian rats became infected but remained asymptomatic. Entero-epithelial stages (schizonts, gamonts) were found in cats fed tissue cysts. Large (up to 40 x 50 μm) first generation schizonts developed in the lamina propria of the small intestine of cats. A second generation of small sized (8 μm) schizont containing 4-8 merozoites was detected in enterocytes of small intestine. Gamonts and oocysts were seen in goblet cells of the small intestinal epithelium. Tachyzo- ites were present in mesenteric lymph nodes of cats. Phylogenetic analysis indicated that the wood rat Bes- noitia was related to other Besnoitia species from rodents, rabbits, and opossums. The woodrat Besnoitia is distinct from the three other species of Besnoitia, B. wallacei, B. darlingi, and B. oryctofelisi that utilize cats as a definitive host.

Session 12 - Interesting clinical cases 70 Equine abortion caused by Encephalitozoon cuniculi Karen F. Snowden1*, Travis Heskitt2, Barbara Sheppard2 1Veterinary Pathobiology, Texas A&M University College of Veterinary Medicine, College Station, TX, 2Infec- tious Diseases and Pathology, University of Florida, Gainesville, FL

The eukaryotic single-celled microsporidian parasite, Encephalitozoon cuniculi, is well documented as a pathogen in rabbits and a variety of rodents. The organism has occasionally been documented in a variety of other mammals, most importantly in domestic dogs, fur foxes and humans. Transplacental transmission of parasites has been shown experimentally in rabbits and dogs. In this case we document an early stage abortion in a horse caused by this parasite. A thoroughbred mare spontaneously aborted a fetus within the amniotic sac approximately 3 months after the last breeding date. Gross necropsy findings included yellow- brown fluid within the amnionic membrane and within the chorion. The body of the fetus, weighing 287 gm appeared to be moderately autolyzed. Histological evaluation of multiple layers of the fetal membranes showed marked necrosis, vasculitis and inflammation and included intracellular gram positive organisms consistent with microsporidial spores. Histological sections of fetal brain and a section of fetal skin also contained the organisms. The identity of the intracellular organisms was confirmed as Encephalitozoon based on ultrastructural morphology. Additionally the parasite was confirmed asE. cuniculi genotype III using PCR amplification and sequencing of portions of the parasite ribosomal RNA genes. Interestingly, this genotype has previously been documented only in dog and rare human infections. This is the fifth reported case of Encephalitozoon-associated abortions in horses.

69 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

71 Unusual Cause of Ectoparasitic Pruritus in a Horse Araceli Lucio-Forster1*, Mary C. Smith2, Ann Georgi Leonard3, Dwight D. Bowman1 1Microbiology and Immunoloy, Cornell University, Ithaca, NY, 2Population Medicine and Diagnostic Sci- ences, Cornell University, Ithaca, NY, 3Ithaca Artisan, Freeville, NY

Ectoparasites including biting lice (Damalinia equi), sucking lice (Haematopinus asini), and mites (e.g. Cho- rioptes bovis), among others, are relatively common causes of pruritus in horses. On occasion the chicken mite, Dermanyssus gallinae, has been associated with dermatitis in horses. Here we present a suspected case of Dermanyssus gallinae dermatitis in an 8 year old mare. The mare was one of five horses stabled in the same barn, but was the only animal affected. The observant owner noted the presence of roosting chickens over the animal’s stable and contacted her veterinarian regarding the possibility that poultry mites could be involved. Clinical signs consisted of severe pruritus; no hair loss or self-inflicted lesions were present. A skin scraping was performed the day after signs were first noticed by the owner. Upon examination poultry mites were confirmed, however, these were identified asOrnithonyssus sylviarum, and not D. gallinae. A single treatment with an insecticide-repellent solution containing pyrethrin, permethrin and piperonyl butoxide technical, along with the removal of the chickens from the rafters above the horse’s stable were successful in resolving the infestation.

72 Aberrant migration of Dirofilaria immitis in three dogs - case report Rhonda Pinckney1*, Tara E Paterson2 1Pathobiology, St. George’s University, School of Veterinary Medicine, True Blue, 2Clinical Studies, St. George’s University, School of Veterinary Medicine, True Blue

Dirofilaria immitisis a pathogenic filarial nematode that infects many mammalian species, with the infection in dogs and cats being of greatest importance to the small animal veterinarian. The worms most commonly reside in the pulmonary arteries where they complete their development into mature adults. In rare cases, the larvae migrate aberrantly through the body, ending up in sites other than the cardiovascular system. This report discusses three unusual cases of D. immitis aberrant migration in dogs that presented to St. George’s University School of Veterinary Medicine on the island of Grenada, West Indies. Two cases involved identification of D. immitis worms in the peritoneal cavity during routine ovariohysterectomy. In the third case, two immature male worms were expressed from the bladder. Additionally, there was a case of an adult worm being expelled in inflammatory respiratory exudates by a dog undergoing heartworm treatment with melarsomine dihydrochloride. While this is not a true case of aberrant migration, it is uncommon and noteworthy. Identification of live heartworms following successful aberrant migration is rare. To the author’s knowledge, this is the first report of aberrant heartworm migration in dogs from the Caribbean – an endemic heartworm region.

73 Autochtonal Dirofilaria repens in a dog in The Netherlands Paul A.M. Overgaauw1*, Evert P. Van Dijk2 1Veterinary Public Health, Institute for Risk Assessment Sciences, Utrecht University, The Netherlands, Utrecht, 2Veterinary Clinic De Arker, Nijker, The Netherlands

In November 2008 a 1.5 year old female English Bulldog was presented with a painful skin lump of 1 cm diameter on her leg. The nodule contained a few white small (ca. 2-4 mm) worms of approximately 10 cm length. The dog had never been abroad, but was taken for camping 6 months earlier in an area in the middle of The Netherlands where many mosquitoes were present. Microscopic examination showed nematodes

70 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA filled with oval eggs with moving larvae. Based on the morphologic characters the diagnosis wasDirofilaria repens. This was confirmed by PCR of the worm. Microfilaria were absent in the blood.D. repens is a zoonotic filarial of the subcutaneous tissue of carnivores (dogs, cats, foxes) in Southern and Eastern Europe, Africa and Asia. Another subgenus is D. immitis (heartworm). D. repens is ovoviviparous and the larvae are transferred as microfilaria through blood-sucking mosquitoes. The disease can cause cutaneous disorders, such as pruritus, dermal swelling and subcutanous nodules containing the parasites. The prepatent period in the dog is 180 to 240 days. Our patient probably acquired the infection during camping. Adult worms can be removed by surgical excision. To prevent infection, mosquitoes should be controlled and dogs treated with anti-filarial anthelmintics, such as moxidectin, selamectin, and milbemycin. The number of zoonoticD. repens infections has increased in the last few decades and the infection may be considered as an emerging zoonosis. The infection is spreading probably as a consequence of climatic changes, together with an increase in the movement of cats and dogs across Europe. This is the first autochtonal case of D. repens infection in The Netherlands, which may be a warning for other North European countries that the infection may be spreading in this direction.

74 Uncommon Complications in a Dog with Dirofilaria immitis Jennifer E. Carter1*, Guillaume Chanoit2, Cheryl Kata3 1Clinical Sciences, Ross University School of Veterinary Medicine, St. Kitts, 2Clinical Sciences, North Caro- lina State University, Raleigh, NC, 3Molecular and Biomedical Sciences, North Carolina State University, Raleigh, NC

Case description- A 7-year-old 23-kg (50.6-lb) spayed female Border Collie with a history of heartworm disease was evaluated for respiratory distress due to bilateral pneumothorax.

Clinical findings-Computed tomography imaging revealed possible pulmonary bullae or blebs and, based on those findings, a tentative diagnosis of bullous emphysema was made. Exploratory median sternotomy revealed gross pathology in the right caudal lung lobe. Upon excision of the lung lobe, a nematode segment was noticed both in the dog and in the removed lobe, indicating inadvertent surgical transection. At this time, the dog experienced an anaphylactoid reaction. The reaction was successfully treated with fluid therapy, antihistamines, and steroids. During the post-operative period the dog developed a hemothorax and was returned to surgery. As no obvious cause was seen at the second surgery the dog was treated for a potential coagulopathy. Parasitological examination of the nematode segment confirmed the diagnosis of Dirofiliaria immitis.

Treatment and outcome- Lung lobectomy of the right caudal lobe was performed with clinical resolution of the bilateral pneumothorax. A second surgical procedure and treatment with fresh frozen plasma provided clinical resolution of the hemothorax. The dog was discharged 5 days post-operatively and treated with 30 days of doxycycline and monthly heartworm preventative.

Clinical relevance- This report describes pneumothorax secondary to heartworm disease as well as the management of an anaphylactoid reaction associated with the accidental dissection of the adult heart worm. These are unusual complications associated with Dirofilaria immitis infection in the dog.

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75 Babesia canis rossi Infection in a Texas Dog Mason V. Reichard1*, Robin W. Allison1, Todd J. Yeagley2 1Department of Veterinary Pathobiology, Oklahoma State University, Stillwater, OK, 2Veterinary Clinical Sci- ences, Oklahoma State University, Stillwater, OK

A 5-month old intact male Boerboel canine, imported from South Africa one week previously, was presented to a Texas veterinarian for lethargy, anorexia and labored breathing. The dog was febrile (104.5°F; 40.3°C), anemic (HCT 23%) and leukopenic (3400/µL), with an increased BUN (46 mg/dL). An enzyme immunoassay (SNAP-4Dx, IDEXX Laboratories) was negative for Dirofilaria immitis antigen and antibodies against Ehrlichia canis, Borrelia burgdorferi, and Anaplasma phagocytophilum. An EDTA blood sample was sent to Oklahoma State University Center for Veterinary Health Sciences, where a CBC revealed non-regenerative anemia (HCT 19%, reticulocytes 40,000/µL), neutropenia (1900/µL) and large protozoal piroplasms within 0.7% of erythrocytes. Piroplasms were 2-5 µm long and varied in shape from round to oval to piriform, and merozoites were also observed extracellularly. A nested PCR was performed on DNA extracted from blood using primers that amplify the 18s rRNA gene from all known canine Babesia species, and the product sequenced. BLAST analysis of the 432 base sequence revealed 99-100% similarity to B. canis rossi, 93% similarity to B. canis canis, and 92-93% similarity to B. canis vogeli. The dog responded well to treatment with imidocarb. PCR analysis of a second blood sample 2 weeks later was negative for Babesia spp. DNA. To our knowledge, this is the first diagnosis of B. canis rossi infection in the United States.

76 Hepatic alveolar echinococcosis in a dog in British Columbia, Canada Andrew S. Peregrine1*, Emily Jenkins2, Brian Barnes3, Shannon Johnson4, Janet Hill2, Lydden Polley2 Ian K. Barker1, Bruno Gottstein5 1Pathobiology, Ontario Veterinary College, University of Guelph, Guelph, ON, 2Veterinary Microbiology, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK, 3Westview Veteri- nary Hospital, Powell River, BC, 4Idexx Laboratories, Markham, ON, 5Institut for Parasitologie, UniversitSt Bern, Bern

In July 2009 a 3.5-year old neutered male Shi Tzu/Bichon cross presented to a small animal practice in Powell River, British Columbia (BC), with a 10-day history of intermittent vomiting and lethargy. Palpation revealed a large firm mass in the cranial aspect of the abdomen. Ultrasound demonstrated a 12-13cm diameter hepatic mass with mixed echogenicity, cavitated lesions within, and an irregular surface. Explor- atory laparotomy revealed a discrete multi-lobulated firm mass that appeared to originate in the left medial liver lobe, was firmly adherent to the greater curvature of the stomach, and was adherent to the left lateral liver lobe, spleen and omentum. No abnormalities were detected in other abdominal organs. In total, a 570 g mass was resected that involved the left medial liver lobe. Following surgery, the dog made an uneventful recovery. In light of the following diagnosis, life-long treatment with albendazole was recommended for the dog.

Grossly, the hepatic mass contained multiple coalescing white nodules that were poorly demarcated and appeared to infiltrate adjacent hepatic tissue. Histologically, the mass contained multilocular cystic struc- tures surrounded by fibrosis; multiple intraluminal protoscolices were present that were occasionally surrounded by a thin membrane. Sequence data of PCR amplificates for the mitochondrial 12S rRNA gene, and RFLP analysis of mitochondrial NADH dehydrogenase 1 and ribosomal DNA genes, indicated that the parasite responsible for the mass was Echinococcus multilocularis. Direct immunofluorescence with MAbG11-FITC confirmed this diagnosis. Comparison of sequences at multiple mitochondrial loci with published sequences worldwide identified the BCE. multilocularis isolate as European in origin. Since the dog

72 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA had never travelled outside BC, this suggested that the infection must have been acquired within BC. This would appear to be the first diagnosis of E. multilocularis in BC. Furthermore, the genotype data suggest that a European strain has established within the province.

77 The use of ponazuril to treat a Toxoplasma gondii outbreak in a zoo setting Jennifer A. Spencer1*, Leah A. Kuhnt1, John F. Roberts1, Byron L. Blagburn1, Bishop Thompson2 1Pathobiology, College of Veterinary Medicine, Auburn, AL, 2Sparks State Diagnostic Laboratory, Auburn, AL

Toxoplasma gondii is a highly virulent protozoan parasite in lemurs, Australian marsupials and New World monkeys. Several outbreaks with high mortality rates have been reported in zoological parks. Affected animals can exhibit lethargy, respiratory signs, neurological deficits or die before clinical signs become apparent. Animals can die before producing antibodies that can be detected by conventional means, making antemortem diagnosis difficult. Outbreaks are not usually detected until the pathology results on the first case are received. Treatment of toxoplasmosis with traditional drugs has met with varying degrees of success as neither sulfadiazine-pyrimethamine or clindamycin is able to completely eliminate the rapidly dividing tachyzoite stages. Deaths have been reported in both juvenile and adult susceptible animals with or without treatment.

Recently, the Gulf Coast Zoo in Alabama suffered the loss of six animals over a four week period. The species affected included two kangaroos, two marmosets, one Patagonian cavy and one wallaby. Tissue samples were sent to the Thompson-Bishop-Sparks State Diagnostic Laboratory, where a tentative diagnosis of Toxoplasma gondii was made based on histopathological findings. These findings were confirmed by polymerase chain reaction. We decided to attempt to treat all susceptible species with the off-label use of ponazuril (Marquis®, Bayer) and this was done with the approval of their veterinarian and the director of the zoo. All susceptible species were treated twice a day with 10mg/kg for 14 days. After treatment com- menced, there were no further deaths due to T. gondii. This is a possible management tool that can be implemented in other zoos experiencing T. gondii outbreaks.

78 Parasitologic Pet Peeves Craig R. Reinemeyer1*, Wendell L. Davis2 1East Tennessee Clinical Research, Inc., Rockwood, TN, 2Animal Health, Bayer HealthCare LLC, Shaw- nee, KS

Like all responsible scientists, veterinary parasitologists strive to communicate clearly and accurately. Ha- bitual, erroneous word usage, however, often prevents us from achieving this lofty standard. Imprecise parasitologic communication occurs in many forms, but the four categories to be discussed in this presentation include: 1) pseudotaxonomy, 2) incorrect word forms, 3) inappropriate word substitutions, and 4) anthropomorphism in veterinary literature. Examples of pseudotaxonomy include inappropriate use of common terms, such as “roundworms” as a synonym for members of the order Ascaridida. All nematodes are arguably “roundworms”, having cylindrical bodies and being circular in cross section. Other examples of pseudotaxonomy include fabricated names for various parasitic organisms at the genus and family level. Selected, incorrect word forms will be identified and examined, including the use of “preventative” for “preventive” and “euthanize” for “euthanatize”. Another abuse occurs when terms that are not synonyms are used interchangeably (e.g., “EPG” substituted for “fecal egg counts”; and “dose”, “dose rate” or “dose level” used as synonyms for “dosage”). Lastly, reviewers and editors of veterinary journals constantly remind authors that certain human medical terms cannot be adopted for veterinary medicine. One example is that “symptoms” are subjective; what the patient experiences during an illness, and as such are not usu-

73 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA ally visible to others. In contrast, “signs” can be felt, heard, seen, or measured by a diagnostician. Clearly, veterinarians can detect signs in animals, but not symptoms. Similar distinctions can be made for the terms “depression” vs. “lethargy”. With these examples as a starting point, it is hoped that critical evaluation of our speaking and writing habits will improve the standards of communication in veterinary parasitology.

Session 13 - Anthelmintic efficacy /resistance 79 Pharmacological Characterization of New Cholinergic Anthelmintics in C. elegans Timothy G. Geary1*, Charles Viau2, Elizabeth Ruiz Lancheros1, Abdel Francis3, Tita N. Walter1 1Institute of Parasitology, McGill University, Ste-Anne-de-Bellevue, QC, 2Department of Microbiology and Immunology, McGill University, Ste-Anne-de-Bellevue, QC, 3McGill University, McGill University, Ste-Anne- de-Bellevue, QC

The nematode Caenorhabditis elegans is a useful model for characterizing anthelmintic pharmacology. Currently used and in-development anthelmintics have various modes of action, including the ability to bind to nicotinic acetylcholine receptors (nAChRs). Classic anthelmintic agonists of these receptors (nicotine, levamisole, pyrantel and bephenium) caused intact specimens of C. elegans to undergo contracted paralysis. The nAChR antagonist mecamylamine had flaccid paralytic effects on intact worms and also blocked the actions of the agonists. The potency and time to onset of effect of these drugs were enhanced when the worms were bisected (‘cut worm’ preparation). In contrast to these observations, the novel anthelmintic cholinergic antagonist 2-desoxoparaherquamide (derquantel) was only weakly active in intact specimens of C. elegans at concentrations of 50 uM over several days. No antagonism of nAChR agonists was observed with this drug in intact worms. However, 2-desoxoparaherquamide had direct effects on motility in cut worms and blocked the effects of the nAChR agonists in this preparation. In contrast, a representative of the novel amidoacetonitrile derivatives (AAD) anthelmintic class also paralyzed cut worms, but these effects were not antagonized by 2-desoxoparaherquamide. Instead, cut worms were paralyzed faster in the presence of both drugs than in the presence of either alone. These data confirm that 2-desoxoparaherquamide is a nicotinic antagonist in C. elegans and suggest that cuticular permeability properties of this free-living nematode may be different from those of adult parasitic species. These results also suggest that these two new cholinergic anthelmintics, one an agonist and one an antagonist, may not be therapeutically incompatible.

80 Anti-Parasitic Efficacy of Herb Extracts on Ovine, Equine, and Canine Strongylid Eggs and Larvae Heejeong Youn1*, Kyonghee Kim1, Yeongsuk Lim1, Kyongeun Lee1, Bongkyun Park1, Janice L. Liotta2, Clement Alawa3, Araceli Lucio-Forster2, Dwight D. Bowman2 1College of Veterinary Medicine, Seoul National University, Seoul, South Korea, 2Microbiology and Im- munoloy, Cornell University, Ithaca, NY, 3National Animal Production Research Institute, Ahmadu Bello University, Shika-Zaria, Nigeria

This worked examined herbal materials as alternatives to various existing anti-helminthic products for the control of nematodiasis in domestic animals. Efficacy of extracts from 9 different herbs, Artemisia annua (An), Artemisia asiatica (As), Bupleurum chinense (Bc), Dichroa febrifuga (Df), Gleditsia japonica (Gj), Pulsatila koreana (Pk), Sinomenium acutum (Sa), Sophora flavescens (Sf) and Torillis japonica (Tj), were tested against ovine, equine, and canine strongylid eggs and larvae. Strongylid eggs were collected from

74 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA ovine, equine, and canine fecal samples, and larvae were cultured as needed from these eggs. Adult Hae- monchus contortus were also collected for eggs from the abomasa of sheep from a local slaughter facility. At high concentrations (1/400 – 1/800) of As (68hrs), Bc (40hrs), Df (88hrs), Pk (64hrs) and Sf (40hrs), the ovine and equine strongylid eggs and larvae died or became inactive before those of control groups (120 hrs). At low concentrations (1/1,600 – 1/3,200) of An (164hrs), Gj (332hrs) and Tj (264hrs), these eggs and larvae lived longer, and larvae were more active than those of control groups. Hookworm larvae were more resistant to the medicinal herb extracts; dying within 1 day at the 1/120 dilution, but surviving over 9 days in 1/240 dilution of all herb extracts. Further research on the above herbal materials will be carried out after HPLC fractionation and chemical analysis of the extracts.

81 On the Mode of Action of Tribendimidine Alan P. Robertson1*, John A Carr2, Sreekanth Puttachary1, Santosh Pandey2, Richard J. Martin1 1Biomedical Sciences, Iowa State University, Ames, IA, 2Electrical and Computer Engineering, Iowa State University, Ames, IA

Tribendimidine is a new anthelmintic from China and has been licensed for use in humans. Its mode of action in parasitic nematodes has yet to be fully elucidated. The mode of action has been partially characterized using molecular experiments in the free living nematode C. elegans. In C. elegans tribendimidine is an agonist of a nicotinic acetylcholine receptor subtype on somatic muscle, as are other established anthelmintics like levamisole and pyrantel. Previous studies from our lab have demonstrated that the nAChRs on nematode muscle show important differences between species. In brief, the nAChR on C. elegans muscle has different pharmacological and single-channel properties to the nAChRs on Ascaris muscle. We have investigated the mode of action of tribendimidine using functional studies in Ascaris suum and Oesopha- gostomum dentatum. We used standard muscle contraction, current-clamp and larval migration inhibition assays to investigate the effect of this drug. In Ascaris muscle strips tribendimidine produced concentration- dependent increases in muscle tension similar to those produced by levamisole, although tribendimidine was more potent with an EC50 of 0.2 µM. Using the current-clamp technique, tribendimidine produced muscle cell depolarization and increases in input conductance- similar in nature to those produced by levamisole and pyrantel. The EC50 for tribendimidine induced depolarization was 0.7 µM. The membrane potential and input conductance effects of tribendimidine were antagonized by mecamylamine confirming it as a nicotinic acetylcholine receptor agonist. We have also observed that tribendimidine inhibits larval migration in O. dentatum larvae and this effect is under investigation. Current work is focusing on determining the specific selectivity of tribendimidine for parasite nAChR subtypes.

82 Investigating Candidate Resistance Genes in an Ivermectin-resistant Isolate of Haemon- chus contortus. Sally M. Williamson1*, Gerald C. Coles2, Samantha McCavera3, Adrian J. Wolstenholme1 1Infectious Diseases, University of Georgia College of Veterinary Medicine, Athens, GA, 2University of Bris- tol, Bristol, 3Biology & Biochemistry, University of Bath, Bath

Anthelmintic resistance is well documented in veterinary parasites, and resistance to the macrocyclic lactones in Haemonchus contortus, a gastrointestinal parasite of ruminants, is a particularly acute problem. The genetic polymorphism exhibited by Haemonchus populations found in the field has created considerable difficulties when attempting to make a direct comparison of drug resistant and susceptible parasites at the molecular level. In this study we have used a Haemonchus contortus isolate selected for ivermectin resistance from a susceptible ISE isolate in only 4 generations, minimising the confounding effects of genetic variability which can complicate analysis of candidate resistance genes. An interesting and unexpected finding of this study showed that the ivermectin-resistant population was also resistant to thiabendazole

75 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA in the egg-hatch test and possessed an increased level of benzimidazole resistance-associated alleles (17.7% vs 0.3 % in the parent population) at position 200 of the β-tubulin gene. The mechanism by which this cross-resistance could arise is unknown, as the parasites were not exposed to benzimidazoles at any point during the selection process. To investigate potential molecular changes associated with ivermectin resistance in the Haemonchus isolate, we have focused on candidate resistance genes identified in other studies, namely the P-glycoproteins and the ligand-gated chloride channels. We used the genome sequence of Haemonchus contortus to search for P-glycoprotein sequences, which were then assigned a putative identity based on sequence similarity to PGP genes in C. elegans. Partial P-glycoprotein sequences were amplified, and expression levels of PGP mRNA were compared between the resistant isolate and the parent ISE isolate using real-time quantitive PCR. Full-length sequences of ligand-gated chloride channels implicated in ivermectin sensitivity and resistance were also amplified from both resistant and susceptible parasites; these sequences were directly compared to identify any sequence changes in the resistant isolate which may be linked to ivermectin resistance.

83 Multiple Anthelmintic Resistance on a Llama Farm in the Southeastern United States Bob E. Storey1*, Sue B. Howell2, Anand N. Vidyashankar3, Lisa H. Williamson4, Ray M. Kaplan1 1Department of Infectious Diseases, University of Georgia, Athens, GA, 2Infectious Diseases, College of Veterinary Medicine, Athens, GA, 3Department of Statistics, George Mason University, Fairfax, VA, 4Depart- ment of Large Animal Medicine, University of Georgia CVM, Athens, GA

Anthelmintic resistance on goat and sheep farms in the southeastern United States is widely documented; however, very little is known about anthelmintic resistance on camelid farms. A study was conducted on a llama farm in Florida to investigate the resistance status of fenbendazole, levamisole, ivermectin, and moxidectin using both the fecal egg count reduction test (FECRT) and DrenchRite® larval development assay (LDA). Seventy-two llamas were allocated randomly into six treatment groups (n=12/group) balanced by fecal egg count (FEC). Coprocultures were performed before and after treatment in all groups to identify species of nematodes present. Statistical analysis was carried out using logistic and probit regression after accounting for overdispersion. Model adjusted efficacies were calculated for each species separately and overall. Analysis using overall FEC showed no statistical difference between untreated control, fenbendazole, and ivermectin treatment groups, however species-specific analysis demonstrated 100% efficacy for fenbendazole in Trichostongylus colubriformis (Tc). Levamisole yielded efficacies of 96.5%, 81.3%, and 96.2% against Haemonchus contortus (Hc), Tc and Nematodirus spp. (Ns), respectively. Moxidectin oral had efficacies of 83.1%, 91.3%, and 79.3% against Hc, Tc, and Ns respectively; whereas, moxidectin injectable demonstrated efficacies of 82.4%, 87.5%, and 83.7% against Hc, Tc, and Ns, respectively., Confidence intervals for efficacy for oral and subcutaneous moxidectin overlapped indicating there were no significant differences between routes of administration. Using WAAVP guidelines, these results indicate resistance in one or more species to all drugs tested. The LDA data indicated resistance for benzimidazoles and ivermectin, and sensitivity to levamisole and moxidectin. These findings confirm the presence of multiple anthelmintic resistance on a llama farm in the southeastern US. Furthermore, the incongruent results for moxidectin in the LDA and FECRT suggest that the dose applied likely is not adequate. Further research is required to assess the prevalence of anthelmintic resistance on camelid farms in US.

76 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

Session 14 – Helminth efficacy / diagnosis/ impact 84 A Pilot Study on the Effect of an Integrated Control Program of Fasciola hepatica in Caja- marca, Peru Francisco Raunelli1*, Sergio Gonzalez1, Jorge Guerrero2 1Foncreagro, Cajamarca2Department of Pathobiology, University of Pennsylvania, Philadelphia, PA

Fasciolosis is recognized as the major health problem of dairy cattle production in the Cajamarca region of Peru. This disease produces the highest economic losses on cattle productivity as well as severely impacts the health of the rural human population of that area. In order to determine the effects of an integrated parasite control program on parasite prevalence and dairy cattle production a pilot study was conducted in this northern region of Peru. The study was conducted over a 2-year period in 21 rural settlements in the dis- trict of La Encañada, province of Cajamarca, and involved 1727 dairy cattle producers and 8000 heads of cattle. This program involved 3 oral triclabendazole treatments per year, strategically timed according to the epidemiological cycle of the parasite. Treatments were administered to all cattle at each of the participating sites together with snail control activities aimed at modifying the angles of irrigation ditches and paddock drainage. The treatment program was supplemented by training and a technical assistance program aimed to educate producers. The initial prevalence of F hepatica in the cattle was 63.2%. At the end of the study only 13.6% of the cattle tested were infected, representing a reduction of more than 49% in prevalence. The mean live weight of the animals in the study increased from 245.60 to 339.09 kg, an increase of 38%. Analysis of milk production was performed on milk samples collected from all cattle in the study one month following each treatment. Monthly milk production increased from a total of 125,453 kg to 219,689 kg, an increase of 75% over the 2 years of the study.

85 Coprological Evaluation of Pour-on and Injectable Formulation of Moxidectin in Beef Cattle J.G. Powell1*, S.A. Gunter2, C.A. Tucker3, J.L. Reynolds3, Z.B. Johnson3 1Animal Science Department, University of Arkansas, Fayetteville, AR, 2USDA ARS, Woodward, OK, 3Uni- vesity of Arkansas, Fayetteville, AR

Two, 28d studies were conducted with naturally infected beef calves to assess the effect of possible allo- grooming behavior on fecal egg count reductions and coproculture counts following treatment with moxidectin. The first study was conducted at the USDA Southern Plains Experimental Range near Ft. Supply, OK (n=137; initial BW=253±19 kg), and the second study was conducted at the University of Arkansas Stocker Cattle Unit in Fayetteville, AR (n=80; initial BW=226±38kg). Calves at both locations were allocated to one of three treatments: injectable moxidectin, pour-on moxidectin, and a negative control. Calves were also assigned to one of five different pasture arrangements consisting of: 1) control calves only, 2) injectable calves only, 3) pour-on calves only, 4) pour-on and control calves, and 5) injectable and control calves. Stocking rates were 1hd/1.25ha, and 1hd/0.15ha at the Oklahoma and Arkansas locations, respectively. At each location, fecal egg and coproculture counts were similar between groups on d 0; however, there was a difference for location effect (P<0.0001), indicating cattle at the Arkansas location exhibited higher FEC on days 0, 14 and 28 compared to calves at the Oklahoma location. Results from each site indicate that control calves pastured with pour-on treated calves did not exhibit reduction in FEC from possible allo-grooming.

77 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

However, at both locations, regardless of pasture assignment, fecal egg count reductions were greatest for pour-on treated calves (P<0.05). Also at both locations, calves that were treated with moxidectin developed higher (P<0.05) percentages of coproculture L3 as Cooperia compared to control calves. 86 The Effects of Cooperia punctata on cattle productivity Bert E. Stromberg1*, Louis C. Gasbarre2, Audie Waite3, David T. Bechtol3, Michael S. Brown4, Nicholas Robinson5, Erik Olson5, Harold Newcomb6 1College of Veterinary Medicine, University of Minnesota, St. Paul, MN, 2Gasbarre Consulting, Buffalo, WY, 33Agri Research Center, Inc., Canyon, TX, 4West Texas A&M University, Canyon, TX, 5University of Min- nesota, St. Paul, MN, 6Intervet/Schering-Plough Animal Health, DeSoto, KS

Cooperia spp. have become the most prevalent parasite in US cow/calf operations as observed in the NAHMS Beef Cow/Calf survey in 2008. The effects of Cooperia sp. on cattle productivity are largely unknown, and this study was conducted to assess such effects. Two hundred calves (average weight 209kg) were acquired from Northwestern Arkansas and Northeastern Oklahoma and were vaccinated and dewormed upon arrival. Animals were preconditioned for approximately one month and fed a standard growing ration. At 4 weeks, all calves were dewormed, re-vaccinated, moved to pens equipped with GrowSafe® system feed bunks and given an additional week to acclimate. Calves were randomly divided into two groups (n=80) and each group was further divided into two replicate pens (n=40). On day 0 and 14, two pens were orally inoculated with a gavage of ~1X105 Cooperia punctata infective larvae, with the control pens receiving tap water. Data collected included biweekly fecal egg counts, daily individual feed con- sumption and weight gain over the 60 day test period. The presence of Cooperia punctata (>99%) was confirmed by necropsy on days 35 and 60 post infection (PI). Egg counts were positive by day 14 PI and remained at levels similar to values seen in field studies. The sham-treated group gained weight 7.4% more rapidly (p=0.02) than inoculated animals (3.24 lb vs 3.0 lb ADG, respectively). The inoculated animals also consumed 1.5 lb (DMI) less compared to sham treatment (p = 0.02). Data suggested that C. punctata has a deleterious affect on both appetite and nutrient uptake or utilization. Mesenteric lymph nodes were increased in size and significant gut thickening and mucus production were noted. Treatment of one group (40) of infected calves with an endectocide did not remove the parasites (FECRT = 8.8%), while treatment with a benzimidazole was effective (FECRT = 98.1%).

87 Modification and further evaluation of a fluorescein-labeled peanut agglutinin test for identification ofHaemonchus contortus ova in fecal samples and findings of note since the inception of offering the diagnostic test Janell K. Bishop-Stewart1*, ME Jurasek2, JA Hall2, JM O’Hara2, J. Snyder3, ML Kent4 1Veterinary Diagnostic Lab Bacteriology, Oregon State University College of Veterinary Medicine, Corvallis, OR, 2Oregon State University College of Veterinary Medicine, Corvallis, OR, 3Myrtle Veterinary Hospital, Myrtle Point, OR, 4Oregon State University College of Veterinary Medicine, Dept. of Micrbiology, Corvallis, OR

Gastrointestinal nematodes of the family Trichostrongylidae are recognized to be some of the parasites that are most pathogenic to sheep, goats and other ruminants worldwide. A method to distinguish Haemonchus ova from other trichostrongyle-type ova was adapted from previous work by Palmer and colleagues. We employed their method of staining eggs with a peanut lectin conjugated to fluorescein isothiocyanate, which has been proven to be very specific for the surface ofHaemonchus eggs. However, we established a much less labor-intensive method of collecting the ova for this test. This has allowed this diagnostic test to be offered to clients through the Oregon State University College of Veterinary Medicine Veterinary Diagnostic Laboratory and University of Georgia Veterinary Diagnostic Laboratory for approximately a year now. Some

78 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA intriguing observations have been made with samples from sheep, goats and llamas in Oregon and other northern areas of the United States. Haemonchus is generally regarded as a “seasonal” parasite, with little evidence of active infection during the colder months of the year in northern climates. Our data from clinical specimens and research animals collected through out the year indicates that Haemonchus has less seasonality in these more northern areas than previously thought.

88 Comparison of a McMasters Chamber with increased detection sensitivity to the Stoll and Modified Wisconsin Fecal Egg Count Methods Sue B. Howell*, Bob E. Storey, Ray M. Kaplan Department of Infectious Diseases, University of Georgia, Athens, GA

The current methods of choice for performing fecal egg counts (FEC) on equine, bovine and camelid species when sensitive detection is desired are either the Stoll or Wisconsin methods. However, these methods require one or two centrifugation steps respectively, making these procedures more time consuming when compared to the McMaster’s technique. Modification of the traditional McMaster’s counting chamber to include a third chamber, and increasing the area of the counting grid can greatly increase the detection sensitivity. Recently, at the request of our laboratory, such a chamber was manufactured by Chalex Corporation. Using fecal samples from horses, cattle and llamas, we evaluated the consistency of data and the time necessary to perform FEC using this new modified McMaster chamber at a detection level of 8.3 eggs per gram (epg) as compared to the Stoll and Wisconsin techniques at a detection level of 5 epg. Experiments were repeated three times, and 5 FEC were performed on each subsample of feces, yielding 15 FEC for each species and method. Results indicate equal or improved detection sensitivity on both horse and camelid samples with the new McMaster’s chamber. The time required to perform the centrifugation methods ranged from 35-45 minutes per sample, compared to 15 minutes for the McMaster’s. The McMaster yielded reduced detection with cattle feces, but this appeared to be due a severe darkening of the fluid resulting from reaction of the sodium nitrate with an unknown component in the feces, causing reduced egg visibility. Our study demonstrated that egg detection and consistency between individual FEC measurements using this new chamber was equal to or improved as compared to the more time consuming centrifugation methods.

Session 15 - President’s symposium 89 Molecular Mechanisms of Resistance : Nicotinic and Macrocyclic Lactone Anthelmintics Richard J. Martin*, Alan P. Robertson Biomedical Sciences, Iowa State University, Ames, IA

The classes of anti-nematode drugs include: 1) nicotinic agonists (levamisole, pyrantel, bephenium, oxantel, monepantel); 2) nicotinic antagonists (derquantel); 3) Macrocyclic lactones/GluCl allosteric modulators (avermectins: ivermectin, milbemycins: moxidectin); 4) β-tubulin ligands (benzimidazoles: thiabendazole, mebendazole, flubendazole, albendazole); 5) Slo-1 K channel activators (emodepside); 6) GABAago- nists (piperazine); 7) Chitinase inhibitors/ionophores (closantel); 8) Pyruvate oxireductase inhibitors (ni- tazoxanide). Our knowledge of the molecular mechanisms of resistance to the anthelmintics of classes 1-4 has increased recently. We will review molecular mechanisms of resistance to nicotinic anthelmintics

79 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA and overlapping areas of macrocyclic lactone anthelmintics. The development of resistance in parasitic nematodes, which may be polygenic, has been associated with the loss of redundant anthelmintic-sensitive receptor subtypes and increased excretion via drug transporters.

Different nicotinic acetylcholine receptor channel (nAChR) subtypes, sensitive to different nicotinic anthelmintics, occur on nerves and muscle of nematode parasites. There is a B-subtype on muscle, which is more sensitive to bephenium and derquantel; there is the L-subtype nAChR on muscle, which is more sensitive to levamisole; there is the N-subtype on muscle, which is more sensitive to oxantel; and there are the DEG-3 subtypes that are more sensitive to monepantel. A reduced number of functioning anthelmintic sensitive subtypes can explain resistance in parasitic nematodes; these subtypes are, apparently, redundant receptor subtypes. For resistance to levamisole there may be loss of the L-subtype without loss of other subtypes. The reduction in the number of functioning sensitive receptor subtypes has been associated with expression of truncated of alternatively spliced receptor channel subunits or reduced levels of RNA transcripts of receptor channel subunits. The identified genes include:unc-63, unc-29, unc-38 and acr-8 for levamisole and mpt-1, des-2H and deg-3H for monepantel.

The macrocyclic lactones appear to target a number of different ligand-gated chloride channels of parasitic nematodes, including some gated by glutamate, GABA and dopamine. A number of channel subunit genes: avr-14, avr-15, glc-1 of the GluCls receptors, lgc-37 a GABA subunit, and ggr-3 a dopamine chloride channel may be involved in resistance. In parasitic nematodes, P-glycoprotein multi-drug transporters have also been implicated in anthelmintic resistance. Several ABC-transporter genes, including pgp-1 and pgp-2 as well as genes involved in glutathione synthesis can increase their expression following exposure to ivermectin and moxidectin.

In summary, the molecular mechanism of resistance depends upon the anthelmintic and is likely to be polygenic. Molecular diagnosis may require looking for a number of specific changes as well as expression levels and combinations of different subtype selective anthelmintics are predicted to slow the development resistance.

90 Molecular mechanisms of resistance to benzimidazole and macrocyclic lactone anthelmintics Roger K. Prichard Institute of Parasitology, McGill University, Ste-Anne-de-Bellevue, QC

Anthelmintic resistance in nematode parasites of livestock is becoming an increasing problem. Resistance results from the genetic selection of parasites able to survive exposure to an anthelmintic and pass on their resistance genes, while susceptible parasites are killed and unable to pass on their genes to offspring. With repeated treatment the frequency of resistance alleles increases until a resistance phenotype can be observed. Understanding the genes responsible for resistance, or linked with resistance, can allow us to monitor for the development of resistance before it affects production and spreads. An understanding of the resistance genes can also explain the resistance, so that actions can be taken to overcome the resistance or slow its selection and allow rational choices of alternative anthelmintics, drug combinations and treatment rotations.

Changes in β-tubulin are responsible for resistance to benzimidazole (BZ) anthelmintics. Resistance to macrocyclic lactones (MLs) is less clearly understood and appears to be multigenic. There is strong evidence that some ABC transporter genes may be involved in ML resistance. There are many ABC transporter genes (P-glycoproteins, Multidrug Resistance Proteins, Half transporters) in nematodes and not all of them will be involved in ML resistance. Furthermore, different MLs are transported to different extents by a transporter, so that there can be differences between MLs in resistance development and mechanisms.

80 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA

MLs act on ligand-gated chloride channels (LGCCs). However, this class of cys-loop receptors is also quite diverse in nematodes. Again different MLs will affect different LGCCs. Changes in relevant LGCCs may also contribute to ML resistance. Finally, there is evidence that MLs select on β-tubulin. The genetic changes involved in ML resistance can be due to changes in expression level or to loss of sensitivity. Further work is needed to fully understand the mechanisms and genetics of resistance to different MLs.

91 Cattle Internal Parasitism and Deworming Effectiveness from the 2008 USDA NAHMS Beef Cow/Calf Study Bert E. Stromberg1*, Louis C. Gasbarre2, Lora R. Ballweber3, David A. Dargatz4, Judy M. Rodriguez4, Dante S. Zarlenga5 1College of Veterinary Medicine, University of Minnesota, St. Paul, MN, 2Gasbarre Consulting, Buffalo, WY, 3Colorado State University, Fort Collins, CO, 4VS Centers for Epidemiology and Animal Health, USDA, APHIS, Fort Collins, CO, 5Animal Parasitic Diseases Lab, USDA, ARS, Beltsville, MD

During the USDA National Animal Health Monitoring System’s (NAHMS) Beef 2007-08 study 567 producers from 24 states were offered the opportunity to collect fecal samples from weaned calves for evaluation of parasite burden (Phase 1). There was also an opportunity to evaluate the response to treatment with an anthelmintic product (Phase 2). Producers choosing to participate in one or both parts were provided with instructions and materials to collect fecal samples. Up to 20 fresh fecal samples from weaned beef calves were collected prior to treatment (Phase 1) and for those that participated in Phase 2, again approximately 2 weeks after anthelmintic treatment. Fresh fecal samples were submitted to one of 3 laboratories for evaluation. In the laboratories all samples were processed in a similar manner using the Modified Wisconsin technique for the enumeration of strongyle, Nematodirus, and Trichuris eggs, and the notation of the presence or absence of coccidian oocysts and tapeworm eggs. In submissions where the eggs per gram value for strongyle eggs exceeded 30, aliquots from 2-6 animals were pooled for extraction of DNA from the eggs. Extracted DNA was subjected to PCR analysis for the presence of Ostertagia, Cooperia, Haemonchus, and Oesophagostomum. In Phase 1 85.6% of the operations had calves shedding strongyle type eggs. Overall, for pooled samples evaluated, 88% had Cooperia, 79% Ostertagia, 52% Haemonchus, 38% Oesophagos- tomum, 18% Nematodirus and 7% Trichuris. In Phase 2, 31.2% of all the operations tested had efficacies less than 80%, 13.1% had efficacies between 80 and 90% and 55.7% had efficacies greater than 90%. After treatment the proportion of Cooperia positive pooled samples increased to 95%, while the proportion of samples positive for other genera all decreased. Lack of efficacy may have been due to improper and incomplete treatment or reduced efficacy of the compounds utilized.

American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA Author Index Abstract Abstract Abstract A C Eversole, Rob...... 27 Ahn, KyuSung...... 39 Campbell, Tyler A...... 1 Aksel, Sharp...... 62 Carr, John A...... 81 F Fayer, Ronald...... 66 Alawa, Clement...... 80 Carter, Jennifer E...... 74 Fletcher, JA...... 18 Aliaga-Leyton, Andrea...... 63 Chanoit, Guillaume...... 74 Foster, Joyce G...... 24 Alkhalife, Ibrahim...... 62 Chapman, Hilary David...... 11 Francis, Abdel...... 79 Allen, Kelly...... 19, 20 Charles, Roxanne A...... 28 Friendship, Robert M...... 63 Allison, Robin W...... 75 Chavez, Deanna J...... 46 Fuller, A. Lorraine...... 12 Ambrose, Dana...... 7 Cho, Kyung-Oh...... 39 Arther, Robert...... 32 Chungpivat, Sudchit...... 41 Coles, Gerald C...... 82 G B Conboy, Gary...... 44, 52, 54 Gadde, Ujvala Deepthi...... 11 Gasbarre, Louis C...... 86, 91 Ball, Cathy...... 33 Conner, Mike...... 6 Geary, Timothy G...... 56, 79 Ballweber, Lora R...... 26, 46, 91 Cook, Joshua O...... 7 Gerhold, Richard W...... 9, 12 Barker, Ian K...... 76 Correa, Maria T...... 55 Gleim, Elizabeth R...... 6 Barnes, Brian...... 76 Cosgrove, Sallie...... 42 Gonzalez, Sergio...... 84 Barr, Stephen C...... 21 Cruthers, Larry...... 32 Goodwin, David G...... 21, 23 Baum, Kristen A...... 50 Cruz-Reyes, Alejandro...... 62 Goolsby, John A...... 1 Beall, Melissa J...... 57 Cyr, Tracy L...... 47 Goring, Jonas K...... 40 Bechtol, David T...... 86 Gottstein, Bruno...... 76 Beck, Margaret...... 45 D Gould, Jessica C...... 30 Beckmen, Kimberlee B...... 26 Dargatz, David A...... 91 Gourley, Michelle L...... 27 Beckstead, Robert...... 9 Davey, Ronald B...... 1 Gross, Sheila...... 33 Bellosa, Mary L...... 41 Davis, Cheryl D...... 45 Grove, Daniel M...... 5 Beringer, Jeff...... 5 Davis, Wendell L...... 78 Guerrero, Felix D...... 1 Bevins, Sarah...... 5 Davis, Wendell...... 32 Guerrero, Jorge...... 84 Bienhoff, Stephen E...... 58 de Leon, Beto Perez...... 1 Gunter, S.A...... 85 Bishop-Stewart, Janell K...... 87 De Wolf, Bradley D...... 22 Blagburn, Byron L...... 32, 33, del Mar Nieto, Prixia...... 43 37, 56, 77 Dewey, Cate...... 63 H Blizzard, Emily L...... 45 Dillon, Ray...... 57 Hall, JA...... 87 Boardman, David S...... 55 Dimon, Danielle E...... 15 Heise, Stephanie R...... 4 Boonmars, Thidarat...... 29 Dryden, M...... 34, 49 Heise, Stephanie...... 20 Bourguinat, Catherine...... 56 Dubey, J. P...... 21, 41, 51, 69 Henke, Scott...... 45 Bowles, Joy V...... 33 Duhaime, Roberta...... 1 Heskitt, Travis...... 70 Bowman, Dwight D...... 13, 41, 57, Duncan, Natalie D...... 55 Hewitt, David G...... 1 68, 71, 80 Dzimianski, Mike T...... 17 Hill, Janet...... 76 Bowman, Dwight Douglas...... 62 Hobson, M...... 49 Brake, Danett K...... 1 E Holman, Patricia J...... 48 Holman, Patricia...... 1 Brown, Holly M...... 5 Edwards, Amy...... 54 Houk, Alice E...... 21 Brown, Michael S...... 86 Ellis, Angela E...... 28 Howell, Sue B...... 83, 88 Burke, Casey...... 24 Ellis, Dee...... 1 Hrubec, Terry...... 23 Burke, Joan M...... 25 Elsheikha, Hany M...... 64, 65 Hughes, Daymond W...... 5 Butfiloski, Jay...... 5 Erf, Gisela F...... 11 Butler, Jamie M...... 32, 33 Evans, Christopher Charles...... 17

83 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA Author Index Abstract Abstract Abstract Lin, Rui-Qing...... 64 J Lindsay, David S...... 21, 23, 67 N Jansen, Jocelyn T...... 22 Linton, Ashley...... 26 Nettles, Victor F...... 5 Jenkins, Emily...... 76 Liotta, Janice L.. 41, 57, 62, 68, 80 Neumann, Brian...... 8 Jenkins, Mark C...... 66 Little, Susan E...... 19 Neumann, Norwood R...... 13 Johnson, Eileen M...... 19 Little, Susan...... 20, 52, 54 Newcomb, Harold...... 86 Johnson, Shannon...... 76 Liu, Miao-Miao...... 64 Johnson, Z.B...... 85 Lockhart, Mitchell...... 5 O Jones, Andria Q...... 22 Lohmeyer, Kimberly H...... 1 O’Brien, Celia...... 66 Jurasek, ME...... 87 Lollis, Lori A...... 10 O’Hara, JM...... 87 Long, David B...... 45 O’Hara, Todd M...... 26 K Lorentzen, Leif...... 41 Oh, DaeSung...... 39 Kaltenboeck, Bernhard...... 33 Lucio-Forster, Araceli...41, 57, 68, Oh, Seok-Il...... 39 Kammlah, Diane M...... 1 71, 80 Okyere, Britta A...... 68 Kang, Kyuhyung D...... 68 Lutzy, Amanda...... 61 Olfenbuttel, Colleen...... 5 Kania, Steven...... 42 Lynn, Elizabeth Carolyn...... 9 Olson, Erik...... 86 Kapil, Sanjay...... 19 Ortega, J. Alfonso...... 1 Kaplan, Ray M...... 7, 14, 15, 16, M Overgaauw, Paul A.M...... 73 17, 83, 88 Macarasin, Dumitru...... 66 Overvelde, Sebastien...... 40 Karns, Jeffrey...... 66 Mackenzie, Charles...... 27, 3 Karpoff, Marissa...... 68 MacTavish, Jennifer...... 22 P Kata, Cheryl...... 74 Malone, Jb...... 18 Panciera, Roger...... 20 Keller, Kathy...... 56 Malone, John B...... 29, 43 Pandey, Santosh...... 81 Kent, ML...... 87 Manthei, Dawn E...... 13 Paras, Kelsey...... 20 Ketzis, Jennifer...... 32 Marcotte, Lynne...... 40 Park, Bongkyun...... 80 Khampoosa, Panita...... 29 Martin, Richard J...... 81, 89 Pas, An...... 51 Kim, Kyonghee...... 80 McBride, A...... 34, 49 Paterson, Tara E...... 72 King, Vickie...... 42 McCarroll, Jc...... 18 Patton, Laura L...... 5 Klein, Bradley...... 23 McCarroll, Jennifer C...... 29 Patton, Sharron...... 61 Knowles, Donald...... 2 McCavera, Samantha...... 82 Payne, P...... 34, 49 Kok, Dawid J...... 59, 60 McDougald, Larry R...... 12, 9 Peek, Matthew...... 5 Krecek, Tammi...... 52 Meeus, Patrick...... 42 Peregrine, Andrew S.....22, 63, 76 Kuhnt, Leah A...... 77 Menzies, Paula I...... 22 Peterson, David S...... 5 Kulsantiwong, Jutharat...... 29 Messenger, Matthew...... 1 Phillips, Pamela L...... 1 Miller, James E...... 25 Pinckney, Rhonda...... 72 L Miller, Michael W...... 46 Piratae, Supawadee...... 29 Lagrow, Cari...... 42 Miller, Robert J...... 1 Polley, Lydden...... 76 Lancheros, Elizabeth Ruiz...... 79 Mischler, Paula D...... 38 Pornmingmas, Pitcha...... 41 Lee, Alice C.Y...... 13, 41, 57, 62 Mohammed, Hussni O...... 68 Pound, J. Mathews...... 1 Lee, Kyongeun...... 80 Moorehead, Andy R...... 17 Powell, J.G...... 85 Leonard, Ann Georgi...... 71 Moorhead, Andrew R.....7, 14, 55 Prado, Julio C...... 31 LeVan, Ivy...... 46 Mosjidis, Jorge A...... 25 Prange, Suzanne...... 5 Levin, Michael...... 6 Mosley, Courtney...... 8 Prasopdee, Sattrachai...... 29 Li, Andrew Y...... 1 Mount, Jane D...... 33 Prichard, Roger K...... 56, 90 Li, Bo...... 68 Murdock, Jessica H...... 7 Pulaski, Cassan...... 18 Lim, Yeongsuk...... 80 Murphy, Staci M...... 5 Puttachary, Sreekanth...... 81

84 American Association of Veterinary Parasitologists 55thAnnual Meeting Loews Atlanta Midtown Hotel, Atlanta, GA Author Index Abstract Abstract Abstract Storey, Bob E...... 16, 17, 83, 88 Williamson, Sally M...... 82 R Strobl, Jeannine...... 23 Wilson, Sherry...... 33 Racelis, Alex E...... 1 Stromberg, Bert E...... 86, 91 Wolstenholme, Adrian J...... 82 Rajendran, C...... 51 Su, C...... 51 Wongsaroj, Thitima...... 29 Rathinam, Thilakar...... 11 Sukchan, Somsak...... 29 Raunelli, Francisco...... 84 Sukhumavasi, Woraporn...... 41 Reichard, Mason V...... 50, 75 Y Suwannatrai, Apiporn...... 29 Yabsley, M.J...... 28, 5, 6, 69 Reichard, Mason...... 20 Suwannatrai, Kulwadee...... 29 Yabsley, Michael J...... 45 Reinemeyer, Craig R...... 31, 78 Suwanwerakamtorn, Rasamee.29 Yager, Stephanie B...... 68 Reinemeyer, Craig...... 42 Yeagley, Todd J...... 75 Reiskind, Michael...... 20 Youn, Heejeong...... 62, 68, 80 Rendon, Daniel A. Zarate...... 16 T Tarbsripair, Pairat...... 29 Yuan, Zi-Guo...... 64 Reynolds, J.L...... 85 Taylor, Amanda...... 44 Ritchie, D...... 34 Teel, Pete...... 1 Roberts, John F...... 77 Z Temeyer, Kevin B...... 1 Robertson, Alan P...... 81, 89 Zajac, Anne M...... 8, 21, 23, 24 Terrell, Thomas H...... 25 Robinson, Nicholas...... 86 Zarlenga, Dante S...... 91 Tesana, Smarn...... 29 Rodriguez, Judy M...... 91 Zercher, Adrienne B...... 7 Thammasiri, Chalida...... 29 Rogers, Ashley M...... 28 Zhu, Xing-Quan...... 64 Thompson, Bishop...... 77 Rohrbach, Barton...... 61 Todd, Cory...... 63 Rossano, Mary G...... 30 Towner, Jennifer N...... 14 Rust, Michael K...... 37 Tucker, C.A...... 85 S Salman, Mo D...... 26 U Ulrich, Michael A...... 13 Santin-Duran, Monica...... 66 Usher, Ruth D...... 55 Schenker, Rudolph...... 56, 59, 60 Schroeder, Iris...... 32, 37 Schuster, Greta...... 1 V Settje, Terry...... 32 Vaala, Wendy E...... 31 Sheppard, Barbara...... 70 Van Dijk, Evert P...... 73 Shin, SungShik...... 39 Varner, Kevin P...... 1 Shock, Barbara C...... 5 Viau, Charles...... 79 Shock, Philip M...... 5 Vidyashankar, Anand N...... 83 Smith, Katherine F...... 28 Villeneuve, Alain...... 40 Smith, Mary C...... 71 Smith, V...... 34, 49 W Snowden, Karen F... 52, 53, 54, 70 Wagner, G. Gale...... 1 Snyder, Daniel E...... 35, 36 Waite, Audie...... 86 Snyder, J...... 87 Walter, Tita N...... 79 Soliz, Liza...... 1 Wang, Chengming...... 33 Spencer, Jennifer A...... 77 West, Misti...... 20 Spencer, TB...... 18 Weston, Sarah...... 37 Stallknecht, David...... 7 Whitley, Niki C...... 25 Starkey, Lindsay...... 20 Wikel, Stephen...... 1 Stewart, Tonya A...... 44 Williamson, Lisa H...... 83

MEMBERSHIP DIRECTORY 2010

Membership Directory

Elizabeth M. Abbott Kelly E. Allen Robert G. Arther Lora Rickard Ballweber Abbott Associates Dene Oklahoma State University, Bayer Health Care, Animal Coloradoi State University, Lodge, Husbands Bosworth Coll. Vet. Med. Health Coll. Vet. Med Lutterworth, UK LE17 6NL 250 McElroy Hall, CVHS 12809 Shawnee Mission 1619 Campus Delivery 44 185 888 1050 Stillwater, OK 74078 Parkway Fort Collins, CO 80523 44 185 888 1151 405-744-3236 Shawnee, KS 66216 970-491-5015 emabbott@ecoanimalhealth. kelly.allen.warren@okstate. 913-268-2503 [email protected] com; ema@abbottassoc. edu 913-268-2541 demon.co.uk [email protected] Akinsanya Bamidele Lynn Allen Univ. of Lagos, Dept. Zoology Albert Abraham Novartis Animal Health Marjory Artzer P.O. Box 216 Bayer HealthCare 1722 SW 300 Kansas State University Akoka, Yaba, Lagos, Nigeria P.O. Box 390 Kingsville, MO 64061 1800 Denison, Coleo Hall 2341 Shawnee Mission, KS 66201 816-597-3497 Manhattan, KS 3182 234 080 2345 9087 913-268-2860 816-597-3497 785-532-4611 [email protected] [email protected] [email protected] [email protected] Stephen C. Barr Alex D.W. Acholonu Maria Sonia Almeria Seyed M. Atighechi Cornell University, Coll. Vet. Alcorn State University Autonomous University Y Disinfection Research Center Med. 1000 ASU Drive, #843 Barcelona, Veterinary School, No. 18 Abshoori Ave, Shahr- Dept. Clinical Sciences Alcorn State, MS 39096 Parasitology ara Str. Ithaca, NY 14853 601-877-6236 Campus UAB, Edifici V, 08193 Tehran, Iran 14456 607-253-3043 601-877-2328 Bellaterra [email protected] 607-253-3534 [email protected] Barcelona, Spain 8193 [email protected] [email protected] 011 34 649 808414 Rupinderjit Avapal 011 34 93 587 2006 Veterinary Parasitology Verona A. Barr Benjamin AduAddai [email protected] (P.A.U.) Heartland Community College Michigan State University 28 Sestina Crt. 1500 W. Raab Road Pathobiology & Diagnostic Mohammad Al-Sabi Brampton, Ontario, Canada Normal, IL 61761 Investigation Section of Zoology L6P1R9 309-268-8667 A42 Veterinary Medical center Throvaidsensvej 40 905-789-6158 [email protected] East Lansing, MI 48824 Copenheagen, Denmark 1871 [email protected] 517-353-9070 453 533-2672 John R. Barta 517-432-5836 [email protected] David G. Baker University of Guelph, Ontario [email protected] Louisiana State University Vet. Coll. Ulla Andersen 1424 Westchester Dr. Dept. Patholobiology Dalen Agnew Royal Vet. And Agriculture Baton Rouge, LA 70810 Guelph, Ontario, Canada N1G Michigan state University, University 225-578-9643 2W1 DCPAH Klostermarken 69, Skjem 225-578-9649 519-824-4120, Ext. 54017 4125 Beaumont Road, Room Denmark, The Netherlands [email protected] jbarta@uoguelph,ca 152B 6900 East Lansing, MI 48810 [email protected]; ullava@ Norman F. Baker William Barton 517-432-5806 life.ku.dk 7586 Nunes Road Teva Animal Health, Inc. 517-432-5836 Dixon, CA 95620 3915 S. 48th St. Terrace [email protected] Prema Arasu 707-675-5007 St. Joseph, MO 64503-4711 North Carolina State Univer- 816-676-6152 Armando Aguilar-Caballero sity, Coll. Vet. Med. Norman L. Baldwin 816-676-6874 Universidad Autonoma de 4700 Hillsborough Street Baldwin Aquatics, Inc [email protected] Yucatan Raleigh, NC 27606 12480 Pedee Creek Road 17 no. 93 X 18 y 20 919-513 6530 Monmouth, OR 97361-9541 Dieter Barutzki KanasinXX, Mexico 97370 919-513 6455 503-838-2600 Veterinay Laboratory Freiburg +52 999 9423200 [email protected] 503-838-2600 Wendlinger Str. 34 +52 999 9423205 norman.baldwin@bigfoot. Freiburg 1.BR. 79111 [email protected] James J. Arends com;norm@irvecastdesign. (49761) 459-9780 S&J Farms Animal Health com [email protected] Lesel Ali-De Silva 2340 Sanders Road P.O. Box 1323 Willow Springs, NC 27592 Cathy A. Ball Sophie Beaulieu Port-of-SpainXX, Mexico 919-894-5684 Summit VetPharm 9 Rang Du Rocher 868-766-0272 919-894-9010 301 Route 1704, Floor 12 St-Étienne de Bolton, Quebec, 868-522-4908 [email protected] Rutherford, NJ 7070 Canada J0E 2E0 [email protected] 201-242-8412 877-918-2948 201-585-9525 [email protected] [email protected]

89 Brenda Beerntsen Byron L. Blagburn Emanuele Brianti Joseph W. Camp, Jr. University of Missouri Auburn University, Coll. Vet. University of Messina, Sch. Purdue University, Veterinary 209 Connaway Hall Med., Dept. Pathobiology Vet. Med. Pathobiology Columbi, MO 65211 122 Green Hall Polo Universitario Bella 725 Harrison St. 573-882-5033 Auburn University, AL 36849- Annunziata, Messina, Italy West Lafayette, IN 47906 [email protected] 5519 98168 765-496-2463 334-844-2702 0039 090 355922 [email protected] Thomas R. Bello 334-844-2652 [email protected] Sandhill Equine Center [email protected] Bill R. Campbell POB 1313 Mary Lou Brongo Piedmont Pharmaceuticals, Southern Pines, NC 28388 David L. Bledsoe 617 Kenmore Road LLC 910-692-6016 Farnam Companies, Inc. Brick, NJ 8723 204 Muirs Chapel Rd, Suite 910-949-4559 301 West Osborn Rd. 732-477-9440 200 [email protected] Phoenix, AZ 85013-3997 [email protected] Greensboro, NC 27410 602-664-1249 336-544-0320, X203 Mary L. Bellosa 602-207-2145 Dave Bromert 336-544-0322 Cornell University [email protected] Intervet Schering-Plough bill.campbell@piedmontphar- College of Vet Med 649 Butternut Court ma.com Ithaca, NY 14853 Venkata Boppana Liberty, MO 64068 518-926-9095 University Conn. Health [email protected] W. C. Campbell [email protected] Center Drew University 211 Main Street, Apt D3 Holly Brown Dept. Biology, 36 Madison Ave Gerald W. Benz Farmington, CT 6032 University of Georgia Madison, NJ 7940 830 Gabbettville Rd. 860-751-8882 310 King Ave 973-408-3096 LaGrange, GA 30240 [email protected] Athens, GA 30606 973-408-3504 706-645-1999 [email protected] [email protected] 706-645-1999 Dwight D. Bowman [email protected] Cornell University, Coll. Vet. Martha Brown Mary Ellen Carino Med. Pfizer Animal Health University of Georgia Harold Berger C4-119 VMC/Micro & Immunol 601 W. Cornhusker Hwy. 235 Milledge Terrace 54 Dawson Lane Ithaca, NY 14853-6401 Malcolm, NE 68521 Athens, GA 30606 Jamesburg, NJ 08831-2662 607-253-3406 402-441-2165 [email protected] 607-253-4077 402-441-2782 Christine Berthelin-Baker [email protected] [email protected] Doug Carithers Merial Ltd Merial 115 Transtech Drive Louise A. Box-Hutchinson Michelle Bryant 3239 Satellite Blvd, Bldg. 500 Athens, GA 30601 M.D. University of Wyoming Duluth, GA 30096 [email protected] P.O. Box 29349 PO Box 764 678-638-3837, Mobile 770- San Antonio, TX 78229-0349 Casper, WY 82602 331-6069 Everett D. Besch 210-684-0060 307-259-8044 678-638-3817 1453 Ashland Drive [email protected] [email protected] Baton Rouge, LA 70806-7838 Richard E. Bradley 225-927-6352 4981 Ganung Drive Jagdeep K. Buch Justin Carter Morganton, NC 28655 Bayer Health Care TRS Labs, Inc Troy A. Bettner 828-430-8695 PO Box 390 PO Box 5112 Elanco Animal Health [email protected] Shawnee, KS 66210-0390 Athens, GA 30604 PO Box 708 913-268-2035 706-549-0764 Greenfield, IN 46140 Matt Brewer 913-268-2541 706-353-3590 317-433-2870 University of Wisconsin [email protected] [email protected] 317-276-4471 107 ½ Washington St Eau Claire, WI 54701 William W. Burdett Lori Carter Frederic Beugent 715-460-0258 Intervet Inc. Stillmeadow, Inc Merial [email protected] 8066 N. 130th Road 12852 Park One Drive 26 Av Tony Garnier Cairo, NE 68824 Sugar Land, TX 77478 Lyon, France 69007 Holly E. Brianceau 308-379-3261 281-240-8828 3368 774-8983 Intervet, Inc. 308-485-4321 281-240-8448 [email protected] 29160 Intervet Lane, P.O. Box [email protected] [email protected] 318 Ellen Binder Millsboro, DE 19966-0318 Casey Burke Richard J. Cawthorn Virginia Tech, VMRCVM 302-934-4232 Virginia Tech University University of Prince Edward Duckpond Dr., Ph 2, Rm 214 302-933-4008 159 Liberty Lane Island, Atlantic Vet. Coll. Blacksburg, VA 24061 [email protected] Pembroke, VA 24136 AVC Lobster Sci. Ctr. 540-231-6471 540-778-3055 Charlotteetown, PEI, C1A 4P3 [email protected] [email protected] Canada 902-566-0584 902-566-0851 [email protected]

90 Karen C. Ceragioli (Cun- Sreekumar Chirukandoth Gerald C. Coles Roberto Cortinas ningham) APDL, ANRI, BARC-EAST, University of Bristol, Dept. University of Minnesota, Dept. Merial Ltd Bldg. 1001 Clin. Vet. Sci. of Vet. & Biomedical Sciences 3239 Satellite Blvd 10300 Baltimore Ave. Langford House 1971 Commonwealth Ave, Duluth, GA 30096 Beltsville, MD 20705-2350 Langford House, Bristol, Vet. Science 205 678-638-3138 301-504-5999 United Kingdom BS40 5DU St. Paul, MN 55108 karen.cunningham@merial. 301-504-9222 44-117-928-9418 612-625-8078 com [email protected] 44-117-928-9504 612-625-5203 [email protected] [email protected] H. David Chapman Bill Chobotar University of Arkansas, Dept. Andrews University Douglas D. Colwell Charles H. Courtney Poultry Sciences Dept. Biology, Price Hall, Rm Agriculture Canada University of Florida, Coll. Vet. 1260 W. Maple 216 5403 1st Ave S. Med. Fayetteville, AR 72701 Berrien Springs, MI 49104 Lethbridge, AB, Canada T1J Box 100125 479-575-4870 269-471-3262 4B1 Gainesville, FL 32610-0125 479-575-4202 269-471-6911 403-317-2254 352-392-4700, ext.5111 [email protected] [email protected] 403-382-3156 352-392-8351 [email protected] [email protected]. Roxanne Charles Annapoorani Chockalingam edu; [email protected] University of Georgia Cornell University Gary A. Conboy 225 Appleby Drive, Apt 231 520 The Parkway University of Prince Edward April Covington Athens, GA 30605 Ithaca, NY 14850 Island, Atlantic Vet. Coll. Merial Ltd 323-809-6180 607=343-0606 550 University Avenue 2195 Station Village Way, Apt. [email protected] [email protected] Charlottetown, PEI, Canada 1404 C1A 4P3 San Diego, CA 92108 Samuel Charles Les Choromanski 902-566-0965 619-417-8787 Bayer Health Care Pfizer, Inc. 902-566-0851 678-638-8635 P.O. Box 390 7000 Portage Rd [email protected] [email protected] Shawnee Mission, KS 66201 Kalamazoo, MI 49001 913-268-2520 269-833-2503 George A. Conder Bobby Cowles 913-268-2541 269-833-4255 Pfizer Animal Health (Retired) Pfizer Animal Health [email protected] leszek.j.choromanski@pfizer. 9244 Weathervain Trail 18503 NW 41st Ave com Galesburg, MI 4903 Ridgefield, WA 98642 Rajendran Chellaiah 269-665-4948 360-606-1443 USDA, Animal Parasitic Dis- Jeffrey N. Clark [email protected] [email protected] eases Laboratory JNC Consulting Services, Inc. 10300 Edmonston Road 38 Wild rose Lane Donald P. Conway Thomas M. Craig Beltsville, MD 20705 Pittsboro, NC 27712 Conway Associates Texas A&M University, Coll. 301-504-5111 919-942-9222 2 Willever Road Vet. Med. [email protected] deposit^[email protected] Asbury, NJ 8802 Dept. Vet. Pathobiol. 908-730-8823 College Station, TX 77843- John M. Cheney Bill C. Clymer [email protected] 4467 P.O. Box 1905 Consultant to Animal Health 979-845-9191 Greenwood, AR 72936 13351 East FM 1151 Camille Coomansingh 979-862-2344 970-377-0482 Amarillo, TX 79118 St. George University [email protected] [email protected] 806-335-3338 P.O. Box 7 806-335-3531 St. George, Grenada Luiz G. Cramer Jose W. Chernitzky [email protected] 473-444-4175 Merial Limited Facultad de Medicina Veteri- 473-439-5068 3239 Satellite Blvd naria y Zoot, UNAM Nicole Colapinto [email protected] Duluth, GA 30024 Newton 283 Depto. 702, [email protected] Colonia Polanco Chapultepec, Rebecca A. Cole Helder Cortes Mexico DF CP 11560 USGS Nat’l Wildlife Health Evora University Larry R. Cruthers (5255)52030310 Research Ctr. Lab of Parasitology, Victor Professional Laboratory & Re- [email protected] 6006 Schroeder Rd. Caeiro, Mitra center search Services, Inc. (PLRS) Madison, WI 53711 Evora, Portugal 7000 1251 NC 32 North M.B. Chhabra 608-270-2468 35198 501-7554 Corapeake, NC 27926 New Delhi 608-270-2415 [email protected] 252-465-8686 D11/2518, Vasant Vihar [email protected] 252-465-4493 New Delhi, India 110070 Manuel R. Cortinas [email protected], [email protected] University of Illinois, Dept. Vet. [email protected] Pathobiol., MC-002 2001 S. Lincoln Ave. Urbana, IL 61820 217-244-3020 217-244-7421 [email protected]

91 Tracy L. Cyr Bart De Leeuw Dan T. Domingo J. P. Dubey Texas A&M University, Coll. Pfizer Animal Health Pfizer Animal Health, USDA, ARS, APDL Vet. Med. PO Box 37, Capelle A/D ‘jssel 150/39/11 Bldg. 1001, BARC-East Dept. Vet. Pathobiology, 4467 The Netherlands 2900AA 150 E. 42nd St. Beltsville, MD 20705-2350 TAMU 31-10-406-4600 New York, NY 10017 301-504-8128 College Station, TX 77843- 31-10-406-4293 212-573-3050 301-504-9222 4467 [email protected] 212-309-0428 [email protected] 979-458-3276 [email protected] 979-862-2344 B. Joe Dedrickson Vivien G. Dugan [email protected] Merial William A. Donahue, Jr. University of Georgia, SC- 1409 Silver Fox Run Sierra Research Laboratories WDS Patricia Daly Woodstock, GA 30188-5628 5100 Parker Road Coll. Vet. Med. Bayer Health Care 678-428-7405 Modesto, CA 95357 Athens, GA 30602-4393 39 Waterbury St., Apt 1 678-638-8870 209-521-6380 706-542-1741 Saratoga Springs, NY 12866 [email protected] 209-521-6380 706-542-5865 518-581-3276 [email protected], bdona- [email protected] [email protected] Nicole DeFraeye [email protected] Pfizer Animal Health Karen L. Duncan John B. Dame 5310 Pain Court Line John M. Donecker Intervet, Inc. University of Florida, Coll. Vet. Pain Court, Ontario, Canada Pfizer Animal Health 29160 Intervet Lane Med N0P 120 707 Parkway Blvd Millsboro, DE 19966 Box 110880 519-351-5514 Reidsville, NC 27320 302-934-4368 Gainesville, FL 32611 519-355-1247 336-552-6027 302-934-4203 352-392-4700 [email protected] 866-590-0323 [email protected]. 352-392-9704 [email protected] com [email protected] Kimberley DeLisi Virginia Tech Ann R. Donoghue Donya D. Dupree Sriveny Dangoudoubiyam 1410 Prices fork Rd Donoghue Consulting, LLC South East Texas Animal University of Kentucky, Gluck Blacksburg, VA 24060 150 N Country Rd 3 Emergency Clinic Equine Research Center [email protected] Fort Collins, CO 80524 2813 Independence Rd 1435 Nicholasville Road 970-214-5999 Westlake, LA 70669 Lexington, KY 40546 Bradley DeWolf 970-482-2044 225-931-8703 859-257-4757, ext. 81167 University of Guelph [email protected] [email protected] [email protected] 8066 Heritage Line Wallaceburg, Canada N8A4L3 Mary E. Doscher Michael T. Dzimianski Craig Datz 519-824-4120 Retired Michael Dzimianski, DVM University of Missouri, Coll. [email protected] 147 Gary Dr. 452 Pony Trail Vet. Med. Trenton, NJ 8690 Nicholson, GA 30565 900 E. Campus Drive Steven D. Diehl 609-586-3185 706-546-8520 Columbia, MO 65211 Bayer Animal Health [email protected] 706-546-8520 573-882-7821 1612 Redwood Hill Rd [email protected] 573-884-7563 Santa Rosa, CA 95404 Frank W. Douvres [email protected] 707-545-7540 538 Shipley Road Jenifer Edmonds 707-545-7548 Linthicum Heights, MD 21090 Johnson Research, LLC Gabriel Davila [email protected] 24007 Highway 2026 University of Florida Jason Drake Parma, ID 83660 2929 SW 39th Ave Danielle Dimon Novartis Animal Health 208-722-5829 Gainesville, FL 32608 University of Georgia, Coll. 7809 Charles Place Drive 208-722-7371 352-281-1124 Vet. Med. Kernersville, NC 27284 jedmonds@johnsonre- [email protected] Dept Infectious Diseases 336-387-1027 searchllc.com Athens, GA 30602 [email protected] Wendell L. Davis 404-646-1764 Matthew Edmonds Bayer Health Care [email protected] Robert Dressler Johnson Research, LLC P.O. Box 390 Pfizer Animal Health 24007 Highway 2026 Overland Park, KS 66201 Sandra K. Dixon Box 250 Parma, ID 83660 913-268-2079 Sandra Dixon, DVM Barksdale, TX 78828 208-722-5829 913-268-2040 P.O. Box 192 830-234-4100 208-722-7371 [email protected] Grayson, GA 30017 830-234-4103 medmonds@johnsonre- 770-235-5768 [email protected], searchllc.com Theresa de Blieck [email protected] [email protected] University of Minnesota Amy Edwards 3883 Co Rd 74 Cynthia Doffitt Michael W. Dryden Oklahoma State University Saint cloud, MN 56302 Mississippi State University, Kansas State University 1818 w. Adnmiral 612-868-8643 Coll. Vet. Med. 1800 Denison Ave, Coles Hall Stillwater, OK 74074 [email protected] 1 Spring St Manhattan, KS 66506 [email protected] Mississippi State, MS 39762 785-532-4613 662-325-1154 785-532-4039 [email protected] [email protected]

92 Jessica Edwards Michael Endrizzi Maarten Eysker Glenn R. Frank University of Georgia Pfizer Animal Health University of Utrecht, Div. Heska Corp. 335 Spring Lake Ct 16036 Eagle River Way Para. & Trop. Vet. Med., 3760 Rocky Mountain Ave Athens, GA 30605 Tampa, FL 33624 Yalelaan 1 Loveland, CO 80538-9833 404-558-0438 336-210-9760 P.O. Box 80.165, 3508 TD 970-493-7272 [email protected] [email protected] Utrecht, The Netherlands 970-619-3003 31-30-253-1223 [email protected] Kristine Edwards Christian Epe 31-30-254-0784 Mississippi State University, Novartis Centre de Recherche [email protected] Jeanne M. Freeman Howell Dept. Entomology Sanite Animal USDA/ARS/KBUSLIRI 1855 Oktoc Road St. Aubin, Switzerland CH- Silvina Fernandez 2700 Fredericksburg Rd Starkville, MS 39759 1566 University of Guelph, Ontario Kerrville, TX 78028 662-323-5443 +41 26 6791507 Vet. Coll. 830-792-0332 [email protected], kted- +41 26 6791228 Organic Ag Centre of Canada 830-792-0314 [email protected] christian.epe@tiho-hannover. (OACC), c/o Dept. Pathobiol. [email protected] de Guelph, Ontario, Canada N1G Amal K. El-Gayar 2W1 Marguerite K. Frongillo Texas A&M Lynn F. Erdman 519-824-4120, Ext. 54819 University of South Carolina 200 Charles Haltom, #7 Raleigh Hills Veterinary Clinic 519-824-5930 3600 Beverly Drive College Station, TX 77840 7980 S.W. 87th Avenue [email protected] Columbia, SC 29204 [email protected], ael- Portland, OR 97223 803-743-0731 [email protected] 503-292-9227 Raymond H. Fetterer [email protected] 503-292-8487 USDA-ARS, APDL, BARC- Selene Elizondo [email protected] East Catalina Fung Berengena no. 15 Bldg. 1040 P.O. Box 21958 Xatapa, Mexico 91157 Andrew K. Eschner Beltsville, MD 20705 West Palm Beach, FL 33416 228 1456950 Merial Ltd 301-504-8762 [email protected] [email protected] 4 Pepper Place 301-504-5306 Gansevoort, NY 12831 [email protected]. Ujvala Deepthi Gadde Siobhan Ellison 518-580-0459 gov University of Arkansas Pathogenes, Inc. 518-580-0460 7 S Duncan Ave PO Box 970 drandrew.eschner@merial. Michael G. Fletcher Fayetteville, AR 72701 Fairfield, FL 32634 com Y-Tex Corporation, PO Box 479-966-9834 352-591-3221 1450 [email protected] 352-591-4318 Christopher Evans 1825 Big Horn Ave. [email protected] University of Georgia, Coll. Cody, WY 82414 Alvin Gajadhar Vet. Med. 307-587-5515, Ext. 115 Centre for Animal Parasitol- Siobhan Ellison Dept Infectious Diseases 307-527-6433 ogy, Canadian Food Inspec- Pathogenes, Inc. Athens, GA 30602 [email protected] tion Agency P.O. Box 970 480-747-3028 116 Veterinary Road Fairfield, FL 32634 [email protected] James R. Flowers Saskatoon, SK, S7N 2R3 352-591-3221 North Carolina State Univer- Canada 352-591-4318 Kevin Evans sity, Coll. Vet. Med. 306-975-5344 [email protected] Pfizer Animal Health, 4700 Hillsborough Street 306-975-5711 150/40/14 Raleigh, NC 27606 [email protected] Stacey Elmore 150 East 42nd St 919-513-6404 Colorado State University New York, NJ 10017 [email protected] Rajshekhar Gaji 914 Cherry Street 212-733-0910 Univ. Michigan Fort Collins, CO 80521 [email protected] Akande Foluke Adedayo 1647 Beal Ave, Apt 15 303-579-6667 University Ann Arbor, MI 48105 [email protected] Richaed Evans Alogi St. [email protected] Pacific Marine Mammal Abeokuta, Nigeria 234 David Elsemore Center 234(0)8035008607 H. Ray Gamble IDEXX Laboratories 22501 Chase #3102 [email protected] National Academy of Sciences One Idexx Drive Aliso Viejo, CA 92656 500 Fifth Street NW, Room Westbrook, ME 4106 949-494-3050 Fleeta Fore 570 207-556-3337 949-494-2802 306 Whispering Hills Washington, DC 20001 [email protected] [email protected] Hot Springs, AR 71901 202-334-2787 [email protected] 202-334-2759 Richard G. Endris Sidney A. Ewing [email protected] Intervet Schering Plough Oklahoma State University, William J. Foreyt Animal Health Dept. Vet. Pathobiol, Washington State University Robert D. Garrison 56 Livingston Ave 250 McElroy Hall Dept. Vet. Micro. and Path. Texas Corners Animal Hos- Roseland, NJ 7068 Stillwater, OK 74078-2007 Pullman, WA 99164-7040 pital 862-245-5133 405 744-8177 509-335-6066 7007 w. Q Ave. 862-245-3654 405 744-5275 509-335-8529 Kalamazoo, MI 49009 [email protected]. [email protected] [email protected] 269-375-3400 com [email protected]

93 Louis C. Gasbarre Richard W. Gerhold Jonas Goring Matt Griffin USDA-ARS, RM 110 University of Georgia, SC- IDEXX Laboratories, Inc. P.O. Box 197 Bldg 1002, BARC-East WDS, Coll. Vet. Med. One IDEXX Drive Stoneville, MS 38776 Beltsville, MD 20705 501 D.W. Brooks Dr. Westbrooke, ME 4092 [email protected] 301-504-8509 Athens, GA 30602-4393 613-697-5689 301-504-6426 706-542-1741 [email protected] Nicole Grosjean [email protected] 706-542-5865 Michigan State University [email protected] Michelle Gourley 1720 S. Michigan Rd Carol L. Geake Western Michigan University Eaton Rapid, MI 48827 Hidden Spring Veterinary H. C. Gibbs 4273 Vine St [email protected] Clinic 588 Kennebec Road Bridgman, MI 49106 48525 W. Eight Mile Road Hampton, ME 4444 219-393-9655 Frank Guerino Northville, MI 48167 207-862-3578 [email protected] Intervet Schering-Plough 248-349-2319 Animal Health 248-349-2517 John Gilleard David E. Granstrom 556 Morris Ave [email protected] University of Calgary American Veterinary Medical Summit, NJ 07901-1330 16, Edenstone View Association 908-473-3118 James Geary Edgemont, Canada 1931 N. Meachum Rd, Suite 908-473-3654 Michigan State University 403-375-0844 1000 [email protected] Dept Pthobiology & Diagnostic [email protected] Schaumberg, IL 60173 Investugation 847-285-6674 Jorge Guerrero East Lansing, MI 48824 Keith P. Goldman 847-925-1329 University of Pennsylvania 269-903-8099 The Hartz Mountain Corp [email protected] 10 North Riding Drive [email protected] 400 Plaza Drive Pennington, NJ 8534 Secaucus, NJ 7094 Sara Ellen Green 609-510-1337 Timothy G. Geary 201-271-4800, Ext. 2457 2455 S. Roby Farm Road 609-737-6793 McGill University, Institute of 973-429-0699 Rocheport, MO 65279 [email protected] Parasitology [email protected] 573-445-5034 21,111 Lakeshore Road [email protected] Gagan D. Gupta Ste-Anne de Bellevue, PQ Daniel Golden University of Missouri, Coll. H9X 3V9, Canada Merial Spencer Greenwood Vet. Med. 54-398-7612 3239 Satellite Blvd University of Prince Edward 205 Connaway Hall [email protected] Duluth, GA 30096 Island Columbia, MO 65211 678-638-3703 Dept. Path. & Micro., 550 573-884-2732 Claudio Genchi [email protected] University Ave 573-884-5414 University of Milan, Fac, Vet. Charlottetown, PEI, Canada [email protected] Med., Luis A. Gomez-Puerta C1A 4P3 Via Celoria 10 Universidad National Mayor 902-566-6002 Nicole J. Guselle 20133 Milano, Italy de San Marcos, Facultad de 902-566-0851 Atlantic Veterinary College 3.9025E+11 medicine Veterinaria [email protected] Dept. Pathobiology & Micro., 3.90258E+11 J. Jose Gabriel Chariarse 296 550 University Ave. [email protected] San Antonio, Miraflores, Lima Ellis C. Greiner Charlottetown, PEI, Canada Peru University of Florida, Coll. Vet. C1A 4P3 Jinming Geng (511)436-8938 Med. 902-566-0595 Idexx (511)436-5780 X 106 Dept. Pathobiol, Box 110880 [email protected] 1 Idexx Drive [email protected] Gainesville, FL 32610 Westbrook, ME 4092 352-294-4161 Aliessia Guthrie 207-556-6809 David G. Goodwin 352-392-9704 University of Guelph [email protected] Virginia Tech, Coll. Vet. Med., [email protected] 302-63 Queen St. Lab 203 Guelph, ON, Canada NTE Jay & Marion Georgi 1410 Prices Fork Rd John H. Greve 4RP 132 Starr Stanton Hill Rd. Blacksburg, VA 24060 334 24th Street [email protected] R.D. #2 540-231-7074 Ames, IA 50010 Freeville, NY 13068 540-231-3426 515-232-3520 Richard J. Hack 607-844-8507 [email protected] [email protected] Elanco Animal Health 2001 West Main Street, P.O. David C. Gerdon Velmurugan Gopal Viswa- Robert B. Grieve Box 708 Merial Limited natahn Heska Corporation Greenfield, IN 46140 3239 Satellite Blvd USDA, ARS, ANRI 3760 Rocky Mountain Avenue 317-371-8614 Duluth, GA 30041 4601 Tonquil St. Loveland, CO 80538-9833 317-277-4288 678-638-3865 Beltsville, MD 20705 970-493-7272 [email protected] 678-638-3873 301-504-5999 970-619-3003 [email protected] [email protected] [email protected], [email protected] Khurram Goraya Faisalabad, Pakistan 9.23087 [email protected]

94 Bruce Hammerberg Stephanie Heise Sally I. Hirst Daniel K. Howe North Carolina State Univer- Oklahoma State University Editor, Trends in Parasitology University of Kentucky sity, Coll. Vet. Med. 2207 W. Sherwood Ave Elsevier, 32 Jamestown Road 108 Gluck Equine Research 4700 Hillsborough St. Stillwater, OK 74074 London, UK, WC1X 8RR Center Raleigh, NC 27606 405-762-6438 44-20-7611-4128 Lexington, KY 40546-0099 919-785-0332 [email protected] 44-20-7611-4470 859-257-4757, Ext. 812113 bruce_hammerberg@ncsu. [email protected] 859-257-8542 edu Klaus Hellmann [email protected] Klifovet AG Jessica S. Hoane Qian Han Geyerspergerstr. 27 University of Kentucky Sue Howell Virginia Tech Munchen, Germany D-80689 108 Gluck Equine Research University of Georgia 204 Engel Hall +49-89-58 00 82 Center 84 Keys Court Blacksburg, VA 24061 +49-89-56 00 82 7 Lexington, KY 40546-0099 Braselton, GA 30517 540-231-5779 [email protected] 859-257-4757, Ext. 81213 [email protected] [email protected] 859-257-8542 W. Lance Kemsarth [email protected] Armando Hung Aaron Harmon The Hartz Mountain Corp. Universidad Peruana Cay- Novartis 192 Bloomfield Avenue Eric P. Hoberg etano Heredia, Laboratorio de 1447 140th St. Bloomfield, NJ 7003 USDA, ARS US National Biología Molecular Facultad Larchwood, IA 51241 201-271-4800 Parasite Collection, Animal de Veterinaria y Zootecnia, 712-47-2811, ext. 2230 [email protected] Parasitic Disease Laboratory Universidad Peruana Cay- [email protected] BARC-East, #1180, 10300 etano Heredia Douglas I. Hepler Baltimore Ave. Av. Honorio Delgado 430, San Elizabet Harp Kado Enterprises Beltsville, MD 20705-2350 Martin Colorado State University 815 Cliff dr. 301-504-8588 Lima, Peru L12 Campus Mail 1878 (Biology) Mcleansville, NC 27310 301-504-8979 51 1 3190030 – anexo 2259 Fort Collins, CO 80523 336-708-2842 [email protected] 51 1 3190039 970-203-5462 336-697-7296 [email protected]; ar- [email protected] [email protected] Johan Hoglund [email protected] Ulls v Theresa A. Hartwell Dr. Harry Herlich Uppsala, SE 750 07 Frank Hurtig Stillmeadow, Inc. 1025 Oakridge-D +46 18 674156 Merial 12852 Park One Dr. Deerfield Beach, FL 33442- [email protected] 468 E. Ridge Dr. Sugar Land, TX 77478 1958 Eagle, ID 83616 281-240-8828 954-419-9547 Patricia Holmer 314-409-2760 281-240-8442 Texas A&M 208-938-5060 [email protected] James A. Higgins 13009 Tall Timber Dr. [email protected] USDA-ARS, Bldg 173 College Station, TX 77845 John M. Hawdon 10300 Baltimore Blvd 979-845-4202 Douglas E. Hutchens George Washington University Beltsville, MD 20705 [email protected] Bayer Animal Health Medical Center, 725 Ross Hall 301-504-6443 P.O. Box 390 2300 Eye St., NW 301-504-6608 Joe A. Hostetler Shawnee Mission, KS 66201 Washington, DC 20037 [email protected] Bayer Health Care 678-638-3206 202-994-2652 1467 N. 300 Road 678-638-8877 202-994-2913 Michael B. Hildreth Baldwin City, KS 66006 douglas.hutchens.b@bayer. [email protected] South Dakota State Univer- 913-268-2010 com sity, Dept. Biol. & Micro., SNP 913-268-2878 James A. Hawkins - Rm 252 [email protected] John P. Hutcheson Merial Ltd College of Agriculture & Bio- Intervet Schering Plough 118 Quail Run Dr. logical Sciences Alice Houck Animal Health Madison, MS 39110 Brookings, SD 57007 Virginia Tech 7101 Red Rock Rd 601-853-8442 605-688-4562 VA/MD Reg. Coll. Vet. Med. Amarillo, TX 79118 678-638-8875 605-688-5624 Blacksburg, VA 24061-0442 806-622-1080 [email protected] [email protected] 540-231-7074 806-622-1086 [email protected] [email protected] Terence J. Hayes Cherly Ann Hirschlein Hoffman-La Roche, Inc. Merial Ltd. Vikki Howard Marianna Ionita 340 Kingsland Street 5970 Sycamore Rd. Novartis University of Agronomical Sci- Nutley, NJ 7110 Buford, GA 30518 10 Charles Hill Road ences & Veterinary Medicine 973-235-2120 678-638-3479 Kittery Point, ME 3905 Splaiul Independental, No. 975-235-2981 678-638-3074 603-498-9305 105, Sector 5 [email protected] [email protected], cheryl. [email protected] Bucharest, Romania 50097 [email protected] (40)21-318.04.69 (40)21-318.04.98 [email protected]

95 Robert S. Isenstein Mitchell A. Johnson Maxine F. Kellman Thomas R. Klei FSIS, USDA Intervet 9418 Riverbrink Court Louisiana State University, Building 318C, BARC-East 35500 West 91st Laurel, MD 20723 PBS Beltsville, MD 20705 DeSoto, KS 66018 301-317-9859 Sch. Vet. Med. [email protected] 301-317-9859 Baton Rouge, LA 70803 Ahmed A. Ismail [email protected] 225-578-9900 Sudan University of Science Sandra S. Johnson 225-578-9916 Khartoum, Sudan Pfizer Animal Health Thomas J. Kennedy [email protected] +249 9122149209 7000 Portage Road Central Life Sciences, Inc., [email protected] Kalamazoo, MI 49001 301 West Osborn St. Ronald D. Klein 269-833-2660 Phoenix, AZ 85013-3997 Dancing Turtle, Farm & Dairy Abdo Jassim M. 269-833-2769 800-720-0032, Ext. 2269, 9721 South 6th Street 11 Ailol [email protected] 602-664-1258 Schoolcraft, Mi 49087 Dohuk, Iraq 44 602-207-2145 269-375-3073 075 04143224 Colin Johnstone [email protected] [email protected] [email protected] University of Pennsylvania. 416 Clinton Road. Michael Kent Bruce Klink Philippe Jeannin Brookline, MA 02445-4167 Oregon State University Bayer Health Care, LLC Merial 508-413-2107 220 Nash Hall 2508 Greenbrook Parkway Parc Industriel de la Plaine de [email protected] Corvallis, OR 97331 Mattews, NC 28104 l’Ain Allée des Cyprès 541-737-8652 704-957-5589 F- 01150 SAINT-VULBAS, Helen E. Jordan michael.kent@oregonstate. 704-844-6725 France Oklahoma State University edu [email protected] 33 04 72 72 33 95 2923 Fox Ledge Drive 33 04 72 72 29 57 Stillwater, OK 74074 Stephanie Keroack Roland Klober 405-372-6420 Pfizer Animal Health Pfizer, Inc. Emily Jenkins [email protected] 370 Missisquol N 55 North Broadway, No. 2-23 University of Saskatchewan Bromont, Canada J2L 2N5 White Plains, NY 10601-1640 Dept. Vet. Microbiology Richard Kabuusus 514-606-6831 212-733-8248 Saskatchewan, Canada S7N St. George’s University stephanie.keroack@pfizer. 212-573-2821 584 School of Vet. Med. com [email protected] 306-966-2569 Grenada [email protected] 1473 439 2000 Andrea Ketschek Don Knowles [email protected] University of Pennsylvania, USDA, ARS, WSU Mark Jenkins Sch. Vet. Med. 3005 ADBF, ADRU USDA, ARS, APDI Malika Kachani RM 212, 3800 Spruce St. Pullman, WA 99164-6630 Building 1040, BARC-East Western University, Coll. Vet. Philadelphia, PA 19104 509-335-6022 Beltsville, MD 20705 Med. 215-573-8092 509-335-8328 301-504-8054 CVM 309 E. 2nd Street 215-573-7023 [email protected] [email protected] Pomona, CA 91766 [email protected] 909-469-5302 April Knudson Edward G. Johnson [email protected] Jennifer K. Ketzis Merial Ltd Johnson Research, LLC Charles River Biolabs Europe 12056 Avenida Sivaita 24007 Highway 2026 Ray M. Kaplan Ltd San Diego, CA 92128 Parma, ID 83660 University of Georgia, Coll. Carrentrila, Ballina, Co Mayo 619-427-8787 208-722-5829 Vet. Med. Ireland 678-638-8636 208-722-5119 Dept Infectious Diseases 353.96.20800 [email protected] [email protected] Athens, GA 30602 353.96.22517 706-542-5670 [email protected] Dennis E. Kobuszewski Eileen Johnson 706-542-0059 Summit VetPharm LLC Oklahoma State University, [email protected] Muhammad Tanveer Khan 8740 Iron Horse Dr. Coll. Vet. Med., Center Vet. Pakistan Irving, TX 75063 Hlth. Sci. Kevin R. Kazacos University of Vet. & Animal 804-614-8358 Dept. Patholbiology, 250 Purdue University, Dept. Vet. Sciences dkobuszewski@summitveph- McElroy Hall Pathobiology Lahore, Pakistan arm.com Stillwater, OK 74078-2007 725 Harrison St. 35 39620856 405-744-8549 West Lafayette, IN 47907- [email protected] Katherine M. Kocan 405-744-5275 2027 Oklahoma State University, [email protected] 765-494-7556 Jan M. Kivipelto Coll. Vet. Health Sci. 765-494-9830 University of Florida, Dept. Dept. Vet. Pathobiology, 250 Kathy Johnson [email protected] An. Sci. McElroy Hall Purdue University 459 Shealy Drive Stillwater, OK 74078-2007 4125 Trilithon Court Suzanne Keeys Gainesville, FL 32611 405-744-7271 West Lafayatte, IN 47906 Pac Behavior 352-392-3342 405-744-5275 774-263-3825 333 James Jackson Ave 352-392-7652 [email protected] [email protected] Cary, NC 27513 [email protected] 919-468-8301 [email protected]

96 Dorsey L. Kordick Ken-ichi Kusano Alice Lee Janice Liotta Idexx Pharmaceuticals, Inc Sankyo Yell Yakuhin Co., Inc. Cornell University Cornell University 4249-105 Piedmont Parkway 3-10 Takamatsu, Toshima C4183 VMC, Campus Rd. C4-114 VMC Tower Rd Greensboro, NC 27320 Tokyo, 171-0042, Japan Ithaca, NY 14853 Ithaca, NY 14853 336-834-6523 03 3955 5354 607-253-3305 607-253-3394 336-834-6525 03 3955 5354 [email protected]; [email protected] dorsey.kordick@idexxpharma. [email protected] com William G. Kvasnicka Susan E. Little 7131 Meadowview St Chung G. Lee Oklahoma State University, Rosina C. (Tammi) Krecek Shawnee, KS 66227 Chonnam National University, Center Vet. Hlth. Sci. Ross University, School of 913-441-7946, cell: 775-530- Coll. Vet Med. 250 McElroy Hall, Dept. Vet. Vet. Med. 2068 Kwangju Pathobiology 499 Thornall St., Suite 1101 913-441-0937 Republic of South Korea Stillwater, OK 74070-2007 Edison, NJ 8837 [email protected] 500-757 405-744-8523 869-465-2405 Ext. 119, mo- 062 5302870 405-744-5275 bile: 869-665-3196 Dongmi Kwok 062 5302874 [email protected] 869-465-1203 1370 Sankyuk 3-dong [email protected] [email protected] Bukgu, Daegy, Korea 70270 1 John E. Lloyd [email protected] Thomas Letonja University of Wyoming, Dept. Eva Maria Kruedewagen USDA APHIS VS 3354, Renewable Resources Opladener Straude 117 Elise LaDouceur 1219 James Rifle Ct. NE 1000 E. University Ave Monheim Germany Tufts University Leesburg, VA 20176 Laramie, WY 82071 492173, 2036468 1 Arch St. 301-734-0817 307-766-2234 eva.kruedewagen@t-online. Westborough, MA 1581 301-734-3652 307-766-5025 de 443-629-8883 [email protected]. [email protected] [email protected] gov Ewa Kuczynska J. Mitchell Lockhart 3004 Reed CT, Apt 2B Rolando Landin Michael G. Levy Valdosta State University, Merrillville, IN 46410 P.O. Box 22993 North Carolina State Univer- Biology Dept. 815-295-1509 West Palm Beach, FL 33416 sity, Coll. Vet. Med. 1500 N. Patterson St. [email protected] [email protected] 4700 Hillsborough Street Valdosta, GA 31698 Raleigh, NC 27606 229-333-5767 Raymond E. Kuhn Carlos E. Lanusse 919-513-6293 229-245-6585 Wake Forest University, Dept. Lab de Farmacologia, Fac de 919-513-6464 [email protected] Biology Cien.Veter [email protected] PO Box 7325 Campus Universitario, Univer- Linda L. Logan Winston-Salem, NC 27109 sidad Nacional J. Ralph Lichtenfels Texas Animal Health Com- 336-758-5022 Tandil, 7000 - ARGENTINA USDA-ARS-ANRI mission 336-758-6008 5.42293E+11 12311 Whitehall Dr. 2105 Kramer Lane [email protected] 5.42293E+11 Bowie, MD 20715-2350 Austin, TX 78758 [email protected] 301-262-9496 800-550-8242 Daniel Kulke 301-262-9496 512-719-0719 Heinrich Heine Universitaet Diane Larsen [email protected] Universitaetstr. 1, Institut fuer Merial Joyce A. Login Zoomorphologie und Parasit. 3239 Satellite Blvd David R. Lincicome Bayer Animal Health Duesseldorf, NRW, Germany Duluth, GA 30519 11 Falls Road 9133 Taylor Court 40225 678-638-3633 Roxbury, CT 6783 W. Windsor, NJ 8550 49 2173 384232 678-638-3636 860-355-1031, 240-286-2438 877-513-5222 daniel.kulke@uni-duesseldorf. [email protected] [email protected], 609-716-8455 de [email protected] [email protected] Mette Larsen Suresh Kumar Royal Vet & Agricultural Univ., David S. Lindsay Lori Lollis Allahabad Agriculture Uni- Vallborg All 17, 1.1 v Virginia Tech, Center for Mo- University of Georgia versity Dept. Large An. Sci., Dyrlæ- lecular Med. & Inf. Dis., V-M 3939 Quail Hollow Road 50, a-First floor, Tampale Rd., gevej 88, Frederiksberg Regional Coll. Vet. Med. Albany, GA 31721 Jung Pura Bhogal Copenhagen, Denmark, Valby, 1410 Prices Fork Road 229-869-7855 New Delhi, India 110014 XX DK-2500 Blacksburg, VA 24061-0342 [email protected] 011-24374576 [email protected] 540-231-6302 011-24374545 540-231-3426 Alvin F. Loyacano [email protected] Jeffrey R. Laursen [email protected] Louisiana State University, Eastern Illinois University Ag. Center, LAES, Dean Lee Dept. Biol. Sci., 600 Lincoln Ashley Linton Res. Station Ave. Colorado State University 8105 Tom Bowman Dr. Charleston, IL 61920 Fort Collins, CO Alexandria, LA 71302 217-581-6390 [email protected] 318-473-6523 217-581-7141 318-473-6535 [email protected] [email protected]

97 Aaron S. Lucas Claire Mannella John Mathew Anne McKee Virginia Tech, Dept. Vet. Med. Novartis Animal Health Merck & Co. Sci. 1545 N. Geronimo Rd. 203 River Road Mary E. McKenzie 833 Claytor Sq. Apache Junction, AZ 85219 Somerville, NJ 7950 Pfizer Animal Health Blacksburg, VA 24060 [email protected] 908-685-6823 150 E. 42nd Street 540-230-7917 908-722-7433 New York, NY 10017 [email protected] Linda S. Mansfield [email protected] 212-573-5438 Michigan State University 212-573-5348 Joan K. Lunney B43 Food Safety Toxicology Martin Matisoff m.elizabeth.mckenzie@pfizer. USDA, ARS, ANRI, APDL Bldg Kentucky State University, com Bldg. 1040, Room 107, East Lansing, MI 48808 Honey Bee Laboratory BARC-East 517-432-3100, Ext. 119 8606 Cool Brook Court Kelsey McNally Beltsville, MD 20705 517-432-2310 Louisville, KY 40291 Mississippi State University 301-504-9368/8373 [email protected] 502-243-7637 Coll. Vet. Med. 301-504-5306 [email protected]. Mississippi State, MS 39762 [email protected] Abdelmoneim Mansour edu, martinmatisoff@gmail. 662-325-1345 TRS Labs com [email protected] Elizabeth Lynn 295 Research Dr. University of Georgia, Coll. Athens, GA 30605 Mark E. Mazaleski Tom McTier Vet. Med. 706-354-1531 Pfizer, Inc. Pfizer Animal Health Dept Infectious Diseases [email protected] 8437 Harvest Ave 7000 Portage Rd Athens, GA 30602 Richland, MI 49083 Kalamazoo, MI 49001 912-237-0760 Kenny Marbury 269-833-3203 269-833-3499 [email protected] Novartis Animal Health [email protected] [email protected] 538 Wyatt Dr. Eugene T. Lyons St. Peters, MO 63376 William McBeth Lisa Meader University of Kentucky, Dept. 636-928-3601 Pfizer, Inc. 11705 West Main Vet. Sci. 636-447-5690 812 Springdale Drive Whitmore Lake, MI 48187 Gluck Equine Res. Center [email protected] Exton, PA 19341 [email protected] Lexington, KY 40546-0099 610-968-3558 859-257-4757, Ext. 81115 Alan A. Marchiondo [email protected] America Mederos 859-257-8542 Pfizer Animal Health University of Guelph [email protected] 7000 Portage Road John W. McCall 54 University Ave Kalamazoo, MI 49001 University of Georgia, Coll. Guelph, ON, Canada N1G Ana E.B. Maciel 269-833-2674 Vet. Med., Dept. Inf. Diseases, 1N7 Merial 269-833-3231 TRS Labs, Inc. [email protected] 3239 Satellite. Blvd [email protected] PO Box 5112 Duluth, GA 30096-4640 Athens, GA 30605 Patrick F.M. Meeus 678-638-3331 Sara E. Marley 706-542-5684 Pfizer-VMRD 678-638-3322 Bayer Animal Health 706-542-5771 7000 Portage Road, Bldg 300 [email protected] 20831 Brandt Road [email protected] Kalamazoo, MI 49001 Tonganoxie, KS 66086 269-833-2661 A. J. MacInnis 913-268-2041 Scott McCall 269-484-4615 University of California – Los [email protected] TRS Labs, Inc [email protected], Angeles, Dept. Biology PO Box 5112 meeusjordan@jasnetworks. PO Box 951606 Antoinette Marsh Athens, GA 30605 net Los Angeles, CA 90095-1606 1950 Westwood Ave 706-549-0764 310-825-3069 Columbus, OH 43212 706-353-3590 Shelley Mehlenbacher 310 206-3987 614-282-1154 [email protected], Bayer Healthcare [email protected] [email protected] [email protected] P.O. Box 83158 Portland, OR 97283 Charles D. MacKenzie Randell J. Martin James Mcclanahan 503-247-9108 Michigan State University Schering-Intervet What A Relief shelley.mehlenbacher.b@ 11649 Jarvis Highway 10116 Tapestry Street 8726 E. Solano Dr. bayer.com Dimondale, MI 42881 Forth Worth, TX 76244 Scottsdale, AZ 95250 517-432-3644 [email protected] 602-317-8443 Norbert Mencke [email protected]; mack- 480-626-7295 Bayer Animal Health [email protected] Richard J. Martin [email protected] R&D/Animal Expertise Center Iowa State University, Coll. / Preclinic John B. Malone, Jr. Vet. Med. Christine M. McCoy Leverkusen, Germany, DE- Louisiana State University, #2008, Dept. Biomed. Sci. Elanco Animal Health 51368 Sch. Vet. Med. Ames, IA 50011 2001 W. Main St., PO Box 49 2173 384921 Pathobiological Sciences 515-294-2470 708 49 2173 3894921 Baton Rouge, LA 70803 515-294-2315 Greenfield, IN 46140 norbert.mencke@bayerhealth- 225-578-9692 [email protected] 317-651-4377 care.com 225-578-9701 317-277-4522 [email protected] [email protected]

98 Christina Mertens Ioan Liviu Mitrea Eva Nace Thomas J. Nolan Intervet Innovation University of Agronomical Sci- CDC University of Pennsylvania, Zur Propstei enses & Veterinary Medicine 4770 Buford Hwy., F-22 Sch. Vet. Med. 55270 Schwagenheim, Splaiul Independental No. Atlanta, GA 30342 3800 Spruce St Germany 105, Sector 5 770-488-4414 Philadelphia, PA 19104-6050 49 6130 948211 Bucharest, Romania 50097 [email protected], ebk5@cdc. 215-898-7895 4.96131E+11 0040-21-318.04.69 gov 215-573-7023 christina.mertens@intervet. 0040-21-318.04.98 [email protected] com [email protected] Soraya Naem Urimia University, Fac. Vet. Robert A. Norton Jeffrey A. Meyer Daniel Moncol Med., Nazloo Campus Auburn University Poultry Sci- Eli Lilly and Company 1219 Brooks Avenue PO Box 1177 ence Dept. PO Box 708, GL21, 2001 Raleigh, NC 27607 Urmia, Iran 37135 236 Upchurch Hall West Main St. 919-782-1025 98(441)2776142 Auburn, AL 36849-5416 Greenfield, IN 46140 98 (441)3443442 334-844-2604 317-651-3233 Andrew R. Moorhead [email protected] 334-844-2649 317-277-4788 University of Georgia [email protected] [email protected] Dept. Inf. Diseases, Coll. Vet. Sundar Natarajan Med. 10403 46th Ave, Apt #302 Bruce Nosky Daniel K. Miller 501 D.W. Brooks Drive Ath- Beltsville, MD 20705 Merial 2209 Mayer Ave. ens, GA 301-443-2376 P.O. Box 25485. Lancaster, PA 17603 30602 [email protected] Macdonald campus, Arizona 717-393-7016 706-542-8168 85255 [email protected] Tom Nelson 480-747-3193 James E. Miller American Heartworm Society [email protected] Louisiana State University, Jean Mukherjee 719 Quintard Ave Sch. Vet. Med. 200 Westboro Road Anniston, AL 36201 Tom O’Connor Dept. Pathobiological Sci- North Grafton, MA 1536 256-236-8387 IDEXX Laboratories ences 508-887-4756 256-236-5888 One Idexx Drive Baton Rouge, LA 70803 508-839-7911 [email protected] Westbrook, ME 4092 225-578-9652 [email protected] 207-556-4428 225-578-9701 Harold Newcomb [email protected] [email protected] Thomas Murphy Intervet Central Veterinary Research 200 Watt St. Ryan O’Handley Thomas A. Miller Laboratory Batesville, MS 38606 Murdoch University PO Box 4142 Backweston Campus, Young’s harold.newcomb@intervet. 54 Dotterel Way Sequim, WA 98382 Crossing com Yangebup, WA, Australia 6150 360-582-0985 Celbridge, Co. Kildare, Ireland +61 8 9360 2457 360-582-0986 Co. Kildare Brain Newman +61 8 9310 4144 [email protected] 3531615-7141 Virginia Tech [email protected] tom.murphy@agriculture. VA/MD Reg. Coll. Vet. Med. Zachary T. Mills gov.ie Blacksburg, VA 24061-0442 Pia U. Olafson Merial Limited, Bldg. 500, 362 240-893-8473 USDA, ARS, Knipling-Bush- 3239 Satellite Blvd K. Darwin Murrell [email protected] land US Livestock Insects Duluth, GA 30096 Royal Veterinary & Agricultural Research Lab 678-638-3834 University, Center for Exp. Martin K. Nielsen 2700 Fredericksburg Rd 678-638-3873 Parasitol. Royal Vet & Agricultural Univ. Kerrville, TX 78028 [email protected] 5126 Russett Road Dept. Large An. Sci., Dyrlæ- 830-792-0322 Rockville, MD 20853 gevej 88, Frederiksberg 830-792-0314 Pamela S. Mitchell 301-460-9307 Copenhagen, Denmark DK- [email protected] Novartis Animal Health US [email protected] 1870 3200 Northline Ave, Suite 300 45 35 28 28 42 Itziar Olea Greensboro, NC 27408 Rainer K. Muser 45 35 28 28 38 Oxoid S.A. 800-447-2391, Ext. 1691 18 Montgomery Ave. [email protected] Calle Jose Bergamin, 40, 8°D 504-891-8716 Rocky Hill, NJ 8553 Madrid, Spain, 28030 pamela.mitchell@novartis. 609-924-1802 Robert HR Nixon 34913711644 com [email protected] Pfizer Animal Health Canada [email protected] 17300 Trans-Canada Hwy Sheila M. Mitchell Gil Myers Kirkland QC H9J 2M5 Merle Olson Scynexis, Inc. Gil Myers, Ph.D., Inc. Canada MBEC Bio Products, Univer- Durham, NC 3289 Mt Sherman Road 519-432-5502 sity of Calgary 540-231-7074 Magnolia, KY 42757 514-693-4272 Room 025, Bio Sciences, 540-231-3426 270-324-3811 [email protected], shan- 2500 University Ave NW [email protected] 270-324-3811 [email protected] Calgary, Alberta, T2N 4N1 [email protected] Canada 403-220-6835 403-270-0954 [email protected]

99 Dan A. Ostlind Sarah Parker Maria Pena Kathleen M. Picciano Merck & Co, (Retired) CFIA, Saskatoon Lab Louisiana State University, Manor College 94 Dearhook Rd 116 Veterinary Road Lab Res. Branch, Hansen’s 700 Foxchase Rd Whitehouse Station, NJ 8889 Saskatoon, SK, Canada S7N Disease Program Jenkintown, PA 19046 908-236-9238 2R3 Sch. Vet. Med., Skip Bertman 215-885-2360 908-236-9238 306-975-5996 Dr. 856-358-7315 [email protected] 306-975-5711 Baton Rouge, LA 70803 [email protected] [email protected] 225-578-9860 Jelena Ostojic [email protected] Rhonda D. Pinckney Iowa State University, Dept. Mukund Parkhie St. George’s University, Sch. Vet. Pathobiology Regulatory Consultant Andrew S. Peregrine Vet. Med., Dept. Paraclinical 1789 College Vet. Med. 19353 Elderberry Terrace University of Guelph, Ontario Studies Ames, IA 50011 Germantown, MD 20876-1647 Veterinary College True Blue 515 294-0959 301-972-5251 Dept. Pathobiol. Grenada, West Indies [email protected] [email protected] Guelph, Ontario N1G 2W1, 473-444-4175. Ext. 2448 CANADA 473-439-5068 Nadia Ouellette Jim C. Parsons 519-824-4120 Ext. 54714 [email protected] Atlantic Veterinary College Agriculture Victoria 519-824-5930 550 University Ave. 475 Mickleham Road [email protected] Edward G. Platzer Charlottetown, PEI, Canada Attwood, Victoria, Australia University of California - C1A 4P3 3049 Adalberto A Perez de Leon Riverside 902-569-6096 61-3 92174275 ARS, USDA Dept. Nematology, 4205 [email protected] 61-3 92174161 2700 Fredericksburg Road Carney Court [email protected] Kerrville, TX 78028 Riverside, CA 92507 Paul A.M. Overgaauw 830-792-0304 951-827-4352 Consultant Vet. Parasitol. & Tara Paterson [email protected]. [email protected] Zoonoses St. George’s University gov Molecaten 57 P.O. Box 7, True Blue Kenneth C. Plaxton Barneveld, The Netherlands Granada Gabriela Perez Tort Elsevier 3772 LJ 473-435-2900 University of Buenos Aires Molenwerf 1 31-342-421533 473-435-2997 Jose p. Varela 3152 Amsterdam, The Netherlands 31-342-419794 [email protected] Buenos Aires, Argentina 1417 1014 AG [email protected] 00541º145032095 31 20 485 3332 Sharon Patton [email protected], gabri- 31 20 485 3249 Brooke Pace University of Tennessee, Coll. [email protected] [email protected] Pfizer Animal Health Vet. Med., Dept. Comparative 812 Springdale Drive med., Rm A205 Priscilla M. Peterson Linda Marie Pote Exton, PA 19341 2407 River Dr. P.O. Box 1364 Mississippi State University, 800-366-5288 Knoxville, TN 37996-4543 Janesville, WI 53547-1364 Coll. Vet. Med. 866-590-1149 865-974-5645 608-756-0799 Box 6100, Wise Center Spring [email protected] 865-974-5640 [email protected] St. [email protected] Mississippi State, MS 39762 Azhahainambi Palavesam Raffaele Petragli 662-325-1130 18763 Nathan Place Michael Paul Via F.lli Rosselli, 27 662-325-8884 Montgomeryville, MD 20866 CAPC Montescudaio, Italy PXX [email protected] [email protected] 1210 Vance Ct. 56040 Bel Air, MD 21014 [email protected] Kerrie Powell Kathleen G. Palma 510-315-0550 Scynexis. Inc. Kado Enterprises 410-838-8530 John E. Philliips P.O. Box 12878 815 Cliff Dr. [email protected]; mapaul. Pfizer Animal Health Research Triangle Park, NC McLeansville, NN 27301 [email protected] 2200 NW 23rd Way 27709 336-708-2842 Boca Raton, FL 33431 919-206-7223 336-697-7296 Patricia A. Payne 561-995-1648 [email protected] [email protected] Kansas State University, Coll. 561- 995-0574 Vet. Med. [email protected] Roger K. Prichard John A. Pankavich 3005 Payne Dr. McGill University, Macdonald American Cyanamid (Retired) Manhattan, KS 66503 Celine Picard Campus 513 Thompson Avenue 785-236-9081 Bayer, Inc. 21111 Lakeshore Rd, Ste LeHigh Acres, FL 33972-4104 [email protected] 22 Brisson St., Box 235 Anne De Bellevue Limoges, Ontario, Canada Quebec, H9X 3V9 Canada Carla C. Panuska K0A 2M0 514-398-7729 Mississippi State University, 613-443-7664 514-398-7594 Coll. Vet Med. 613-443-5364 [email protected] P.O. Box 6100 [email protected] Mississippi State, MS 39762 662-325-1198 662-325-1031 [email protected]

100 Helen Profous-Juchelka Robert H. Rainer David M. Reinitz Carol K. Robertson- Plouch Merck Sharp & Dohme Corp. 10720 Club Chase USGS National Wildlife Health Eli Lilly 126 East Lincoln Ave Fishers, IN 46038 Center 5736 E. 400 S. Rahway, NJ 7065 317-598-9120 6006 Schroeder Rd Greenfield, IN 46140 732-594-5569 317-598-9120 Madison, WI 53711 317-433-1708 732-594-3624 [email protected] 608-270-2487 317-433-0448 [email protected] 608-270-2415 [email protected] Brenda J. Ralston [email protected] Frank L. Prouty Alberta Agriculture, Food & David W. Rock Pfizer Animal Health Rural Development Robert S. Rew Fort Dodge Animal Health 9225 Indian Creek Parkway, 909 Irricana Rd NE Rewsearch Consulting 590 Montgomery Rd Suite 400 Airdrie, Alberta, Canada T4A 400 N. Wawaset Rd Hillsborough, NJ 8844 Overland Park, KS 66210 2G6 West Chester, PA 19382 908-369-6413 913-664-7041 403-948-8545 610-918-1248 908-369-4163 913-217-6872 403-948-2069 484-356-0452 [email protected] [email protected] [email protected] [email protected] RaffaeleA. Roncalli John H. Pruett Siva Ranjan GC Richie 29 Louise Road Retired Pfizer Novartis Animal Health Milltown, NJ 8850 P.O. Box 379 8 Wheatston Court 6741 Layton Road 732-940-4070 Milano, TX 76556 Princeton, NJ 8550 Liberty, NC 27298 732-940-7881 334-826-8473 609-799-7746 336-402-4831 [email protected] 334-826-8473 [email protected] [email protected] [email protected] Douglas Ross Vijayaraghara Rao Robert K. Ridley Bayer HealthCare, Animal Joseph B. Prullage McGill University Kansas State University, Coll. Health Division Merial, Inc 2111 Lakeshore, Ste. Anne Vet. Med. 12809 Shawnee Mission 5498 Jade Road de Bell. Coles Hall Parkway Fulton, MO 65251 Montreal, Canada H9X 349 Manhattan, KS 66506 Shawnee, KS 66216 573-642-5977, Ext. 1168 vijayaraghara.rao@mail. 785-532-4615 9132682828 573-642-0356 mcgill.ca 785-532 4851 9132682541 [email protected] [email protected] [email protected] Lisa Rascoe David G. Pugh Auburn University, Coll. Vet. Sherri L. Rigby Mary G. Rossano Fort Dodge Animal Health Med. Bayer Animal Health University of Kentucky P.O. Box 26 151 Greene Hall 804 Greentree Arch 255 Handy’s Bend Rd Waverly, AL 36879 Auburn, AL 36849 Virginia Beach, VA 23451 Wilmore, KY 40390 334-826-8473 251-554-8948 888-833-3124 859-257-7552 334-826-8473 334-844-2652 757-491-8752 [email protected] [email protected] [email protected] [email protected] Alexa Rosypal Trisha Putro Thilaker Rathinam John L. Riner University of North Carolina Bristol-Myers Sqibb University of Arkansas KMG Animal Health at Chapel Hill, Dept. Path./ Route 206 & Provinceline Rd 900 N. Leverett Ave #120 502 South 15th St., POB 80 Lab. Med. Princeton, NJ 8543 Fayetteville, AR 72701 Elwood, KS 66024 CB #7525, 805 Brinkhous- 609-252-5557 479-575-7230 913-365-5215, x301 Bullitt 609-252-6896 479-575-3026 913-365-5275 Chapel Hill, NC 27599-7525 [email protected] [email protected] [email protected] 919-966-4294 919-966-0704 Tariq Qureshi Mason Reichard Edward L. Roberson [email protected] Schering-Plough Oklahoma State University 190 Sunnybrook Drive 490 Franklin Circle Coll. Vet. Med., Dept. Pathobi- Athens, GA 30605 Harvey L. Rubin Yardley, PA 19067 ology, 250 McElroy Hall 706-543-1680 P.O. Box 421836 908-629-3707 Stillwater, OK 74078-2007 706-542-0059 Kissimmee, FL 34782 [email protected] 405-744-8159 407-847-8162 405-744-5275 Alan P. Robertson 407-847-0127 Alex Raeber [email protected] Iowa State University Prionics AG 2058 Coll. Vet. Med. Tony Rumschlag Wagistrasse 27A Craig R. Reinemeyer Ames, IA 50011 Merial Schlieren, Zurich, Switzerland East Tennessee Clinical 515-294-1212 614 White Pine Drive 8952 Research 515-294-2315 Noblesville, IN 46062-8533 4144 200-2000 80 Copper ridge Farm Rd. [email protected] 317-773-5181 [email protected] Rockwood, TN 37854 317-773-1224 865-354-8420 [email protected] 865-354-8421 [email protected]

101 Nicholas C. Sangster Rudolf Schenker Barbara Shock Owen Slocombe Charles Sturt University Novartis Animal Health, WRO- University of Georgia University of Guelph, Ontario School of Agricultural and 1033.3.19 313 Oak Grove Dr Vet. Coll. Veterinary Sciences, Locked CH-4002 Grantsville, WV 26147 29 Oak Street Bag 588 Basel, Switzerland 706-542-1741 Guelph, Ontario, N1G 2N1 Wagga Wagga, N.S.W. Aus- 41 61 697 4776 [email protected] Canada tralia 2678 41 61 697 6788 519-821-7222 02 6933 4107 [email protected] Rick Sibbel 519-824-5930 02 6933 2812 Intervet/Schering-Plough AH [email protected] [email protected] Thomas Schnieder 1401 NW Campus Dr. Institute for Parasitology, Uni- Ankeny, IA 50023 Robyn L. Slone Monica Santin-Duran versity of Veterinary Medicine, 515-249-2111 PLRS USDA, ARS, ANRI, EMSL Hannover [email protected] 1251 NC 32 North Bldg. 173 Barc-East Buenteweg 17 Corapeake, NC 27926 10300 Baltimore Ave. Hannover, Germany D-30559 Gary Sides 252-465-8686 Beltsville, MD 20705 +49 0511-953- 8711 Pfizer Animal Health 252-465-4493 301-504-6774 +49 0511-953-8870 13355 CR 37 [email protected], 301-504-6608 thomas.schnieder@tiho- Sterling, CO 80751 [email protected] [email protected], hannover.de 970-520-5953 monica.santin-duran@ars. [email protected] Gary Smith usda.gov Philip J. Scholl Sch. Vet. Med, Univ. of Penn- Marques do Herval 315, Apto Christian Eric Simmons sylvania Raj Saran 602, P.O. Box 675 4011 Cordova Dr. NBC, 382 West St. Road MGK Porto Alegre, RS, Brazil, Austin, TX 78759 Kennett Square, PA 19348 8010 Tenth Ave North Richland Center, WI 90570- 610-444-3129 Minneapolis, MN 55427 140, 53581 Everett E. Simmons 610-925-8130 [email protected] 011-55-51-3346-9202 Burnet Road Animal Hospital [email protected] [email protected] 8511 Burnet Rd. Mohamed Z. Satti Austin, TX 78759 Harry J. Smith St. Matthew’s University, Bettina Schunack 512-452-7606 15 Princess St School of Vet. Medicine Novartis Animal Health, Inc. [email protected] Sackville, New Brunswick, 6504 Kenneaw Rd Schwarzwaidalle 215 E4L 4E9 Canada Canton, MI 48187 Basel, Switzerland 4058 Anthony Simon 506-536-1445 345-927-5752 4179 8260307 Pfizer Animal Health [email protected] bettina.schunack@ 150 East 42nd St Larry L. Smith novartis,com New York, NY 10017 Larry L. Smith DVM, R&D, John Schaefer 646-509-9140 Inc. Cornell University George C. Scott [email protected] 108 Davis Street 258 Bundy Road 800 Hessian Circle Lodi, WI 53555 Ithaca, NY 14850 West Chester, PA 19382 Greg L. Simons 608-592-3275 607-229-7186 610-793-1852 Novartis Animal Health 608-592-5118 [email protected] [email protected] 11205 W. 140th Place [email protected] Overland Park, KS 66221 James H. Schafer R. Lee Seward 800-447-2391 Stephen A. Smith Schafer Veterinary Consul- Seward Farms LLC 913-851-0472 Virginia Tech, VA-MD Reg. tants PO Box 6 [email protected] Coll. Vet. Med. 800 Helena Ct. Eaton, CO 80615-0006 Phase II, Duck Pond Dr. Fort Collins, CO 80524 970-834-0216 Hal R. Sinclair Blacksburg, VA 24061-0442 970-224-5103 970-834-0216 Teva Animal Health, Inc 540-231-5131 970-224-5882 [email protected] 8609 N.W. Shannon Ave. 540-231-6033 jschafer@schaferveterinary. Kansas City, MO 64153 [email protected] com Thomas A. Shelton 816-676-6108 Intervet, Inc. 816-676-6877 Karen Snowden Peter M. Schantz 1085 Chama Court [email protected] Texas A&M University, Coll. 2856 Woodland Park Washington, Utah 84780 Vet. Med. Atlanta, GA 30345 435-986-1853, cell 208-867- Frank Slansky Dept. Vet. Path., Mail Stop 404-982-0591 3502 University of Florida, Dept. 4467; Texas A&M [email protected] 435-986-1853 Ento. & Nematology College Station, TX 77843- [email protected] POB 110620 4467 Roland Schaper Gainesville, FL 32611 979-862-4999 Bayer Animal Healh SungShik Shin 352-332-2001 979-862-2344 Bayer Health Care AG Chonnam National University [email protected] [email protected] Leverkusen, Germany 57368 Coll. Vet. Med., 300 Yong- 4.92773E+11 bong-dong roland.schaper@bayerhealth- Gwangju, South Korea 500- care.com 757 [email protected]

102 Daniel E. Snyder David G. Stansfield Roger Stich Sivapong Sungpradit Elanco Animal Health Pfizer Animal Health University of Missouri, Dept. Depsrtment of Molecular Mail Drop GL 14, P.O. Box 3200 Northline Ave, Suite 300. Pathobiology Biology 708, 2001 W. Main St. Greensboro, NC 27408 209 Connaway Hall 615 N. Wolfe Greenfield, IN 46140 336-255-0201 Columbia, MO 65211 Baltimore, MD 21205 317-277-4439 336-387-1030 573-882-3148 410-955-3442 312-277-4522 [email protected], david. [email protected] [email protected] [email protected] [email protected] Heather D. Stockdale Xun Suo Cheryl D. Sofaly Lindsay Starkey University of Florida, Coll. Vet. China Agricultural University, US Army Oklahoma State University Med. Coll. Vet. Med. 572 Acorn Ln 250 McElroy Hall 2484 Royal Point Drive Yuanmingyan West Road 2#, Killeen, TX 76542 Stillwater, Ok 74074 Green Cove, FL 32043 Haidian Disrict 254-288-2894 405-744-3236 352-294-4142 Bejing, China 100193 254-287-4676 [email protected] [email protected] 861061 273-4325 [email protected]. [email protected] army.mil James H. Steele Bob Storey University of Texas – Houston, UGA Ritesh Tandon Mark D. Soll Sch. Public Health 1224 McGinnis Chandler Rd University of Georgia, Coll. Merial Ltd. Box 20186 Astrodome Stn Commerce, GA 30530 Vet. Med., #2212 3239 Satellite Blvd Houston, TX 77225 [email protected] 501 DW Brooks Drive Duluth, GA 30096-4640 713-500-9361 Athens, GA 30602 678-638-3605 713-500-9364 B. E. Stromberg 706-542-0742 678-638-3668 University of Minnesota 706-542-0059 [email protected] Nina R. Steenhard 1971 Commonwealth Ave [email protected] Centre of Experimental Para- St. Paul, MN 55108 Smaragda Sotiraki sitology 612-625-7008 Patrick Tanner NAGREF-VRI 100 Dyrlaegevej 612-625-4734 Merial NAGREF Campus Thermi 1870 Frederiksberg, Copen- [email protected] PO Box 661 Thessaloniki, Greece 57001 hagen, Denmark Crawford, GA 30630 30231 936-5373 45 35 282788 John A. Stuedemann 706-759-3859 [email protected] 4534282774 USDA, ARS [email protected] [email protected] 1420 Experiment Station Rd Leticia M Souza-Dantas Watkinsville, GA 30677-2373 Cynthia M. Tate Universidade Federal Flumi- Michael Stegemann 706-769-5631; Ext. 247 University of Georgia, SC- nense Pfizer Ltd, Animal Health 706-769-8962 WDS, Coll. Vet. Med. Rua Maria da Ajuda Medeiros, Group [email protected] Wildlife Health Building 31/casa 1 – Loteamento Boa IPC 896, VMR&D, Ramsgate Athens, GA 30602-4393 Vista, Itaipu Road Chunlei Su 706-542-1741 Niteroi, RJ - Brazil 24.340-170 Sandwich, England CT13 9NJ University of Tennessee, [email protected] 55 21 2609 3155 44-1304-646121 Knoxville [email protected], 44-1304-656257 701 Dawson Creek LN Tiffany Taylor [email protected] michael.stegemann@pfizer. Knoxville, TN 37922-9007 Tuskegee University com 612-625-7008 90 Neal Street Jennifer Spencer 612-625-5203 Crawford, GA 30602 Auburn University, Coll. Vet. Joyce Steinbock [email protected] [email protected] Med. BioMed Diagnostics, Inc. Dept. Pathobiology, 166 1388 Antelope Rd Miguel T. Suderman Stephan Tegarden Greene Hall White City, OR 97503 Cell Systems 3-D, LLC Intervet/Schering Plough Auburn University, AL 36849 541-830-3000 426 Starborrough Drive 22694 W. 267th Street 334-844-2701 541-830-3001 League City, TX 77573-5937 Paola, KS 66071 334-844-2652 joyce@biomeddiagnostics. 281-389-1861 913-422-6828 [email protected] com 281-334-2459 stephan.tegarden@ msuderman@cellsystems3D. sp.intervet.com Earl J. Splitter Jay Stewart com Apt. 2403 CAPC Kevin B. Temeyer 9221 W. Broward Blvd P.O. Box 230 Woraporn Sukhumavasi, USDA, ARS, Knipling-Bush- Plantation, FL 33324 Aumsville, OR 97325 Chulalongkorn University, land US Livestock Insect Res. 503 448 1855 Faculty of Veterinary Science, Lab. Madeleine S. Stahl [email protected] Parasitology Unit, Dept. of 2700 Fredericksburg Rd. Intervet, Inc. Pathology Kerrville, TX 78028 29160 Intervet Lane, PO Box T. Bonner Stewart Henri-Dunant Rd., Pathum- 830-792-0330 318 Louisiana State University wan 830-792-0314 Millsboro, DE 19966 5932 Forsythia Ave. Bangkok, Thailand 10330 [email protected] 302-934-4362 Baton Rouge, LA 70808 662-218-9664 302-934-4281 225-766-8327 668-5-351-1080 [email protected]. [email protected] [email protected] com Jerold H. Theis Ba Cuong Tu Andrea S. Varela-Stokes Bradley J. Waffa University of California-Davis, Modern Veterinary Therapeu- Mississippi State University, University of Tennessee, Sch. of Medicine tics, LLC Coll. Vet. Med. Sewanee Dept. Med. Micro., 1 Shields 18301 SW 86 Avenue Dept of basic Sciences, Wise 735 University Ave Ave. Miami, FL 33157 Center Sewanee, TN 37383 Davis, CA 95616 305-232-6645, cell: 786-200- Mississippi State, MS 39762 630-484-3435 530-752-3427 2495 662-325-1345 [email protected] 530-752-8692 [email protected] , [email protected] [email protected] cuong.tuba@modernveterin- Kenneth A. Waldrup arytherapeutics.com Julie Vargas Texas Animal Health Com- V. J. Theodorides University of Georgia mission 1632 Herron Lane Mary E. Ulrick 233 Sleepy Creek Dr. P.O. Box 3041 West Chester, PA 19380 Cheri-Hill Kennel & Supply, Athens, GA 30606 Cleburne, TX 76033 610-696-5493 Inc. 478-461-3185 817-783-6493 610-701-6395 17190 Polk Rd [email protected] 817-783-6493 [email protected] Stanwood, MI 49346 [email protected] 231-823-2392 Adriano Vatta Patricia Thomblison 231-823-2925 Ross University Tracy I. Ward 2711 SW Westport Dr. [email protected] P.O. Box 334 Bayer Animal Health Topeka, KS 66614 Basseterre, Saint Kitts 99999 47 Meadow Pointe Drive 785-271-6005 Michael A. Ulrick 869-763-7875 DeWinton, Alberta, TOL OXO 785-273-9505 Cheri-Hill Kennel & Supply, [email protected] Canada [email protected], Inc. 403-201-4712 [email protected] 17190 Polk Rd Jozef Pieter Vercruysse 403-201-3643 Stanwood, MI 49346 Ghent University, Faculty Vet. [email protected] Philip R. Timmons 231-823-2392 Med., Lab Parasitol Scynexis, Inc. 231-823-2925 Salisburylaan 133 Andre Weil 3501C Tricenter Blvd. [email protected] Merelbeke, BELGIUM B 9820 Avogadro Durham, NC 27713 32-9-264 7390 185 Alewife Brook Parkway 919-549-5233 Govind G. Untawale 32-9-2647496 Cambridge, MA 2138 919-549-5240 Hartz Mountain Corp. [email protected] 403-201-4712 [email protected] 281 Farmingdale Road 403-201-3643 Wayne, NJ 7470 Isabelle Verzberger andre.weil@avogadro-lab Gabriela Perez Tort 201-271-4800, Ext. 7760 Atlantic Vet. Coll. Jefe de Clin.: Hosp. Vet de 201-271-0357 550 University Ave Henry Weinberg Virreyes; J.T.P.: Parasitología [email protected] Charlottetown, PEI, CANADA Pfizer Animal Health y Enfermedades Parasitarias. C1A 4P3 812 Springdale Dr Fac Cs. Vet. UBA Joseph F. Urban Jr. 902-566-0843 Exton, PA 19341 Acceso Norte 2502,San Fer- USDA, ARS, BHNRC, NRFL [email protected] 610-363-3273 nando, pcia de Buenos Aires, Bldg. 307 C, BARC-East [email protected] Argentina Beltsville, MD 20705 John M. Vetterling Buenos Aires 301-504-5528 Parasitologic Services Stephen K. Wikel 301-504-9062 P.O. Box 475 University of Connecticut Argentina [email protected] Fort Collins, CO 80522-0475 Health Center, Sch. of Medi- 00 54 11 4745-8612; 00 54 11 970-484-6040 cine, Dept. Immunology 4580-2820 Jennifer Uribe 970-221-0746 263 Farmington Avenue, [email protected] University of Illinois [email protected] MC3710 1506 S. Jefferson St Farmington, CT 6030 Jennifer Towner Lockport, IL 60441 Eric Villegas 860-679-3369 190 Round Table Ct. 630-220-8824 Environmental Protection 860-679-8130 Athens, GA 30606 [email protected] Agency [email protected] 912-441-9921 26 W. MLK Dr. (MS:320) [email protected] Wendy Vaala Cincinnati, OH 45268 Heike Williams Schering Plough - Intervet [email protected] Intervet Joseph P. Tritschler 51476 Pleasant View Road [email protected] Virginia State University, Alma, WI 54610 Alain Villeneuve Agriculture Research 608-685-4560 University of Montreal James C. Williams PO Box 9061 [email protected] 3200 Sicotte, C.P. 5000 5214 N. Chalet Ct. Petersburg, VA 23842 St. Hyacinthe, Quebec J2S Baton Rouge, LA 70808-4843 804-524-5957 Frank van Knapen 7C6 CANADA 225-766-4728 804-524-5186 Utrecht University 450-773-8521, Ext 8405 [email protected] [email protected] P.O. Box 80175, 3508 TD 450-778-8116 Utrecht, the Netherlands 3508 alain.villeneuve@umontreal. 31-30-253-5367 ca 31-30-253-2365 [email protected]

104 Jeffrey F. Williams Michael Yabsley Dante Zarlenga Haolsource, Inc. University of Georgia USDA, ARS, ANRI, Bovine 1631 220th Street, Suite 100 SCWDS, coll. Vet. Med. Func. Genomics Lab Bothell, WA 98021 Athens, GA 30602 Bldg. 1180, BARC-East 425-974-1949, 208-390-7178 706-542-1741 Beltsville, MD 20705 425-882-2476 [email protected] 301-504-8754 [email protected], 301-504-8979 www,halosource.com Tom A. Yazwinski [email protected] University of Arkansas, Dept. Andrea Wilson of Animal Science Guan Zhu Novartis AH Canada 1120 W. Maple Texas A&M University, Dept. 2000 Argentia Rd., Plaza 3, Fayetteville, AR 72701 Vet. Pathobiology Suite 400 479-575-4398 4467 TAMU Mississauga, ON, L5N 1V9, 479-575-5756 College Station, TX 77843- Canada [email protected] 4467 905-814-4897 979-845-6981 905-567-0221 Sekar Yokananth 979-845-9972 [email protected]. University of Alberta [email protected] com 2148 Michener Park NW Edmonton T6H4M5 Gary L. Zimmerman John B. Winters 780-436-7930 Zimmerman Research Beverly Hills Sm. An. Hospital [email protected] 1106 W. Park, PMB 424 353 N. Foothill Road Livingston, MT 59047 Beverly Hills, CA 90210 Timothy Yoshino 406-223-3704 310-276-7113 Dept. Pathobiological Sci- 406-223-7829 310-859-0411 ences [email protected] [email protected] 2115 Ob Madison, WI 53706 Erich Zinser Adrian Wolstenholme 608-263-6002 Pfizer Animal Health University of Georgia [email protected]. 7000 Portage Road Dep. Infectious Diseases, edu Kalamazoo, MI 49001 Coll. Vet. Med. 269-833-2401 Athens, GA 30603 David R. Young [email protected] [email protected] Young Veterinary Research Services Ming M. Wong 7243 East Avenue University of California – Turlock, CA 95380-9124 Davis 209-632-1919 Vet. Path. Micro. & Immun. 209-632-1944 Davis, CA 95616 [email protected] 916-756-0460 916-756-0460 Silene Young [email protected] Bayer HealthCare P.O. Box 390 Jerry Woodruff Shawnee Mission, KS 66201- 8182 Moran Canyon Road 0390 North Platte, NE 64101 913-268-2867 [email protected] 913-268-2878 [email protected] Debra Woods Pfizer Animal Health Cole Younger VMRDCIPCD883 Ramsgate Stillmeadow, Inc. Rd. [email protected] Sandwich, Kent, CT139NJ, U.K. Anne M. Zajac 4413046473 Virginia Tech 4.41305E+11 VA/MD Reg. Coll. Vet. Med. [email protected] Blacksburg, VA 24061-0442 540-231-7017 Dehai Xu 540-231-6033 USDA Agricultural Research [email protected] Services P.O. Box 952 Aquatic Animal Daniel Zarate Rendon Health Research University of Georgia Auburn, AL 36831-0952 160 Dudley Drive, Apt 505 334-887-3741 Athens, GA 30606 334-887-2983 706-255-1544 [email protected] [email protected]

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