Proceedings AAVP Membership Directory

Transcription

Photo Credit: Rich Grant & VISIT DENVER
Photo Credit: Bob Ash & VISIT DENVER
Proceedings AAVP
American Association of Veterinary Parasitologists
Emerging Issues
Photo Credit: Rich Grant & VISIT DENVER
Membership Directory
59th Annual Meeting
July 26-29, 2014 t Denver, CO
Animal Health
A SANOFI COMPANY
59th Annual Meeting, 26-29 July 2014
The Curtis Hotel, Denver, CO
2014 AAVP-Merial Distinguished Veterinary Parasitologist
Timothy G. Geary, Ph.D.
Tim Geary was born in Grand Rapids, Michigan. One of 6 siblings raised by a school teacher and a
(then) stay at home mom, he enjoyed a quintessential baby boomer childhood. There wasn’t a lot of
money but there was always enough food and hand-me-down clothes that were just fine. Seasonal
sports were a constant source of fun, even though he wasn’t particularly accomplished at any of
them. Coming from these parents, academic achievement was expected and attained. High school
for Tim happened during a time of great change in America and around the world; it made a
significant impact on him which is evident today (just check the hair style). He also learned to
work, beginning as a janitor cleaning toilets after school at a local elementary school at the age of
14. This experience prepared him well for his eventual evolution into a parasitologist. But it was his
position in a pharmacy that determined his career trajectory and led to a life-long fascination with
drugs (as medicines, mind you). He was fortunate to be able to attend the University of Notre
Dame, alma mater to his father and 4/5 of his siblings. He majored in Biology and had enough
credits (not all the right ones) to minor in chemistry, graduating cum laude in 1975. He learned a lot
there, but perhaps the most important lesson was that there are a lot of smart people in the world.
It’s not so rare or special to do well academically; it’s what you do as a result that counts. He also
developed a deep interest in parasitology through the teaching of Howard Saz and Rob Rew, then a
post-doc in Howard’s lab.
From ND, Tim continued his Midwestern sojourn by pursuing a PhD in Pharmacology at the
University of Michigan, studying the pharmacogenetics of cytochrome P450 enzymes involved in
drug metabolism under the supervision of a terrific mentor, Dr. Bert La Du. It was at that point in
his life that the parasitology infection fully developed. Tim joined the parasitology group at
Michigan State University (continuing the Midwestern theme), then under the direction of the
eminent veterinary parasitologist Dr. Jeff Williams. Tim joined the lab of Dr. Jim Jensen to work
on malaria parasites, particularly from the perspective of chemotherapy. Under Jeff’s guidance,
MSU had developed into a dynamic and flourishing mecca of parasitology. Tim was committed. It
was at MSU that he met his wife and partner in love for life Alma, who was working in the
Williams lab. Once she got over the fact that Tim was a Michigan grad, things developed rapidly.
2
After 5 years at MSU, it was tim e to move on. Tim was fortunate to find a position at the Upjohn
Company in Kalamazoo (the Midwest magnet at work again) in the animal health division. Once
again, he was f ortunate to be able to work with outstanding parasitologi sts like Bud Folz and
George Conder, along with m any other excellen t friends and colleagues and great m anagers,
especially Bill Maxey, Steve Nelson, Dave Lowery, John Welser and George Gunn. Kalam azoo
was a great place to raise a family (Jim and then Rose) and the Upjohn Company was a great place
to work. In particular, Tim formed a great team with Dr. Dave Thompson, another MSU product.
At Upjohn, Tim was able to forg e a link betw een basic and appl ied research, leading to new
discoveries about neuropharm acology and participating in the discovery and developm ent of
2-desoxoparaherquamide (derquantel) as a new anthelmintic. The group actively participated in the
currency of science, regularly publishing and pres enting their work at na tional and international
conferences. He developed any close friendships and collaborations with veterinary parasitologists
that still thrive today.
The acquisition of Upjohn (by then Pharmacia) by Pfizer brought a lot of changes. On the positive
side was the opportunity to work with another set of terrific colleagues, exemplified by Dr. Debra
Woods. However, Tim was m oved out of research into a business devel opment role, and this
probably was not the best place for him (certainly based on hi s habits of groom ing and dress
alone!). When he was asked to interview for a position as a professor and Canada Research Chair at
the Institute of Parasitology at McGill University, the allure of a return to research was too hard to
resist. Farewell for now, Midwest!
Since joining McGill, Tim has been f ortunate to work yet again with another eminent veterinary
parasitologist, Roger Prichard, and to continue his work on drug discovery and development. This
work is focused on finding non-peptide ligands
for invertebrate peptide G protein-coupled
receptors, a quest of almost 30 years duration (now with treasured colleague Dr. Eliane Ubalijoro).
In a separate project, Tim is working on the development of flubendazole as a macrofilaricide (with
close friend and colleag ue Charles Mackenzie of MSU, who originated the project, along with
collaborators at Janssen Research and Development). He is still working on macrocyclic lactones,
including how they work as filaricides and how resistance to th em has developed in heartworm
populations (with Roger Prichard). He is also returning to basic research and is studying how the
molecules secreted by parasites influence host i mmune responses. He has been Director of the
Institute of Parasitology since 2008 (but has perfected the role of Col. Blake to Shirley Mongeau’s
Radar O’Reilly, so this is not much of a burden) and has thoroughly enjoyed teaching and
supervising excellent graduate students, including 3 PhD student s who have earned their degrees
with him: Yovany Moreno, Vanessa Dufour and Elizabeth Ruiz Lancheros. It has been a privilege
to come back to research. His output of research articles and book chapters has reached almost 200
along with a number of patents. He regrets being far away from family and old friends, especially
from his grandchildren Olivia and Isaac back in Kalamazoo. When he is forcibly removed from the
Institute, Tim likes to fish and play golf (and wonder what adventure lies ahead).
3
AMERICAN ASSOCIATION OF VETERINARY PARASITOLOGISTS
AWARDS HISTORY
AAVP-Merial Distinguished Veterinary Parasitologist Award
1985
1986
1987
1988
1989
1990
1991
1992
1993
1994
1995
1996
1997
1998
1999
2000
2001
2002
2003
2004
2005
2006
2007
2008
2009
2010
2011
2012
2013
Jitender P. Dubey*
Norman D. Levine
E. J. Lawson Soulsby
Jeffrey F. Williams
K. Darwin Murrell
William C. Campbell
Jay Hal Drudge and Eugene T. Lyons
Gilbert F. Otto
Thomas R. Klei
Peter M. Schantz
James C. Williams
T. Bonner Stewart
J. Owen D. Slocombe
J. Ralph Lichtenfels
Roger K. Prichard
Edward L. Roberson
Byron L. Blagburn
Sidney A. Ewing
Louis C. Gasbarre
David S. Lindsay
Jorge Guerrero
John W. McCall
Ronald Fayer
Dwight D. Bowman
Ellis C. Greiner
George A. Conder
Thomas M. Craig
James E. Miller
Dante Zarlenga
*National Academy of Sciences 2010
2014 AAVP-Merial Distinguished Veterinary Parasitologist Award Winner
Timothy G. Geary
AAVP Distinguished Service Award
1976
1983
1987
1988
1994
1997
2006
2008
Rurel R. Bell
Terance J. Hayes
Norman F. Baker
Donald E. Cooperrider
S. D. “Bud” Folz
Honorico Rick Ciordia
Raffaele “Raf” Roncalli
Anne M. Zajac
4
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
Scott T. Storandt
1996
A. Lee Willingham III
1997
Carla C. Siefker
1998
Ryan M. O’Handley
1999
John S. Mathew
20001
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
20082
Heather D. Stockdale
2009
Kelly E. Allen
2010
Stephanie R. Heise
2011
Aaron S. Lucas
20123
Flavia A. Girao Ferrari
2013
Lindsay A. Starke
1
2000, award renamed the “AAVP/Intervet Graduate Student Research Award”
2
2008, award renamed the “AAVP/Schering-Intervet Graduate Student Research Award”
3
2012, award renamed the “AAVP/Merck Animal Health Graduate Student Research Award”
2014 AAVP/Merck Animal Health Graduate Student Research Award
Alice Che Yu Lee
AAVP-Companion Animal Parasite Council (CAPC) Graduate Student Award in Zoonotic Disease
2008
David G. Goodman
2009
Stephanie R. Heise
2010
Sriveny Dangoudoubiyam
2011
Jessica Edwards
2012
Lindsay Starkey
2013
Gail M. Moraru
2014 AAVP-CAPC Graduate Student Award in Zoonotic Disease
Anne Barrett
5
Founded 1956
Officers 2013-2014
President
Dwight D. Bowman
Cornell University
Ithaca, NY
President-Elect
Andrew S. Peregrine
University of Guelph
Guelph, ON, Canada
Vice-President
Ray M. Kaplan
University of Georgia
Athens, GA
Secretary/Treasurer
Robert G. Arther
Bayer HealthCare, Animal Health
Shawnee, KS
Immediate Past-President
Alan A. Marchiondo
Zoetis
Kalamazoo, MI
2013-2014 AAVP Committee Chairs
Program - Andrew S. Peregrine, Guelph, ON, Canada
Archives - Tom J. Nolan, Philadelphia, PA
Historian - Raf Roncalli, Milltown, NJ
Awards - Dan Snyder, Greenfield, IN
Constitution/Bylaws - Wendell L. Davis, Overland Park, KS
Education - Gary Conboy, Charlottetown, PEI, Canada
Finance - Andrew R. Moorhead, Athens, GA
Newsletter - Jenifer Edmonds, Parma, ID
Nominations - James E. Miller, Baton Rouge, LA
Outreach/Research - Lora R. Ballweber, Fort Collins, CO
Publications/Internet - Tariq Qureshi, Shawnee, KS
Student Representatives - Melissa Beck, Lethbridge, Alberta, Canada &
Javier Garza, Baton Rouge, LA
List Serve Manager - Bert Stromberg, St. Paul, MN
6
PAST PRESIDENTS
OF THE
AMERICAN ASSOCIATION OF
VETERINARY PARASITOLOGISTS
1956-1958
1958-1960
1960-1962
1962-1964
1964-1966
1966-1968
1968-1970
1970-1972
1972-1973
1973-1975
1975-1977
1977-1979
1979-1981
1981-1983
1983-1985
1985-1986
1986-1987
1987-1988
1988-1989
1989-1990
1990-1991
1991-1992
1992-1993
1993-1994
1994-1995
1995-1996
1996-1997
1997-1998
1998-1999
1999-2000
2000-2001
2001-2002
2002-2003
2003-2004
2004-2005
2005-2006
2006-2007
2007-2008
2008-2009
2009-2010
2010-2011
2011-2012
2012-2013
2013-2014
L. E. Swanson
Fleetwood R. Koutz
Wendell H. Krull
Saeed M. Gaafar
E. D. Besch
George C. Shelton
John H. Greve
Harold J. Griffiths
Demetrice L. Lyles
Harold J. Smith
Norman F. Baker
Edward L. Roberson
Jeffrey F. Williams
John B. Malone
Robert M. Corwin
K. Darwin Murrell
Thomas R. Klei
Harold C. Gibbs
Bert E. Stromberg
Roger K. Prichard
J. Owen D. Slocombe
Ronald Fayer
George A. Conder
Charles H. Courtney
Byron L. Blagburn
Peter M. Schantz
James C. Williams
Louis C. Gasbarre
Robert S. Rew
Thomas J. Kennedy
Anne M. Zajac
Joseph F. Urban, Jr.
Craig R. Reinemeyer
Linda S. Mansfield
Ann R. Donoghue
Daniel E. Snyder
David S. Lindsay
Susan E. Little
Lora R. Ballweber
Karen Snowden
Patrick F.M. Meeus
Alan A. Marchiondo
Dwight D. Bowman
7
PAST SECRETARY-TREASURERS OF THE
Wendell H. Krull
Edward G. Batte
Rurel Roger Bell
Terence J. Hayes
Vassilios J. Theodorides
S.D. “Bud” Folz
Thomas J. Kennedy
Daniel E. Snyder
Alan A. Marchiondo
Robert G. Arther
1956-1959
1960
1961-1969
1969-1977
1978-1983
1983-1986
1987-1992
1993-1998
1998-2004
2004-2010
2010-2014
2014 AAVP Secretary Treasurer
Doug Carithers
PAST AAVP ANNUAL MEETINGS
1956
1957
1958
1959
1960
1961
1962
1963
1964
1965
1966
1967
1968
1969
1970
1971
1972
1973
1974
1975
1976
1977
1978
1979
1980
1981
1982
1983
1984
1985
1986
1st Annual Meeting – SAN ANTONIO, TX 16 OCT
2nd Annual Meeting – COLUMBUS, OH 17 AUG
3rd Annual Meeting – PHILADELPHIA, PA 18 AUG
4th Annual Meeting – KANSAS CITY, MO 23 AUG
5th Annual Meeting – DENVER, CO 14 AUG
6th Annual Meeting – WEST LAFAYETTE, IN 20 AUG
7th Annual Meeting – MIAMI BEACH, FL
12 AUG
8th Annual Meeting – NEW YORK CITY, NY 28 JUL
9th Annual Meeting – CHICAGO, IL 19 JUL
10th Annual Meeting – PORTLAND, OR
11 JUL
11th Annual Meeting – LOUISVILLE, KY
13-14 JUL
12th Annual Meeting – DALLAS, TX
9 JUL
13th Annual Meeting – BOSTON, MA
21 JUL
14th Annual Meeting – MINNEAPOLIS, MN 13 JUL
15th Annual Meeting – LAS VEGAS, NV 22 JUN
16th Annual Meeting – DETROIT, MI 18 JUL
17th Annual Meeting – NEW ORLEANS, LA 17 JUL
18th Annual Meeting – PHILADELPHIA, PA 15 JUL
19th Annual Meeting – DENVER, CO
21 JUL
20th Annual Meeting – ANAHEIM, CA 13 JUL
21st Annual Meeting – CINCINNATI, OH 19 JUL
22nd Annual Meeting – ATLANTA, GA 11 JUL
23rd Annual Meeting – DALLAS, TX
17 JUL
24th Annual Meeting – SEATTLE, WA
22-24 JUL
25th Annual Meeting – WASHINGTON, D.C. 20-22 JUL
26th Annual Meeting – ST. LOUIS, MO
19-20 JUL
27th Annual Meeting – SALT LAKE CITY, UT 18-19 JUL
28th Annual Meeting – NEW YORK, NY
17-18, JUL
29th Annual Meeting – NEW ORLEANS, LA 15-17 JUL
30th Annual Meeting – LAS VEGAS, NV
22-24 JUL
31st Annual Meeting – ATLANTA, GA
20-22 JUL
8
1987
1988
1989
1990
1991
1992
1993
1994
1995
1996
1997
1998
1999
2000
2001
2002
2003
2004
2005
2006
2007
2008
2009
2010
2011
2012
2013
19-21 JUL
32nd Annual Meeting – CHICAGO, IL
33rd Annual Meeting – PORTLAND, OR
17-18 JUL
34th Annual Meeting – ORLANDO, FL
16-18 JUL
35th Annual Meeting – SAN ANTONIO, TX
21-24 JUL
36th Annual Meeting – SEATTLE, WA
28-30 JUL
2-4 AUG
th
38 Annual Meeting – MINNEAPOLIS, MN
17-20 JUL
39th Annual Meeting – SAN FRANCISCO, CA 9-12 JUL
40th Annual Meeting – PITTSBURGH, PA
6-10 JUL
(Joint meeting with the American Society of Parasitologists)
41st Annual Meeting – LOUISVILLE, KY
20-23 JUL
42nd Annual Meeting – RENO, NV 19-22 JUL
43rd Annual Meeting – BALTIMORE, MD
25-28 JUL
44th Annual Meeting – NEW ORLEANS, LA
10-13 JUL
45th Annual Meeting – SALT LAKE CITY, UT
22-25 JUL
14-17 JUL
47th Annual Meeting – NASHVILLE, TN 13-16 JUL
48th Annual Meeting – DENVER, CO 19-23 JUL
49th Annual Meeting – PHILADELPHIA, PA
24-28 JUL
(Joint meeting with the American Society of Parasitologists)
50th Annual Meeting – MINNEAPOLIS, MN 16-19 JUL
51st Annual Meeting – HONOLULU, HI 15-18 JUL
52ndAnnual Meeting – WASHINGTON, DC 14-17 JUL
53rd Annual Meeting – NEW ORLEANS, LA
19-22 JUL
54th Annual Meeting – CALGARY, CANADA
9-13 AUG
(Joint meeting with the World Association for the Advancement of
Veterinary Parasitology and the International Commission on Trichinellosis)
55th Annual Meeting – ATLANTA, GA
31 JUL – 2 AUG
56th Annual Meeting – ST. LOUIS, MO
16-19 JUL
(Joint meeting with the Livestock Insect Workers Conference and the
International Symposium of Ectoparasites of Pets)
57th Annual Meeting – San Diego, CA 4-7 AUG
58th Annual Meeting – Chicago, IL
20-23 JUL
The AAVP claims copyright privileges to the non-abstract porti ons of proceedings booklets and
acknowledges that the copyright assignment for each abstract remains with the submitting author.
If a company, an institution or an individual wish es to reproduce and dist ribute our proceedings
(even as an internal document), they must obtain copyright permission for each and every abstract
in the book, as well as perm ission from the AAVP for the non-abstract portions of the book.
Alternatively, they may purchase a copy of the proceed ings from the AAVP for each individu al
who may wish to utilize the content of the book.
9
AAVP 59th ANNUAL MEETING
The Curtis Hotel
REGISTRATION
Corridor next to Peek-A-Boo Ballroom
Saturday, 26 July 2014, 1:00 PM – 5:00 PM
Sunday, 27 July 2014, 8:00 AM – 5:00 PM
Monday, 28 July 2014, 10:00 AM – Noon
SOCIAL PROGRAMS
Welcoming Reception – Bayer HealthCare, Animal Health
Four Square Ball Room - Saturday, 26 July 2014, 7:30 PM – 9:30 PM
Merial Ltd Social
Marco Polo Ball Room - Sunday, 27 July 2014, 7:00 PM – 9:00 PM
Elanco Animal Health Social
Marco Polo Ball Room - Monday, 28 July 2014, 7:00 PM – 9:00 PM
AAVP COMMITTEE MEETINGS
Breakfast Sponsored by Novartis
Peek-A-Boo Ball Room & Patty-Cake – Sunday, 27 July 2014, 7:45 AM – 8:30 AM
AAVP BREAKFAST
Breakfast Sponsored by Novartis
Peek-A-Boo Ball Room & Patty-Cake – Monday, 28 July 2014, 7:45 AM – 8:30 AM
STUDENT FUNCTIONS
AAVP Student Member Meeting & Parasite Jeopardy
Lunch Sponsored by Novartis
Patty-Cake Room - Saturday, 26 July 2014, Noon – 1:15 PM
AAVP Student Luncheon & Elections
Patty-Cake Room - Sunday, 27 July 2014, Noon – 1:00 PM
AAVP Student Luncheon / Careers in Parasitology
Patty-Cake Room - Monday, 28 July 2014, Noon – 1:30 PM
10
AAVP-NCVP PARASITOLOGY CLICKER CASES
Boxed Lunches Sponsored by the National Center for Veterinary Parasitology (NCVP)
Peek-A-Boo Ballroom – Tuesday, 29 July 2014, 10:30 AM – 12:30 PM
11
59th ANNUAL MEETING SPONSORS
SOCIAL EVENT SPONSORS
BAYER HEALTHCARE, ANIMAL HEALTH
MERIAL LTD
ELANCO ANIMAL HEALTH
AWARDS SPONSORS
BAYER HEALTHCARE, ANIMAL HEALTH1
COMPANION ANIMAL PARASITE COUNCIL2
MERCK ANIMAL HEALTH3
MERIAL, LTD.4
1
AAVP Best Student Paper Presentation Awards
Honorarium for AAVP-CAPC Graduate Student Award – Zoonotic Diseases
3
Honorarium for AAVP-Merck Animal Health Graduate Student Award - Research
4
Honorarium for AAVP-Merial Distinguished Veterinary Parasitologist Award
2
GENERAL MEETING SPONSORSHIP
PARTNER LEVEL
BAYER HEALTHCARE, ANIMAL HEALTH
ELANCO ANIMAL HEALTH
MERIAL, LTD
NOVARTIS ANIMAL HEALTH – Breakfast and Student Lunches
SCYNEXIS – Refreshment Breaks
PLATINUM LEVEL
CLINVET
ZOETIS
GOLD LEVEL
IDEXX LABORATORIES, INC
MERCK ANIMAL HEALTH
MPI RESEARCH
SILVER LEVEL
CENTRAL STATES (MVS)
CHERI HILL KENNELS
ECTO DEVELOPMENT
HESKA
JOHNSON RESEARCH, LLC
RESEARCH MANAGEMENT GROUP
12
NOTES
13
NOTES
14
15
AAVP Merial
Presentation,
Sunday
July 27, 2014
NexGard™ (afoxolaner) Chewables for Dogs:
Overview of Safety and Efficacy Against Fleas and Ticks
Featuring Dr. Marlene Drag / Dr. Diane Larsen
(Followed by the Merial AAVP Social)
IMPORTANT SAFETY INFORMATION: For use in dogs only. The most common adverse reaction is vomiting.
Other adverse reactions reported are dry/flaky skin, diarrhea, lethargy, and anorexia. The safe use of NexGard in
pregnant, breeding, or lactating dogs has not been evaluated. Use with caution in dogs with a history of seizures.
CAUTION: Federal (USA) law restricts this drug to use by or on the order of a licensed veterinarian.
Description:
NEXGARD™ (afoxolaner) is available in four sizes of beef-flavored, soft chewables for oral
administration to dogs and puppies according to their weight. Each chewable is formulated to
provide a minimum afoxolaner dosage of 1.14 mg/lb (2.5 mg/kg). Afoxolaner has the chemical
composition 1-Naphthalenecarboxamide, 4-[5- [3-chloro-5-(trifluoromethyl)-phenyl]-4, 5-dihydro-5(trifluoromethyl)-3-isoxazolyl]-N-[2-oxo-2-[(2,2,2-trifluoroethyl)amino]ethyl.
Indications:
NEXGARD kills adult fleas and is indicated for the treatment and prevention of flea infestations
(Ctenocephalides felis), and the treatment and control of Black-legged tick (Ixodes scapularis),
American Dog tick (Dermacentor variabilis), and Lone Star tick (Amblyomma americanum)
infestations in dogs and puppies 8 weeks of age and older, weighing 4 pounds of body weight or
greater, for one month.
Dosage and Administration:
NEXGARD is given orally once a month, at the minimum dosage of 1.14 mg/lb (2.5 mg/kg).
Dosing Schedule:
BodyAfoxolaner Per
WeightChewable (mg)
Chewables
Administered
4.0 to 10.0 lbs.
11.3
One
10.1 to 24.0 lbs.
28.3
One
24.1 to 60.0 lbs.
68
One
60.1 to 121.0 lbs.
136
One
Over 121.0 lbs.
Administer the appropriate combination of chewables
NEXGARD can be administered with or without food. Care should be taken that the dog
consumes the complete dose, and treated animals should be observed for a few minutes to
ensure that part of the dose is not lost or refused. If it is suspected that any of the dose has been
lost or if vomiting occurs within two hours of administration, redose with another full dose. If a
dose is missed, administer NEXGARD and resume a monthly dosing schedule.
Flea Treatment and Prevention:
Treatment with NEXGARD may begin at any time of the year. In areas where fleas are common yearround, monthly treatment with NEXGARD should continue the entire year without interruption.
To minimize the likelihood of flea reinfestation, it is important to treat all animals within a
household with an approved flea control product.
Tick Treatment and Control:
Treatment with NEXGARD may begin at any time of the year (see Effectiveness).
Contraindications:
There are no known contraindications for the use of NEXGARD.
Warnings:
Not for use in humans. Keep this and all drugs out of the reach of children. In case of accidental
ingestion, contact a physician immediately.
Precautions:
The safe use of NEXGARD in breeding, pregnant or lactating dogs has not been evaluated. Use
with caution in dogs with a history of seizures (see Adverse Reactions).
Adverse Reactions:
In a well-controlled US field study, which included a total of 333 households and 615 treated dogs
(415 administered afoxolaner; 200 administered active control), no serious adverse reactions were
observed with NEXGARD.
Over the 90-day study period, all observations of potential adverse reactions were recorded.
The most frequent reactions reported at an incidence of > 1% within any of the three months of
observations are presented in the following table. The most frequently reported adverse reaction was
vomiting. The occurrence of vomiting was generally self-limiting and of short duration and tended to
decrease with subsequent doses in both groups. Five treated dogs experienced anorexia during the
study, and two of those dogs experienced anorexia with the first dose but not subsequent doses.
Table 1: Dogs With Adverse Reactions.
Treatment Group
Afoxolaner
Vomiting (with and without blood)
Dry/Flaky Skin
Diarrhea (with and without blood)
Lethargy
Anorexia
N1
17
13
13
7
5
% (n=415)
4.1
3.1
3.1
1.7
1.2
Oral active control
N2
25
2
7
4
9
% (n=200)
12.5
1.0
3.5
2.0
4.5
1
Number of dogs in the afoxolaner treatment group with the identified abnormality.
2
Number of dogs in the control group with the identified abnormality.
In the US field study, one dog with a history of seizures experienced a seizure on the same day
after receiving the first dose and on the same day after receiving the second dose of NEXGARD.
This dog experienced a third seizure one week after receiving the third dose. The dog remained
enrolled and completed the study. Another dog with a history of seizures had a seizure 19 days
after the third dose of NEXGARD. The dog remained enrolled and completed the study. A third dog
with a history of seizures received NEXGARD and experienced no seizures throughout the study.
To report suspected adverse events, for technical assistance or to obtain a copy of the MSDS,
contact Merial at 1-888-637-4251 or www.merial.com/nexgard. For additional information about
adverse drug experience reporting for animal drugs, contact FDA at 1-888-FDA-VETS or online at
http://www.fda.gov/AnimalVeterinary/SafetyHealth.
Mode of Action:
Afoxolaner is a member of the isoxazoline family, shown to bind at a binding site to inhibit insect
and acarine ligand-gated chloride channels, in particular those gated by the neurotransmitter
gamma-aminobutyric acid (GABA), thereby blocking pre- and post-synaptic transfer of chloride
ions across cell membranes. Prolonged afoxolaner-induced hyperexcitation results in uncontrolled
activity of the central nervous system and death of insects and acarines. The selective toxicity
of afoxolaner between insects and acarines and mammals may be inferred by the differential
sensitivity of the insects and acarines’ GABA receptors versus mammalian GABA receptors.
Effectiveness:
In a well-controlled laboratory study, NEXGARD began to kill fleas four hours after initial
administration and demonstrated >99% effectiveness at eight hours. In a separate well-controlled
laboratory study, NEXGARD demonstrated 100% effectiveness against adult fleas 24 hours postinfestation for 35 days, and was ≥ 93% effective at 12 hours post-infestation through Day 21, and
on Day 35. On Day 28, NEXGARD was 81.1% effective 12 hours post-infestation. Dogs in both the
treated and control groups that were infested with fleas on Day -1 generated flea eggs at 12- and
24-hours post-treatment (0-11 eggs and 1-17 eggs in the NEXGARD treated dogs, and 4-90 eggs
and 0-118 eggs in the control dogs, at 12- and 24-hours, respectively). At subsequent evaluations
post-infestation, fleas from dogs in the treated group were essentially unable to produce any eggs
(0-1 eggs) while fleas from dogs in the control group continued to produce eggs (1-141 eggs).
In a 90-day US field study conducted in households with existing flea infestations of varying
severity, the effectiveness of NEXGARD against fleas on the Day 30, 60 and 90 visits compared
with baseline was 98.0%, 99.7%, and 99.9%, respectively.
Collectively, the data from the three studies (two laboratory and one field) demonstrate that
NEXGARD kills fleas before they can lay eggs, thus preventing subsequent flea infestations after
the start of treatment of existing flea infestations.
In well-controlled laboratory studies, NEXGARD demonstrated >94% effectiveness against
Dermacentor variabilis and Ixodes scapularis, 48 hours post-infestation, and against Amblyomma
americanum 72 hours post-infestation, for 30 days.
Animal Safety:
In a margin of safety study, NEXGARD was administered orally to 8- to 9-week-old Beagle puppies
at 1, 3, and 5 times the maximum exposure dose (6.3 mg/kg) for three treatments every 28 days,
followed by three treatments every 14 days, for a total of six treatments. Dogs in the control group
were sham-dosed. There were no clinically-relevant effects related to treatment on physical
examination, body weight, food consumption, clinical pathology (hematology, clinical chemistries,
or coagulation tests), gross pathology, histopathology or organ weights. Vomiting occurred
throughout the study, with a similar incidence in the treated and control groups, including one dog
in the 5x group that vomited four hours after treatment.
In a well-controlled field study, NEXGARD was used concomitantly with other medications, such
as vaccines, anthelmintics, antibiotics (including topicals), steroids, NSAIDS, anesthetics, and
antihistamines. No adverse reactions were observed from the concomitant use of NEXGARD with
other medications.
Storage Information:
Store at or below 30°C (86°F) with excursions permitted up to 40°C (104°F).
How Supplied:
NEXGARD is available in four sizes of beef-flavored soft chewables: 11.3, 28.3, 68 or 136 mg
afoxolaner. Each chewable size is available in color-coded packages of 1, 3 or 6 beef-flavored
chewables.
NADA 141-406, Approved by FDA
Marketed by: Frontline Vet Labs™, a Division of Merial Limited.
Duluth, GA 30096-4640 USA
Made in Brazil.
1050-4493-02
Rev. 4/2014
™NexGard and FRONTLINE VET LABS are trademarks of Merial.
©2014 Merial. All rights reserved. NEX14AAVPEVENTAD (6/14).
16
xng225947_AAVPEventAd8.375x10.75_rsg.indd 1
6/17/14 1:10 PM
Elanco is a proud sponsor of
the 2014 AAVP Annual Meeting
Join us for a
special social event
AAVP members and guests are welcome!
Monday, July 28
7-9 p.m.
Marco Polo Ballroom, The Curtis Double Tree Hotel
©2014 Elanco CAH1752
17
The National Center for Veterinary Parasitology
Invites you to attend a special session of
Parasitology Clicker Cases
a case-based clinical parasitology experience combining fun with
parasites and audience participation (anonymously – it’s clickers!)
Tuesday, July 29th
10:30 am – 12:30 pm
Join your parasitology colleagues for some entertaining and challenging
clinical case quiz questions. Whether you are looking forward to
brushing up on your clinical veterinary parasitology skills, arguing
with your colleagues, or just want to relax and enjoy some entertaining
clinical case discussions with fellow parasitophiles at the AAVP
meeting, this session is sure to delight.
Questions and answers have been vetted by leading AAVP
parasitologists and include material on parasites of small animals,
ruminants, horses, and wildlife.
So please plan to stay for the morning, and bring your diagnostic
expertise!
Box lunches provided to all session attendees courtesy of the
NCVP.
18
Final logos
19
MISSING
SOMETHING?
. More complete.*
For ferrets†
HEARTWORMS
MICROFILARIA
Dogs only
FLEAS
ROUNDWORMS
HOOKWORMS
WHIPWORMS
Dogs only
SARCOPTIC MANGE
Dogs only
EAR MITES
Cats only
FLEAS
HEARTWORMS
See the difference at bayerdvm.com/multi
*Based on label indications: spectrum of species, parasites (dog) and life stages (dog and cat).
†
Advantage Multi® for Cats (imidacloprid + moxidectin) (0.4 mL) is indicated for ferrets that weigh at least 2 lbs.
CAUTION: Federal (U.S.A.) law restricts Advantage Multi® for Dogs (imidacloprid + moxidectin) to use by or on the order of a licensed veterinarian. WARNING: DO NOT
ADMINISTER THIS PRODUCT ORALLY. For the first 30 minutes after application ensure that dogs cannot lick the product from application sites on themselves or other
treated animals. Children should not come in contact with the application sites for two (2) hours after application. (See Contraindications, Warnings, Human Warnings,
and Adverse Reactions, for more information.) CONTRAINDICATIONS: Do not use this product on cats.
CAUTION: Federal (U.S.A.) law restricts Advantage Multi® for Cats to use by or on the order of a licensed veterinarian. WARNINGS: Do not use on sick or debilitated cats
or ferrets. Do not use on underweight cats. (see ADVERSE REACTIONS). Do not use on cats less than 9 weeks of age or less than 2 lbs body weight. Do not use on ferrets
less than 2 lbs body weight. PRECAUTIONS: Avoid oral ingestion. HUMAN WARNINGS: Children should not come in contact with the application site for 30 minutes
after application.
©2014 Bayer HealthCare LLC, Animal Health Division, Shawnee Mission, Kansas 66201
Bayer, the Bayer Cross and Advantage Multi are registered trademarks of Bayer.
AM14535
20
Advantage Multi® for Dogs and for Cats
HUMAN WARNINGS: Not for human use. Keep out of the reach of children. Dogs: Children should not
come in contact with the application sites for two (2) hours after application. Cats: Children should not
come in contact with the application site for 30 minutes after application.
Causes eye irritation. Harmful if swallowed. Do not get in eyes or on clothing. Avoid contact with skin.
Wash hands thoroughly with soap and warm water after handling. If contact with eyes occurs, hold
eyelids open and flush with copious amounts of water for 15 minutes. If eye irritation develops or persists,
contact a physician. If swallowed, call poison control center or physician immediately for treatment advice.
Have person sip a glass of water if able to swallow. Do not induce vomiting unless told to do so by the
poison control center or physician. People with known hypersensitivity to benzyl alcohol, imidacloprid or
moxidectin should administer product with caution. In case of an allergic reaction, contact a physician. If
contact with skin or clothing occurs, take off contaminated clothing. Wash skin immediately with plenty
of soap and water. Call a poison control center or physician for treatment advice. The Material Safety
Data Sheet (MSDS) provides additional occupational safety information. For a copy of the Material Safety
Data Sheet (MSDS) or to report adverse reactions call Bayer Veterinary Services at 1-800-422-9874. For
consumer questions call 1-800-255-6826.
PRECAUTIONS: Do not dispense dose applicator tubes without complete safety and administration
information. Use with caution in sick, debilitated or underweight animals. The safety of Advantage Multi
for Dogs has not been established in breeding, pregnant, or lactating dogs. The safe use of Advantage
Multi for Dogs has not been established in puppies and dogs less than 7 weeks of age or less than 3 lbs.
body weight. Advantage Multi for Dogs has not been evaluated in heartworm-positive dogs with Class 4
heartworm disease.
Cats may experience hypersalivation, tremors, vomiting and decreased appetite if Advantage Multi for
Cats is inadvertently administered orally or through grooming/licking of the application site. The safety
of Advantage Multi for Cats has not been established in breeding, pregnant, or lactating cats. Use of
this product in geriatric cats with subclinical conditions has not been adequately studied. Ferrets: The
safety of Advantage Multi for Cats has not been established in breeding, pregnant, and lactating ferrets.
Treatment of ferrets weighing less than 2.0 lbs. (0.9kg) should be based on a risk-benefit assessment.
The effectiveness of Advantage Multi for Cats in ferrets weighing over 4.4 lbs. (2.0 kg) has not
been established.
ADVERSE REACTIONS: Heartworm Negative Dogs: the most common adverse reactions observed during
field studies were pruritus, residue, medicinal odor, lethargy, inappetence and hyperactivity. Heartworm
Positive Dogs: the most common adverse reactions observed during field studies were cough, lethargy,
vomiting, diarrhea, (including hemorrhagic), and inappetence. Cats: The most common adverse reactions
observed during field studies were lethargy, behavioral changes, discomfort, hypersalivation, polydipsia
and coughing and gagging. Ferrets: The most common adverse reactions observed during field studies
were pruritus/scratching, scabbing, redness, wounds and inflammation at the treatment site, lethargy
and chemical odor.
For a copy of the Material Safety Data Sheet (MSDS) or to report adverse reactions call Bayer Veterinary
Services at 1-800-422-9874. For consumer questions call 1-800-255-6826.
(imidacloprid + moxidectin)
BRIEF SUMMARY: Before using Advantage Multi® for Dogs (imidacloprid+ moxidectin) or Advantage
Multi ® for Cats (imidacloprid +moxidectin), please consult the product insert, a summary of which follows:
CAUTION: Federal (U.S.A.) Law restricts this drug to use by or on the order of a licensed veterinarian.
Advantage Multi for Dogs:
WARNING
• DO NOT ADMINISTER THIS PRODUCT ORALLY.
• For the first 30 minutes after application ensure that dogs cannot lick the product from application
sites on themselves or other treated animals.
• Children should not come in contact with the application sites for two (2) hours after application.
(See Contraindications, Warnings, Human Warnings, and Adverse Reactions for more information.)
INDICATIONS:
Advantage Multi for Dogs is indicated for the prevention of heartworm disease caused by Dirofilaria
immitis and the treatment of Dirofilaria immitis circulating microfilariae in heartworm-positive
dogs. Advantage Multi for Dogs kills adult fleas and is indicated for the treatment of flea infestations
(Ctenocephalides felis). Advantage Multi for Dogs is indicated for the treatment and control of sarcoptic
mange caused by Sarcoptes scabiei var.canis. Advantage Multi for Dogs is also indicated for the treatment
and control of the following intestinal parasites species: Hookworms (Ancylostoma caninum) (Uncinaria
stenocephala), Roundworms (Toxocara canis) (Toxascaris leonina) and Whipworms (Trichuris vulpis).
Advantage Multi for Cats is indicated for the prevention of heartworm disease caused by Dirofilaria
immitis. Advantage Multi for Cats kills adult fleas (Ctenocephalides felis) and is indicated for the
treatment of flea infestations. Advantage Multi for Cats is also indicated for the treatment and control of
ear mite (Otodectes cynotis) infestations and the intestinal parasites species Hookworm (Ancylostoma
tubaeforme) and Roundworm (Toxocara cati).Ferrets: Advantage Multi for Cats is indicated for the
prevention of heartworm disease in ferrets caused by Dirofilaria immitis.Advantage Multi for Cats kills
adult fleas (Ctenocephalides felis) and is indicated for the treatment of flea infestations in ferrets.
CONTRAINDICATIONS: Do not administer this product orally. (See WARNINGS). Do not use the Dog
product (containing 2.5% moxidectin) on Cats.
WARNINGS:
Advantage Multi for Dogs: For the first 30 minutes after application: Ensure that dogs cannot lick
the product from application sites on themselves or other treated dogs, and separate treated dogs
from one another and from other pets to reduce the risk of accidental ingestion. Ingestion of this
product by dogs may cause serious adverse reactions including depression, salivation, dilated pupils,
incoordination, panting, and generalized muscle tremors. In avermectin sensitive dogsa, the signs
may be more severe and may include coma and deathb.
a
Some dogs are more sensitive to avermectins due to a mutation in the MDR1 gene. Dogs with this
mutation may develop signs of severe avermectin toxicity if they ingest this product. The most common
breeds associated with this mutation include Collies and Collie crosses.
b
Although there is no specific antagonist for avermectin toxicity, even severely affected dogs have
completely recovered from avermectin toxicity with intensive veterinary supportive care.
Advantage Multi for Cats: Do not use on sick, debilitated, or underweight cats. Do not use on cats less
than 9 weeks of age or less than 2 lbs. body weight. Do not use on sick or debilitated ferrets.
Advantage Multi is protected by one or more of the following U.S. patents: 6,232,328 and 6,001,858.
NADA 141-251,141-254 Approved by FDA
© 2013 Bayer HealthCare LLC
Bayer, the Bayer Cross, Advantage Multi are registered trademarks of Bayer.
GHG060414
Made in Germany.
21
18726
Elanco Companion Animal
Health enables veterinarians
to help pets live longer,
healthier, higher-quality lives
Building on an outstanding legacy in animal health, Elanco
Companion Animal Health strives to develop the latest
product innovations to enable veterinarians to help pets
live longer, healthier, higher-quality lives. To learn more
about Elanco’s support of the veterinary profession, please
visit Elanco booth or visit elancovet.com.
©2014 Elanco CAH1734
22
It’s a soft chew.
Kills both fleas and ticks.
It’s prescription only.
Now a
pprov
to kill m ed
ore
ticks!
NexGardTM (afoxolaner) is the protection you
asked for, and patients will beg for.
NexGard is FDA-approved to kill fleas, prevent flea infestations, and kill Black-Legged (deer) ticks,
Lone Star ticks and American Dog ticks. NexGard is available only with a veterinarian’s prescription,
and features anti-diversion technology monitored by Pinkerton® Consulting & Investigations.
NexGard and FRONTLINE VET LABS are trademarks
of Merial. ®PINKERTON is a registered trademark of
Pinkerton Service Corporation. ©2014 Merial Limited,
Duluth, GA. All rights reserved. NEX14TTRADEAD (06/14).
TM
xng222320_AAVP-8.375x10.75_rsg.indd 1
IMPORTANT SAFETY INFORMATION: For use in dogs only. The most
common adverse reaction is vomiting. Other adverse reactions
reported are dry/flaky skin, diarrhea, lethargy, and anorexia. The
safe use of NexGard in pregnant, breeding, or lactating dogs has not
been evaluated. Use with caution in dogs with a history of seizures.
23
6/16/14 1:57 PM
T:7.375”
THERE’S A NEW
CHEWABLE IN TOWN.
Introducing new Sentinel Spectrum (milbemycin oxime/lufenuron/praziquantel)
chewables — protection against six parasites, instead of only three.
®
®
T:9.75”
Dogs should be tested for heartworm prior to use. Mild hypersensitivity
reactions have been noted in some dogs carrying a high number of
circulating microfilariae. Treatment with fewer than 6 monthly doses
after the last exposure to mosquitoes may not provide complete
heartworm prevention. Visit sentinelpet.com for more information.
*A. caninum **Prevents flea eggs from hatching; is not an adulticide.
© 2014 Novartis Animal Health US, Inc. Sentinel and Spectrum are registered trademarks of Novartis AG.
®Heartgard and the Dog & Hand logo are registered trademarks of Merial.
Proof #:
JOB #: 54559
Print Scale: None
Bleed: 8.625” x 11”
Cyan
Date: 5-20-2014 9:11 AM
VE
24
GCD: -
BUILDING ALLIANCES TO BRING VALUE TO OUR PARTNERS
Bringing Innovation To
Parasiticide Discovery
Animal
Health
SCYNEXIS
Difference
• Turn-key solutions for your
parasiticide discovery needs
• Multidisciplinary teams:
• Outlicensing opportunities for
internal assests
• Accessing compound collections
to bring value to our partners
• Biology
• Medicinal Chemistry
• DMPK
• Process Development
• Analytical Chemistry
• GMP Manufacturing
FIND THESE SCYNEXIS REPRESENTATIVES AT THE AAVP CONFERENCE
Kerrie Powell, Business Development Director
Itta Bluhn-Chertudi, Business Development Specialist
Bakela Nare, Manager, Parasitology
SCYNEXIS, Inc.
Research Triangle Park, NC
business.development@scynexis.com
w w w. s c y n e x i s . c o m
25
All trademarks are the property of Zoetis Inc., its affiliates and/or its licensors. ©2013 Zoetis Inc. All rights reserved. GCA13151
ONE
NEW
NAME.
HERE
FOR
YOU.
Today we’re Zoetis, a company with a singular focus on animal health
committed to supporting you and your operation. We’re still home to
the people and products you’ve come to trust for more than 60 years as
Pfizer Animal Health, dedicated to providing veterinarians and producers
with the medicines, vaccines, and services you need. We look forward to
working with you in ever better ways. Meet us at ZoetisUS.com
FOR ANIMALS. FOR HEALTH. FOR YOU.
26
The Companion Animal Parasite Council (CAPC) recommends
annual blood and fecal testing for every patient, every year. For
the health of your patients rely on IDEXX tests, every time.
The SNAP® 4Dx® Plus Test
Broadest available pet-side screen provides results for six pathogens
from five vectors with just one blood sample.
© 2014 IDEXX Laboratories, Inc. All rights reserved. • 104846-00 • All ®/ TM
marks are owned by IDEXX Laboratories, Inc. or its affiliates in the United States
and/or other countries. The IDEXX Privacy Policy is available at idexx.com.
Every
canine patient
annual visit
parasitic screening
To order or for more
information, contact your
authorized IDEXX distributor
or call 1-800-355-2896.
Fecal Panels with Whipworm Antigen ELISA
The new Whipworm Antigen ELISA is paired with the CAPC-recommended
fecal centrifugation method. Find more whipworm positives with this panel.
104846-00_rz1 Parasite Screen DSR ad_7375x475.indd 1
4/21/14 1:11 PM
BEYOND EXPECTATIONS
MPI Research is a proud sponsor of the American Association of
Veterinary Parasitologists (AAVP) annual meeting. MPI Research strives
to maintain our status as the industry’s leading provider of contract
research services. Our experienced staff diligently upholds the regulatory
measures needed to sustain our high standard, and has the knowledge
to ensure the proper conduct of studies that test the efficacy of products
against a variety of external parasites. We work closely with our
Consultants and Sponsors to expand capabilities and techniques
using both ectoparasite and endoparasite models.
To learn more, stop by our exhibit or visit us at www.mpiresearch.com
27
ADD
– ADD POUNDS
Important Safety Information:
Internal Parasites Can Cost You Over $190/Head
1
Internal parasites in cattle reduce average daily gain (ADG), reduce feed intake and impair the immune
system. Safe-Guard cuts economic losses by killing internal parasites in the gut where they live.
Cooperia is now confirmed as the most prevalent internal parasite in U.S.
cattle herds (NAHMS 2009, 2010)
Internal parasite (Cooperia) resistance to macrocyclic lactone compounds
(endectocides) was confirmed in 2004 and continues to grow2
RESIDUE WARNING: Cattle must not be
slaughtered within 13 days following
last treatment. For dairy cattle, the milk
discard time is zero hours. A withdrawal
period has not been established for this
product in pre-ruminating calves. Do not
use in calves to be processed for veal.
Safe-Guard eliminates resistant parasites3
(98.1% effective)
Add Safe-Guard to your parasite management
program and add pounds to your calves.
2
3
RESIDUE WARNING: Cattle must not be
slaughtered within 11 days following last
treatment. A withdrawal period has not
been established for this product in preruminating calves. Do not use in calves to
be processed for veal.
Safe-Guard mineral, feed through
products and liquid feed:
In a peer-reviewed published study, calves infected with macrocyclic
lactone-resistant Cooperia had 7.4% less average daily gain and 5.4% less
feed intake3
1
Safe-Guard block:
Safe-Guard drench and paste:
RESIDUE WARNING: Cattle must not
be slaughtered within 8 days following
last treatment. For dairy cattle, the milk
discard time is zero hours. A withdrawal
period has not been established for this
product in pre-ruminating calves. Do not
use in calves to be processed for veal.
Economic analysis of pharmaceutical technologies in modern beef production, John D. Lawrence and Maro A. Ibarburu, Iowa State University, 2007.
Gasbarre, L.C., Smith, L.L., Lichtenfels, J.R., Pilitt, P.A., 2004. The identification of cattle nematode parasites resistant to multiple classes of anthelmintics in
a commercial cattle population in the US. Proceedings of the 49th American Association of Veterinary Parasitologists. Philadelphia, July 24-28 (Abstract 44).
Stromberg, B.E., et al., Cooperia punctata: Effect on cattle productivity? Vet. Parasitol. (2011), doi: 10.1016/vetpar.2011.05.030
556 Morris Avenue • Summit, NJ 07901 • merck-animal-health-usa.com • 800-521-5767
Copyright © 2014 Intervet Inc., d/b/a Merck Animal Health, a subsidiary of Merck & Co., Inc. All rights reserved. 1/13 BV-SG-48786
48786AAVP_SG_FastFacts_030114 V2.indd 1
3/13/14 9:14 AM
Drives fleas to extinction
with indoxacarb
Both patients and clients want a product that really gets rid of fleas.
Activyl®, the latest innovation in monthly spot-on flea control, features
indoxacarb and bioactivation — a mode of action that uses enzymes
inside the flea to activate Activyl®’s full flea-killing power.
Available strictly through veterinarians.
Protected by Merck Animal Health
Track & Trace™ technology.
Quick-Drying • Fragrance-Free • Waterproof
us.activyl.com
Copyright © 2014 Intervet Inc., a subsidiary of Merck & Co., Inc.
All rights reserved. Intervet Inc. d/b/a Merck Animal Health,
Summit, NJ 07901
US/ACT/0214/0012
MAH_ACT_A43554_GradAD.indd 1
28
5/14/14 2:08 PM
For companies in or entering the animal health and nutrition markets who have information, analytical or personnel needs, Brakke Consulting provides
a menu of services which are designed to provide results quickly, accurately and reliably. Unlike firms with no industry specialization or single person
consultancies, our work is based upon broad and deep, ongoing, long-term involvement in the animal health and nutrition markets by a team of talented
individuals.
Helping Parasiticide Drug Research Teams…
•
AAALAC accredited BSL -2 facilities
Established in 1986, Brakke Consulting has specialized in the veterinary, pet, animal health, and bio-pesticide industry segments. The firm provides a
•
In-vitro and In-vivo testing platforms available
menu of services that covers six important business areas: marketing intelligence, general consulting, professional recruiting services, transaction and
investment services, sales• training
and coaching,
and Propagation
veterinary practice of
development.
Screening
and
field isolates of ruminant
nematodes
Website: www.brakkeconsulting.com
Email: info@brakkeconsulting.com
•
Experienced
Corporate office: 972.243.4033
•
•
staff
Works diligently with the customers to achieve goals
Short turn around time
17190 Polk Rd.
17190 Polk Rd. Stanwood, MI 49346
Stanwood, MI 49346
E-mail: misiu@frontier.com
Tel: (231) 823-2392
Tel: (231) 823-2392 Fax: (231) 823-2925
Fax: (231) 823-2925
 Conducting canine & feline parasite research since 1990.
 Conducting canine
feline
research
1990.areas:
 & GLP
& parasite
GCP studies
in thesince
following
 GLP & GCP studies
the following
areas: efficacy studies
 inEndo
and Ectoparasite
 Endo and Ectoparasite
efficacy
studies
 Safety Studies utilizing our homozygous mdr -1 gene Collie colony
our homozygous
mdr -1 gene Collie colony
17190 Polk Rd. Safety Studies utilizing
 Pharmakokinetic
studies
 Pharmakokinetic studies
Stanwood, MI 49346
Nutrition/weight loss studies
 Nutrition/weight loss Palatability
studies
studies
Tel: (231) 823-2392
 Palatability studies
 Provider of canine and feline blood products
Fax: (231) 823-2925








yoursince
research
involves dogs or cats, contact us to assist you with your research needs.
Conducting canine & feline parasite When
research
1990.
When your research involves dogs or cats, contact us to assist you with your research needs.
GLP & GCP studies in the following areas:
Endo and Ectoparasite efficacy studies
Safety Studies utilizing our homozygous mdr -1 gene Collie colony
Pharmakokinetic studies
Palatability studies
Provider of canine and feline blood products
Ecto
DEVELOPMENT
CORPORATION
Healthier animals…through science
www.ectodev.com
30
29
17190 Polk Rd.
17190 Polk Rd. Stanwood, MI 49346
Stanwood, MI 49346
Tel: (231) 823-2392
Tel: (231) 823-2392 Fax: (231) 823-2925
Fax: (231) 823-2925
 Conducting canine & feline parasite research since 1990.
 Conducting canine
feline
research
1990.areas:
 & GLP
& parasite
GCP studies
in thesince
following
 GLP & GCP studies
the following
areas: efficacy studies
 inEndo
and Ectoparasite
 Endo and Ectoparasite
efficacy
studies
 Safety Studies utilizing our homozygous mdr -1 gene Collie colony
our homozygous
mdr -1 gene Collie colony
17190 Polk Rd. Safety Studies utilizing
 Pharmakokinetic
studies
 Pharmakokinetic studies
Stanwood, MI 49346
 Nutrition/weight loss Palatability
studies
studies
Tel: (231) 823-2392
 Palatability studies
Fax: (231) 823-2925








yoursince
research
involves dogs or cats, contact us to assist you with your research needs.
Conducting canine & feline parasite When
research
1990.
GLP & GCP studies in the following areas:
Endo and Ectoparasite efficacy studies
Safety Studies utilizing our homozygous mdr -1 gene Collie colony
Pharmakokinetic studies
Palatability studies
Provider of canine and feline blood products
30
30
DiagramoftheCurtisHotel,Denver,CO 31
59th Annual Meeting of the American Association of Veterinary Parasitologists
CONFERENCE PROGRAM
Saturday July 26, 2014
AAVP Executive Committee Meeting
All AAVP officers and committee chairs please plan to attend
nd
Room: Jax (2 floor)
AAVP Student Member Meeting & Parasite Jeopardy
Lunch sponsored by Novartis
All AAVP students and post-docs please plan to attend
nd
Room: Patty-Cake (2 floor)
CONFERENCE REGISTRATION
nd
Outside Peek-A-Boo ballroom (2 floor)
Break
PLENARY SESSION
Opening remarks
President: Dwight D. Bowman
President-Elect and Program Chair: Andrew S. Peregrine
nd
Room: Peek-A-Boo ballroom (2 floor)
PLENARY SESSION: EMERGING ISSUES – ANTHELMINTIC RESISTANCE
Moderator: Andrew S. Peregrine
8.00-13.15
12.00-13.15
13.00-17.00
13.15-14.00
14.00-14.15
1.
14.15-15.45
The path of least (anthelmintic) resistance: yellow brick road or highway to hell?
Morgan McArthur
New Berlin, Wisconsin
2.
15.45-16.15
16.15-17.30
16.15-16.45
16.45-17.30
17.30-19.30
19.30-21.30
Macrocyclic lactone-resistant Dirofilaria immitis: where do we go from here?
Tim Geary
McGill University
Room: Peek-A-Boo ballroom
Scynexis Coffee Break (outside Peek-A-Boo ballroom)
AAVP AWARDS
2014 AAVP-Merck Animal Health Outstanding Graduate Student
Moderator: Dan Snyder
Awarded to Alice Lee
3. Antibody-mediated trapping of Toxocara canis larvae within the murine
liver upon challenge infection
2014 AAVP-Merial Distinguished Veterinary Parasitologist
Moderator: Doug Carithers
Awarded to Tim Geary
Break
BAYER SOCIAL
rd
Room: Four Square Ballroom (3 floor)
32
Sunday July 27, 2014
7.45-8.30
8.30-12.00
8.30-10.00
AAVP Committee meetings (breakfast sponsored by Novartis)
Rooms: Peek-A-Boo ballroom & Patty-Cake
AAVP STUDENT COMPETITION PRESENTATIONS
CONTROL
IMMUNOLOGY
Room: Keep Away
Moderators: Melissa Beck & Ann Donoghue
Moderators: Dana Pollard & Wenbin Tuo
8.30-8.45
4. Objective evaluation of two deworming regimens in
young thoroughbreds: parasitological parameters and
growth rates
Jennifer Bellaw
University of Kentucky
10. Evaluation of immune responses during Haemonchus
contortus worm expulsion in Native and Suffolk sheep
Javier Garza
Louisiana State University
8.45-9.00
5. Transabdominal ultrasonography as a tool for monitoring
Parascaris spp. in foals
Maci Stephens
9.00-9.15
9.15-9.30
9.30-9.45
9.45-10.00
10.00-10.30
10.30-12.00
10.30-10.45
10.45-11.00
11.00-11.15
6. Evaluation of worm replacement as a means to reverse
the Impact of multiple-anthelmintic resistant Haemonchus
contortus on a sheep farm
Melissa Miller
7. Anthelmintic activity of Melilotus alba (white sweet
clover) on Haemonchus contortus infective juveniles and its
cytotoxicity on bovine ileal epithelial cells
Anwar Sarah
South Dakota State University
8. Treatment and control of Giardia duodenalis in a dog
colony used for veterinary instruction
Meriam Saleh
Virginia Tech
9. Demonstration of flotation of Toxocara canis eggs in
commercial bleach (5.25% sodium hypochlorite solution)
and effects of bleach treatment on embryonation
of T. canis eggs
Alice Houk
Virginia Tech
11. Delayed immune responses of parasite-susceptible
sheep during Haemonchus contortus infection are
associated with greater larval burden
Jesica Jacobs
West Virginia University
12. Direct and indirect effects of Copper Oxide Wire
Particles on Haemonchus contortus infections in small
ruminants
Vicky Kelly
13. Effects of peripheral blood mononuclear cells on larval
motility of Haemonchus contortus in vitro
Rush Holt
14. Modulation of vaccine-induced responses by
anthelmintic treatment in ponies
Emily Rubinson
15. Evaluation of the systemic inflammatory response to
anthelmintic treatment in ponies
Alejandra Betancourt
EPIDEMIOLOGY & MOLECULAR
WILDLIFE & PARASITE BIOLOGY
Room: Keep Away
Moderators: Meriam Saleh & Doug Colwell
Moderators: Rachel Curtis & Sue Howell
16. County scale distribution of Amblyomma americanum in
Oklahoma: addressing local deficits in tick maps based on
passive reporting
Anne Barrett
Oklahoma State University
17. Confirmation of Borrelia burgdorferi and Anaplasma
phagocytophilum in established populations of Ixodes
scapularis in southwestern Virginia
Brian Herrin
22. A survey of the parasites of urban Red Foxes (Vulpes
vulpes) in Charlottetown, Prince Edward Island, Canada
based on fecal analysis
William Robbins
University of Prince Edward Island
18. Prevalence of vector-borne pathogens in dogs from Haiti
Lindsay Starkey
33
23. Detection and characterization of haemosporidia in
whooping cranes and sandhill cranes
Miranda Bertram
Texas A&M University
24. Detection of circulating cathodic antigen and circulating
anodic antigen in raccoons naturally infected with
Heterobilharzia americana and comparison of test
performance with fecal sedimentation and fecal PCR
Jessica Rodriguez
Sunday July 27, 2014 (continued)
11.15-11.30
19. Comparative patterns of recruitment, reproduction, and
size of a generalist trematode, Dicrocoelium dendriticum,
in sympatric hosts
Melissa Beck
University of Lethbridge
25. Association between nymphal engorgment weight and
sex of adult Amblyomma americanum, Amblyomma
maculatum, Dermacentor variabilis,
and Rhipicephalus sanguineus
Yoko Nagamori
11.30-11.45
20. Development of a ‘Nemabiome” sequencing assay for
the relative quantitation of parasitic nematode species in
cattle fecal samples
Russell Avramenko
University of Calgary
26. Investigating the suitability of beagle dogs as an animal
model for the human parasite Brugia malayi
Erica Burkman
11.45-12.00
21. Ascaris suum ACR-21 forms a homomeric nicotinic
acetylcholine receptor with novel pharmacology
Shivani Choudhary
Iowa State University
27. Inactivation of helminth eggs with short- and mediumchain fatty acids alone and in combinations at naturally
occurring concentrations
Dan Zhu
Cornell University
12.00-13.00
13.00-14.00
14.00-14.15
14.15-15.15
Lunch on your own
AAVP Students – Novartis Lunch & Elections (Patty-Cake)
PLENARY SESSION: Center for Veterinary Medicine (CVM)
Discussion moderators: Jenifer Edmonds & Martin Nielsen
28. FDA's Center for Veterinary Medicine's Antiparasitic Resistance Management Strategy
Anna O’Brien & Michelle Kornele
Office of New Animal Drug Evaluation, FDA-CVM
Break
STUDENT COMPETITION PRESENTATIONS
GENERAL SESSION PRESENTATIONS
MOLECULAR & EPIDEMIOLOGY
CATTLE
Room: Keep Away
Moderators: Jessica Rodriguez & Karen Snowden
14.15-14.30
14.30-14.45
14.45-15.00
15.00-15.15
15.15-15.45
15.45-17.15
15.45-16.00
29. Assessment of the genetic variability of Cytauxzoon felis
rRNA Internal Transcribed Spacers
Dana Pollard
30. Diversity of Trypanosoma cruzi strain types in vector and
host populations throughout Texas
Rachel Curtis
31. The public health significance of macrocyclic lactone
resistance in Dirofilaria immitis
Cassan Pulaski
32. Epidemiological factors effecting the prevalence of
Entamoeba histolytica in different populations of dogs
Muhammad Nazir
Bahauddin Zakariya University
Moderators: Javier Garza & Morgan McArthur
33. Treatment of stocker calves at turnout with an
eprinomectin extended-release injection or a combination
of injectable doramectin and oral albendazole
Thomas Yazwinski
University of Arkansas
34. Managing anthelmintic resistance in cattle parasites in
New Zealand – choice of anthelmintic product
Dave Leathwick
AgResearch
35. Use of conventional PCR as a tool to monitor
gastrointestinal strongyle populations in US cattle
Ashley McGrew
Colorado State University
36. Utilization of computer processed high definition video
imaging for measuring motility of microscopic nematode
stages on a quantitative scale: “The Worminator”
Bob Storey
VECTOR-BORNE DISEASES & DIAGNOSIS
HORSES & HISTORY
Room: Keep Away
Moderators: Brian Herrin & Jean Tsao
Moderators: Jennifer Bellaw & Craig Reinemeyer
37. Ectoparasites of free-roaming domestic cats (Felis catus)
presented to a spay-neuter clinic in Oklahoma
Jennifer Thomas
43. Strongylus vulgaris and colic – a retrospective casecontrol study
Martin Nielsen
34
Sunday July 27, 2014 (continued)
16.00-16.15
38. Evaluation of culture conditions for the cultivation of the
Cytauxzoon felis erythrocytic stage
Dana Pollard
16.15-16.30
39. Factor selection for utilization in maps of four vectorborne diseases
Dwight Bowman
Cornell University
16.30-16.45
40. Identifying relevant canine heartworm factors
Dongmei Wang
Clemson University
16.45-17.00
17.00-17.15
17.15-18.00
18.00-19.00
19.00-21.00
41. Factors influencing the seroprevalence of canine
anaplasmosis in the United States
Christopher McMahan
Clemson University
42. What’s trending now: what parasites are we most
commonly seeing, through the snapshot of diagnosis by the
Veterinary Diagnostic Laboratory at the Oregon State
University College of Veterinary Medicine?
Janell Bishop-Stewart
Oregon State University
44. Comparison of a single dose of moxidectin and a five-day
course of fenbendazole to reduce and suppress
cyathostomin fecal egg counts in a commercial recipient
mare herd
Ray Kaplan
45. Parascaris equorum is dead. Long live
Parascaris univalens!
Martin Nielsen
46. Serum Strongylus vulgaris-specific antibody responses to
anthelmintic treatment in naturally infected horses
Martin Nielsen
47. Performance of a new duplex real-time PCR assay for
Theileria equi and Babesia caballi in horses
Vladislav Lobanov
Canadian Food Inspection Agency
48. Glimpses of parasitology history through
commemorative medals
Raffaele Roncalli
N/A
Break
Merial Symposium
NexGard (Afoxolaner) Chewables for Dogs: Overview of Safety and Efficacy Against
Fleas and Ticks
Dr. Marlene Drag / Dr. Diane Larsen
Merial Limited
MERIAL SOCIAL
rd
Room: Marco Polo Ballroom (3 floor)
35
Monday July 28, 2014
7.45-8.30
8.30-17.15
8.30-10.00
Breakfast sponsored by Novartis
Rooms: Peek-A-Boo ballroom & Patty-Cake
HEARTWORM
NOVEL CASES
Room: Keep Away
Moderators: Alice Lee & Roger Prichard
8.30-8.45
8.45-9.00
9.00-9.15
9.15-9.30
9.30-9.45
9.45-10.00
10.00-10.30
10.30-12.00
12.00-13.30
13.30-15.00
49. Administration of 10% imidacloprid-1% moxidectin
protects cats from subsequent, repeated infection with
Dirofilaria immitis
Susan Little
50. Protection of dogs against canine heartworm infection
28 days after 4 monthly treatments with
Advantage Multi® for Dogs
Dwight Bowman
Cornell University
51. Comparative efficacy of four commercially available
heartworm preventive products against the JYD-34
laboratory strain of Dirofilaria immitis
Byron Blagburn
Auburn University
52. Forecasting annual canine heartworm prevalence in the
United States
Robert Lund
Clemson University
53. Prevalence of Dirofilaria immitis antigen in feline
samples after heat treatment
Susan Little
54. False negative antigen tests in dogs infected with
heartworm and placed on macrocyclic lactone preventives
Susan Little
13.45-14.00
14.00-14.15
55. Treatment of a Mammomonogamus spp.
infection in a domestic cat
Jennifer Ketzis
Ross University
56. Identification of Trichuris serrata in cats on St. Kitts
Jennifer Ketzis
Ross University
57. Ocular onchocercosis in a dog
Gary Conboy
58. Trypanosoma theileri associated with a second
trimester aborted Holstein fetus
Dana Pollard
59. Spectacular rhabdiasid infections in captive-bred Ball
pythons from Wisconsin and Virginia
Araceli Lucio-Forster
Cornell University
60. Five cases of horse blindness due to the aberrant
migration of Setaria digitata into the aqueous humor
SungShik Shin
Chonnam National University
AAVP Business Meeting & Awards
AAVP Executive Committee, Student Officers, Corporate Sponsor representatives,
and Awardees stay for pictures
Lunch on your own
AAVP Students – Novartis Lunch & Careers in Parasitology (Patty-Cake)
DOG & CAT - DIAGNOSIS
MOLECULAR & IMMUNOLOGY
Room: Keep Away
Moderators: William Robbins & Jennifer Ketzis
13.30-13.45
Moderators: Jesica Jacobs & Anne Zajac
61. ELISA detection of Trichuris serrata
coproantigen in cats
Jinming Geng
IDEXX Laboratories, Inc.
62. ELISA detection of ascarid and hookworm
coproantigen in dogs and cats
David Elsemore
IDEXX Laboratories, Inc.
63. Evaluation of fecal samples from dogs with naturally
acquired gastrointestinal nematodes for coproantigen of
Trichuris vulpis, Toxocara canis, and Anyclostoma caninum
Chris Adolph
36
Moderators: Russell Avramenko & John Gilleard
67. Not just GluCls (Glutamate-gated Chloride channels):
abamectin has potent effects on nicotinic acetylcholine
receptors of nematode parasites
Richard Martin
68. Electrophysiological effects of cholinergic anthelmintics
in two species of filarial parasite
Saurabh Verma
69. Characterization of a homomeric nicotinic acetylcholine
receptor, Ascaris suum ACR-16,
in Xenopus laevis oocytes
Alan Robertson
Monday July 28, 2014 (continued)
14.15-14.30
14.30-14.45
14.45-15.00
15.00-15.30
15.30-17.15
64. Prevalence of internal parasites in shelter cats based on
centrifugal fecal flotation
Byron Blagburn
Auburn University
65. Prevalence of internal parasites in shelter dogs
based on centrifugal fecal flotation
Byron Blagburn
Auburn University
66. Assessing the speed of kill of Ancylostoma caninum by
Advantage Multi® for Dogs using endoscopic methods
Alice Lee
Cornell University
WILDLIFE & SCYNEXIS
SMALL RUMINANTS & EDUCATION
Room: Keep Away
Moderators: Melissa Miller & Dave Leathwick
15.30-15.45
15.45-16.00
16.00-16.15
16.15-16.30
16.30-16.45
16.45-17.00
17.00-17.15
17.15-19.00
17.30-18.30
19.00-21.00
70. Apyrases in Ostertagia ostertagi; controlling the
inflammatory response and longevity of
L4 larvae in the gastric glands
Dante Zarlenga
USDA, ARS
71. Characterization of Ostertagia ostertagi macrophage
migration inhibitory factor
Wenbin Tuo
USDA, ARS
72. Host immune responses to Eimeria adenoeides
infection in turkey poults
Thilakar Rathinam
73. Challenges in assessing anthelmintic resistance and the
use of FAMACHA in sheep on St. Kitts
Jennifer Ketzis
Ross University
74. Opportunities for molecular diagnosis of infection and
anthelmintic resistance in parasitic nematodes
Roger Prichard
McGill University
75. Competitive effects between genetically diverse
Haemonchus contortus strains during
experimental co-infection
John Gilleard
76. Worm burdens of Haemonchus contortus in alpacas and
sheep following experimental infection
Anne Zajac
Virginia Tech
77. Development of OSCEs for assessment of
parasitologic skills
Karen Snowden
78. Advancing veterinary parasitology education and
knowledge through online discussions - the
VIN experience
Craig Datz
University of Missouri
79. Defining core competencies in veterinary parasitology
Karen Snowden
Moderators: Miranda Bertram &
Janell Bishop-Stewart
80. Epidemiology of Haemonchus contortus in a zoological
park giraffe enclosure in Florida
Sue Howell
81. Efficacy of Duddingtonia flagrans on gastrointestinal
nematode larvae in feces of exotic artiodactylids
James Miller
82. Influence factors on infection with Taenia hydatigena
larvae of wild ruminants in the National Park Losiny
Ostrov (Elk Island), Moscow, Russia
Nina Samoylovskaya
All-Russian K.I. Skryabin Scientific Research Institute
of Helminthology
83. Rickettsial and ehrlichial infection in small mammals
from northeast Brazil
Solange Gennari
University of São Paulo
84. PK/PD approaches to anti-parasitic drug discovery: are
we there yet with filarial parasites?
Bakela Nare
SCYNEXIS, Inc.
Break
DACVM meeting (Jax)
ELANCO SOCIAL
rd
Room: Marco Polo Ballroom (3 floor)
37
Tuesday July 29, 2014
PRESIDENT’S SYMPOSIUM
EMERGING ISSUES – TICKS AND TICK-BORNE DISEASES
Moderator: Dwight D. Bowman
8.30-10.00
85. Tick-borne infections in the southern United States – changing patterns
Susan Little
86. The emergence of Ixodes scapularis and Lyme disease in North America: how differences in
ecology may affect regional patterns
Jean Tsao
Michigan State University
10.00-10.30
10.30-12.30
12.30
AAVP-NCVP Parasitology Clicker Cases
Moderator: Andrew Peregrine
Boxed lunches for attendees (sponsored by NCVP)
Meeting Adjourns – Happy Trails
38
2014 ABSTRACTS
The AAVP claims copyright privileges to the non-abstract portions of this proceedings booklet and
acknowledges that the copyright assignment for each abstract remains with the submitting author.
If a company, an institution or an individual wishes to reproduce and distribute our proceedings
(even as an internal document), they must obtain copyright permission for each and every abstract
in the book, as well as permission from the AAVP for the non-abstract portions of the book.
Alternatively, they may purchase a copy of the proceedings from the AAVP for each individual
who may wish to utilize the content of the book.
39
40
ABSTRACTS
Plenary Session – Anthelmintic Resistance
1
The path of least (anthelmintic) resistance: yellow brick road or highway to hell?
Morgan McArthur*. Self-employed consultancy, New Berlin, WI.
Three decades of heavy and often indiscriminate reliance on the macrocyclic lactone (ML)
family of anthelmintics for worm control in cattle has favored the development of parasites that
are resistant to these drugs. As anthelmintic resistance (AR) becomes more commonplace in
the US a simple question must be asked: Who Cares?
Parasitologists study the epidemiology and genetics of nematodes, devise and apply
measurement methods and know the ramifications of AR. We worry that the worms are winning
the war. Producers may not share the same concern. In cattle, morbidity due to internal
parasites is subtle and mortality is rare. If MLs seem to be working and are getting cheaper, why
worry? ‘If it ain’t broke, don’t fix it.’
As we endeavor to develop and deliver effective management strategies for AR the first form of
resistance that has to be dealt with is human resistance - to change. There is considerable
complacency about parasite management in the producer community. The intrinsic potency of
MLs, convenient modes of application and blind faith in their efficacy has supplanted the need
for critical thinking about parasites. In a recent NAHMS survey very few producers reported
using fecal testing to make deworming decisions.
Recommendations for managing AR will not be an easy sell to producers who do not perceive
that they have an efficacy problem. Shifting the balance from productivity toward sustainability
with fecal egg count monitoring, creating refugia, and using dewormers more strategically
involves cost and compromise. Again, the question: Who Cares? None of these changes can
occur without effective education. Anthelmintic sustainability may be influenced as much by
psychology as parasitology. It will be important to consider leadership and behavioral change
strategies as well as the use of specific best practices to manage anthelmintic resistance.
2
Macrocyclic lactone-resistant Dirofilaria immitis: where do we go from here?
Tim Geary*. McGill University, Institute of Parasitology, Ste-Anne-de-Bellevue, QC, Canada.
Decades of reliance on macrocyclic lactone (ML) endectocides for prophylactic control of
heartworm infections in dogs and cats have led to profound changes in veterinary practice in
endemic areas. Safe, convenient and fully effective ML regimens were of tremendous benefit for
clients, practitioners and veterinarians. They also led to a sense of complacency and a
reduction in research on the discovery and development of new drugs or vaccines for this
indication. However, recent evidence conclusively shows that populations of Dirofilaria immitis
resistant to ML prophylaxis have arisen in the USA. This evidence is based on break-through
41
phenotypes of several strains against multiple MLs in dogs exposed to L3 stages while under
normally effective ML regimens. These strains harbor genetic signatures consistent with a single
point of origin, consistent with drug selection.
The existence of ML-resistant parasites poses significant therapeutic challenges for heartworm
control. Molecular markers can be used to monitor the epidemiology of resistant strains and to
determine their fitness relative to susceptible parasites. The spread of resistant parasites may
be limited to some degree by treatment of infected animals with doxycycline, which is reported
to prevent the development of infective capacity in exposed microfilariae. Advocacy of
intermittent doxycycline treatment as a routine substitute for MLs in areas of risk is unwise in
light of the possibility of encouraging the development of antibiotic-resistant bacterial pathogens.
New investment in alternative drugs for heartworm control is an urgent need, but it may be
difficult to recapture the benefits of the MLs. In lieu of alternatives, and in the large areas in
which ML-resistant parasites have not been reported, continued reliance on these drugs is the
best solution at present.
2014 AAVP-Merck Animal Health Outstanding
Graduate Student Presentation
3
Antibody-mediated trapping of Toxocara canis larvae within the murine liver upon
challenge infection.
Alice Lee*, Janice Liotta, Samantha Peneyra, Michelle Florentine, Dwight Bowman. Cornell
University, College of Veterinary Medicine, Microbiology & Immunology, Ithaca, NY.
In the United States, 14% of people are infected by Toxocara larvae. Larval migration within
people can cause symptoms including fever, abdominal pain, dyspnea, and seizures; however,
the majority of cases are asymptomatic. In mice, repeat infections with Toxocara canis results in
the trapping of a portion of the larvae in the liver. This has been postulated to help protect
sensitive organs like the brain from migration-associated damage, and may be one reason why
cerebral toxocariasis is not more common. Building on work of previous investigators, the
immunologic basis of larval trapping was further elucidated. Female BALB/c mice were orally
inoculated twice with T. canis eggs 4 weeks apart and euthanized one week later. Hepatic T
cells were characterized by flow cytometry, showing a predominant CD4+ population with
significantly more T helper 2 cells in the livers of infected mice compared to controls. The
importance of CD4+ T cells in effecting trapping was assessed using the same infection model
described above. In vivo depletion of CD4+ T cells was performed at various time points. Mice
depleted at the time of secondary infection had a similar number of larvae in their livers as nondepleted control mice. In contrast, mice depleted during the primary infection or throughout the
experiment failed to trap larvae to any significant degree. Measurement of T. canis-specific
serum antibodies by ELISA demonstrated a positive relationship between IgG levels and
hepatic larval counts. Altogether, these results suggest that CD4+ T cells – most probably the
Th2 subpopulation – is necessary at the time of primary T. canis infection to generate strong
humoral immunity, which then mediates migratory arrest of larvae in the liver during a
subsequent infection. The next step is to explore how this dynamic may be altered when a host
is infected with more than one parasite.
42
Student Competition - Control
4
Objective evaluation of two deworming regimens in young thoroughbreds:
parasitological parameters and growth rates.
Jennifer Bellaw1*, Joe Pagan2, Steve Cadell3, Eileen Phethean2, John Donecker4, Martin
Nielsen1. 1M. H. Gluck Equine Research Center, University of Kentucky, Veterinary Science,
Lexington, KY, 2Kentucky Equine Research, Versailles, KY, 3Hallway Feeds, Lexington, KY, and
4
Zoetis, Outcomes Research, Reidsville, NC.
Parasitic helminths of equids are capable of causing general ill-thrift, clinical disease, and death.
Parascaris spp. nematodes are arguably the most pathological parasites, primarily infecting
horses under one year of age. Although this is the most intensively treated cohort of horses and
anthelmintic resistance is an emerging problem, deworming regimens are rarely evaluated in
young horses. The goal of this study was to objectively evaluate the impact of deworming
regimen on the growth of young Thoroughbreds and to characterize parasite populations on
managed farms. Forty-nine Thoroughbred foals from three Central Kentucky farms were
randomly allocated to an interval dose program or daily deworming regimen. Those in the
interval dose program were treated bi-monthly beginning at two months of age, rotating between
pyrantel pamoate and ivermectin. The daily deworming group received oxibendazole at two
months of age, daily rations of pyrantel tartrate feed additive throughout the study, and
moxidectin treatments at 9.5 months of age. These horses, born February - May 2013, were
followed from two months of age and will continue to be monitored until the 2014 September
sales. Parasitological data, including monthly pre- and post-treatment fecal egg counts (FEC) of
Parascaris spp. and strongyle family parasites and gel/paste dewormer efficacies, were
collected. Additionally, monthly weighing data were collected. As of yet, deworming regimens do
not differ in strongyle parasite mean FECs; however, the interval dose program exhibited a peak
Parascaris spp. mean FEC 200 eggs per gram higher than the daily deworming regimen.
Pyrantel pamoate and ivermectin efficacies are 23.8%, 46.4% and 80.9-99.9%, 36.9-31.9% for
strongyles and ascarids respectively. Also, foal growth rates do not differ between the two
regimens. Despite differences in ascarid egg shedding levels, deworming regimen did not
appear to affect growth rates, leading to the conclusion that good management may
compensate for parasitism.
5
Transabdominal ultrasonography as a tool for monitoring Parascaris spp. in foals.
Maci Stephens1*, Clara Fenger2, Eugene T. Lyons1, John M. Donecker3, Martin K. Nielsen1.
1
M.H. Gluck Equine Research Center, University of Kentucky, Veterinary Science, Lexington,
KY, 2Equine Medical Medicine Consulting, Lexington, KY, and 3Zoetis Outcomes Research,
Reidsville, NC.
Parascaris spp. are pathogenic parasite nematodes universally infecting foals under a year of
age. Increasing levels of anthelmintic resistance in this parasite across the world pose a threat
for accumulating worm burdens and increased risk for life-threatening ascarid impactions and
rupture of the small intestine in foals. This condition is often triggered by anthelmintic treatment
of foals with large worm burdens, but there are currently no antemortem quantitative diagnostic
43
methods available for this parasite. The goal of this study was to develop and evaluate a
transabdominal ultrasonographic method for monitoring ascarid burdens in healthy foals. Ten
naturally infected foals were monitored with biweekly ultrasound examinations between August
2013 and April 2014, and euthanatized sequentially with subsequent ascarid worm count
throughout the study. The ultrasonographic examinations were performed in three different
transabdominal areas on the foal; the xiphoid area, the umbilicus and midway between the two.
Obtained images were scored on a scale from 1 to 4 according to the number of ascarid objects
encountered. A number of two by two tables were constructed evaluating different cutoffs for
high ascarid burden and positive test results. With two repeated examinations, diagnostic
predictive values of 100% were obtained with a Chi square analysis yielding a P-value below
0.05. The value of a single examination was evaluated by analyzing the three most recent
examinations as individual data points (n=27) prior to each necropsy. This analysis also
returned a significant Chi square analysis (P<0.05) with positive and negative predictive values
of 100% and 62.5%, respectively. Transabdominal ultrasonography could be useful as a
monitoring tool for Parascaris spp. in foals with more than 10 worms.
6
Evaluation of worm replacement as a means to reverse the impact of
multiple-anthelmintic resistant Haemonchus contortus on a sheep farm.
Melissa Miller*, Sue Howell, Adriano Vatta, Bob Storey, Ray Kaplan. The University of Georgia,
Department of Infectious Diseases, Athens, GA.
In Spring 2011 we confirmed a population of Haemonchus contortus infecting a flock of sheep in
Georgia were highly resistant to benzimidazoles, ivermectin, and moxidectin, and borderline
resistant to levamisole. Since the sheep were being moved to a new location not previously
grazed by livestock, worm replacement with a fully drug-susceptible laboratory isolate was
implemented to intervene in this desperate situation. Using a multiple-day regimen of a
levamisole/albendazole combination, FEC were reduced by 98.7%. In September 2011, each
animal was inoculated with 5000 L3 of a fully drug-susceptible laboratory isolate. DrenchRite
larval development assays (LDA) and fecal egg count reduction tests (FECRT) were completed
at several time points throughout the study to determine in vitro and in vivo resistance status.
After successful establishment of the susceptible isolate, the flock was moved to the new
location. Initially post-replacement, albendazole treatment yielded 97.0% FECR and LDA
showed reversion to susceptibility for all drugs, confirming successful replacement, though a
small resistant subpopulation persisted. After replacement, targeted selective treatments (TST)
based on FAMACHA were done using albendazole only. In April 2013, FECRT was performed
using albendazole yielding a FECR of -55.6%. FECRT were then performed on the other drugs,
yielding FECR of 98.8%, -200.0%, and 59.2% for levamisole, ivermectin, and moxidectin,
respectively. LDA also showed reversion to resistance for all except levamisole, which showed
borderline resistance. In May 2014, FECRT was performed using levamisole yielding a FECR of
40.7%. These data reveal that reversion to resistance was rapid, suggesting that lab isolates
may lack fitness to compete with field isolates. Future analysis of benzimidazole resistanceassociated SNP frequencies in beta-tubulin, and genetic analysis using microsatellite markers
should provide important insights for explaining what occurred on this farm.
44
7
Anthelmintic activity of Melilotus alba (white sweet clover) on Haemonchus contortus
infective juveniles and its cytotoxicity on bovine ileal epithelial cells.
Anwar Sarah1*, Larry Holler1, Sue Holler1, R. Neil Reese2, Michael Hildreth1,2. 1South Dakota
State University, Department of Veterinary & Biomedical Sciences, Brookings, SD, and 2South
Dakota State University, Department of Biology & Microbiology, Brookings, SD.
There is growing interest in finding natural plant products for use as alternatives to commercial
anthelmintics for controlling Haemonchous contortus in pastured sheep and goats. A previous
study revealed that methanol extracts from Melilotus alba (white sweet clover) possessed
anthelmintic activity in egg-hatch and larval migration assays involving H. contortus. However,
no anthelmintic activity was identified when freshly harvested, mature second-year M. alba
plants (predominantly stems and pods) were fed to yearling ewes naturally infected with H.
contortus. To understand these conflicting results, the anthelmintic activity of methanol extracts
from different sweet clover plant parts (i.e. leaves, stems, pods) were measured using a larval
migration assay involving unsheathed third-stage H. contortus larvae. Stems and pods showed
no anthelmintic activity, while 97.3% migration inhibition was measured in the leaf extract at 30
mg/ml. Inhibition was higher in aqueous solutions at an acidic pH (e.g. pH of 3 and 5) than at a
neutral pH (e.g. 7.4). An aqueous extract of the leaves (concentration of 670 mg/ml water)
inhibited migration by 98% after 24 hrs, and no motility was observed after 48 hrs. Cytotoxicity
of the aqueous leaf extract was measured with unpolarized bovine ileal epithelial cells at 5
differing concentrations (670, 340, 134, 67, and 17 mg/ml) of the extract using an absorbancebased AlamarBlue assay. After 48 hr treatments, extract concentrations higher than 134 mg/ml
were 100% lethal to cells; 11.4% of the cells died at 17 mg/ml. These in vitro studies indicated
that M. alba leaves contain one or more compounds with anthelmintic activity that is soluble in
methanol and water, but that these compounds are also cytotoxic to unpolorized intestinal cells.
Future in vivo studies should focus on young first-year plants that are composed primarily of
leaves, but ewes should also be carefully monitored for any toxicity problems.
8
Treatment and control of Giardia duodenalis in a dog colony used for veterinary
instruction.
Meriam Saleh1*, Alexandra D. Gilley2, Meghan Byrnes2, Anne M. Zajac1. 1Virginia Tech,
Department of Biomedical Sciences and Pathobiology, Blacksburg, VA, and 2Virginia Tech,
Department of Academic Affairs, Blacksburg, VA.
Infection with the protozoan parasite Giardia duodenalis is a common cause of canine diarrhea.
Infections are more common in dogs housed in kennels. Metronidazole and fenbendazole are
commonly used for treatment. Controlling infection can be difficult and should include cleaning
and disinfection. Twenty-two mixed breed dogs were acquired for instruction and added to the
existing colony of 12 dogs in August 2013. They were dewormed with Panacur® (fenbendazole
50 mg/kg) two weeks prior to arrival. In September several dogs developed diarrhea. Zinc
sulfate fecal flotations on pooled samples were positive for Giardia. All dogs were treated with a
7-day course of metronidazole, but no change in clinical signs was observed. Subsequently, all
dogs were tested by ZnSO4 flotation and 47% were positive for Giardia, the remaining 53%
were positive by immunofluorescence assay (IFA). We attempted to eliminate Giardia from the
colony using an integrated approach that included: 1) a 10 day course of fenbendazole (50
mg/kg), 2) removing dogs from kennels on treatment day 5 (D5) for bathing, 3) kennel
45
disinfection with Labsan 256CPQ (quaternary ammonium disinfectant) and drying (80°F for 24
hours), and 4) fecal pickup when walked outside. One dog also received metronidazole (30
mg/kg) D11-D17 because of loose stool. All dogs were IFA negative for Giardia on D8 and D13.
On D20 one cyst was detected in one dog. Additional samples from all dogs were examined by
IFA on days 20, 27, 34, 41, 62, 83, 104, 125, 146, 167,188 and were all negative for Giardia. A
sample collected before the integrated treatment was genotyped and found to be Assemblage
C. Over half of the dogs were adopted by D188, and another sample collection is scheduled for
the remaining dogs on D209. This integrated approach appears to have eliminated Giardia from
this colony, despite their outdoor access and frequent handling.
9
Demonstration of flotation of Toxocara canis eggs in commercial bleach (5.25% sodium
hypochlorite solution) and effects of bleach treatment on embryonation of T. canis eggs.
Alice Houk1*, Alexa Rosypal2, Anne Zajac3, David Lindsay1. 1Virginia-Maryland Regional
College of Veterinary Medicine, Virginia Tech, Department of Biomedical Sciences and
Pathobiology, Blacksburg, VA, 2Johnson C. Smith University, Department of Natural Sciences
and Mathematics, Charlotte, NC, and 3Virginia-Maryland Regional College of Veterinary
Medicine, Virginia Tech, Department of Biomedical Science and Pathology, Blacksburg, VA.
Toxocara canis is a common intestinal nematode of young dogs. Puppies contaminate the
environment with large numbers of eggs that embryonate and become infective in less than a
month. The eggs are environmentally resistant and able to persist from months to years under
optimal conditions. These embryonated eggs are infectious for humans and other paratenic
hosts. The seroprevalence of T. canis is estimated to be 14% in the US and infections in
humans are generally asymptomatic but the disease may have ocular, visceral, or neurological
presentations based on the migration of the larvae. Toxocariasis is 1 of 5 diseases the CDC has
labeled as neglected parasitic infections needing further study. Studies have found that T. canis
eggs are highly resistant to harsh environmental conditions and routinely used chemical
disinfectants. The objective of this study was to evaluate the effects of full-strength commercial
bleach (5.25% sodium hypochlorite solution) on embryonation of T. canis eggs for various timepoints and to report our incidental finding that T. canis eggs can be extracted from canine feces
using bleach flotation. T. canis eggs were collected in feces from weaned puppies and mixed
with full-strength bleach for 15 min, 30 min, 60 min and 120 min. Eggs were washed 3 times in a
cell culture medium containing 10% (v/v) fetal bovine serum and resuspended in embryonation
solution which was 0.1 N sulfuric acid. We examined samples daily for 18 days for development
of T. canis eggs after exposure to bleach treatments. Embryonation of eggs was observed in all
treatments and in eggs continuously exposed for 18 days. Our results indicate that bleach may
not be an appropriate disinfectant for dog kennels, cages, or laboratory utensils and work
surfaces. T. canis eggs are resistant to bleach treatment and continue to pose a risk for canine
and human infections.
46
Student Competition - Immunology
10
Evaluation of immune responses during Haemonchus contortus worm expulsion in
Native and Suffolk sheep.
Javier Garza*, James Miller. Louisiana State University, Pathobiological Sciences, Baton
Rouge, LA.
Sheep breeds are known to have varying levels of susceptibility or resistance to gastrointestinal
nematode (GIN) parasitism. While it is known that these differences are immunologically based,
they have yet to be fully elucidated. The immune response to H. contortus worm expulsion was
examined in Native and Suffolk lambs. Fifty-four Native and Suffolk lambs were allowed to
acquire a natural GIN infection in pasture. After being moved to parasite free housing for 2
months, lambs were given a challenge infection of 20,000 H. contortus L3 and were euthanized
at 0, 1, 3, and 7 days post infection (DPI). At necropsy abomasal mucosa, prescapular and
abomasal lymph node samples were taken for quantitative PCR analysis of IL-4, IL-5, IL-13,
IFN-γ, IL-10, and Foxp3 gene expression. Differences in Th2 cytokine (IL-5, and IL-13) gene
expression were seen in the Native lambs at 7 DPI compared to 0 DPI lambs but not in Suffolk
lambs. No differences in IFN-γ, IL-10, and Foxp3 gene expression in the abomasal mucosa
were found between breeds or time points. While there were no differences in IL-4 gene
expression between breeds, increases in expression at 3 and 7 DPI compared to 0 DPI were
noted in both breeds. IL-5 gene expression similarly increased over time in both breeds,
however, Suffolk lambs showed higher levels of expression at 3 DPI. Additionally, a two-fold
difference in IL-5 gene expression was observed in Native lambs compared to Suffolk lambs at
1 DPI. At 3 DPI, a two-fold difference in IL-5 was observed in the Suffolk lambs compared to the
Native lambs. Similarly, IL-13 gene expression increased significantly at 3 and 7 DPI compared
to 0 DPI in both breeds and a two-fold difference in the Native lambs compared to Suffolk lambs
was observed at 7 DPI.
11
Delayed immune responses of parasite-susceptible sheep during Haemonchus contortus
infection are associated with greater larval burden.
Jesica Jacobs*, Karen Sommers, Scott Bowdridge. West Virginia University, Division of Animal
and Nutritional Sciences, Morgantown, WV.
Previous data have indicated that interleukin-4 (IL-4) expression increases from day 3 to 7 and
is associated with local lymph node hypertrophy in challenged St. Croix lambs whereas no IL-4
expression or lymph node hypertrophy was observed in parasite-susceptible lambs during this
time. Immune responses occurring from day 7 to 14 during a Haemonchus contortus infection
are not well characterized. Therefore, the current study examines immune events occurring at
day 10 following a challenge H. contortus infection. Twenty-four lambs (12 St. Croix and 12
Suffolk crossbred) were randomly assigned to naïve or challenged infection groups. Gene
expression analysis of abomasal mucosa revealed IL-4 expression in challenged Suffolk
crossbred lambs on day 10, yet no detection in St. Croix lambs. Lymph node weight was not
different between breeds of challenged lambs as both groups averaged 4g, but were
significantly different from naïve lambs. In comparison to previous studies, a 7 day delay in IL-4
47
expression and a 5 day delay in lymph node hypertrophy of parasite-susceptible sheep were
associated with differences in worm burden. St. Croix lambs at day 10 had an average of 16
larvae in the abomasum compared to Suffolk crossbred lambs averaging 530 (P<0.05).
Reduction of IL-4 and lymph node weight in St. Croix sheep by day 10 indicates a resolution of
infection while increases observed in Suffolk crossbred lambs indicate an initiation of immune
response. Taken together these data indicate a significant role for IL-4 and local lymph node
hypertrophy in the rapid and early reduction of larvae in a parasite-resistant host. Thus, a delay
in the generation of host protective immune responses in parasite-susceptible sheep permit
establishment of adult worms thereby increasing pathology resulting from H. contortus infection.
12
Direct and indirect effects of Copper Oxide Wire Particles on Haemonchus contortus
infections in small ruminants.
Vicky Kelly*, James Miller. Louisiana State University School of Veterinary Medicine,
Pathobiological Sciences, Baton Rouge, LA.
Copper is an essential heavy metal required by all animals for life. Heavy metals have been
known to influence immune responses. Heavy metals such as lead and cadmium can alter the
isotope switching of antibodies from an IgM or IgG predominant manner to an IgE isotope and
alter lymphocyte differentiation, favoring a Th2 response over a Th1 response. Previous studies
have shown that copper, in the form of copper oxide wire particles (COWP), aids significantly in
the expulsion of Haemonchus contortus. To further assess the direct effect that COWP may
have on H. contortus, three Gulf Coast Native-Suffolk crossbred lambs received 15,000 L3 H.
contortus and were treated four weeks post infection with 2.0 g of COWP. SEM was
subsequently performed on adult worms one day after treatment and cuticular differences were
noted. Previous preliminary work demonstrated that COWP resulted in physical changes to the
cuticle of H. contortus using TEM. SEM results from this study indicated an increase in damage
to the cuticle of H. contortus from treated animals in the form of torn synlophe and an increased
occurrence of pock mark like damage. To assess possible immune responses to absorbed
copper, tissue samples were collected from the abomasum, small intestine, and select lymph
nodes for histopatholgy. Blood was collected to determine pack cell volume (PCV), total white
blood cell (WBC) counts, and WBC differentials. Preliminary results suggest that COWP has a
direct effect on H. contortus as well as an indirect effect on the immune response.
Histopathology suggested an increase in eosinophilia of the abomasum and small intestine, and
lymphoid hyperplasia of the abomasum and prescapular lymph nodes. Further work needs to be
completed to confirm these initial findings.
13
Effects of peripheral blood mononuclear cells on larval motility of Haemonchus
contortus in vitro.
Rush Holt1*, Amanda G. Ammer2, Scott A. Bowdridge1. 1West Virginia University, Division of
Animal and Nutritional Sciences, Morgantown, WV, and 2West Virginia University, Robert C.
Byrd Health Science Center, Morgantown, WV.
Parasite resistant hair sheep have been shown to generate elevated cellular immune responses
both peripherally and at local site of infection yet little is known of their specific activity in
parasite expulsion or direct larval killing in parasite-resistant or susceptible hosts. Culturing
48
peripheral blood mononuclear cells, derived from parasite-resistant St. Croix or parasitesusceptible Suffolk sheep, with Haemonchus contortus L3 stage larvae revealed differences in
location of PBMC binding. Suffolk-derived PBMC tend to bind posteriorly whereas St. Croixderived PBMC bind to both anterior and posterior portions of the parasite. To determine
differences in larval motility as a result of PBMC binding and infection status of the sheep, cells
were collected from 10 lambs of each breed, 5 of which were naïve and 5 were challenge
infected with H. contortus. To quantify larval motility a MIF Nikon Sweptfield microscope and
NIS Elements AR software were able to track parasite movement analyzing path length (µm),
velocity (µm/s) and acceleration (µm/s2). After 18 h of incubation differences in path were
observed as Suffolk naïve lambs had the greatest path distance (1862.85 µm) and velocity
(392.27 µm/s) compared to larvae exposed to cells from challenged animals of both breeds and
naïve St. Croix sheep (P < 0.01). Differences in acceleration were only detected between larvae
exposed to challenged St. Croix PBMC (10.91 µm/s2) and those of naïve Suffolk sheep (µm/s2)
(P=0.035). These data indicate no differences in larval mobility as a result of exposure to PBMC
derived from either challenge group or naïve St. Croix sheep. While PBMC binding affected
larval mobility, infectivity of PBMC-exposed H. contortus larvae is still unknown and will be
addressed in future experiments.
14
Modulation of vaccine-induced responses by anthelmintic treatment in ponies.
Emily Rubinson1*, Thomas Chambers1, David Horohov1, Stine Jacobsen2, Bettina Wagner3,
Alejandra Betancourt1, Stephanie Reedy1, Martin Nielsen1. 1Maxwell H. Gluck Equine Research
Center, University of Kentucky, Lexington, KY, 2Department of Large Animal Sciences-Medicine
and Surgery, University of Copenhagen, Copenhagen, Denmark, and 3Department of
Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell
University, Ithaca, NY.
Cyathostomins cause a type 2 helper T cell (Th2) response, which modulates inflammation. In
addition, anti-inflammatory effects of ivermectin have been found in rodent models. Conversely,
commonly used vaccines cause a marked acute phase inflammatory response. Anthelmintics
and vaccines are commonly given concurrently in routine equine management, but it is
unknown to what extent the interaction between the two exists. This study evaluated whether
the acute phase inflammatory response, the systemic gene expression of pro-inflammatory
cytokines, and vaccine-specific titers induced by three different vaccines (West Nile virus,
keyhole limpet hemocyanin, and Equine Herpes Rhinopneumonitis) would be modulated by
concurrent administration of ivermectin or pyrantel pamoate in cohorts of ponies naturally
infected with cyathostomins. Mixed-breed yearling ponies were blocked by gender and fecal
strongyle egg count, then randomly assigned to three treatment groups: ivermectin (n=8),
pyrantel pamoate (n=8), and control (n=7). All ponies received vaccinations intramuscularly on
days 0 and 29, and anthelmintics were administered on the same days. Whole blood, serum
and plasma samples were collected one, three and 14 days after each vaccination. Samples
were analyzed for inflammatory markers (haptoglobin, serum amyloid A, fibrinogen and iron),
cytokine (IL-1β, IL-4, IL-10, TNF-α and IFN-γ) gene expression and vaccine-specific IgG(a)
titers by ELISA. A marked acute-phase response was noted following both vaccination dates;
however, the increases in serum amyloid A and haptoglobin were less pronounced in the
ivermectin-treated group. West Nile Virus IgG(a) levels were numerically higher in the untreated
control group. Gene expression of the pro-inflammatory cytokines IL-1β and TNF-α increased
after the first vaccination, but decreased after the second vaccination in the anthelmintic-treated
49
groups, which could be related to a reduced parasite burdens in these. Taken together, the
study suggests that ivermectin has a modulatory effect on systemic inflammation, whereas
cytokine expression might be affected by the worm burden.
15
Evaluation of the systemic inflammatory response to anthelmintic treatment in ponies.
Alejandra Betancourt1*, Martin Nielsen1, Eugene Lyons1, David Horohov1, Stine Jacobsen2.
1
University of Kentucky, Lexington, KY, and 2Faculty of Health Sciences, University of
Copenhagen, Department of Large Animal Science, Taastrup, Denmark.
Grazing horses are widely exposed to infection with strongyle type parasites, infections which
are largely controlled with administration of anthelmintic formulations to avoid parasitic disease.
However, anthelmintic treatment can inadvertently induce inflammatory reactions and clinical
disease. Very little research has been performed evaluating the inflammatory response to
anthelmintic treatment, but one study indicates that treatment with moxidectin causes less of an
inflammatory reaction than treatment with other drugs. Within the scope of this study, we aimed
to explore the differences in inflammatory response following treatment with three different
anthelmintic drugs: moxidectin, pyrantel pamoate, and oxibendazole. A population (n=30) of
healthy, naturally parasitized ponies were allocated into the three treatment groups, based on
age and worm fecal egg counts. All ponies were weighed and received the labeled anthelmintic
dosage. Treatment efficacy was evaluated using the fecal egg count reduction test over a period
of eight weeks, with weekly egg counts. The inflammatory response was assessed at four
measuring points during the 14 days following treatment. Measurements involved
characterization of cytokine gene expression and systemic inflammatory reaction. The objective
of this study was to determine the effect of de-worming treatment on pro-inflammatory cytokine
gene expression in the peripheral blood, and to evaluate any correlation between the
expression of inflammatory cytokines with levels of acute phase proteins and inflammatory
markers. Fecal egg counts from the study confirmed resistance levels in the parasite population.
Treatment with oxibendazole and pyrantel pamoate was unsuccessful in the elimination of
luminal parasites. Moxidectin, however, was very effective and egg counts of zero persisted for
several weeks. Analysis of cytokine gene expression data and evaluation of acute phase protein
levels suggests that treatment with these anthelmintics provokes minimal peripheral
inflammatory responses.
Student Competition – Epidemiology & Molecular
16
County scale distribution of Amblyomma americanum in Oklahoma: addressing local
deficits in tick maps based on passive reporting.
Anne Barrett1*, Bruce Noden2, Eileen Johnson1, Susan Little1. 1Oklahoma State University,
Department of Veterinary Pathobiology, Center for Veterinary Health Sciences, Stillwater, OK,
and 2Oklahoma State University, Department of Entomology and Plant Pathology, College of
Agricultural Sciences and Natural Resources, Stillwater, OK.
50
Lone star ticks (Amblyomma americanum) transmit a wide variety of human and animal
pathogens in North America, including spotted fever group Rickettsia spp. and Ehrlichia spp.,
but geographic distribution records for this tick in the published literature are incomplete,
preventing accurate disease risk assessments. Indeed, a recent meta-analysis of the literature
showed a dearth of reports for A. americanum throughout much of Oklahoma where this tick is
widely considered to occur commonly. To address this lack and document the presence of A.
americanum in available habitats throughout the state, we are using dry ice traps and tick drags
to systematically collect adult and nymphal ticks from each county. To date, lone star ticks have
been collected from 18 counties from which this species had not been previously reported to be
established; A. americanum accounted for the great majority of all adult ticks found although
small numbers of Ixodes scapularis, Dermacentor variabilis, and A. maculatum were also
collected. A subset of A. americanum found at each site was retained for molecular detection of
Rickettsia spp. infection via PCR. Taken together, these data suggest that A. americanum ticks
and the disease agents they transmit are much more widespread in Oklahoma than reflected in
the published literature, a phenomenon likely repeated throughout the geographic range of this
tick in the eastern half of North America, and provides a straightforward model for field
entomologists to correct local and regional deficits in tick distribution reports that could
contribute to underestimation of the public and veterinary health importance of tick-borne
infections.
17
Confirmation of Borrelia burgdorferi and Anaplasma phagocytophilum in established
populations of Ixodes scapularis in southwestern Virginia.
Brian Herrin1*, Anne Zajac2, Mauricia Shanks3, Susan Little1. 1Oklahoma State University Center
for Veterinary Health Sciences, Veterinary Pathobiology, Stillwater, OK, 2Virginia-Maryland
Regional College of Veterinary Medicine, Department of Biomedical Sciences and Pathobiology,
Blacksburg, VA, and 3M&J Pet Grooming, Pearisburg, VA.
In recent years, the geographic range of Ixodes scapularis, which transmits Borrelia burgdorferi
and Anaplasma phagocytophilum to people, dogs, and horses in the northeastern and upper
Midwestern United States, has dramatically increased in both latitude and altitude. To determine
the prevalence of these infections in a newly established population of I. scapularis in the
mountainous region of southwestern Virginia, questing adult Ixodes spp. ticks were collected
and the identity and infection status of each tick individually confirmed by PCR. A total of 364
adult ticks were tested from three locations, one a public park bordered by woody edge habitat
and a river (Site A), the second a wooded, low-density residential area (Site B), and the third a
recreational lake area (Site C). All ticks were morphologically and molecularly confirmed to be
Ixodes scapularis. Borrelia burgdorferi sensu stricto was detected by sequence-confirmed PCR
of flaB in a total of 32/101 (32%) ticks from site A, 49/154 (32%) ticks from site B, and 36/101
(36%) ticks from site C, for a total prevalence rate of 33% (117/356), confirming that the agent
of Lyme disease is emergent or endemic in this region. In addition, Anaplasma phagocytophilum
was also detected in a total of 4/364 (1.1%) ticks, two from both Sites A and B. Two of the ticks
positive for A. phagocytophilum were co-infected with B. burgdorferi. The prevalence of both
pathogens in ticks is similar to that reported from areas considered endemic for Lyme disease
and/or human granulocytic anaplasmosis in the United States and provides further evidence of
range expansion for both the tick vector and the public and veterinary health risk for these
diseases.
51
18
Prevalence of vector-borne pathogens in dogs from Haiti.
Lindsay Starkey1*, Kassie Newton2, Jill Brunker2, Kelly Crowdis3, Emile Jean Pierre Edourad4,
Pedro Meneus5, Susan Little1. 1Oklahoma State University, Department of Veterinary
Pathobiology, Stillwater, OK, 2Oklahoma State University, Department of Veterinary Clinical
Sciences, Stillwater, OK, 3Christian Veterinary Mission, Bon Repos, Haiti, 4Ministère de
l'Agriculture des Ressources Naturelles et du Dèveloppment Rural, Port Au Prince, Haiti, and
5
Ministère de l'Agriculture des Ressources Naturelles et du Dèveloppment Rural, Cap-Haitien,
Haiti.
Vector-borne pathogens are of concern to people and dogs on some Caribbean islands,
including Haiti, where survey data for canine vector-borne infections are lacking. To determine
the prevalence of a variety of vector-borne pathogens in dogs from Haiti, we conducted a
molecular and serologic survey of whole blood collected from 210 owned dogs in 2013, 28
(13.3%) of which were infested with Rhipicephalus sanguineus ticks at the time of blood
collection. No other tick species were identified on these dogs. A commercially available ELISA
(SNAP® 4Dx® Plus) was utilized on all 210 dogs; 37 (17.6%) had antibodies to Anaplasma
spp., 69 (32.9%) had antibodies to Ehrlichia spp., and 0 (0%) had antibodies to Borrelia
burgdorferi. Antigen to Dirofilaria immitis was detected in 55 dogs (26.2%). To date, blood from
183 dogs has been tested by PCR for vector-borne pathogens. Babesia canis vogeli was
detected in 14 (7.7%) dogs and Hepatozoon canis was detected in 37 (20.2%). Wolbachia spp.
DNA was present in 39 (21.3%) dogs. Ehrlichia canis was detected in 14 (7.7%) dogs with an
additional 16 awaiting sequence confirmation. At least 5 (2.7%) samples were positive for A.
platys (sequence confirmed), but the nested PCR used will also detect Wolbachia spp., and
sequence confirmation is pending on 24 samples. None of the 183 canine samples tested were
PCR-positive for B. gibsoni, E. chaffeensis, E. ewingii, or H. americanum. Co-infection with 2 or
more tick-borne pathogens was detected in at least 4 (2.2%) dogs. Evidence of past or current
infection with at least one vector-borne pathogen was detected in 138 (65.7%) dogs in this
study. The common nature of these pathogens, some of which are zoonotic, suggests that
canine and public health in Haiti would benefit from vector control programs to prevent infection.
19
Comparative patterns of recruitment, reproduction, and size of a generalist trematode,
Dicrocoelium dendriticum, in sympatric hosts.
Melissa Beck1*, Cameron Goater1, Douglas Colwell2. 1University of Lethbridge, Biological
Sciences, Lethbridge, AB, Canada, and 2Agriculture and Agri-Food Canada, Livestock
Parasitology, Lethbridge, AB, Canada.
Our understanding of epidemiological processes of generalist parasites is poor, in part because
there is little information available on the relative performance of individual worms within multihost systems. In this study we used a combination of experimental exposures in cattle and
sheep, field collections of livers from hunter-shot elk, and assessments of per capita worm
fecundity in artificial media to compare the recruitment, reproduction and size of adult
Dicrocoelium dendriticum, a generalist trematode that has emerged in grazing hosts in Cypress
Hills Provincial Park, Alberta and other regions of northern North America. Overall, there were
no remarkable differences in the recruitment of metacercariae within artificially exposed cattle
and sheep. Fluke body surface area (BA) and body length, and also egg output were
approximately equal in all three host species, although infrapopulations in sheep and cattle
52
tended to contain slightly larger worms than elk. Fluke from sheep also produced eggs slightly
earlier than those in cattle. Although all worms evaluated in this study (N=168) were gravid,
reproductive inequality between individual worms was high (range = 1-5820 eggs/day/worm)
within each infrapopulation, and approximately equal among the 3 host species. Nested ANOVA
analyses indicated that most of the observed total variation in worm BA and reproduction could
be explained by differences between host individuals, not between host species. These data
support the idea that D. dendriticum is an extreme host generalist, with worm fitness
approximately equal among at least three potential definitive hosts. The generalist life-history
strategy of this trematode, which is known to extend to other stages of its life cycle, likely has
contributed to its invasion history outside of its native range in Europe.
20
Development of a “Nemabiome” sequencing assay for the relative quantitation of
parasitic nematode species in cattle fecal samples.
Russell Avramenko1*, Elizabeth Redman1, Roy Lewis2, James Wasmuth1, John Gilleard1.
1
University of Calgary, Veterinary Medicine, Calgary, AB, Canada, and 2Merck, Westlock,
Canada.
Parasitic nematode infections are a major cause of production loss and disease in cattle
worldwide. Nematodes often occur as mixed species infections, which vary in pathogenicity and
drug sensitivity. Fecal Egg Counts (FECs) are used to assess parasite burdens and drug
efficacy. However, eggs must be cultured to L3 larvae and carefully examined by microscopy to
determine species identity. This is labour intensive and requires specialist expertise. Molecular
approaches have been used for species identification using the rDNA ITS-2 locus for a number
of years. However standard or real time PCR approaches are at best semi-quantitative and can
only detect pre-defined species. They also have repeatability and scalability problems. We have
developed an approach, in which the rDNA ITS-2 region is simultaneously sequenced from
thousands of parasites (L3 larvae or eggs) per sample at once using “NextGen” sequencing
approaches. This is akin to ‘microbiome’ sequencing of bacterial communities and thus have
called it “nemabiome” sequencing. The assay provides digital data, allowing accurate
assessment of relative species quantities per sample. Multiple samples can be sequenced
simultaneously in a single Illumina MiSeq run. We have established the protocol from sample
preparation to data analysis and have verified the sensitivity, specificity and quantitative
accuracy using artificially created larval pools from the major cattle nematodes. This assay will
provide a powerful tool to conduct epidemiological studies, surveillance and improve drug
efficacy assessments in cattle parasites. We have applied the assay to investigate parasite
species prevalence in Canadian cattle. We have observed regional differences in the
distribution of nematodes. Cooperia oncophora and Ostertagia ostertagi are the most prevalent
species across Canada, while Cooperia punctata is present in the eastern provinces. We have
observed distinct shifts in parasite species proportions following anthelmintic treatment,
indicating which species are surviving anthelmintic drug treatment.
53
21
Ascaris suum ACR-21 forms a homomeric nicotinic acetylcholine receptor with novel
pharmacology.
Shivani Choudhary1*, Melanie Abongwa1, Samuel K. Buxton2, Ciaran J. McCoy3, Alan P.
Robertson4, Saurabh Verma4, Richard J. Martin5. 1Graduate Student, Biomedical Sciences,
Ames, IA, 2Post-Doc, Biomedical Sciences, Ames, IA, 3Graduate Student, Biological Sciences,
Belfast, Northern Ireland, United Kingdom, 4Research Associate Professor, Biomedical
Sciences, Ames, IA, and 5Major Professor, Biomedical Sciences, Ames, IA.
Parasitic infections impose a substantial burden on the economy by affecting the health and
productivity in animals as well as humans. The use of highly effective and relatively inexpensive
chemotherapeutic drugs like nicotinic agonists has been central to combatting these infections
in both human and veterinary medicine. The nicotinic agonists specifically target nicotinic
acetylcholine receptors (nAChRs) found at the nematode neuromuscular junction, causing
depolarisation and hypercontraction of the parasite. Unfortunately, heavy reliance and extensive
use of anthelmintic compounds has led to the inevitable emergence of resistance. There is,
thus, an urgent need to develop new agents to address this issue. With the advent of genome
sequencing, a large number of distinct nAChR subunits have been described. ACR-21 is an α-9
like nAChR subunit that has been identified in the Ascaris suum and Caenorhabditis elegans
transcriptome. Using molecular techniques, we have cloned an ACR-21 subunit gene from A.
suum and analysed the expression and pharmacology of the receptor formed by the subunit
using two electrode voltage-camp electrophysiological techniques. The ACR-21 subunit has
been reported to be present in the ventral nerve cord neurons. Based on our RT-PCR results
we demonstrate its presence in muscle flap, pharynx, gut, head region and ovijector of A. suum.
This indicates the widespread expression of the subunit throughout the worm. We also found
that ACR-21 forms a homopentameric receptor and requires an ancillary factor, RIC-3 for its
functional expression in Xenopus laevis oocytes. Our results show that the receptor expressed
by ACR-21 subunit, is more sensitive to acetylcholine than to nicotine and is insensitive to
levamisole and pyrantel. This pharmacological and physiological identification of the previously
uncharacterised ACR-21 subunits offers a possibility for this receptor subunit to be used as a
potential target site for developing new anthelmintic compounds.
Student Competition – Wildlife & Parasite Biology
22
A survey of the parasites of urban red foxes (Vulpes vulpes) in Charlottetown, Prince
Edward Island, Canada based on fecal analysis.
William Robbins1*, Gary Conboy2, Raphael Vanderstichel3, Spencer Greenwood4, Sheldon
Opps5, Marina Silva-Opps1. 1University of Prince Edward Island, Biology, Charlottetown, PE,
Canada, 2Atlantic Veterinary College, Pathology and Microbiology, Charlottetown, PE, Canada,
3
Atlantic Veterinary College, Health Management, Charlottetown, PE, Canada, 4Atlantic
Veterinary College, Biomedical Sciences, Charlottetown, PE, Canada, and 5University of Prince
Edward Island, Physics, Charlottetown, PE, Canada.
The red fox (Vulpes vulpes) is the most widely distributed of all carnivores globally and has
recently become increasingly prevalent in urban areas in many regions where it occurs.
Currently, 42 red fox den sites have been identified within Charlottetown, Prince Edward Island
54
(PEI) – a small urban center. Red foxes share a susceptibility to the same helminth and
protozoan parasites as dogs and in some cases with cats. The movement of red foxes into
human inhabited areas has the potential to increase the parasite infection exposure risk to pets
and in the case of zoonotic parasites, to humans through environmental contamination. In order
to determine the specific parasites involved and the seasonality of parasite shedding, red fox
scat samples were collected on a weekly basis from the vicinity of 16 den sites scattered
throughout the city. Both Zinc Sulphate centrifugal flotation and a modified Baermann technique
were used to examine samples. From July 2, 2013 to March 20, 2014, 163 scat samples were
obtained and analyzed. Parasites detected included: Uncinaria stenocephala (55.6%),
Crenosoma vulpis (45.6%), Alaria spp. (31.5%), Eucoleus boehmi (27.8%), Eucoleus aerophilus
(26.5%), Isospora canis (19.75%), Aonchotheca putorii (19.1%), Isospora ohioensis-complex
(11.7%), Toxocara canis (11.7%), Cryptocotyle lingua (9.3%), Demodex spp. (5.6%),
Sarcocystis spp. (6.2%), Physaloptera spp. (0.6%) and Taenia spp. (0.6%). Spurious parasites
detected included: Monocystis spp. (58%), Eimeria spp. (7.4%), Citotaenia spp. (4.3%),
Baylisascaris procyonis (0.6%), Rodent Pinworm (6.2%), Strongyle-type (2.5%), and Callodium
hepaticum (0.6%). One or more canid parasites were detected in nearly all of the scat samples
collected (90.2%) indicating a high prevalence of infection in the urban red fox population. The
environmental contamination resulting from the urban red foxes is likely to increase the
exposure and risk of parasitic infections to pets and humans in Charlottetown.
23
Detection and characterization of haemosporidia in whooping cranes and sandhill
cranes.
Miranda R. Bertram1*, Gabriel L. Hamer2, Barry K. Hartup3, Sarah A. Hamer1. 1Texas A&M
University, Department of Veterinary Integrative Biosciences, College Station, TX, 2Texas A&M
University, Department of Entomology, College Station, TX, and 3International Crane
Foundation, Baraboo, WI and University of Wisconsin, Department of Surgical Sciences,
Madison, WI.
The population of the endangered whooping crane (Grus americana) is not increasing as
quickly as desired for recovery efforts, and we hypothesize that disease may be limiting the
population growth. While parasite infection can cause direct pathology and disease, it can also
indirectly reduce fitness. For example, studies have documented that passerines infected with
avian malaria (Plasmodium sp.) have reduced breeding success and population recruitment.
The goal of our study was to quantify the prevalence of blood parasites, including Plasmodium
sp. and Haemoproteus sp., in cranes as a first step in determining whether these parasites may
be impacting population growth. In addition to analyzing whooping crane blood, we are also
employing a surrogate species approach through the analysis of sandhill cranes (Grus
canadensis). The sandhill crane is an appropriate surrogate because it is common in our study
region, co-mingles with whooping cranes, and is actively hunted in regions of the state, thereby
allowing us to necropsy a large number of sandhill cranes. Whooping cranes were captured
between December 2009 and January 2013 and blood was collected as part of a health
assessment and spatial telemetry study of the population. Blood was also collected from
sandhill cranes at necropsy between November 2012 and January 2014. Samples were
subjected to PCR analysis and DNA sequencing to detect Plasmodium sp. and Haemoproteus
sp. Preliminary results show 59% of whooping cranes (n=46) and 75% of sandhill cranes (n=16)
were infected with haemosporidia parasites. A phylogenetic analysis shows that most of these
haemosporidia lineages group with previously reported parasites. However, one lineage of
55
Plasmodium collected in several whooping cranes falls into a unique clade. This study
documents that whooping cranes and sandhill cranes are exposed to a suite of Diptera-borne
haemosporidia. Future work will determine the impact of haemosporidia infection on crane
population health.
24
Detection of circulating cathodic antigen and circulating anodic antigen in raccoons
naturally infected with Heterobilharzia americana and comparison of test performance
with fecal sedimentation and fecal PCR.
Jessica Rodriguez1*, Govert van Dam2, Barbara Lewis3, Karen Snowden1. 1Texas A&M
University College of Veterinary Medicine and Biomedical Sciences, Veterinary Pathobiology,
College Station, TX, 2Leiden University Medical Centre, Parasitology, Leiden, Netherlands, and
3
Texas Veterinary Medical Diagnostic Laboratory, College Station, TX.
Heterobilharzia americana, a trematode (family: Schistosomatidae), infects a wide range of wild
and domestic mammals in the southeastern United States. Infected dogs present with a
spectrum of clinical disease ranging from diarrhea to end-stage liver failure. Currently
commercially available diagnostic tests include fecal saline sedimentation and fecal PCR.
Because these tests rely on the presence of eggs/parasite DNA in feces, which can be
intermittent, other methods of diagnosis may aid in properly identifying infected patients. A
commercially available immunodiagnostic test (POC CCA; Rapid Medical Diagnostics, Pretoria,
South Africa) detects circulating cathodic antigen (CCA) in urine of Schistosoma mansoni
infected humans. An experimental quantitative immunodiagnostic assay, detects circulating
anodic antigen (CAA) of schistosome parasites in the serum and urine of infected humans and
animals. This study evaluated the ability of these two tests, along with fecal saline
sedimentation and fecal PCR, to detect H. americana in naturally infected raccoons. Sixteen
raccoons were sampled secondarily from a planned group hunt. Using the identification of H.
americana eggs on histopathologic evaluation of tissues as a gold standard, 9 animals were
infected. A complete set of samples (urine, feces, small intestine, and liver) were available from
6 of the positive raccoons. H. americana was detected in 4/6 animals by urine POC CCA; 3/6 by
urine quantitative CAA; 4/6 by fecal saline sedimentation; and 5/6 by fecal PCR. In 2/6 cases, all
four tests agreed in the diagnosis of H. americana. In 1/6 cases, no test detected infection.
These data indicate that the CCA and CAA of S. mansoni are similar to that of H. americana
and may be useful in immunodiagnostic tests. More samples are required to estimate the
sensitivity and specificity of these tests in raccoons. Test performance may differ in dogs due to
potentially lower levels of parasitemia and lower prevalence rates.
25
Association between nymphal engorgment weight and sex of adult Amblyomma
americanum, Amblyomma maculatum, Dermacentor variabilis, and Rhipicephalus
sanguineus.
Yoko Nagamori1*, Jennifer Thomas1, Lisa Coburn2, Mason Reichard1. 1Oklahoma State
University, Veterinary Pathobiology, Stillwater, OK, and 2Oklahoma State University,
Entomology & Plant Pathology, Stillwater, OK.
The life cycle of ixodid ticks includes four stages: eggs, larvae, nymphs, and adults. Only adults
are sexually dimorphic, and differentiation between male and female ticks is morphologically
56
impossible as eggs, larvae, and nymphs. The purpose of our experiment was to determine if
engorgement weight of replete nymphs differed between those ticks that molted to males and
females. Laboratory-reared nymphs of Amblyomma americanum, Amblyomma maculatum,
Dermacentor variabilis, and Rhipicephalus sanguineus were fed to repletion on sheep, weighed
individually to the nearest mg, held separately according to their weights in humidity chambers,
and allowed to molt to adults. The mean engorgement weight of replete A. americanum nymphs
that molted to males was 8.4 mg (n = 327). The mean body weight of those nymphs that molted
to females was 13.2 mg (n = 354). Similarly, the mean nymphal body weights of A. maculatum,
D. variabilis, and R. sanguineus that became males were 17.8 mg (n = 155), 12.8 mg (n = 376),
and 4.9 mg (n = 307) respectively. The mean body weights of engorged nymphs that became
female were 20.5 mg (n = 304), 15.5 mg (n = 386), and 5.7 mg (n = 306) respectively.
Comparison of nymphal engorgement weights of each tick species showed that engorgement
weight of those nymphs that molted to females was significantly greater (P<0.001; respectively)
than those of males in these four species of ticks. Our findings indicate that there is a
relationship between nymphal engorgement weight and sex of adult A. americanum, A.
maculatum, D. variabilis, and R. sanguineus.
26
Investigating the suitability of beagle dogs as an animal model for the human parasite
Brugia malayi.
Erica Burkman*, Christopher C. Evans, Molly R. Riggs, Michael T. Dzimianski, Andrew R.
Moorhead. University of Georgia, Infectious Diseases, Athens, GA.
Lymphatic filariasis (LF) is a mosquito-borne disease caused by the parasitic nematodes
Wuchereria bancrofti and Brugia malayi. These parasites are a major cause of morbidity
globally, with an estimated 120 million people infected. Brugia malayi is the preferred laboratory
model for LF due to W. bancrofti requiring the use of primate hosts. For over 30 years, the
domestic cat has been utilized as the primary non-rodent animal model for B. malayi. However,
due to their low rate of patent infections and requirement for anesthesia during blood collection,
a more suitable non-primate laboratory animal is desired. Because the domestic dog is
permissive to infection with other filarial species, we decided to investigate the suitability of the
dog as a definitive host for B. malayi. Eight adult male and three adult female, heartwormnegative, beagle dogs were infected by subcutaneous injection of 500 or 1,000 B. malayi thirdstage larvae (L3). Beginning six months post-infection, animals were checked weekly for
microfilaremia using the modified Knott test. The first set of four dogs infected yielded two
animals that became patent and maintained a microfilaremia for approximately six weeks with a
peak level of 375 microfilariae (mf)/ml. The second set of seven dogs infected yielded three
patent animals with a peak level of 1,250 mf/ml. Two of these dogs have maintained a relatively
medium-to-low parasitemia for approximately six months, although the microfilaremia is too low
for blood-feeding mosquitoes. While the dog is not a suitable laboratory host for maintenance of
the B. malayi life cycle, the dog’s suitability as a model for the pathogenesis of LF remains to be
determined.
57
27
Inactivation of helminth eggs with short- and medium-chain fatty acids alone and in
combinations at naturally occurring concentrations.
Dan Zhu1*, Janice L Liotta2, Dwight D Bowman2. 1Cornell University, Biological and
Environmental Engineering, Ithaca, NY, and 2Cornell University, Microbiology & Immunology,
Ithaca, NY.
This study assessed the inactivation activity of short- and medium-chain fatty acids against
Ascaris suum eggs, which are routinely used as surrogates to study the ovicidal activity of
various manure and biosolids disinfection methods. Previous work with high acid concentrations
demonstrated eggs are killed when the pH of the acid solution is below the pKa of the acid;
under these conditions most of the acid is in its undissociated form. Expanding on this earlier
work, acetic acid, butyric acid, valeric acid, and caproic acid alone or in combination in a
complete block design were tested for their inactivation ability at 37°C at pH 4 with the acids at
concentrations generated naturally in a prototype of a dry toilet system where the fecal matter
undergoes acidic anaerobic digestion. There were 16 groups of eggs, 15 treated groups and
one untreated control group. Single acids had no significant effect on viability: ethanoic (acetic)
acid (87.84% viable), butanoic (butyric) acid (93.99% viable), pentanoic (valeric) acid (92.82%
viable), and hexanoic (caproic) acid (94.21% viable). Out of the 6 pairs of acids examined, only
one pair of acids (butanoic and hexanoic) caused a significant decrease in egg viability (0%
viable). Of the four combinations of triple acids, there were two triplet groups, ethanoic,
butanoic, and hexanoic acids & butanoic, pentanoic, and hexanoic acids, that caused significant
reductions (100% reduction, 0% viable) in egg viability. In addition there was a significant
decrease in egg viability (0% viable) when all four acids (ethanoic, butanoic, pentanoic and
hexanoic acids) were used in combinations at the given concentrations. Overall, viabilities were
reduced to zero in four sets of eggs at 37°C for 48 hours. The next work will examine these
concentrations in wetted sand, silt, and clay.
Plenary Session – Center for Veterinary Medicine
28
FDA's Center for Veterinary Medicine's Antiparasitic Resistance Management Strategy.
Michelle Kornele*, Anna O'Brien*. FDA-CVM, Office of New Animal Drug Evaluation, Rockville,
MD.
In September 2012, the FDA’s Center for Veterinary Medicine launched the Antiparasitic
Resistance Management Strategy (ARMS) to promote sustainable use of anthelmintic drugs in
grazing livestock species in the United States. The impetus for this initiative was CVM’s public
meeting held in March 2012 titled, “Antiparasitic Drug Use and Resistance in Ruminants and
Equines.” This meeting hosted seven national and international veterinary parasitologists and
pharmacologists. A paradigm shift from complete parasite elimination to parasite management
within herds was emphasized and CVM viewed the promotion of sustainable use of
anthelmintics as an opportunity for proactive leadership. Within the past year and a half, ARMS
projects have included educational outreach, speaking engagements, communication with
national and international regulatory bodies, and updating and developing applicable internal
regulatory policies. As the ARMS initiative looks ahead, we would like to encourage
58
conversation on antiparasitic resistance in livestock species among veterinarians and livestock
producers, and to increase education of sustainable parasite management practices. The
ARMS initiative also looks to professional veterinary organizations to assume leadership roles
on these issues. The small ruminant and equine industries have taken steps to educate
producers and veterinarians about antiparasitic resistance. Continued research is needed in
numerous aspects, particularly investigations into the practicality of refugia in the cattle industry
and gathering of prevalence data for antiparasitic resistance in small ruminants, equines, and
cattle in the US. Ultimately, promoting the responsible use of approved anthelmintics, so that
they will remain effective, should be a shared goal between CVM and all stakeholders, including
pharmaceutical firms, veterinarians, livestock producers and owners, researchers, and
academics.
Student Competition – Molecular & Epidemiology
29
Assessment of the genetic variability of Cytauxzoon felis rRNA Internal Transcribed
Spacers.
Dana Pollard1*, Mason Reichard2, Leah Cohn3, Andrea James1, Patricia Holman1. 1Texas A & M
University, College Station, TX, 2Oklahoma State University, Stillwater, OK, and 3University of
Missouri, Columbia, MO.
Cytauxzoon felis, a tick-borne hemoparasite, causes cytauxzoonosis in domestic cats in the
United States. Historically, the outcome from this devastating disease was nearly 100% fatal;
however, an increasing number of reports with cats overcoming cytauxzoonosis or remaining
subclinically infected has suggested that there are strains of C. felis that vary in pathogenicity.
The objective of this study was to use C. felis-infected blood samples of domestic cats from
Oklahoma, Arkansas, Kansas, Missouri, and Texas to assess the genetic variability of the
parasite first and second ribosomal RNA internal transcribed spacer (ITS-1, ITS-2) regions for
any spatial or clinical outcome associations. Cloned C. felis amplicons revealed several ITS-1
sequences that were identical to those previously reported, while the remaining ITS-1
sequences were unique. A previously undescribed 23-bp insert was found within the C. felis
ITS-1 sequence from a domestic cat from Oklahoma. Within the ITS-2 region, there were
sequences that were identical to those previously reported as well as some that were unique. A
40-bp insert in the ITS-2 region of several cloned C. felis amplicons from different domestic cats
was similar to the 40-bp insert previously found in C. felis from domestic and wild felids.
Combined ITS-1 and ITS-2 sequences found in this study corresponded to previously described
C. felis genotypes. Due to the identification of several unique ITS-1 and ITS-2 sequences, there
were also C. felis genotypes that were not previously described. From the data, genetically
variable strains of C. felis do exist but cannot be solely used in discriminating between less and
more pathogenic strains.
59
30
Diversity of Trypanosoma cruzi strain types in vector and host populations throughout
Texas.
Rachel Curtis1*, Gabriel Hamer2, Sarah Hamer1. 1Texas A&M University, Veterinary Integrative
Biosciences Department, College of Veterinary Medicine, College Station, TX, and 2Texas A&M
University, Department of Entomology, College of Agriculture and Life Sciences, College
Station, TX.
Trypanosoma cruzi is the etiologic agent of Chagas disease; infection with this zoonotic vectorborne protozoal parasite can cause cardiac disease or acute death in humans and dogs.
Various wildlife species serve as reservoirs for the parasite, and blood-feeding triatomine bugs
maintain sylvatic transmission and bridge transmission to humans. Chagas disease is endemic
across Latin America, where estimates indicate over 8 million people are infected. There is
increasing awareness for Chagas disease in the southern US, and human and veterinary cases
of Chagas disease became reportable in Texas in 2013. We aimed to characterize the T. cruzi
genetic strain diversity in vectors and mammalian hosts across Texas, because different
parasite strains may be associated with different vectors, hosts, and disease outcomes.
Beginning in March 2013, we trapped bugs and implemented a public submission program to
collect kissing bugs from diverse ecoregions. Mammalian tissue samples originated from
hunter-donated wildlife and client-owned dogs previously diagnosed with Chagas disease. We
screened all samples using real-time PCR and subjected positive samples to amplification and
sequencing of the TcSC5D gene in order to characterize the genetic strain type. Additionally,
selected tissue samples were subjected to histological preparation and examination for
visualization of parasites and cellular response to infection. We collected over 1200 bugs of six
species, and preliminary results indicate infection prevalence of bugs exceeds 50%. Our
preliminary results from wildlife tissues demonstrate higher infection prevalence in raccoons
than other species such as coyotes, foxes, bobcats, and feral hogs. The DNA sequencing
revealed that our Texas samples include strains TcI and TcIV, which are two of the six
recognized T. cruzi discrete typing units that occur throughout the Americas. Our future work is
aimed at characterizing variation in ecology, epidemiology, and pathogenic outcomes
associated with these particular parasite strains in vector and mammalian populations across
Texas.
31
The public health significance of macrocyclic lactone resistance in Dirofilaria immitis.
Cassan Pulaski1*, James Diaz2, John Malone1. 1Louisiana State University School of Veterinary
Medicine, Pathobiological Sciences, Baton Rouge, LA, and 2Louisiana State University Health
Sciences Center School of Public Health, Environmental and Occupational Health Sciences,
New Orleans, LA.
Strains of Dirofilaria immitis suspected of lack of efficacy (LOE) to macrocyclic lactone (ML)
preventive drugs have been reported in dogs by practicing veterinarians in the Lower
Mississippi Delta region. The author’s previous experimental infection studies provided in vivo
evidence of the existence of ML drug resistance in dogs infected by D. immitis L3 from suspect
field LOE cases in the region. If not controlled in the early stages, the emergence of ML drug
resistance threatens to become a widespread problem in the US that may limit the effectiveness
of current preventive drug treatment. This may lead to an increased prevalence of canine
dirofilariasis, specifically in southern states, as well as the possibility of a similar trend in human
60
dirofilariasis cases. Human infection with D. immitis is possible wherever the parasite is
endemic in dogs, and infection has been reported in many countries around the world. The
current review is aimed at describing the increasing prevalence rates of animal and human
dirofilariasis worldwide, as well the misconceptions regarding the pathophysiology and clinical
outcomes of animal versus human dirofilariasis.
32
Epidemiological factors effecting the prevalence of Entamoeba histolytica in different
populations of dogs.
Muhammad Mudasser Nazir1*, Muhammad Azhar Alam2, David Scott Lindsay3. 1Faculty of
Veterinary Sciences, Bahauddin Zakariya University, Pathobiology, Multan, Punjab, Pakistan,
2
University of Veterinary and Animal Sciences, Parasitology, Lahore, Punjab, Pakistan, and
3
Virginia – Maryland Regional College of Veterinary Medicine, Virginia Tech, Department of
Biomedical Sciences and Pathobiology, Blacksburg, VA.
Entamoeba histolytica is a ubiquitous protozoan parasite that affects humans, domestic animals
and wildlife all over the world and has been enlisted as a waterborne parasitic pathogen. The
aim of study was to examine the prevalence of amoebiasis in three different dog populations.
600 fecal samples were collected and processed for E. histolytica-like cysts by using triple fecal
test (microscopy) and fecal antigens of E. histolytica were detected using fecal antigen ELISA.
Samples from 281 household dogs without clinical signs, 122 samples from dogs exhibiting
clinical signs and 197 stray dogs were examined. The overall prevalence of positive samples
were 15.6% (94/600) by triple fecal test and 11% (66/600) by fecal antigen ELISA. Significant
difference (P < 0.05) of positivity was found between three populations. Forty-five (16.01%) and
32 (11.38%) of 281 fecal samples from household dogs without clinical signs were positive by
triple test and by antigen ELISA, respectively. Twenty-nine (23.8%) and 23 (18.8%) of 122 fecal
samples from household dogs with clinical signs were positive by triple test and by antigen
ELISA, respectively. Twenty (10.15%) and 11 (5.6%) of 197 fecal samples from stray dogs were
positive by triple test and by antigen ELISA, respectively. Dogs from the age group 1-3 years
were more likely to be E. histolytica antigen positive than were dogs from other groups; a
significant difference (P=0.012) was observed between different age groups. Prevalence was
significantly higher (P<0.05) in the summer season and no significant (P>0.05) difference was
seen in male and female dogs. These findings suggest that dogs may play an important role in
the epidemiology of this pathogen.
Cattle
33
Treatment of stocker calves at turnout with an eprinomectin extended-release injection
or a combination of injectable doramectin and oral albendazole.
Thomas A. Yazwinski1*, Paul Beck2, Chris Tucker1, Eva Wray1, Christine Weingartz1, Hannah
Gray2, Jeremy Powell1, Andrew Fidler1, Linda Jones1, Alan A. Marchiondo3, Himabindu B.
Vanimisetti3, Susan Holzmer3, Adriano F. Vatta3. 1University of Arkansas, Department of Animal
Science, Fayetteville, AR, 2University of Arkansas, Southwest Research and Extension Center,
Hope, AR, and 3Zoetis, Inc., Veterinary Medicine Research and Development, Kalamazoo, MI.
61
All phases of cattle production present unique challenges relative to nematode control, with the
stocker phase arguably the most problematic. Repeated as well as combination anthelmintic
treatments have long been recommended in Arkansas for optimal performance of stocker
calves. On 22 May 2013, 128 stocker calves (previously, randomly allocated to treatment after
blocking by body weight and pasture location, with fecal egg counts being stratified within the
weight blocks) were treated and grazed on treatment-specific, similar pastures (4 animals per
0.8 hectare pasture) for 119 days. Pasture was the experimental unit for treatment. Treatment
groups were saline-injected (8 pastures), 0.2 mg/kg doramectin (Dectomax Injectable Solution,
Zoetis Inc.) combined with 10 mg/kg albendazole (Valbazen Suspension, Zoetis Inc.) (12
pastures) and 1 mg/kg eprinomectin (LongRange Extended Release Injection, Merial Limited)
(12 pastures). Coprology and animal weighings were conducted throughout the study. At the
end of grazing, the animal from each pasture with the FEC closest to the pasture mean was
drylotted and then sacrificed for nematode quantifications. Over the 119-day grazing period,
average daily gains±SE were 1.24±0.06a, 1.48±0.05b and 1.56±0.05b lb/day for the saline,
combination and eprinomectin-treated groups, respectively (P<0.05). Relative to the salinetreated group, statistically significant reductions of FECs were noted for up to 58 days posttreatment (combination group) and up to 30 days PT (eprinomectin-treated group). Nematode
populations present at necropsy with sufficient nematodes in the controls to allow for statistical
analysis included adult Haemonchus placei, Ostertagia ostertagi, O. lyrata and Cooperia
punctata and early and late fourth larval stages of Ostertagia spp. No significant differences
between treatments were seen except that the combination group had fewer EL4 stages than
the controls. Judging from the above data, the one-time treatment of stocker calves in the spring
in Arkansas was not sufficient for adequate parasite control regardless of the treatment
employed.
34
Managing anthelmintic resistance in cattle parasites in New Zealand – choice of
anthelmintic product.
Dave Leathwick*. AgResearch, Animal Nutrition and Health, Palmerston North, Manawatu, New
Zealand.
Resistance to the macrocyclic lactone (ML) class of anthelmintics, in Cooperia oncophora, is
present on virtually every cattle farm in New Zealand. In addition, resistance to ivermectin has
now been confirmed in Ostertagia ostertagi from two farms. Historically, the anthelmintic market
in New Zealand has been dominated by pour-on products containing a ML, but evidence is
mounting that there are significant disadvantages from delivering anthelmintics by this route. A
study comparing the efficacy of moxidectin delivered by the oral, injectable or pour-on routes
found that against ML-resistant C. oncophora, the oral route resulted in significantly higher and
less variable efficacy than either the pour-on or injection. This presumably reflects the amount of
drug reaching the target worms in the gut. Modelling the effect of variation in the dose of drug
reaching the target worms indicates that both lower mean dose and/or a more variable dose is
likely to accelerate the development of resistance.
Comparable data against O. ostertagi in cattle is not currently available but equivalent studies
against ML-resistant Ostertagia spp. in deer show that while the injectable route is marginally
better than the oral (both giving similar efficacies) the pour-on route results in significantly lower
efficacy than the other routes. Given these data, the optimal anthelmintic to optimise worm
control and minimise the further development of resistance is an oral combination containing at
62
least two broad-spectrum actives. The continued use of pour-on products is likely to exacerbate
an already serious problem with anthelmintic resistance. Therefore, in New Zealand, the use of
oral combination drenches is highly recommended while the use of pour-ons, especially single
active products, is actively discouraged.
35
Use of conventional PCR as a tool to monitor gastrointestinal strongyle populations in
US cattle.
Ashley K. McGrew1*, Dante S. Zarlenga2, Lora R Ballweber1. 1Colorado State University,
College of Veterinary Medicine and Biomedical Sciences, Veterinary Diagnostic Laboratory, Fort
Collins, CO, and 2USDA, ARS, Animal Parasitic Diseases Lab, B1180, BARC-East, Beltsville,
MD.
It is impractical to think that gastrointestinal (GI) strongyles can be eradicated in cattle.
Therefore, determining which ones are present in a herd, as well as which ones remain after
anthelmintic treatment, will allow for more optimally designed control programs. This is
particularly important in light of the emergence of drug resistance among bovine strongyles,
where research has shown Cooperia and Haemonchus to be the common genera left behind
after treatment. Using a conventional PCR, with genera-specific primer sets capable of
differentiating the five major genera of bovine strongyles (Ostertagia, Cooperia, Haemonchus,
Trichostrongylus, Oesophagostomum), we are screening fecal samples for the presence of
these nematodes. Samples from 30 untreated groups, of up to 5 animals/group, have been
screened thus far. All five genera were present in 16/30 groups (53%). Haemonchus, Cooperia
and Oesophagostomum were found in 90% of the groups. Ostertagia (83%) and
Trichostrongylus (53%) were also common. A subset of 15 groups had post-treatment data
available. These groups were divided into 2 lots. Ostertagia, Cooperia, Haemonchus, and
Oesophagostomum were present in all pre-treatment groups. In the first lot, 6/7 post-treatment
groups had only Cooperia and Haemonchus. In the second lot, all post-treatment groups
contained Haemonchus; Cooperia and Ostertagia were found in 50% of the groups. Other
samples evaluated include 23 unpaired post-treatment groups. For these, Cooperia and
Haemonchus were the most common genera left behind (83% and 100% of the groups,
respectively). Ostertagia (52%), Oesophagostomum (22%), and Trichostrongylus (4%) were
also present after treatment. Our results are in agreement with previous data showing all five
genera are still present in the US, and that Cooperia and Haemonchus still appear to be the
primary genera present post-treatment. In addition, Ostertagia was found in some posttreatment samples, which is of concern should this trend continue.
36
Utilization of computer processed high definition video imaging for measuring motility of
microscopic nematode stages on a quantitative scale: “The Worminator”.
Bob Storey1*, Chris Marcellino2, Melissa Miller1, Mary Maclean1, Eman Mostafa3, Sue Howell1,
Judy Sakanari4, Adrian Wolstenholme1, Ray Kaplan1. 1University of Georgia, Department of
Infectious Diseases, Athens, GA, 2Case Western Reserve University School of Medicine,
Cleveland, OH, 3Zagazig University, Faculty of Medicine, Zagazig, Egypt, and 4University of
California at San Francisco, Center for Discovery and Innovation in Parasitic Diseases, San
Francisco, CA.
63
A major hindrance to evaluating nematode populations for anthelmintic resistance, as well as for
screening existing drugs, new compounds, or other bioactive plant extracts for anthelmintic
properties, is the lack of an efficient, objective and repeatable in vitro assay that is adaptable to
multiple life stages and parasite genera. To address this need we have developed the
“Worminator” system, which objectively measures the motility of microscopic stages of parasitic
nematodes on a quantitative scale. The system is built around the computer application
“WormAssay”, developed at the Center for Discovery and Innovation in Parasitic Diseases at
the University of California, San Francisco. WormAssay was designed to assess motility of
macroscopic parasites for the purpose of high throughput screening of potential anthelmintic
compounds, utilizing high definition video as an input to assess motion of adult stage
(macroscopic) parasites (e.g. Brugia malayi). Resulting motility scores provide a measurement
of drug effectiveness. We leveraged this state-of-the-art assay for use with microscopic
parasites by modifying the software to support a full frame analysis mode that applies the
motion algorithm to the entire video frame. Thus, the motility of all parasites in a given well are
recorded and measured simultaneously. Video is captured through an inverted microscope and
input to an Apple Imac® computer via HDMI™ and Thunderbolt™. Assays performed on
Caenorhabditis elegans, Cooperia spp., and Dirofilaria immitis L3, as well as B. malayi and D.
immitis microfilariae yielded reasonable and repeatable dose responses using the macrocyclic
lactones ivermectin, eprinomectin, doramectin, and moxidectin, as well as the nicotinic agonists,
pyrantel, oxantel, morantel, and tribendimidine. The resulting computer based assay is simpler
to use, does not rely on mesh screens as in the larval migration inhibition assay, or require
subjective scoring of motility by an observer, thereby providing increased efficiency, accuracy,
repeatability, and decreased variability.
Vector-Borne Diseases & Diagnosis
37
Ectoparasites of free-roaming domestic cats (Felis catus) presented to a spay-neuter
clinic in Oklahoma.
Jennifer Thomas1*, Yoko Nagamori1, Jaime Goolsby1, Lesa Staubus2, Mason Reichard1.
1
Center for Veterinary Health Sciences, Oklahoma State University, Department of Veterinary
Pathobiology, Stillwater, OK, and 2Center for Veterinary Health Sciences, Oklahoma State
University, Department of Veterinary Clinical Sciences, Stillwater, OK.
As knowledge of feline vector-borne diseases continues to emerge, a comprehensive
understanding of ectoparasites on domestic cats is needed. Feral and free-roaming domestic
cats presented to a spay-neuter clinic in Oklahoma were surveyed for ectoparasites from
January to May 2014. Cats were examined for infestation with fleas and ticks (Jan–May), lice
(Mar–May), and ear mites (Apr–May). The prevalence of fleas or flea dirt on cats was highest
(p=0.000002) in January (87.2%; 41/47) compared to other months which ranged from 43.2%
(41/95) in Apr to 66.1% (41/62) in May. We sampled 187 fleas from 82 of 212 infested cats. The
majority (94.6%) of the fleas were Ctenocephalides felis followed by Pulex irritans (3.7%). In
each month of the study, ticks were found on at least one cat. However, cats sampled in April
and May had the highest (p=0.001) prevalence (22.6%; 24/106 and 33.3%; 22/66, respectively)
of infestation with ticks. Of 183 ticks collected, the majority (73.3%) were Amblyomma
americanum (68 adults, 67 nymphs, 2 larvae) followed by Ixodes scapularis (12.8%; 24 adults),
Dermacentor variabilis (9.6%; 11 adults, 7 nymphs), and Rhipicephalus sanguineus (2.1%; 4
64
adults). Tick burdens ranged from 1–23 per animal. Hair clippings are currently being evaluated
for infestation with lice. Otodectes cynotis was detected in 23.0% (23 of 100) of cats in April and
10.9% (7 of 64) of cats in May. To understand the variability of ectoparasites on domestic cats
throughout the year, data will continue to be collected throughout the remainder of 2014.
38
Evaluation of culture conditions for the cultivation of the Cytauxzoon felis erythrocytic
stage.
Dana Pollard1*, Mason Reichard2, Leah Cohn3, Patricia Holman1. 1Texas A&M University,
College Station, TX, 2Oklahoma State University, Stillwater, OK, and 3University of Missouri,
Columbia, MO.
Cytauxzoon felis, a tick-borne intraerythrocytic protozoan parasite, is the causative agent of
feline cytauxzoonosis, which may range from clinically normal to causing anorexia and death.
Disease progression is rapid with diagnosis often made during postmortem examination.
Currently, research on C. felis requires the use of infected cats. In vitro cultures of the parasite
would reduce dependence on infected cats for studies, thus the goal of this study is to establish
optimal conditions for continuous in vitro cultivation of C. felis in the erythrocytic stage. Modeled
after the microaerophilous stationary phase cultivation method for Babesia bovis, cultures were
initiated from C. felis infected blood samples from 9 different cats in combinations of 3 different
media and sera from 2 different animal species, with and without varying concentrations of
supplements. The cultures were monitored by daily microscopic examination of Giemsa-stained
cultured erythrocyte smears. Varying percentages of parasitized erythrocytes were initiated in
some cultures after dilution with uninfected erythrocytes or after undergoing different techniques
for isolating parasitized erythrocytes. Cultures were either passaged at a 1:2 or 1:4 subculture
ratio or received fresh erythrocytes on a weekly basis. Parasites persisted at low levels in
culture for as long as 102 days and underwent subculture to as high as 4th passage. HL-1
medium supplemented with fetal bovine serum and hypoxanthine has best supported the
parasite in vitro. Successful cultivation would allow an indefinite laboratory source of this
parasite for future research as well as a reduction in the dependence on using infected cats for
studying C. felis.
39
Factor selection for utilization in maps of four vector-borne diseases.
Dwight D. Bowman*. College of Veterinary Medicine, Cornell University, Department of
Microbiology & Immunology, Ithaca, NY.
On June 9-10 of 2012, the Companion Animal Parasite Council [CAPC] hosted a meeting of
experts to determine potential factors for inclusion in the modeling prevalence rates of
dirofilariasis, borreliosis, anaplasmosis, and ehrlichiosis, using the data that populates the maps
that appear on the CAPC website, http://www.capcvet.org/parasite-prevalence-maps. Three
groups were convened with the charge of identifying and ranking factors for initial inclusion in
models of vector-borne disease prevalence. These models could then be used to predict
observed prevalence rates throughout the United States. The three convened groups dealt with
mosquitoes and heartworm infection, prostriate ticks and Lyme disease and anaplasmosis, and
metastriate ticks and ehrlichiosis. Presented here will be a quick summary of the group
65
members, their deliberations, and a summary of the factors initially identified for inclusion in
models, which have been or are being developed.
40
Identifying relevant canine heartworm factors.
Dongmei Wang1*, Dwight D. Bowman2, Heidi Brown3, Michael Finney1, John B. Malone4,
Christopher S. McMahan1, Julia Sharp1, Robert Lund1. 1Clemson University, Department of
Mathematical Sciences, Clemson, SC, 2College of Veterinary Medicine, Cornell University,
Department of Microbiology & Immunology, Ithaca, NY, 3College of Public Health, University of
Arizona, Division of Epidemiology and Biostatistics, Tucson, AZ, and 4Louisiana State
University, College of Veterinary Medicine, Baton Rouge, LA.
Individual factors that were identified by the heartworm working group at the Companion Animal
Parasite Council [CAPC] meeting were evaluated as potentially useful predictors of prevalence
rates of canine heartworm in the contiguous United States. In particular, a data set provided by
CAPC, which contains county-by-county results of over 9 million heartworm tests conducted
during 2011-2012, were statistically analyzed via logistic regression. Factors that reliably
predicted canine heartworm prevalence rates were determined. These factors include climate
conditions (annual temperature, precipitation, and relative humidity), socio-economic conditions
(population density, household income), local topography (surface water and forestation
coverage, elevation), and vector presence (eight mosquito species). All of the examined factors
show some power for predicting heartworm prevalence. Beyond climatic conditions, the most
important factors appear to be economic. The results from this analysis were used to construct
a map depicting estimated baseline heartworm prevalence rates in the United States.
41
Factors influencing the seroprevalence of canine anaplasmosis in the United States.
Christopher S. McMahan1*, Dwight D. Bowman2, Richard E. Goldstein3, Robert Lund1, Julia
Sharp1, Dongmei Wang1. 1Clemson University, Department of Mathematical Sciences,
Clemson, SC, 2College of Veterinary Medicine, Cornell University, Department of Microbiology &
Immunology, Ithaca, NY, and 3College of Veterinary Medicine, Cornell University, Department of
Clinical Sciences, Ithaca, NY.
Factors identified by the prostriate tick working group, at the Companion Animal Parasite
Council [CAPC] meeting, were analyzed for their ability to explain/predict regional prevalence
rates of canine anaplasmosis within the conterminous United States. This work explores spatial,
temporal, environmental, economic, and vector-born factors that have been suggested to
influence anaplasmosis prevalence rates. The examined factors include climate conditions
(annual temperature, precipitation, and relative humidity), socio-economic conditions (population
density, household income), local topography (surface water, forestation coverage, and
elevation), and vector presence (measures of wildlife/human interaction, such as deer/car
strikes). These factors were then analyzed with anaplasmosis test data, which were obtained
from CAPC, and consist of approximately 2 million testing results conducted during the years of
2011 and 2012. The identified significant factors were then utilized to develop predictive models
for the purpose of forecasting future annual prevalence rates of canine anaplasmosis.
66
42
What’s trending now: what parasites are we most commonly seeing, through the
snapshot of diagnosis by the Veterinary Diagnostic Laboratory at the Oregon State
University College of Veterinary Medicine?
Janell Bishop-Stewart*. Oregon State University College of Veterinary Medicine Veterinary
Diagnostic Laboratory, Bacteriology/Parasitology, Corvallis, OR.
Trending: It’s a popular word these days. Trend analysis is a hot topic. This presentation will be
an evaluation for any trends of parasites, based on samples submitted to the Veterinary
Diagnostic Laboratory (VDL) at Oregon State University’s College of Veterinary Medicine. Due
to the nature of the VDL, it affords a chance to evaluate the parasite trends of a wide variety of
animal species. The focus will mainly be on Oregon, but other areas of the country will also be
discussed. Often, requests are made as to what types of parasites are diagnosed in a particular
area, time of year or over the course of any year. This evaluation will span a multiyear period to
monitor for potential changes in seasonality of parasite diagnosis, as well as shifts in the
species of parasites found. This information on what is trending is important information for
clients, researchers and veterinarians, and provides animals with the best health care.
Horses & History
43
Strongylus vulgaris and colic – a retrospective case-control study.
Martin K Nielsen1*, Stine Jacobsen2, Susanne N. Olsen2, Holli S. Gravatte1, Eric Bousquet3,
Tina Pihl2. 1M.H. Gluck Equine Research Center, University of Kentucky, Department of
Veterinary Science, Lexington, KY, 2University of Copenhagen, Department of Large Animal
Sciences, Taastrup, Denmark, and 3Virbac, Carros, France.
Strongylus vulgaris is regarded the most pathogenic helminth parasite infecting horses. It was
once estimated to be the primary cause of colic in horses and has been termed the horse killer.
Disease is ascribed to thromboembolism caused by larvae migrating in the mesenteric arteries
eventually leading to ischemia and infarction of intestinal segments. This causes a painful colic
with an often fatal outcome. However, this knowledge is derived from studies with experimental
inoculation of parasite-naïve foals and case studies. This documents the pathogenic potential of
the parasite, but does not address its role as risk factor for colic in horse populations. This study
was designed as a retrospective case-control study among equine patients referred to the
University of Copenhagen Large Animal Hospital during 2009-2011. Every referred colic case
was matched with a patient of the same type (pony, warmblooded, coldblooded), age, gender,
and admitted in the same month and year, but for problems unrelated to the gastrointestinal
tract. Serum samples were analyzed for antibodies to migrating S. vulgaris larvae using a
recently developed ELISA. Three case definitions were used; colic sensu latum (n=274),
idiopathic colics (n=48), and strangulating (n=55) vs. nonstrangulating infarctions (n=22). Odds
ratios (OR) revealed no statistical association with ELISA results for the colic sensu latum and
idiopathic colic case definitions. However, nonstrangulating infarctions were strongly associated
with higher S. vulgaris titers, when compared to strangulating infarctions (OR=4.12, P=0.01).
Colic is a very broadly defined symptom complex with numerous possible risk factors
unaccounted for. Horses can harbor S. vulgaris infection without showing clinical symptoms,
67
and a recent study illustrated that a positive ELISA result should be interpreted as exposure to
the parasite within the preceding five months. Nonetheless, the ELISA may be helpful in
evaluating the more severe colic categories involving infarctions in the abdominal cavity.
44
Comparison of a single dose of moxidectin and a five-day course of fenbendazole to
reduce and suppress cyathostomin fecal egg counts in a commercial recipient mare
herd.
Ray Kaplan1*, Maren Mason2, Nathan Voris3, Hunter Ortis4, Amy Geeding4, Deb Amode5.
1
University of Georgia, Department of Infectious Diseases, College of Veterinary Medicine,
Athens, GA, 2University of Georgia, DVM Class of 2015, College of Veterinary Medicine,
Athens, GA, 3Zoetis, Inc., Equine Technical Services, Columbia, MO, 4Equine Medical Services,
Inc., Columbia, MO, and 5Zoetis, Inc., Outcomes Research, Florham Park, NJ.
Cyathostomins are the most prevalent and important internal parasites of adult horses. Infection
can cause a variety of disorders with larval cyathostominosis representing the most severe
manifestation. Effective control presents challenges due to pervasive exposure, anthelmintic
resistance and encysted stages of the cyathostomin life cycle. This study compared the efficacy
of the two larvicidal regimens in reducing and suppressing cyathostomin fecal egg counts (FEC)
in a commercial embryo transfer herd composed of mares from diverse locations across the US.
Given the herd’s geographic diversity, the farm’s cyathostomin population is expected to
represent a broad genetic heterogeneity of the US. Additionally, this farm had never utilized
fenbendzole or moxidectin so no further drug selection for resistance would have occurred.
Eighty-two mares with strongyle FEC ≥200 eggs per gram (EPG) were included. Mares were
ranked by EPG level, blocked, and then assigned randomly within block to treatment with either
a single dose of moxidectin or five consecutive days of fenbendazole at the larvicidal dose. FEC
data were analyzed 14, 45 and 90 days following treatment. Mean FEC reduction was 99.9% for
moxidectin-treated mares and 41.9% for fenbendazole-treated mares 14 days post-treatment.
By 45 days, fenbendazole group mean FEC exceeded pre-treatment levels, while moxidectin
group mean FEC levels remained suppressed throughout the study. Statistically significant
differences were observed between groups 14, 45 and 90 days post-treatment. In this study,
fenbendazole failed to adequately reduce or suppress FEC, suggesting failure to provide
adulticidal and larvicidal effects. In contrast, moxidectin was effective in reducing and
suppressing FEC for an extended period. Given the geographic diversity of mares, it is quite
likely that these results are representative of cyathostomin populations in much of the US.
These data confirm previous findings indicating cyathostomin resistance to fenbendazole is
widespread, and moxidectin remains effective for controlling these important parasites.
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45
Parascaris equorum is dead. Long live Parascaris univalens!
Martin K. Nielsen1*, Jianbin Wang2, Richard Davis2, Jennifer L. Bellaw1, Eugene T. Lyons1, Teri
Lear1, Clara Goday3. 1M.H. Gluck Equine Research Center, University of Kentucky, Department
of Veterinary Science, Lexington, KY, 2University of Colorado School of Medicine, Departments
of Biochemistry and Molecular Genetics and Pediatrics, Aurora, CO, and 3Consejo Superior de
Investigaciones Cientificas, Centro de Investigaciones Biológicas, Madrid, Spain.
The equine ascarid parasite Parascaris equorum is well-known as a ubiquitous parasite
infecting foals. However, very little attention has been given to the existence of a cryptic sibling
species named Parascaris univalens, despite the fact that it was first described over 100 years
ago. P. univalens is unique because it only possesses one chromosome pair as opposed to two
for P. equorum. Apart from their karyotype, the two Parascaris species are considered
morphologically identical. One survey performed in Italy in the 1970s found that 93% of 2,238
parasite specimens examined were P. univalens, and several attempts to identify P. equorum in
Spain in the 1980s failed due to an apparent 100% occurrence of P. univalens. For this study,
adult female worms obtained from the University of Kentucky parasitology horse herd were
dissected and identified using karyotyping techniques. With no exception, all specimens (n=35)
were identified to be P. univalens. Further, karyotyping was carried out on ascarid eggs
collected from 37 Thoroughbred foals from three farms in Central Kentucky in 2013, and P.
univalens was identified in 25 of these, with the remaining samples being inconclusive. Full
genomic sequencing was carried out on one worm identified as P. univalens, and comparison
with available sequence data from NCBI GenBank revealed an average 95-98% sequence
identity with what is published as P. equorum. However, it remains unclear whether the
sequences published in GenBank were unequivocally identified to species level by karyotyping,
and it is possible that this was really P. univalens mislabeled as P. equorum. Taken together,
these data suggest that P. univalens is likely the species now observed in equines and that
perhaps Parascaris spp. should be used unless cytological characterization has confirmed the
species. We are working to develop comprehensive genomic data resources for P. univalens.
46
Serum Strongylus vulgaris-specific antibody responses to anthelmintic treatment in
naturally infected horses.
Martin K. Nielsen1*, Anand N. Vidyashankar2, Jennifer L. Bellaw1, Holli S. Gravatte1, Xin Cao2,
Emily Rubinson1, Craig R. Reinemeyer3. 1M.H. Gluck Equine Research Center, University of
Kentucky, Department of Veterinary Science, Lexington, KY, 2George Mason University,
Department of Statistics, Fairfax, VA, and 3East Tennessee Clinical Research, Inc., Rockwood,
TN.
Strongylus vulgaris is the most pathogenic helminth parasite of horses, causing verminous
endarteritis with thrombo-embolism and infarction. Decades of intensive anthelmintic therapy
have reduced the prevalence of this parasite in managed herds. However, widespread
anthelmintic resistance in other equine parasites has led to recommendations of decreasing
treatment intensity through a surveillance-based approach. Recent research has linked such
strategies with an increased occurrence of S. vulgaris. To better monitor the occurrence of
pathogenic migrating larvae of this parasite, a serum enzyme-linked immunosorbent assay
(ELISA) has been validated for detection of antibodies to an antigen produced by arterial
stages. The aim of this study was to evaluate ELISA responses to anthelmintic treatment in
69
cohorts of naturally infected horses. Fifteen healthy horses harboring patent S. vulgaris
infections were turned out for communal grazing in May, 2013 (Day 0). On Day 55, horses were
ranked according to ELISA titers and randomly allocated to the following three groups: no
treatment followed by placebo pellets daily; ivermectin on Day 60 followed by placebo pellets
daily; or ivermectin on Day 60 followed by daily pyrantel tartrate. Fecal and serum samples were
collected at ~28-day intervals until study termination on Day 231. Increased ELISA values were
observed for the first 53 days following ivermectin treatment. Titers were significantly reduced
80 days after ivermectin treatment. Horses receiving daily pyrantel tartrate maintained lower
ELISA values from 137 days post ivermectin treatment until trial termination. These results
illustrate that a positive ELISA result is indicative of either current or prior exposure to larval S.
vulgaris infection within the previous five months.
47
Performance of a new duplex real-time PCR assay for Theileria equi and Babesia caballi
in horses.
Vladislav Lobanov*, Alvin Gajadhar. Canadian Food Inspection Agency, Centre for Food-borne
and Animal Parasitology, Saskatoon, SK, Canada.
Equine piroplasmosis (EP) is a hemolytic infection of horses and other equids caused by the
tick-borne protozoa, Theileria equi and Babesia caballi. Few regions of the world, including
North America, are considered free of EP. However, recent incursions have prompted renewed
efforts to maintain EP-free status in North America, including the development of reliable
diagnostic methods for EP. We developed a duplex real-time PCR assay for detecting the
causative agents of EP. A previously published real-time PCR assay for B. caballi (Bhoora et
al., Vet. Parasitol. 168:201-211, 2010) was modified by introducing compatible new primers and
a dual labeled fluorescent probe targeting the ema-1 gene of T. equi. After repeated
amplification of standards prepared by ten-fold serial dilution of total DNA extracted from equine
blood infected with either T. equi or B. caballi, the mean amplification efficiency (with standard
deviation) on a 6-log range of template concentrations was 94.5 (±0.85)% for B. caballi, and
93.9 (±1.65)% for T. equi. Assay specificity of 100% was achieved for both parasites when 60
blood samples from serologically negative horses from Canada or USA were tested. Ten copies
of a plasmid containing the ema-1 gene of T. equi were reproducibly detected by the assay in
the presence of total DNA extracted from uninfected equine blood (background DNA). DNA of
B. caballi was detected in up to 100,000-fold dilutions of a positive blood sample with 0.4%
parasitemia, suggesting analytical sensitivity of 4 x 10-6% infected cells. Performance
characteristics of this duplex real-time PCR assay will be compared to those of other molecular
and serological assays for these pathogens using archived diagnostic samples previously tested
by C-ELISA or IFAT and samples obtained from regions known to be endemic for EP.
48
Glimpses of parasitology history through commemorative medals.
Raffaele Roncalli*. Milltown, NJ.
Over the centuries, medals have been coined to celebrate distinguished personages or
historical episodes of note. This is also common in the world of parasitology, where as early as
the 17th century, medals were coined to celebrate distinguished researchers (i.e. Francesco
Redi--1684--et alia) for their discoveries as well as for special events. In more recent years,
70
several organizations have established memorial awards (medals) for distinguished researchers
for their contributions to parasitology i.e.: in the USA the Henry Baldwin Ward Medal by the
American Society of Parasitologists; in the U.K. the C. A. Wright Memorial Medal by the British
Society for Parasitology; in Australia the Bancroft'-Mackerras Medal by the Australian Society
for Parasitology and others. In addition, special medals have been coined to celebrate events of
note; among them two distributed by the American Association of Veterinary Parasitologists in
1993 to celebrate the authors (D. E. Salmon, T. Smith, F. L. Kilborne and C. Curtice) of the
discovery of the agents causing Texas Cattle Fever in 1893 and in 2005 to salute the 50th
anniversary of the first meeting of the association. These medals will assist in perpetuating the
history of parasitology in the years to come.
Heartworm
49
Administration of 10% imidacloprid-1% moxidectin protects cats from subsequent,
repeated infection with Dirofilaria immitis.
Susan Little1*, Joe Hostetler2, Jennifer Thomas1, Keith Bailey3, Anne Barrett4, Kaylynn
Gruntmeir4, Jeff Gruntmeir4, Lindsay Starkey4, Byron Blagburn5. 1Center for Veterinary Health
Sciences, Oklahoma State University, Department of Veterinary Pathobiology, Stillwater, OK,
2
Bayer HealthCare, Animal Health, Shawnee, KS, 3Center for Veterinary Health Sciences,
Oklahoma State University, Department of Pathobiology, Stillwater, OK, 4Center for Veterinary
Health Sciences, Oklahoma State University, Stillwater, NA, and 5College of Veterinary
Medicine, Auburn University, Department of Veterinary Pathobiology, Auburn, AL.
Infection of cats with Dirofilaria immitis causes pulmonary pathology and seroconversion on
antibody tests. Consistent administration of topical 10% imidacloprid-1% moxidectin has been
shown to result in sustained plasma levels of moxidectin in cats after three to five treatments, a
pharmacokinetic behavior known as steady state. To evaluate the ability of moxidectin steady
state to protect cats from subsequent infection with D. immitis, cats (n=10) were treated with the
labeled dose of 10% imidacloprid-1% moxidectin for four monthly treatments. Each cat was
inoculated with 25 third-stage larvae of D. immitis 7, 14, 21, and 28 days after the last treatment;
non-treated cats (n=9) were inoculated on the same days, serving as positive controls.
Measurement of serum levels of moxidectin confirmed steady state in treated cats. Blood
samples were collected from each cat from 1 month prior to treatment until 7 months after the
final inoculation and tested for antibody to, and antigen and microfilaria of, D. immitis. Cats
treated with 10% imidacloprid-1% moxidectin prior to trickle inoculation remained negative on
antibody (ANTECH Diagnostics) and antigen (DiroCHEK®, Zoetis) tests throughout the study
and did not develop gross or histologic lesions characteristic of heartworm infection. A majority
of non-treated cats became positive on antibody (6/9) and, after heat treatment, antigen (5/9)
tests by 3-4 and 6-7 months post infection, respectively. Histologic lesions characteristic of D.
immitis infection, including intimal and medial thickening of the pulmonary artery, were present
in every cat with D. immitis antibodies (6/6), although adult D. immitis were present in only 5/6
antibody-positive cats at necropsy. Microfilariae were not detected at any time. Taken together,
these data indicate that prior treatment with 10% imidacloprid-1% moxidectin protected cats
from subsequent infection with D. immitis, preventing both formation of a detectable antibody
response and development of D. immitis-induced pulmonary lesions.
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50
Protection of dogs against canine heartworm infection 28 days after 4 monthly
treatments with Advantage Multi® for Dogs.
Dwight D. Bowman1*, Joseph A. Hostetler2. 1College of Veterinary Medicine, Cornell University,
Department of Microbiology & Immunology, Ithaca, NY, and 2Bayer HealthCare LLC, Veterinary
Services, Shawnee Mission, KS.
Monthly heartworm preventives are designed to protect dogs by killing heartworms acquired the
month prior to their administration, and after treatment with most products, the drug levels
rapidly dissipate to very low levels. Work with Advantage Multi for Dogs and hookworms
showed protection throughout the month after administration of several monthly doses which
suggested that similar protection might occur with heartworms. This study assessed the amount
of protection afforded to dogs by the administration of a monthly dose of Advantage Multi for
Dogs for four months prior to infection with third-stage heartworm larvae (Dirofilaria immitis) 28
days after the last (fourth) dose. There were 16 purpose-bred dogs in the study that were
divided into two groups, 8 control and 8 treated dogs. Dogs were housed in a manner
preventing contact between animals and groups, and personal protective gear worn by staff
minimized the chance spread of the topically applied product between runs. The dogs in the
treated group received monthly applications of Advantage Multi for Dogs as per label
instructions on Study Days 0, 28, 56, and 84. On Study Day 112, all 16 dogs received 50 thirdstage larvae of D. immitis (“Missouri” isolate) via subcutaneous inoculation in the inguinal
region. The study was terminated on Day 262, and the number of heartworms per dog was
determined at necropsy. All the control dogs had adult worms in their pulmonary arteries (34.25
+ 4.71; range 25-41), and none of the dogs treated 4 times prior to infection harbored worms at
necropsy. Therefore the efficacy of prevention was 100% when the dogs were infected 28 days
after the last monthly treatment. When dogs receive consecutive doses of Advantage Multi for
Dogs as prescribed, heartworm infections will be prevented throughout the monthly dosing
interval after administration of several monthly doses.
51
Comparative efficacy of four commercially available heartworm preventive products
against the JYD-34 laboratory strain of Dirofilaria immitis.
Byron Blagburn1*, Allan Dillon1, Jamie Butler1, Joy Bowles1, Jane Mount1, Tracey Land1, Robert
Arther2, Cristiano Von Simson2, Robert Zolynas2. 1Auburn University, Auburn, AL, and 2Bayer
HealthCare Animal Health, Shawnee Mission, KS.
A controlled laboratory study was conducted to evaluate the efficacy of four commercial
products for the prevention of heartworm disease in dogs caused by the JYD-34 Dirofilaria
immitis strain. Forty–three commercially-sourced Beagle dogs, 10-12 months of age, were used.
On SD -30, each dog was inoculated subcutaneously with 50 infective, third-stage D. immitis
larvae. On SD -2, 40 dogs weighing 17.6-25.2 lbs were ranked by decreasing body weight and
randomized to five groups of eight dogs each. On SD 0, the dogs assigned to Group 1 were
treated with imidacloprid/moxidectin topical solution, Group 2 dogs were treated with selamectin
topical solution, Group 3 dogs were treated orally with milbemycin oxime + spinosad flavored
tablets, and Group 4 dogs were treated orally with ivermectin/pyrantel pamoate chewable
tablets. Group 5 dogs remained non-treated. Dosages for dogs in Groups 1-4 were based on
the individual body weight of each dog and current labeled dose banding for each commercial
product. Dogs in Groups 2, 3 and 4 were retreated with their respective product on SD 31 and
72
60 according to their most current body weight. On SD 94, whole blood was collected and
tested for adult heartworm antigen. On SDs 124-126 the dogs were euthanized and subjected to
necropsy examination for recovery of adult D. immitis. At necropsy, all 8 dogs in the non-treated
group were infected with adult D. immitis (13-32 worms/dog, geometric mean (GM) = 18.4
worms/dog). Three or more adult D. immitis and/or worm fragments were recovered from all 8 of
the dogs in each of Groups 2-4. The respective GM worm burdens/dog for Groups 2-4 was
13.1, 8.8, and 13.1 which resulted in 28.8, 52.2, and 29.0% efficacy, respectively. No worms
were recovered from any of the 8 dogs in Group 1 resulting in 100% efficacy.
52
Forecasting annual canine heartworm prevalence in the United States.
Robert Lund1*, Dwight D. Bowman2, Heidi Brown3, John B. Malone4, Christopher S. McMahan1,
Julia Sharp1, Dongmei Wang1. 1Clemson University, Department of Mathematical Sciences,
Clemson, SC, 2College of Veterinary Medicine, Cornell University, Department of Microbiology &
Immunology, Ithaca, NY, 3College of Public Health, University of Arizona, Division of
Epidemiology and Biostatistics, Tucson, AZ, and 4Louisiana State University, College of
Veterinary Medicine, Baton Rouge, LA.
Two methods for forecasting annual canine heartworm prevalence rates in the contiguous
United States have been compared. Our forecasting techniques make use of spatial, temporal,
environmental, and vector-born factors and can be used to alert the veterinary community to
potentially high levels of heartworm prevalence in advance. A data set provided by the
Companion Animal Parasite Council, which contains county-by-county results of over nine
million heartworm tests conducted in 2011 and 2012, was used to forecast heartworm
prevalence rates in 2013. Our proposed modeling techniques make use of logistic regression
and Bayesian methods. County-by-county factors, including climate conditions (annual
temperature, precipitation, and relative humidity), socio-economic conditions (population
density, household income), local topography (surface water and forestation coverage,
elevation), and vector presence (eight mosquito species) drive each forecast. Our forecasts are
generally in agreement with the observed 2013 prevalence rates, suggesting that our
techniques have adequate predictive capabilities. Forecasts using both models for 2014 are
presented.
53
Prevalence of Dirofilaria immitis antigen in feline samples after heat treatment.
Susan Little1*, Jeff Gruntmeir2, Chris Adolph2, Byron Blagburn3. 1Center for Veterinary Health
Sciences, Oklahoma State University, Department of Pathobiology, Stillwater, OK, 2Center for
Veterinary Health Sciences, Oklahoma State University, Department of Veterinary Pathobiology,
Stillwater, OK, and 3College of Veterinary Medicine, Auburn University, Department of
Pathobiology, Auburn, AL.
Diagnosis of Dirofilaria immitis infection in cats by detection of antigen is complicated, in part, by
formation of antigen-antibody complexes in feline serum samples. Heat treatment of serum prior
to testing destroys these complexes, allowing detection. To determine the degree to which the
presence of blocking antibodies or other inhibitors of antigen detection may interfere with our
ability to detect circulating antigen in feline samples, archived plasma and serum samples (n =
220) collected from cats in animal shelters in the southeastern and southcentral United States
73
were tested for D. immitis antigen before and after heat treatment using a commercial assay
(DiroCHEK, Zoetis). Only 1/220 feline samples was positive before heat treatment, but 13/220
(5.9%) were positive after heat treatment by both colorimetric and spectrophotometric
evaluation. The single sample positive before heat treatment increased in O.D. more than 6 fold
following heating. Paired D. immitis antibody results (SoloStep, Heska) were available for the
samples in which antigen was detected; 10/13 were positive. These data suggest that some
serum and plasma samples from cats in the southern United States contain inhibitors which
may prevent detection of D. immitis antigen. Although additional research is needed to confirm
these findings, surveys conducted based on D. immitis antigen tests alone, without first treating
samples to disrupt immune complexes, may underrepresent the true prevalence of infection,
particularly in cats.
54
False negative antigen tests in dogs infected with heartworm and placed on macrocyclic
lactone preventives.
Susan Little1*, Jason Drake2, Jeff Gruntmeir1, Hannah Merritt3, Lynn Allen2. 1Center for
Veterinary Health Sciences, Oklahoma State University, Department of Veterinary Pathobiology,
Stillwater, OK, 2Novartis Animal Health, Greensboro, NC, and 3Swaim Serum Company,
Oklahoma City, OK.
Some veterinarians place heartworm infected dogs on macrocyclic lactone preventives and
antibiotics long-term rather than treating them with an approved adulticide. Dogs managed with
these “slow-kill” protocols may experience prolonged inflammation which could result in
formation of immune complexes that block detection of antigen on commercial tests. Pretreatment of serum with heat prior to testing for antigen of D. immitis destroys immune
complexes, freeing antigen to allow detection. To determine if administering macrocyclic
lactones and/or antibiotics to D. immitis-infected dogs could result in development of falsenegative antigen tests, we worked with veterinarians in private practice to collect serum samples
from a total of 15 dogs that had been diagnosed with heartworm by antigen detection, with or
without confirmation by detection of D. immitis microfilariae, placed on macrocyclic lactone
preventives (ivermectin or moxidectin) and antibiotics (doxycycline), and that later tested
negative on an antigen test. Serum samples from all 15 dogs were negative for antigen prior to
heating on commercial assay (DiroCHEK®, Zoetis) by colorimetric detection and
spectrophotometry, but after heat treatment, 8/15 (53.3%) samples converted to positive.
Review of the medical records of each dog indicated that, after the heartworm diagnosis, only
7/15 (46.7%) dogs appeared to receive preventive monthly as prescribed, including 3 dogs that
had detectable antigen after heating the sample and 4 dogs that did not have detectable antigen
after heating. Whole blood was available from 9 dogs; microfilariae were detected in 1 sample.
These data suggest that immune complex formation in dogs being managed with slow-kill
protocols can induce false negative antigen test results, misleading veterinarians and owners
about the efficacy of this approach. Moreover, compliance with preventive administration
appears poor, even after a heartworm diagnosis, and the presence of microfilaremia in at least
one dog has implications for selection for resistance.
74
Novel Cases
55
Treatment of a Mammomonogamus spp. infection in a domestic cat.
Jennifer Ketzis1*, Talia Gattenuo2, Linda Shell3. 1Ross University School of Veterinary Medicine,
Biomedical Sciences, Basseterre, St. Kitts and Nevis, 2Ross University School of Veterinary
Medicine, Basseterre, St. Kitts and Nevis, and 3Ross University School of Veterinary Medicine,
Clinical Sciences, Basseterre, St. Kitts and Nevis.
Treatment of Mammomonogamus ierei, a nematode found on many Caribbean islands, is not
well documented. However, it is not unusual to see Mammomonogamus spp. eggs in fecal
examination of feral and non-feral cats from St. Kitts. Normally, affected cats do not have clinical
signs, although chronic inflammation of the nasopharynx, producing mucoid nasal discharge
and sneezing episodes has been associated with infections according to the literature. A 7month-old, female, domestic short-haired, indoor/outdoor cat was presented to the Veterinary
Teaching Hospital at Ross University School of Veterinary Medicine as part of a student training
spay-neuter program. Due to the presence of diarrhea, feces were analysed using double
centrifugation with Sheather’s sugar flotation solution. Mammomonogamus spp., Ancylostoma
spp., Trichuris spp. and Platynosomum spp. eggs were seen. The feces were not examined for
giardia, although an infection was suspected due to the diarrhea. In addition, the owner reported
that the cat had been sneezing and one bout of sneezing was observed in the veterinary clinic.
Fenbendazole (Panacur oral suspension; Intervet) (50 mg/kg bodyweight) was administered for
5 days. Feces were analysed on the first day of treatment (before treatment), on days 3 and 4 of
treatment and 7 and 15 days after the last treatment. The Mammomonogamus spp. egg count
went from 86 eggs per gram of feces (epg) to 138 epg on day 3 of treatment to 0 epg 7 and 15
days post-treatment. Both the diarrhea and the sneezing resolved. Fenbendazole at 50 mg/kg
bodyweight appeared to be effective in treating the Mammomonogamus spp. infection.
56
Identification of Trichuris serrata in cats on St. Kitts.
Jennifer Ketzis*. Ross University School of Veterinary Medicine, Biomedical Sciences,
Basseterre, St. Kitts and Nevis.
Two species of Trichuris in domestic cats, Trichuris serrata and T. campanula, were originally
described by von Listow in 1879 and 1889, respectively. Species differences include the length
of the adults, shape of the vulva (including a finger-like projection of T. serrata), the length of the
spicule sheath and egg size. There are some discrepancies in the descriptions of the two
species and some debate as to whether or not two species actually exist in domestic cats. Many
measurements for the two species overlap and very few of either species have been described.
Given the low occurrence (1%) of Trichuris in much of the world, opportunities to study Trichuris
are limited. However, on St. Kitts up to 71% of feral cats have Trichuris spp. infections,
providing a unique opportunity to study Trichuris. Trichuris were harvested from the large
intestine of 11 cats euthanized between February 2013 and January 2014. In addition, feces
were examined for Trichuris eggs. The size of the eggs in the feces more closely resembled
those of T. campanula. However, eggs harvested from the females more closely resembled
those of T. serrata. Size difference could be related to egg maturity. The size of adult females
75
and males also crossed the description of both species. However, every female adult examined
had a finger-like projection on the vulva. Therefore, it is believed that the species present on St.
Kitts is T. serrata. Based on this study, the best method for distinguishing the species is the
finger-like projection. Further studies are underway to genetically characterize T. serrata and
obtain samples of Trichuris campanula for comparison.
57
Ocular onchocercosis in a dog.
Gary Conboy1*, Fany Marron1, Paul Hanna1, Erin MacDonald2. 1Atlantic Veterinary College,
Pathology and Microbiology, Charlottetown, PEI, Canada, and 2Summerside Animal Hospital,
Summerside, PEI, Canada.
A 3-yr-old, male, toy fox terrier with a presumptive diagnosis of inflammatory bowel disease was
euthanized after 14 months of intermittent vomiting and diarrhea due to a lack of response to
multiple treatments. The dog was born in New Mexico and had traveled to dog shows in the
USA and Canada. It was brought to Prince Edward Island, Canada 7 months prior to its death.
Transient lack of pupillary response and mild auricular dermatitis were noted just prior to
euthanasia. Necropsy revealed gastrointestinal lesions consistent with inflammatory bowel
disease. In addition, multiple sections of nematodes were detected within vessels (lymphatics or
venules) in the periorbital scleral tissue. The worms were adult females, up to 300 microns in
diameter and had weakly developed body wall musculature mainly confined to the ventral and
dorsal regions and flat-broad lateral chords. The cuticle was 2-layered with the inner layer
having striae and outer cuticle surface marked by ridges. Most sections of female worms
contained 2 loops of uterus and a very small intestine. The uterus contained developing eggs
and microfilaria. Small numbers of microfilaria were detected in the periorbital tissue.
Additionally, sections of skin from the auricular dermatitis showed numerous microfilariae in the
mid to deep dermal regions sometimes adjacent to areas with perivascular to interstitial
infiltrates of plasma cells, lymphocytes and eosinophils. Based on host, travel history, organ
location and morphology on histopathology the nematodes were identified as Onchocerca lupi.
Once thought to be aberrant Onchocerca lienialis infections, O. lupi is now recognized as a
pathogen mainly affecting periocular tissues and has been diagnosed in dogs, wolves, cats and
humans in the western states of the USA and parts of Europe and Asia. Infections have been
associated with severe granulomatous nodular lesions in the periorbital tissues resulting in
severe ocular disease.
58
Trypanosoma theileri associated with a second trimester aborted Holstein fetus.
Dana Pollard1*, Delwyn Keane2, Scott Jones3, Craig Radi2, Robert Droleskey4, Patricia J
Holman1. 1Texas A&M University, Veterinary Pathobiology, College Station, TX, 2University of
Wisconsin, Wisconsin Veterinary Diagnostic Laboratory (WVDL), Madison, WI, 3University of
Wisconsin, Wisconsin Veterinary Diagnostic Laboratory (WVDL), Barron, WI, and 4USDA-ARS,
Southern Plains Agricultural Research Center, Food and Feed Safety Research Unit, College
Station, TX.
A Holstein dairy herd in south-central Wisconsin experienced an increase in the number of open
cows and abortions during May to September 2013. Tissues submitted from an aborted second
trimester fetus revealed an acute necrotizing placentitis with myriad intra-lesional organisms
76
resembling trypanosomes. Liver and stomach content tested negative by real time polymerase
chain reaction for leptospira, ureaplasma, bovine viral diarrhea virus, bovine herpes virus Type
1, Neospora, Toxoplasma gondii, and Trichomonas foetus. Bacterial cultures from liver and
stomach content yielded a few colonies of mixed flora, but no salmonella, campylobacter or
listeria were isolated. Trypanosome 18S rDNA was amplified by polymerase chain reaction from
the placenta, fetal liver and maternal blood, and subsequent sequence analysis confirmed T.
theileri in all three tissues.
59
Spectacular rhabdiasid infections in captive-bred Ball pythons from Wisconsin and
Virginia.
Araceli Lucio-Forster1*, Christoph Mans2, Jennifer Dreyfus3, Jennifer Hausmann2, Drury
Reavill4, Mark Rishniw5, Janice Liotta1, Dwight Bowman1. 1Cornell University, Department of
Microbiology and Immunology, Ithaca, NY, 2University of Wisconsin, Department of Surgical
Sciences, Madison, WI, 3University of Wisconsin, Department of Pathobiological Sciences,
Madison, WI, 4Zoo and Exotic Pathology Service., West Sacramento, CA, and 5Cornell
University, Department of Clinical Sciences, Ithaca, NY.
Rhabdiasid nematodes are common parasites of the respiratory tracts of reptiles and
amphibians, yet no members of the Rhabdiasidae have been previously reported from snakes
within the Pythonidae family. Recently, captive-bred Ball pythons (Python regius) from two
separate locales in the United States were determined to have been clinically affected with
ocular, buccal and subcutaneous nematodiasis caused by a rhabdiasid nematode. The breeding
facilities of the three snakes were unrelated and are furthermore, located in different states
(Wisconsin and Virginia). It is unknown how the snakes acquired their infections; bedding was
not processed for nematode recovery, and no free-living nematodes were available for
examination. Examination of histologic sections from the three cases and whole mounts from
nematodes recovered from one snake, yielded compelling evidence that the same parasite was
involved in all cases. The rhabdiasid infection in these Ball pythons is unusual, not only in its
reptile host, but also in its sites of infection within this host. Morphological and molecular data
strongly support this nematode is likely a new species of Rhabdias.
60
Five cases of horse blindness due to the aberrant migration of Setaria digitata into the
aqueous humor.
SungShik Shin1*, Jihun Shin2, Ah-Jin Ahn2, Kyu-Sung Ahn2, Hak-Sub Jeong2, Choung-Seop
Lee3, Byung-Su Kim4, Guk-Hyun Suh5. 1College of Veterinary Medicine, Chonnam National
University, Parasitology, Gwangju, Republic of Korea, 2College of Veterinary Medicine,
Chonnam National University, Department of Parasitology, Gwangju, Republic of Korea,
3
Surabol Equine Veterinary Clinic, Gyeongju, Republic of Korea, 4YeongGwang Veterinary
Clinic, YeongGwang-eup, Jeollanam-do, Republic of Korea, and 5College of Veterinary
Medicine, Chonnam National University, Department of Clinical Medicine, Gwangju, Republic of
Korea.
Adult worms of the genus Setaria are commonly found free within the abdominal cavity of cattle
in the Far East Asia. Larvae often invade other organs such as the brain and spinal cord of
sheep, goat, horse and cattle causing cerebrospinal injuries and lumbar paralysis. Since we
77
reported the first ocular setariasis of cattle in Korea in 2002, many cases of horse blindness with
motile worms in the aqueous humor of the eye have also been diagnosed. Between 2004 and
2013, five horses were found to have motile worms in the aqueous humor. After the affected
animals were properly restrained under sedation and local anesthesia, a 10ml disposable
syringe with a 16-gauge needle was inserted into the anterior chamber of the affected eye to
remove the parasites which were all identified as Setaria digitata. The pathogenesis and
chemotherapy against the infection is reviewed.
Dog & Cat – Diagnosis
61
ELISA detection of Trichuris serrata coproantigen in cats.
Jinming Geng1*, David Elsemore1, Jennifer Ketzis2. 1IDEXX Laboratories, Inc., Rapid Assay,
Westbrook, ME, and 2Ross University School of Veterinary Medicine, Basseterre, St. Kitts,
United Kingdom.
Antibody reagents designed to detect an excreted/secreted antigen from Trichuris vulpis are the
basis of a coproantigen detection ELISA. The ability of these antibody reagents to detect other
Trichuris species was unknown. To answer this question a collection of 35 fecal samples from
cats was assembled from St. Kitts, where prevalence of T. serrata is high. Infection status with
T. serrata was determined by using double centrifugation with sugar flotation solution. In
addition, adult worms were collected from three cats euthanized for health reasons. Here we
demonstrate the whipworm ELISA detects T. serrata antigen. Whole organism extract of T.
serrata used as an assay sample behaves similarly to T. vulpis extract. Detection of 23 of the 28
whipworm egg positive cat samples shows that T. serrata coproantigen is available for
detection. Two egg negative samples were positive on the whipworm ELISA test. Application of
this test to cats from the southernmost United States may be of interest.
62
ELISA detection of ascarid and hookworm coproantigen in dogs and cats.
David Elsemore*, Jennifer Cote, Rita Hanna, Jinming Geng, Phyllis Tyrrell. IDEXX Laboratories,
Inc., Rapid Assay, Westbrook, ME.
Ascarid and hookworm infections in dogs and cats are classically detected by identification of
the eggs by fecal flotation. New technology may shed light on detection of adult ascarids and
hookworm in the absence of positive egg flotation data. Here we describe the development of
two coproantigen capture ELISAs that detect either ascarid or hookworm infections in dogs and
cats. Specific antibody reagents on both the solid and liquid phase of the ELISA allow detection
of infections in experimentally infected dogs. Lack of cross reaction with other experimental
nematode infections demonstrates assay specificity. Identification of field samples positive for
different species within the ascarid or hookworm groups demonstrates assay breadth of
detection within each group of nematodes. Analysis of a 1000 member canine and feline field
fecal sample population shows application of these assays may increase the frequency of
detection. Ascarid flotation egg positive, ELISA negative field samples were further
characterized by a second flotation and PCR. Evidence of exogenous eggs by coprophagy or
environmental contamination is discussed.
78
63
Evaluation of fecal samples from dogs with naturally acquired gastrointestinal
nematodes for coproantigen of Trichuris vulpis, Toxocara canis, and Anyclostoma
caninum.
Chris Adolph1*, David Elsemore2, Jennifer Cote2, Rita Hanna2, Jinming Geng2, Susan Little1.
1
Center for Veterinary Health Sciences, Oklahoma State University, Department of Veterinary
Pathobiology, Stillwater, OK, and 2IDEXX Laboratories, Inc., Rapid Assay, Westbrook, ME.
Dogs are commonly infected with gastrointestinal nematodes, but fecal flotation may not
accurately identify all infections, particularly when worm burdens are low or only immature
stages are present. To determine the utility of fecal ELISA at identifying these infections, we
evaluated fecal samples from 92 adult dogs in which naturally occurring infections with Trichuris
vulpis (36/92; 39.1%), Toxocara canis (9/92; 9.8%), and Ancylostoma caninum (44/92; 47.8%)
were documented at necropsy and compared results of an ELISA (IDEXX Laboratories, Inc.) to
identification of eggs by passive flotation using sodium nitrate and active centrifugation in sugar
solution. Sensitivity of T. vulpis detection was 63.9% (23/36) by passive flotation, 77.8% (28/36)
by active flotation, and 86.1% (31/36) by fecal ELISA. Sensitivity of T. canis detection was
44.4% (4/9) by passive flotation, 55.6% (5/9) by active flotation, and 77.8% (7/9) by fecal ELISA.
Sensitivity of A. caninum detection was 81.8% (36/44) by passive flotation, 84.1% (37/44) by
active flotation, and 97.7% (43/44) by fecal ELISA. Unexpected positives for T. vulpis and A.
caninum, but not T. canis, were also occasionally found by both fecal flotation and ELISA and
may be due to coprophagy or failure to detect nematodes at necropsy. Taken together, these
data suggest that fecal ELISA for coproantigens is a useful adjunct to microscopic examination
of fecal flotation preparations for accurately identifying nematode infections in canine fecal
samples.
64
Prevalence of internal parasites in shelter cats based on centrifugal fecal flotation.
Byron Blagburn1*, Jamie Butler1, Jane Mount1, Tracey Land1, Joe Hostetler2. 1Auburn
University, Auburn, AL, and 2Bayer HealthCare Animal Health, Shawnee Mission, KS.
Few surveys have been conducted on parasite prevalence in free-roaming or feral cats. Data on
prevalence of parasites of shelter cats can aid in justification for parasite control in pet cats. To
determine the prevalence of common internal parasites, we obtained fecal samples from 4,143
cats in shelters from four regions throughout the United States. Five gram fecal specimens were
shipped in climate controlled containers via overnight courier to the Auburn University College of
Veterinary Medicine. Samples were examined using a sucrose centrifugal flotation procedure.
National prevalence of common intestinal parasites were: Toxocara cati – 19.8%; Ancylostoma
tubaeforme – 7.8%; Uncinaria stenocephala – 0.02%. Combinations of parasites in submitted
samples were: T. cati + A. tubaeforme – 3.0%; either T. cati or A. tubaeforme – 24.7%. Other
parasites were also recovered and their prevalence will be presented. Forty (40)% of cats
harbored at least one parasite. These results indicate that parasites remain prevalent in freeroaming or feral cats throughout the United States. Our results support recommendation of
broad spectrum internal parasite control for cats.
79
65
Prevalence of internal parasites in shelter dogs based on centrifugal fecal flotation.
Byron Blagburn1*, Jamie Butler1, Jane Mount1, Tracey Land1, Joe Hostetler2. 1Auburn
University, Auburn, AL, and 2Bayer HealthCare Animal Health, Shawnee Mission, KS.
Control of canine internal parasites remains a principal concern and a substantial effort for
practicing veterinarians. To determine the prevalence of common internal parasites, we
obtained fecal samples from 6,418 dogs in shelters from four regions throughout the United
States. Five gram fecal specimens were shipped in climate controlled containers via overnight
courier to the Auburn University College of Veterinary Medicine. Samples were examined using
a sucrose centrifugal flotation procedure. National prevalence of common intestinal parasites
were: Toxocara canis – 12.5%; Toxascaris leonina – 0.36%; Ancylostoma caninum – 29.8%;
Uncinaria stenocephala – 2.51%; Trichuris vulpis – 18.7%. Combinations of parasites in
submitted samples were: T. canis + A. caninum – 6.3%; T. canis + T. vulpis – 3.2%; A. caninum
+ T. vulpis – 10.6%; T. canis + A. caninum + T. vulpis – 2%. Forty-three (43)% of dogs harbored
at least one of these major internal nematode parasites. Other parasites were also recovered
and their prevalence will be presented. These results indicate that parasites remain prevalent in
dogs throughout the United States. In fact, the prevalence of most major internal parasites of
shelter dogs exceeds those obtained in our 1996 survey. Shelter dogs are beyond the reach of
practicing veterinarians and remain a source of infection for pets. Our results support continued
recommendation of broad spectrum internal parasite control.
66
Assessing the speed of kill of Ancylostoma caninum by Advantage Multi® for Dogs using
endoscopic methods.
Alice Lee1*, Joseph Hostetler2, Dwight Bowman1. 1Cornell University, College of Veterinary
Medicine, Microbiology & Immunology, Ithaca, NY, and 2Bayer HealthCare LLC, Veterinary
Services, Shawnee Mission, KS.
Compared to the oral route, topically administered drugs require more time to reach therapeutic
concentrations in systemic circulation and are therefore thought to be slower to take effect. The
moxidectin component of Advantage Multi® for Dogs (Bayer HealthCare LLC, Shawnee Mission,
KS) reaches maximal plasma concentration 9.3 days after topical application in dogs.
Accordingly, in past trials that tested the efficacy of this medication against intestinal helminths,
necropsy was scheduled 10 days post-treatment rather than the 7 days that is typical for oral
anthelmintics. In the current study, the speed of clearance of Ancylostoma caninum from the
small intestine after treatment with Advantage Multi® for Dogs was assessed endoscopically.
Four adult beagles were experimentally infected with A. caninum larvae in two cohorts of two
dogs each. Advantage Multi® for Dogs was given on day 13 (cohort 1) or day 16 (cohort 2)
post-inoculation. A combination of conventional and capsule endoscopy was performed before
and after product administration to check for the presence of hookworms and any associated
lesions. Prior to treatment, hookworms and the mucosal lacerations they made during feeding –
at times extensive – were observed in all dogs. As early as 24 hours post-treatment, hookworms
were no longer found in the small intestine. By 48 hours, the majority of the hookworm-induced
mucosal bleeding was no longer evident. Results from the present study suggest that
Advantage Multi® for Dogs achieves its anthelmintic effect much faster than previously believed
and is able to rapidly clear hookworms from the small intestine of dogs.
80
Molecular & Immunology
67
Not just GluCls (Glutamate-gated Chloride channels): abamectin has potent effects on
nicotinic acetylcholine receptors of nematode parasites.
Richard Martin*, Melanie Abongwa, Samuel Buxton, Alan Robertson. Iowa State University,
Biomedical Sciences, Ames, IA.
Abamectin and derquantel (StartectR) is a combination of drugs showing synergistic anthelmintic
effects against nematode parasites. We have observed in contraction and electrophysiological
experiments that: 1) derquantel is a potent selective antagonist of muscle nicotinic receptors of
Ascaris suum; and 2) abamectin is an inhibitor of the same nicotinic receptors (Puttachary et al.,
2013). To explore the inhibitory effects of abamectin further, we have cloned muscle nicotinic
receptors of O. dentatum, expressed them in Xenopus oocytes and tested, under voltageclamp, the effects of abamectin on a group of nicotinic receptors activated by pyrantel (Buxton
et al., 2014). We observed that expression of Ode-UNC-63:Ode-UNC-29:Ode-UNC-38
produced receptors that were activated by pyrantel in a concentration-dependent manner above
0.1 µM. These receptors were antagonized by 0.03 µM abamectin in a non-competitive manner
(reduced the Rmax with little change in EC50). The non-competitive antagonism was greater with
0.1 µM abamectin. Interestingly, when we increased the concentration of abamectin to 0.3 µM,
we found that the antagonism decreased and was less than with 0.1 µM abamectin. The biphasic effects of abamectin suggest two allosteric sites of action: one causing antagonism, one
causing potentiation. The site of action of abamectin on the receptors may be in the outer lipid
phase of the membrane (Martin and Kusel, 1992) between M1 transmembrane domain of one
receptor subunit and the M3 domain of the adjacent receptor subunit (Hibbs et al., 2011). Since
the receptor channel is composed of heterogeneous subunits, binding between one pair of
subunits may stabilize closing (negative allosteric modulator) while binding between another
pair of subunits may stabilize opening (positive allosteric modulator). The observations confirm
antagonistic effects of abamectin on nematode nicotinic receptors in addition to GluCl effects,
and relate to use of combination therapies and development of molecular tests for anthelmintic
resistance.
68
Electrophysiological effects of cholinergic anthelmintics in two species of filarial
parasite.
Saurabh Verma1*, Alan Robertson1, Adrian Wolstenholme2, Richard Martin1. 1Iowa State
University, Department of Biomedical Sciences, College of Veterinary Medicine, Ames, IA, and
2
The University of Georgia, Department of Infectious Disease, College of Veterinary Medicine,
Athens, GA.
Filariasis is a serious public and animal health problem. Examples include human lymphatic
filarial nematodes (Wuchereria bancrofti, Brugia malayi and Brugia timori) and canine
heartworm (Dirofilaria immitis). Treatment of filarial infection is directed mainly at prophylaxis
because there are no effective treatments for adult worms. Loss of efficacy and emerging signs
of resistance with widespread use of macrocyclic lactones has produced a strong selective
pressure for resistance in the filarial nematodes. Novel resistance busting cholinergic
81
anthelmintics like tribendimidine and derquantel have increased interest in cholinergic drugs and
their combinations. We have tested and demonstrated that B. malayi can be utilized for single
channel and whole cell patch clamp recordings, enabling us to characterize its nAChR’s. We
have observed responses to the acetylcholine and the cholinergic anthelmintics levamisole,
nicotine, and bephenium. Our results show that the body muscle nAChR receptor of B. malayi
has similarities to those of other nematode parasites. Acetylcholine is a major excitatory
neurotransmitter, and generates the largest response. Acetylcholine induced inward currents
were most potent with dose dependent EC50 of 2.36 µM, whereas levamisole, nicotine and
bephenium EC50 were 3.35, 30.5 and 4.24 µM respectively. Here we also describe whole cell
patch-clamp recordings made from heartworm muscles. We have now identified and studied
nicotinic cholinergic receptors on the muscles of both adult Brugia and Dirofilaria. It will be
interesting to see if these receptors can be exploited for the future development of a filarial
adulticide.
69
Characterization of a homomeric nicotinic acetylcholine receptor, Ascaris suum ACR-16,
in Xenopus laevis oocytes.
Alan Robertson1*, Melanie Abongwa1, Samuel Buxton1, Elise Courtot2, Claude Charvet2, Cedric
Neveu2, Richard Martin1. 1Iowa State University, Biomedical Sciences, Ames, IA, and 2INRA,
Infectiologie Animale et Santé Publique, Nouzilly, France.
Control of nematode parasite infections in both veterinary and human medicine has relied on
the use of anthelminthic drugs, many of which act on nicotinic acetylcholine receptors.
Unfortunately, the widespread use of anthelmintics has given rise to the development of
resistance in parasitic worms of veterinary importance, generating concern that this may extend
to humans. There is an urgent need for the development of new resistance busting drugs which
requires the characterization and validation of novel anthelmintic drug targets. Using molecular
cloning, RT-PCR and the two-electrode voltage clamp electrophysiological technique, we have
cloned and expressed the homomeric nicotinic acetylcholine receptor subunit gene, acr-16, from
Ascaris suum in Xenopus laevis oocytes. Sequence comparison with vertebrate subunits
reveals that acr-16 is most closely related to the vertebrate α-7 subunit. RT-PCR experiments
on A. suum tissue samples demonstrated acr-16 mRNA present in somatic muscle strips,
pharynx, ovijector, head and gut. We found that RIC-3 is required for the functional expression
of ACR-16 in Xenopus oocytes. When ric-3 cRNAs from A. suum, X. laevis or Haemonchus
contortus were co-injected with A. suum acr-16 cRNA, currents were biggest in oocytes
expressing A. suum RIC-3, suggesting species-specific RIC-3 effects. In electrophysiological
experiments we observed that homomeric ACR-16 receptors did not respond to any of the
cholinomimetic drugs available. Nicotine, acetylcholine and cytisine all produced robust inward
currents. Interestingly unlike the vertebrate α-7 receptor, channels formed from ACR-16 did not
respond to genistein and other positive allosteric modulators. We hypothesize that nAChRs
formed from ACR-16 are attractive potential target sites for new anthelmintic compounds.
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70
Apyrases in Ostertagia ostertagi; controlling the inflammatory response and longevity of
L4 larvae in the gastric glands.
Dante Zarlenga*, Wenbin Tuo. USDA, ARS, Animal Parasitic Diseases Lab, Beltsville, MD.
Apyrases comprise a ubiquitous class of glycosylated nucleotidases that hydrolyze extracellular
ATP and ADP to orthophosphate and AMP. Activities on other nucleotides are also common. A
class of calcium-activated, salivary apyrases known to counteract blood-clotting, has been
identified in several hematophagous arthropods. Recently we identified, cloned and expressed a
similar Calcium-dependent protein in Ostertagia ostertagi (rOoapy1-c64) and demonstrated its
ability to dephosphorylate ATP, ADP, UTP and UDP which coincide with the range of
nucleotides involved in signaling across the P2 receptor complex. Immunohistochemical and
Western blot analyses show that the native Ooapy-1 is produced or collects within the glandular
bulb of the esophagus at the esophageal/intestinal junction. It is secreted from the parasite and
is developmentally-regulated appearing only in the L4. Based on the exclusive secretion of the
native Ooapy1 by L4 and the range of substrate activity, we hypothesize that the native protein
is used by the parasite to control the levels of host extracellular nucleotides secreted by locallydamaged and/or infiltrating tissues. In so doing, the L4 could modulate host inflammatory
responses required for early parasite control and expulsion. To better characterize the active
site and provide a target for drug or immunological intervention, we identified a region of the
Ostertagia apyrase that exhibited pseudo-conservation with other apyrases. This 11 amino acid
site was modified through site-directed mutagenesis by inserting a sequence derived from a
similar protein found in Ascaris suum. Results showed that modification of the Ostertagia protein
completely eliminated enzymatic activities on ATP and ADP while modification of the Ascaris
protein with the Ostertagia sequence, which exhibits activity only on GDP and UDP remained
unaffected. Data suggest that attenuating the in vitro activity of native Ooapy1 may positively
influence the host’s ability to control growth and development of the L4 stage.
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Characterization of Ostertagia ostertagi macrophage migration inhibitory factor.
Wenbin Tuo1*, Guanggang Qu1, Raymond Fetterer1, Lin Leng2, Dante Zarlenga1, Richard
Bucala2. 1USDA/ARS, Animal Parasitic Diseases Lab, Beltsville, MD, and 2Yale University
School of Medicine, Department of Medicine, New Haven, CT.
Macrophage migration inhibitory factor (MIF) of Ostertagia ostertagi was characterized in the
present study. There are 3 Oos-MIFs, each encoded by a distinct transcript: Oos-MIF-1.1, OosMIF-1.2, and Oos-MIF-2. Oos-MIF-1.1 and Oos-MIF-1.2 amino acid sequences are similar
(93%) and thus collectively referred to as Oos-MIF-1 when characterized with immunoassays.
Oos-MIF-2 has higher sequence similarity with the Caenorhabditis elegans MIF2. Recombinant
Oos-MIF-1.1 (rOos-MIF-1.1) produced in Escherichia coli is active as a tautomerase. rOos-MIF1.1s carrying a mutation (rOos-MIF-1.1P1G) or duplication of proline-1 (rOos-MIF-1.1P1+P)
resulted in reduced oligomerization and complete loss in tautomerase activity. The tautomerase
activity of rOos-MIF-1.1 was inhibited by a tautomerase inhibitor, ISO-1, and rOos-MIF-1.1specific antibody. Oos-MIF-1 was detected in adult worm excretory/secretory product and all
developmental stages with the highest level in the adult stage. Oos-MIF-1 was localized to the
hypodermis/muscle, reproductive tract and intestine, but not to the cuticle. rOos-MIF-1.1, but not
rOos-MIF-1.1P1G, was able to compete for human CD74, a MIF cell surface receptor, with
human MIF. Furthermore, we also show that rOos-MIF-1.1 inhibited migration of bovine
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macrophages and restored glucocorticoid-suppressed, lipopolysaccharide (LPS)-induced tumor
necrosis factor (TNF)-a and interleukin (IL)-8 in human and/or bovine macrophages. This
functional, secretory, parasite-derived immunomodulator may play a role in parasite-host
interaction and warrants investigation as a vaccine candidate against O. ostertagi infections in
cattle.
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Host immune responses to Eimeria adenoeides infection in turkey poults.
Thilakar Rathinam1*, Ujvala Deepthi Gadde2, Gisela F Erf1, H. David Chapman1. 1University of
Arkansas, Poultry Science, Fayetteville, AR, and 2University of Arkansas, Department of Poultry
Science, Fayetteville, AR.
Coccidiosis is a common enteric disease of turkeys caused by protozoan parasites belonging to
the genus Eimeria. E. adenoeides is one of the most pathogenic and commonly recognized
turkey Eimeria. Infection with Eimeria is known to induce a long lasting protective immunity in
chickens, but nothing is known regarding the acquisition of immunity to Eimeria in turkeys. The
experiments reported here were aimed at investigating the biological and cellular immune
response to E. adenoeides in turkey poults under different conditions of exposure. In experiment
1, 20 day old poults were infected with 12.5 x 103 oocysts and shown to develop immunity when
challenged with a high dose of oocysts at 34 days of age. Changes in peripheral blood
leukocytes following the primary exposure were investigated, and an increase in the number of
total lymphocytes, monocytes, and subsets of lymphocytes (CD4+ and CD8+) was shown in
infected poults.
In experiment 2, local immune activities occurring in the ceca in response to infection were
investigated. An increase in leukocyte infiltration, percent area occupied by CD4+ and CD8+
lymphocytes, and relative expression of cytokines (CXCLi2, IL1β, IFNγ, IL10, IL13) was
observed in the ceca following infection.
In experiment 3, we attempted to model the field situation by investigating the acquisition of
immunity under conditions poults could encounter following placement upon built-up litter.
Results indicated that they had acquired partial protection by day 12 and 18 following challenge.
The leukocyte infiltration score, percent area occupied by CD4+ and CD8+ lymphocytes, and
expression of the cytokines (CXCLi2, IL1β, IFNγ, IL2, IL10, IL12b, IL13, IL18) in the ceca
significantly increased following infection. The changes in the blood and ceca of E. adenoeides
infected poults in terms of infiltration of leukocytes and the relative expression of cytokines
characterize the development of innate and adaptive immune activities.
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Small Ruminants & Education
73
Challenges in assessing anthelmintic resistance and the use of FAMACHA in sheep on
St. Kitts.
Jennifer Ketzis1*, James Robinson2. 1Ross University School of Veterinary Medicine, Biomedical
Sciences, Basseterre, St. Kitts and Nevis, and 2Ross University School of Veterinary Medicine,
Clinical Sciences, Basseterre, St. Kitts and Nevis.
Haemonchus contortus, based on egg cultures, has been shown to be the predominate
trichostrongylid species in sheep on St. Kitts. Given the limited availability of anthelmintic
classes and products on St. Kitts and common use of ivermectin and fenbendazole, anthelmintic
resistance is likely to exist. Several attempts have been made to assess resistance on farms
with limited success. Complications have included: limited number of animals per farm, lack of
identification of the animals, limited ability to weigh animals and, most importantly, low fecal egg
counts. In the largest study conducted, sheep were randomized by FAMACHA score (1, 2, 3
and 4) to treatment groups. Sheep with a score of 5 were treated with the preferred anthelmintic
for the farm and not included in the study. Of 48 sheep, only 15 had fecal egg counts >250, the
preferred limit for conducting a fecal egg count (FEC) reduction test. There appeared to be little
to no relationship between FEC and FAMACHA score, suggesting other causes of anemia and
complicating the ability to randomize animals to treatment groups based on FAMACHA score.
Other potential causes of anemia such as low iron diets, poor nutrition and infectious diseases
such as Babesia spp. are being investigated. Despite these challenges, combining data from
farms indicates emerging resistance to ivermectin, fenbendazole and albendazole. The lowest
efficacy seen was with ivermectin (<20% efficacy) in sheep from intensively managed farms and
fenced pastures. However, ivermectin was 100% efficacious in a group of sheep from a farm
with low inputs and extensive grazing.
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Opportunities for molecular diagnosis of infection and anthelmintic resistance in
parasitic nematodes.
Roger Prichard*. McGill University, Institute of Parasitology, Montreal, QC, Canada.
Anthelmintic resistance has become a serious problem for the control of parasitic nematodes in
livestock. There is also recent evidence that resistance is developing in nematode parasites of
companion animals, such as Dirofilaria immitis and in the human parasites, Onchocerca
volvulus and Trichuris trichiura. Conventional methods for monitoring for anthelmintic
resistance, such as faecal egg count reduction tests are insensitive, expensive and slow and
often only used in research settings. It has been known for some time that mutations in the βtubulin gene can confer benzimidazole resistance. Genetic analyses have been applied to
sheep farms to detect benzimidazole resistance. Recently, we have discovered mutations in the
dyf7 gene confer macrocyclic lactone resistance in Haemonchus contortus and Caenorhabditis
elegans. While the deletion of a 63 bp indel in the acr8 gene causes truncation of the Acr8
subunit of the levamisole receptor in H. contortus and levamisole resistance. This new
information allows us to apply genetic tests to detect resistance to all of the common broad
spectrum anthelmintics used to control parasites in livestock. The availability of new genetic
85
methods for monitoring anthelmintic resistance offers considerable advantages over
conventional in vivo and in vitro methods for assessing anthelmintic resistance. It is also
possible to semi-quantitatively detect the presence of eggs of different species in fecal samples.
The objective of this work is to have protocols for assessing levels of parasite infection and drug
resistance/susceptibility status to all of the broad spectrum anthelmintics in fecal samples from
sheep and cattle.
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Competitive effects between genetically diverse Haemonchus contortus strains during
experimental co-infection.
John Gilleard1*, Neil Sargison2, Elizabeth Redman1, Eric Hoberg3, David Bartley4. 1University of
Calgary, Faculty of Veterinary Medicine, Calgary, AB, Canada, 2University of Edinburgh, Faculty
of Veterinary Medicine, Edinburgh, United Kingdom, 3USDA, Agricultural Research Service,
Animal Parasitic Disease Laboratory, Beltsville, MD, and 4Moredun research Institute,
Edinburgh, Midlothian, United Kingdom.
Relative to the knowledge available for other pathogen groups, there is remarkably little
information on genetic and phenotypic variation between experimental strains and field isolates
of parasitic nematodes. This is an increasingly important issue for trichostrongylid nematode
parasites of livestock due to the need for detailed comparative analysis of isolates and strains
used in anthelmintic resistance, vaccine and other research. There are also increasing
expectations for the pharmaceutical industry to use well characterized strains and field isolates
for clinical studies.
We are studying inter-strain variation in Haemonchus contortus and will present some work on
three genetically and phenotypically diverse Haemonchus contortus strains; MHco3(ISE),
MHco4(WRS) and MHco10(CAVR). We have developed panels of microsatellite markers that
can be used to distinguish individual worms of each of these strains based on multilocus
genotypes. We have undertaken a number of experimental co-infections and used these marker
panels to determine the parentage of individual F1 progeny. We have found that in experimental
co-infections with using equal numbers of infective L3 from each parental strain, one of the two
parental strains produces a disproportionately high number of F1 progeny with respect to those
of the other strain or inter-strain hybrids. In the case of co-infections with MHco3(ISE) and
MHco10(CAVR), the MHco3(ISE) strain predominates and in the case of co-infections with
MHco4(WRS) and MHco10(CAVR), the MHco4(WRS) strain predominates. Our results suggest
that, in addition to the basic phenotypic differences between these strains, there are competitive
effects that occur between strains during co-infections. This has important implications for
parasite population dynamics and response of parasite populations to control measures applied
in the field.
76
Worm burdens of Haemonchus contortus in alpacas and sheep following experimental
infection.
Anne Zajac1*, Sarah Casey1, Stephan Wildeus2. 1Virginia Tech, Biomedical Sciences and
Pathobiology, Blacksburg, VA, and 2Virginia State University, Agricultural Research Station,
Petersburg, VA.
86
Haemonchus contortus is the most important gastrointestinal parasite infecting small ruminants.
Heavy infections may cause anemia and death. The third compartment of the stomach of
alpacas (C3) can also be infected with consequent development of clinical signs. We saw
reduced fecal egg counts (FEC) in alpacas compared to sheep following natural and
experimental Haemonchus infection. In this study we examined whether differences in FEC
reflect a significant difference between hosts in worm burdens. All alpacas and sheep used in
the study were dewormed prior to beginning the study. Infective H. contortus larvae were
administered orally to alpacas as either a bolus infection (20,000 larvae as a single dose, n=8)
or a trickle infection (4,000 larvae/day for 5 days, n=8). Similar bolus and trickle infections were
administered to rams (n=6 per group). Fecal egg counts (FEC) were determined every 5 days
from day 14 to day 49 after infection. All animals were euthanized 49 days after final H.
contortus infection and aliquots of the abomasal or C3 contents of each animal were harvested
for determination of total worm burden. Stomachs were also incubated overnight in saline to
facilitate recovery of larvae and aliquots of the rinsings were collected. Also, the pH of the
rumen and abomasum were measured. Sheep FEC were higher than those of alpacas. Mean
total worm burdens were significantly lower (p<0.0001) in alpacas (bolus=708, trickle=541) than
sheep (bolus=4,895, trickle=3,142). All parasites seen were adults. The mean pH measured by
handheld probe was significantly higher in alpaca C3 (6.2-6.5) than in the abomasum of sheep
(3.7-4.4). Our results provide further support for the hypothesis that alpacas are less susceptible
than sheep to H. contortus infection, but further experimentation is needed to determine the role
of gastrointestinal physiology, immunity and other factors in explaining the differences in host
susceptibility.
77
Development of OSCEs for assessment of parasitologic skills.
Karen Snowden*, Rosina Claudia Krecek, Tom Craig. Texas A&M University, College of
Veterinary Medicine and Biomedical Sciences, Veterinary Pathobiology, College Station, TX.
Based on current AVMA Council on Education standards for accreditation at colleges/schools of
veterinary medicine (CVMs/SVMs), there is an increasing emphasis in documentation of student
skills and competencies beyond the administration of traditional written examinations. The
Objective Structured Clinical Examination (OSCE) is a method of assessment of clinical or
technical skills that has been used extensively in medical education. OSCEs have been adopted
as assessment tools to meet changing curricular needs in an expanding number of CVMs/SVMs
in North America. While multiple recent educational publications describe a selection of OSCE
stations, reviews of the literature and informal communications with parasitology instructors
revealed no OSCE assessments for parasitologic technical skills. Therefore, we developed two
OSCE assessments that were implemented in the laboratory portion of the veterinary
parasitology course in the professional curriculum at Texas A&M University CVM in fall, 2013.
One OSCE assessed microscopy skills used during a fecal flotation assay including items such
as eyepiece, lighting, iris diaphragm and condenser adjustment. The second OSCE assessed
the student's ability to correctly set up a McMaster's quantitative fecal egg count test. These
OSCEs were included in lab exercises where part of the grade was assigned based on correct
identification/quantitation of parasites in fecal samples, and part of the grade was based on a
scoring rubric that was completed by a reviewer who visually assessed student technical
performances while they completed the OSCE tasks. Students were receptive to the concept of
assessment of technical skills in addition to correct microscopic identification of parasite
reproductive products. Conducting the OSCEs as part of laboratory exercises required extra
87
preparatory efforts and increased manpower to visually assess students at the OSCE stations.
Overall, instructors agreed that the use of OSCEs was a valuable assessment tool to evaluate
technical skills needed to complete parasitologic diagnostic tests correctly.
78
Advancing veterinary parasitology education and knowledge through online discussions
- the VIN experience.
Craig Datz*. Royal Canin USA, University of Missouri, College of Veterinary Medicine (adjunct),
Columbia, MO.
Veterinarians in private practice have several options when seeking information about
parasitology. They may use websites such as the CAPC, CDC, or AHS, or anything that comes
up with an online search. Veterinary schools and reference labs usually have parasitologists
available for consults, and pharmaceutical companies can provide practitioners with valuable
information, especially about treatment. An increasingly popular way to ask questions, discuss
cases and topics, access journal articles and Proceedings, and obtain continuing education is
through the Veterinary Information Network (VIN). This is a subscriber-based online platform
which started around 1990 as a community within AOL and in 2000 moved to the Web. The paid
membership has grown from about 6,000 in 2000 to 38,000 today. While VIN has many features
including a comprehensive search function, journal index, Proceedings from veterinary
conferences, rounds, a news service, CE courses, drug formularies, etc. the most-used section
is the Message Boards. There are over 50 folders in the Boards section in which veterinarians
can post questions, introduce discussion topics, ask for ideas and opinions, request information,
and collaborate. Each folder is monitored by Consultants (specialists and experts in the topic
who answer most of the questions), Associate Editors (veterinarians who have an interest in the
field), and Representatives (who track discussions to make sure questions are answered and
follow-ups are completed). The Parasitology folder is currently monitored by Norman Ronald,
MS, PhD, a retired parasitologist in Texas, and Craig Datz, DVM, formerly on the faculty at the
University of Missouri where he taught clinical parasitology to veterinary students. During this
presentation, Dr. Datz will highlight examples of popular topics, frequently asked questions,
controversies, unusual images, educational sessions, and how VIN can be used to advance
knowledge and understanding of parasitology among veterinary practitioners.
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Defining core competencies in veterinary parasitology.
Karen Snowden*. Texas A&M University, College of Veterinary Medicine and Biomedical
Sciences, Veterinary Pathobiology, College Station, TX.
In recent years there has been a trend towards competency based training in medical schools
and in allied health professions. Veterinary medical education in North America is following that
trend. As part of the AVMA accreditation process for college/schools of veterinary medicine
(CVM/SVMs), the AVMA Council on Education (AVMA COE) has broadly defined nine areas of
competency for graduating veterinarians that must be documented at the individual student
level. Therefore, AVMA accredited CVM/SVMs are developing a variety of detailed tracking
systems and skills/knowledge logs or checklists to meet AVMA COE requirements. The AAVP
has been playing an important role in defining core competencies in the subject of veterinary
parasitology. Through the AAVP/Companion Animal Parasite Council sponsored Educator
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Symposia (2011 and 2013), a large group of educators/instructors from 27 CVM/SVMs have
drafted a detailed list of competencies based on the AVMA COE accreditation requirements.
Following organizational protocols, these guidelines will be approved by the AAVP and
published in a peer-reviewed journal. The guidelines can be used by parasitology instructors in
several ways including: 1) defining parasitology competencies in the overall professional
curricula in CVM/SVMs, 2) structuring the course content of professional courses in veterinary
parasitology, and 3) integrating parasitologic topics, knowledge and skills into other courses in
professional curricula. This proactive approach by the AAVP to define core competency
standards in veterinary parasitology establishes a benchmark for other veterinary specialty
groups. The parasitology core competency guidelines should serve as a useful tool for
maintaining the importance and relevance of the subject in veterinary medical education.
Wildlife & Scynexis
80
Epidemiology of Haemonchus contortus in a zoological park giraffe enclosure in Florida.
Sue Howell1*, Tessa Sghiatti1, Katherine Hogan1, Melissa Miller1, Bob Storey1, Adrienne Atkins2,
Nick Kapustin2, Ray Kaplan1. 1University of Georgia, Department of Infectious Diseases,
Athens, GA, and 2Jacksonville Zoo and Gardens, Jacksonville, FL.
Escalating problems in managing infections with Haemonchus contortus in giraffes (Giraffa
camelopardalis) at a zoological park in Florida culminated in the death of a young giraffe due to
haemonchosis in August 2012. In order to improve the ability to manage the problem, an 18month study was initiated to investigate the epidemiology of H. contortus transmission in the
giraffe enclosure. The goals of this study were to determine the relative worm burdens of the
giraffe population, the relative levels of pasture contamination with L3 across different areas of
the enclosure, and how both of these parameters change seasonally over the course of the
year. Rainfall and temperature data were also examined in order to look for trends that may help
explain the seasonal changes in parasitological data that were observed. Fecal samples from
each animal, grass cuttings from 7 different zones within the giraffe enclosure, and soil samples
from 3 zones were collected monthly and shipped to the University of Georgia for analysis.
Fecal egg counts were performed using the modified McMasters method, and grass samples
were washed to recover larvae, which were then identified and counted. Though analysis is still
pending, results suggest that there is a strong association between mean EPG and numbers of
H. contortus L3 recovered from pasture. There also appear to be seasonal trends, with L3
numbers peaking in the winter, decreasing through the spring, spiking briefly in early summer,
and increasing again in mid-fall. Location also seemed to be important, with one zone
consistently having higher than average levels of pasture L3 and another having consistently
lower levels. These data are being used to improve the parasite control program for the giraffe
herd in this zoo. By integrating FEC surveillance, targeted-selective-treatments, pasture
hygiene, and other novel approaches, sustainable integrated control of H contortus is
achievable.
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81
Efficacy of Duddingtonia flagrans on gastrointestinal nematode larvae in feces of exotic
artiodactylids.
James Miller1*, Jenna Terry1, Macy Trosclair1, Deidre Fontenot2, Javier Garza1, Vicky Kelly1.
1
School of Veterinary Medicine Louisiana State University, Pathobiological Sciences, Baton
Rouge, LA, and 2Walt Disney's Parks and Resorts Disney's Animal Kingdom, Department of
Animal Health, E. Bay Lake, FL.
Gastrointestinal nematodes (GIN) are parasites of major concern for domestic and exotic
artiodactylids. Zoological facilities have used anthelmintics as their primary control method, and
challenges in accurate dosing and administration contributed to the development of resistant
GIN populations in those settings. The historic dependency on anthelmintics for control is no
longer a reliable option. Alternatives are urgently needed and one such alternative is the use of
the nematophagous fungus, Duddingtonia flagrans. This study evaluated the efficacy of D.
flagrans (chlamydospores (CS) administered as a powdered mixture incorporated into feed) in
reducing GIN infective larvae (L3) in feces at a dose of 30,000 CS per kg/BW. The CS/feed
mixture was fed daily over an 8 week period to giraffe, antelope and gerenuk. Fecal samples
were collected on a weekly basis for fecal egg count and then cultured to recover and count GIN
L3. Results indicated that D. flagrans effectively reduced GIN L3 in the feces during the period
of feeding by over 80% in all 3 exotic artiodactylid species. The results from this study
demonstrated that the use of D. flagrans in exotic artiodactylids infected with GIN could be a
long term prophylactic tool to reduce forage infectivity. Used in conjunction with other control
methods, D. flagrans could be part of an integrated GIN parasite control program for exotic
artiodactylids in zoological facilities.
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Influence factors on infection with Taenia hydatigena larvae of wild ruminants in the
National Park Losiny Ostrov (Elk Island), Moscow, Russia.
Nina Samoylovskaya*, Aleksandr Uspenskiy, Vladimir Gorohov, Karine Kurochkina, Alexandr
Moskvin. All-Russian K.I. Skryabin Scientific Research Institute of Helminthology, The Russian
Academy of Sciences, Moscow, UZAO, Russian Federation.
Carrying out long-term observations for the state of the ecosystems in general and their single
components in natural areas of preferential protection is the main direction of research. Besides,
routine observations can help to determine changes in ecosystems at an early stage, when their
negative consequences can still be prevented. National Park Losiny Ostrov (Elk Island), with its
area of 12881 ha, is situated in the north-east of Moscow. Its territory is divided into forestparks, namely Mytishchinsky, Losino-pogonny, Shchelkovsky, Alekseevsky, Losinoostrovsky
and Yauzsky forest-parks. The total number of elk is from 45 to 50 specimens. The Losiny
Ostrov belongs to the Russian natural areas of preferential protection. High epizootic and
epidemiological role of dogs in disseminating Сysticercus tenuicollis in wild ruminants (elk and
spotted deer), have been observed in Losiny Ostrov (Elk Island) National Park since the
parasitic research staff of the Institute carried out research in this territory in the period of 2006
and the beginning of 2014. They also found that extensivity of Taenia hydatigena larvae from
the moose fell from 60% (2006-2010) to 27.6% (2011-2014), and the average intensity of
infestations amounted to 4.5 (2006-2010) and 1.7 (2011-2014) instances of the instars of
bubbles accordingly. The sika deer is from 30% (2006-2010) to 21.4% (2011-2014) and the
average intensity of infestations were 10 (2006-2010) and 4 (2011-2014) instances of the
90
instars of bubbles accordingly. Size of instars of bubbles is not much varied as the elk and
reindeers. They reached an average of 5-8 cm. During the years of monitoring of the parasitic
animals of the park only in one case were there 25 specimens of Taenia hydatigena larvae from
the corpse of a spotted deer (female 2.5-3 years), in 2008.
83
Rickettsial and ehrlichial infection in small mammals from northeast Brazil.
Solange M. Gennari*, Marcos G. Lopes, Julia T.R. Lima, Gislene F.S.R. Fournier, Igor C.L.
Acosta, Diego Ramirez, Arlei Macili, Marcelo B. Labruna. Faculty of Veterinary Medicine,
University of São Paulo, Department of Preventive Veterinary Medicine and Animal Health, São
Paulo, SP, Brazil.
The aim of this study was to analyze the serological and molecular occurrence of Rickettsia spp.
and Ehrlichia spp. in small mammals and the ticks present on these animals from two
environmental conservation units in the city of Natal, Rio Grande do Norte State, Brazil. The
collection period was October 2012 and August 2013. Sera were tested against antigens of
Rickettsia amblyommi, R. rhipicephali, R. parkeri, R. rickettsii, R. felis and R. belllii by
immunofluorescence antibody assay (IFA). Tissue samples (spleen and lungs) and ticks were
processed for molecular detection of organisms of the genus Rickettsia and Ehrlichia. Twenty
seven marsupials (23 Didelphis albiventris and 4 Monodelphis domestica) and 4 rodents (2
Necromys lasiurus, 1 Thrichomys apereoides, 1 Rattus rattus) were captured and antibodies
were detected in 6 D. albiventris and in 1 R. rattus (titer >32) to at least one of the tested
rickettsiae, with final titers four times greater for R. amblyommii, R. bellii or R. parkeri compared
with other rickettsia species tested. Two tick species (Amblyomma auricularium and Ixodes
loricatus) were collected from marsupials, and 1 A. auricularium specimen was infected by R.
amblyommii by PCR and DNA sequencing (rickettsial gltA gene). One D. albiventris spleen
yielded PCR amplicons for 2 ehrlichial genes (16S rRNA and dsb). DNA sequencing of the 16S
rRNA amplicon generated a sequence that was closest (99% identity) to several uncultured
Ehrlichia spp. from several parts of the world. The dsb sequence was closest (80-81%) to E.
chaffeensis and E. mineirensis. This is the first detection of Rickettsia spp. and Ehrlichia spp. in
animals from the State of Rio Grande do Norte, Brazil.
84
PK/PD approaches to anti-parasitic drug discovery: are we there yet with filarial
parasites?
Bakela Nare*. SCYNEXIS, Inc., Post Office Box 12878, Research Triangle Park, NC.
Pharmacokinetics (PK) defines the time course of antimicrobial or anti-parasitic concentrations
in the body and pharmacodyamics (PD) focuses on relationships between such concentrations
and the anti-parasitic effect. PK and PD parameters are routinely defined and used to design
dosing regimens in microbiology. Recently, we employed rigorous in vitro PK/PD
experimentation to identify and develop new chemical entities for the treatment of human
African trypanosomiasis (HAT). Availability of advanced tissue culture techniques and predictive
animal models allowed for such a PK/PD driven approach to progress compounds from in vitro
to in vivo efficacy studies. More importantly, these tools are critical in defining dosing regimens
for human clinical studies. The PK/PD tools that we developed for HAT provide some
background and set the stage for expansion into the anthelmintic area.
91
Anti-filarial drug discovery is hampered by lack of suitable in vitro culture conditions and animal
models for compound screening. Consequently, critical issues that define translation of in vitro
to in vivo efficacy have not been defined. Disposition of anti-filarial compounds in plasma and
tissues and the relationships between concentrations and anti-parasitic efficacy are generally
not included in early discovery activities. Recent improvements in tissue culture techniques and
methods for assessment of filarial worm viability through motility and/or metabolic indicators
provide great opportunities for us to start defining some basic in vitro PK/PD parameters to aid
the discovery and development of new anti-filarial agents. The presentation will highlight
opportunities and challenges using specific examples from B. malayi and the canine filarial
worm D. immitis drug discovery activities. Available animal models and their potential role in
making in vitro/in vivo correlates will be explored. Incorporation of PK/PD activities in early
filarial discovery programs is critical and could ultimately be used in defining efficacy and dosing
regimen in target species.
President’s Symposium – Ticks and Tick-Borne
Diseases
85
Tick-borne infections in the southern United States – changing patterns.
Susan Little*. Center for Veterinary Health Sciences, Oklahoma State University, Department of
Veterinary Pathobiology, Stillwater, OK.
Ticks readily feed on humans and other animals, serving as an ideal conduit to move pathogens
between a wide spectrum of potential hosts. As ecological niches flux, opportunities arise for
these disease vectors to interact with different hosts, allowing the infectious agents they
transmit to expand both geographic and host range. Habitat change has been linked to the
emergence of new human and veterinary tick-borne disease agents, and can markedly facilitate
expansion opportunities by allowing existing tick populations to flourish and by supporting the
establishment of new tick-borne pathogen maintenance systems. Recent decades have seen
dramatic increases in the diversity and prevalence of tick-borne infections in people and dogs in
the southern United States, including increased reports of existing diseases such as those
caused by Ehrlichia spp. and spotted fever group Rickettsia spp., as well as recognition of
newer entities like Southern Tick-associated Rash Illness (STARI) and the Heartland virus. Lone
star ticks (Amblyomma americanum) are central to most of these infections, but other tick
species, including Gulf Coast ticks (A. maculatum) and brown dog ticks (Rhipicephalus
sanguineus) play an increasingly important role in disease transmission in this region. Some of
the increased reports appear due to widespread ecologic changes, although improved detection
systems have also greatly facilitated recognition of infections when they occur. In addition, the
presence of several distinct but closely related organisms in the same tick vector system often
complicates understanding of the natural history of these agents, and individual tick species
have been shown to be capable of transmitting a greater variety of infections than previously
thought. Novel strategies will be required to effectively combat tick-borne infections in the future
if we are to succeed in the goal of fostering an environment which supports healthy animals and
healthy people.
92
86
The emergence of Ixodes scapularis and Lyme disease in North America: how
differences in ecology may affect regional patterns.
Jean Tsao*. Michigan State University, Fisheries and Wildlife, Large Animal Clinical Sciences,
East Lansing, MI.
Ixodes scapularis, the blacklegged (=deer) tick, is the vector of several pathogens, including the
etiologic agents of Lyme disease, human granulocytic anaplasmosis, human babesiosis, human
erlichiosis, Powassan encephalitis, and relapsing fever borreliosis. Since the initial populations
were discovered in the mid-20th century, the blacklegged tick has been spreading from two main
foci, one in the Northeast, and one in Upper Midwest. Subsequently, the incidences of diseases
caused by etiologic organisms carried by this tick also have increased. While the abundance of
blacklegged ticks has increased in both regions, 10 of 12 of the states with the highest
incidence of Lyme disease cases occur in the Northeast. It has become more apparent that
differences in the ecologies in the two regions may affect enzootic transmission dynamics and
therefore zoonotic transmission dynamics, disease incidence, and even disease manifestation.
In this talk, I will describe and discuss the potential impacts of one of the most striking ecological
differences between the two regions: the seasonal activity curves of the juvenile stages of the
blacklegged tick. The degree of overlap in the timing of the two stages is influenced by climatic
factors and may have profound effects on the maintenance of pathogens and strains within
pathogens. Furthermore, recently there has been greater attention paid to the role that
passerine birds may play in the dispersal and spread of the blacklegged tick and associated
pathogens. Thus, the seasonality of activity of the various life stages as it coincides with bird
migration timing and direction is very important for predicting future spread of blacklegged ticks
and associated pathogens and therefore disease risk.
93
Author Index
Abstract
Abstract
Abstract
Abongwa, Melanie ......... 21, 67, 69
Acosta, Igor C.L......................... 83
Adolph, Chris ....................... 53, 63
Ahn, Ah-Jin ................................ 60
Ahn, Kyu-Sung .......................... 60
Alam, Muhammad Azhar ........... 32
Allen, Lynn ................................ 54
Ammer, Amanda G.................... 13
Amode, Deb .............................. 44
Arther, Robert ............................ 51
Atkins, Adrienne ........................ 80
Chapman, H. David ....................72
Charvet, Claude .........................69
Choudhary, Shivani....................21
Coburn, Lisa...............................25
Cohn, Leah ..........................29, 38
Colwell, Douglas ........................19
Conboy, Gary .......................22, 57
Cote, Jennifer.......................62, 63
Courtot, Elise .............................69
Craig, Tom .................................77
Crowdis, Kelly ............................18
Curtis, Rachel ............................30
Gajadhar, Alvin .......................... 47
Garza, Javier ....................... 10, 81
Gattenuo, Talia .......................... 55
Geary, Tim ................................... 2
Geeding, Amy ............................ 44
Geng, Jinming................ 61, 62, 63
Gennari, Solange M. .................. 83
Gilleard, John ...................... 20, 75
Gilley, Alexandra D. ..................... 8
Goater, Cameron ....................... 19
Goday, Clara ............................. 45
Goldstein, Richard E. ................. 41
Goolsby, Jaime .......................... 37
Gorohov, Vladimir ...................... 82
Gravatte, Holli S................... 43, 46
Gray, Hannah ............................ 33
Greenwood, Spencer ................. 22
Gruntmeir, Kaylynn .................... 49
Gruntmeir, Jeff ............... 49, 53, 54
A
B
Bailey, Keith .............................. 49
Ballweber, Lora R ...................... 35
Barrett, Anne ....................... 16, 49
Bartley, David ............................ 75
Beck, Melissa ............................ 19
Beck, Paul ................................. 33
Bellaw, Jennifer L. ........... 4, 45, 46
Bertram, Miranda R. .................. 23
Betancourt, Alejandra .......... 14, 15
Bishop-Stewart, Janell............... 42
Blagburn, Byron 49, 51, 53, 64, 65
Bousquet, Eric ........................... 43
Bowdridge, Scott A. ............. 11, 13
Bowles, Joy ............................... 51
Bowman, Dwight D ... 3, 27, 39, 40,
41, 50, 52, 59, 66
Brown, Heidi ........................ 40, 52
Brunker, Jill ............................... 18
Bucala, Richard ......................... 71
Burkman, Erica .......................... 26
Butler, Jamie ................. 51, 64, 65
Buxton, Samuel K.......... 21, 67, 69
Byrnes, Meghan .......................... 8
C
Cadell, Steve ............................... 4
Cao, Xin .................................... 46
Casey, Sarah ............................ 76
Chambers, Thomas ................... 14
D
Dana, Pollard .............................29
Datz, Craig .................................78
Davis, Richard............................45
Diaz, James ...............................31
Dillon, Allan ................................51
Donecker, John M. ...................4, 5
Drake, Jason ..............................54
Dreyfus, Jennifer ........................59
Droleskey, Robert ......................58
Dzimianski, Michael T. ...............26
E
Edourad, Emile Jean Pierre .......18
Elsemore, David.............61, 62, 63
Erf, Gisela F ...............................72
Evans, Christopher C. ................26
F
Fenger, Clara ...............................5
Fetterer, Raymond .....................71
Fidler, Andrew ............................33
Finney, Michael ..........................40
Florentine, Michelle ......................3
Fontenot, Deidre ........................81
Fournier, Gislene F.S.R. ............83
G
Gadde, Ujvala Deepthi ...............72
94
H
Hamer, Gabriel L. ................ 23, 30
Hamer, Sarah A. .................. 23, 30
Hanna, Paul ............................... 57
Hanna, Rita .......................... 62, 63
Hartup, Barry K. ......................... 23
Hausmann, Jennifer................... 59
Herrin, Brian .............................. 17
Hildreth, Michael .......................... 7
Hoberg, Eric ............................... 75
Hogan, Katherine ....................... 80
Holler, Larry ................................. 7
Holler, Sue ................................... 7
Holman, Patricia J.......... 29, 38, 58
Holt, Rush .................................. 13
Holzmer, Susan ......................... 33
Horohov, David .................... 14, 15
Hostetler, Joseph A. ..... 49, 50, 64,
65, 66
Houk, Alice .................................. 9
Howell, Sue ..................... 6, 36, 80
J
Jacobs, Jesica ........................... 11
Jacobsen, Stine ............ 14, 15 , 43
James, Andrea .......................... 29
Jeong, Hak-Sub ........................ 60
Johnson, Eileen ......................... 16
Jones, Linda .............................. 33
Jones, Scott .............................. 58
K
Kaplan, Ray ............... 6, 36, 44, 80
Kapustin, Nick ........................... 80
Keane, Delwyn .......................... 58
Kelly, Vicky .......................... 12, 81
Ketzis, Jennifer ........ 55, 56, 61, 73
Kim, Byung-Su .......................... 60
Kornele, Michelle ....................... 28
Krecek, Rosina Claudia ............. 77
Kurochkina, Karine .................... 82
L
Labruna, Marcelo B. .................. 83
Land, Tracey ................. 51, 64, 65
Lear, Teri ................................... 45
Leathwick, Dave ........................ 34
Lee, Alice .................................... 3
Lee, Choung-Seop .................... 60
Lee, Alice .................................. 66
Leng, Lin ................................... 71
Lewis, Roy ................................. 20
Lewis, Barbara .......................... 24
Lima, Julia T.R. ......................... 83
Lindsay, David S. .................. 9, 32
Liotta, Janice L ................ 3, 27, 59
Little, Susan ..... 16, 17, 18, 49, 53,
54, 63, 85
Lobanov, Vladislav .................... 47
Lopes, Marcos G. ...................... 83
Lucio-Forster, Araceli ................ 59
Lund, Robert ................. 40, 41, 52
Lyons, Eugene T. ............ 5, 15, 45
M
MacDonald, Erin ........................ 57
Macili, Arlei ................................ 83
Maclean, Mary ........................... 36
Malone, John B. ............ 31, 40, 52
Abstract
Abstract
Abstract
Q
Mans, Christoph .........................59
Marcellino, Chris ........................36
Marchiondo, Alan A. ...................33
Marron, Fany..............................57
Martin, Richard J. .....21, 67, 68, 69
Mason, Maren ............................44
McArthur, Morgan ........................1
McCoy, Ciaran J. .......................21
McGrew, Ashley K. ....................35
McMahan, Christopher S. ...40, 41,
52
Meneus, Pedro...........................18
Merritt, Hannah ..........................54
Miller, James ..................10, 12, 81
Miller, Melissa ..................6, 36, 80
Moorhead, Andrew R. ................26
Moskvin, Alexandr......................82
Mostafa, Eman ...........................36
Mount, Jane ...................51, 64, 65
N
Nagamori, Yoko ...................25, 37
Nare, Bakela ..............................84
Nazir, Muhammad Mudasser .....32
Neveu, Cedric ............................69
Newton, Kassie ..........................18
Nielsen, Martin K...4, 5, 14, 15, 43,
45, 46
Noden, Bruce .............................16
O
O'Brien, Anna .............................28
Olsen, Susanne N. .....................43
Opps, Sheldon ...........................22
Ortis, Hunter...............................44
P
Pagan, Joe ...................................4
Peneyra, Samantha .....................3
Phethean, Eileen..........................4
Pihl, Tina ....................................43
Pollard, Dana .......................38, 58
Powell, Jeremy...........................33
Prichard, Roger ..........................74
Pulaski, Cassan .........................31
95
Qu, Guanggang ......................... 71
R
Radi, Craig ................................. 58
Ramirez, Diego .......................... 83
Rathinam, Thilakar..................... 72
Reavill, Drury ............................. 59
Redman, Elizabeth .............. 20, 75
Reedy, Stephanie ...................... 14
Reese, R. Neil.............................. 7
Reichard, Mason...... 25, 29, 37, 38
Reinemeyer, Craig R. ................ 46
Riggs, Molly R............................ 26
Rishniw, Mark ............................ 59
Robbins, William ........................ 22
Robertson, Alan P.... 21, 67, 68, 69
Robinson, James ....................... 73
Rodriguez, Jessica .................... 24
Roncalli, Raffaele....................... 48
Rosypal, Alexa ............................. 9
Rubinson, Emily ................... 14, 46
S
Sakanari, Judy ........................... 36
Saleh, Meriam.............................. 8
Samoylovskaya, Nina ................ 82
Sarah, Anwar ............................... 7
Sargison, Neil ............................ 75
Sghiatti, Tessa ........................... 80
Shanks, Mauricia ....................... 17
Sharp, Julia .................... 40, 41, 52
Shell, Linda ................................ 55
Shin, Jihun ................................. 60
Shin, SungShik .......................... 60
Silva-Opps, Marina .................... 22
Snowden, Karen ............ 24, 77, 79
Sommers, Karen ........................ 11
Starkey, Lindsay .................. 18, 49
Staubus, Lesa ............................ 37
Stephens, Maci ............................ 5
Storey, Bob ...................... 6, 36, 80
Suh, Guk-Hyun .......................... 60
T
Terry, Jenna .............................. 81
Thomas, Jennifer ........... 25, 37, 49
Abstract
Trosclair, Macy .......................... 81
Tsao, Jean ................................ 86
Tucker, Chris ............................. 33
Tuo, Wenbin ........................ 70, 71
Tyrrell, Phyllis ............................ 62
U
Uspenskiy, Aleksandr ................ 82
V
van Dam, Govert ....................... 24
Vanderstichel, Raphael ............. 22
Vanimisetti, Himabindu B. ......... 33
Vatta, Adriano F. ................... 6, 33
Verma, Saurabh .................. 21, 68
Vidyashankar, Anand N............. 46
Von Simson, Cristiano ............... 51
Voris, Nathan ............................ 44
W
Wagner, Bettina ........................ 14
Wang, Dongmei ............ 40, 41, 52
Wang, Jianbin ........................... 45
Wasmuth, James ...................... 20
Weingartz, Christine .................. 33
Wildeus, Stephan ...................... 76
Wolstenholme, Adrian ......... 36, 68
Wray, Eva .................................. 33
Y
Yazwinski, Thomas A. ............... 33
Z
Zajac, Anne M. ............ 8, 9, 17, 76
Zarlenga, Dante S. ........ 35, 70, 71
Zhu, Dan ................................... 27
Zolynas, Robert ......................... 51
96
MEMBERSHIP DIRECTORY
2014
97
98
59th Annual Meeting, 26-29 August 2014
Elizabeth M. Abbott
Abbott Associates
Dene Lodge Husbands Bosworth
Lutterworth, LE17 6NL
United Kingdom
44 185 888 1050
44 185 888 1151
emabbott@ecoanimalhealth.com
ema@abbottassoc.demon.co.uk
Liselotte Aberg
SkagersvÃ¤gen 15
Ã…rsta, 12038
Sweden
+46 709384776
v6liselotte@hotmail.com
Melanie Abongwa
2008 Vet Med
Department of Biomedical Sciences
Ames, IA 50011
US
443-985-1586
mabongwa@iastate.edu
Albert Abraham
Bayer Health Care
P.O. Box 390
Shawnee Mission, KS 66201
United States
913-268-2860
albert.abraham.b@bayer.com
Mathew Abraham
1907 S. Milledge Ave. Apt. C8
Athens, GA 30605
United States
mabrahamvet@gmail.com
Alex D.W. Acholonu
Alcorn State University
1000 ASU Drive, #843
Alcorn State, MS 39096
United States
601-877-6236
601-877-2328
chieffacholonu@yahoo.com
chiefacholonu@alcon.edu
Chris Adolph
6319 S. Elm Pl.
Broken Arrow, OK 74011
United States
918-519-7750
918-449-0485
adolphdvm@tulsacoxmail.com
adolph@okstate.edu
Benjamin AduAddai
Pathobiology and Diagnostic
Investigation A42 Veterinary Medical
Center
East Lansing, MI 48824
United States
517-353-9070
517-432-5836
aduaddai@dcoah.msu.edu
Dalen Agnew
DCPAH 4125 Beaumont Rd. Room
152B
United States
517-432-5806
517-432-5836
agnewd@dcpah.msu.edu
Armando Aguilar-Caballero
Universidad Autonoma de Yucatan
17 No. 93 X 18 y 20
Kanasin, YucatÃ¡n 97370
Mexico
+52 999 9423200
+52 999 9423205
aguilarc@uady.mx
Heather Aitken
Health Canada
11 Holland Ave
Suite 14
Ottawa, Ontario K1A 0K9
Canada
613-960-4638
613-957-3861
heather.aitken@hc-sc.gc.ca
haitken@prescottvet.ca
Lesel Ali-De Silva
P.O. Box 1323
Port-of-Spain,
Trinidad and Tobago
868-766-0272
868-622-4908
leselali@gmail.com
Mohammad Ali-Sabi
Section of Zoology
Throvaidsensvej 40
Copenheagen, DK 01871
Denmark
453 533-2672
nafi@life.ku.dk
Kelly E. Allen
250 McElroy Hall; CVHS OSU
Stillwater, OK 74078
United States
405-744-3236
kelly.allen.warren@okstate.edu
Lynn Allen
Novartis Animal Health
1722 SW 300
Kingsville, MO 64061
United States
816-597-3497
816-597-3497
lynn.allen@novartis.com
Kelly Allen
225 McElroy Hall
USA
405-744-4573
405-744-5275
kelly.allen10@okstate.edu
Jennifer Allen
Bayer HealthCare Animal Health
235 Bateman Avenue
Franklin, TN 37067
USA
615-613-8239
Jennifer.a.allen@bayer.com
Jallendvm99@gmail.com
Maria Sonia Almeria
Autonomous University Y Barcelona
Veterinary School, Parasitology
Campus UAB Edifici V 08193
Bellaterra
Barcelona, Spain 8193
Spain
011 34 649 808414
011 34 935 872006
sonia.almeria@uab.cat
Ulla Andersen
Royal Vet. & Agriculture
University
Klostermarken 69, Skjem
Denmark, 6900
The Netherlands
ullava@gmail.com
ullava@life.ku.dk
Prema Arasu
North Carolina State University
College of Vet. Medicine 4700
Hillsborough St.
Raleigh, NC 27606
United States
919-513 6530
919-513 6455
prema_arasu@ncsu.edu
James J. Arends
S&J Farms Animal Health
2340 Sanders Rd.
Willow Springs, NC 27592
United States
919-894-5684
919-894-9010
parasit@aol.com
99
Allison Arms
Bayer
4614 Fairview
Davis, OK 73030
United States
580-247-0028
Alison.arms@bayer.com
Robert G. Arther
Bayer Health Care
Animal Health, 12809 Shawnee
Mission Parkway
United States
913-268-2503
913-268-2541
bob.arther@bayer.com
Marjory Artzer
Kansas State University
Coleo Hall 1800 Denison
Manhattan, KS 66506
United States
785-532-4611
martzer@vet.ksu.edu
Rupinderjit Avapal
Veterinary Parasitology (P.A.U.)
28 Sestina Crt.
Brampton, ON L6P 1R9
Canada
905-789-6158
rupidoraha@yahoo.com
Russell Avramenko
HSC 2531, 3330, Hospital Drive NW
Calgary, Alberta T2N 4N1
Canada
403-210-6788
403-210-6693
rwavrame@ucalgary.ca
Christine Baker
Merial, Ltd.
115 Transtech Dr.
Athens, GA 30601
United States
706-552-2445
706-543-1667
christine.baker@merial.com
David G. Baker
School of Vet. Medicine 1424
Westchester Dr.
Baton Rouge, LA 70810
United States
225-578-9643
225-578-9649
dbaker@vetmed.lsu.edu
David Baker
School of Veterinary Medicine
USA
225-578-9643
225-578-9649
dbaker6@lsu.edu
Norman L. Baldwin
Baldwin Aquatics, Inc.
12480 Pedee Creek Rd.
Monmouth, OR 97361
United States
503-838-2600
503-838-2600
norman.baldwin@bigfoot.com
norm@truecastdesign.com
Verona A. Barr
Heartland Community College
1500 W. Raab Rd.
Normal, IL 61761
United States
309-268-8667
verona.barr@heartland.edu
Melissa Beck
Box 998
Coaldale, AB T1M 1M8
Canada
403-329-2319
Melissa.thomson2@uleth.ca
Frederic Beugnet
Merial, Ltd.
26 Av Tony Garnier
Lyon, 69007
France
3368 774-8983
frederic.beugnet@merial.com
Virginie Barrere
virginiebarrere@yahoo.fr
Melissa Beck
4401 University Drive
Lethbridge, Alberta T1K3M4
Canada
403.329.2319
m.beck@uleth.ca
Ellen Binder
Virginia Tech University
VMRCVM Duckpond Dr. Ph 2, Room
214
Blacksburg, VA 24061
United States
540-231-6471
ebinder@vt.edu
Anne Barrett
1024 W. Preston Ave.
United States
207-749-6632
Anne.Barrett@okstate.edu
Cathy A. Ball
Summit VetPharm
301 Route 1704 Floor 12
Rutherford, NJ 07070
United States
201-242-8412
201-585-9525
cball@summitvetpharm.com
John R. Barta
Ontario Vet. College Department of
Patholobiology
Guelph, ON N1G 2W1
Canada
519-824-4120, Ext. 54017
jbarta@uoguelph.ca
Lora Rickard Ballweber
College of Veterinary Medicine &
Biomedical Sciences Veterinary
Diagnostic Lab 1644 Campus
Delivery
Fort Collins, CO 80523-1644
United States
970-297-5416
lora.ballweber@colostate.edu
William Barton
IVX Animal Health, Inc.
3915 S. 48th St. Terrace
St. Joseph, MO 64503-4711
United States
816-676-6152
816-676-6874
william_barton@ivax.com
Sharron Barnett
3200 Northline Ave.
Greensboro, NC 27408
United States
336-387-1059
sharron.barnett@novartis.com
Carly Barone
University of Rhode Island
USA
carly_barone@my.uri.edu
Stephen C. Barr
Cornell University
College of Vet. Medicine Dept. of
Clinical Sciences
Ithaca, NY 14853
United States
607-253-3043
607-253-3534
scb6@cornell.edu
Dieter Barutzki
Veterinay Laboratory Freiburg
Wendlinger Str. 34
Freiburg 1.BR., 79111
Germany
49 761 459-9780
barutski@labor-freiburg.de
Bahar Baviskar
250 McElroy Hall
United States
405 334 2397
Drbaharbavis@gmail.com
Melissa Beal
IDEXX Laboratories
4 Mountain View Rd.
Cape Elizabeth, ME 04107-1321
United States
207-556-4746
melissa.beal@idexx.com
Brenda Beerntsen
209 Connaway Hall
Columbi, MO 65211
United States
573-882-5033
beerntsenb@missouri.edu
Jennifer Bellaw
108 Gluck Equine Research Center
Lexington, ky 40546
usa
7407084807
jbe244@l.uky.edu
Thomas R. Bello
Sandhill Equine Center
P.O. Box 1313
Southern Pines, NC 28388
United States
910-692-6016
910-949-4559
bellodvm@earthlink.net
Gerald W. Benz
830 Gabbettville Rd.
LaGrange, GA 30240
United States
706-645-1999
706-645-1999
gb830kiaspdwy@hotmail.com
Miranda Bertram
College Station, TX 77843
mbertram@cvm.tamu.edu
Alejandra Betancourt
Lexington, Ky 40546
United States
859-218-1114
859-257-8542
abeta2@uky.edu
Byron L. Blagburn
Auburn University
College Vet. Medicine Dept.
Pathobiology 122 Green Hall
Auburn University, AL 36849-55
United States
334-844-2702
334-844-2652
blagbbl@vetmed.auburn.edu
David L. Bledsoe
Animal Clinical Investigation, LLC
4926 Wisconsin Ave NW
Washington, DC 20016
United States
2028158918
dbledsoe@animalci.com
DavidDrDad@gmail.com
Venkata Boppana
University Conn. Health Center
211 Main St. Apt. D3
Farmington, CT 06032
United States
860-751-8882
dboppana@yahoo.com
Scott Bowdridge
G050 Ag. Sciences Building
PO Box 6108
Morgantown, WV 26506
USA
(304) 293-2003
(304) 293-2232
scott.bowdridge@mail.wvu.edu
Dwight D. Bowman
Cornell University
College of Vet. Medicine C4-119
VMC/Micro & Immunol.
Ithaca, NY 14853-6401
United States
607-253-3406
607-253-4077
ddb3@cornell.edu
Anastasia Bowman
anaspyder@gmail.com
100
Richard E. Bradley
4981 Ganung Dr.
Morganton, NC 28655
United States
828-430 8695
alla@usit.net
Michael Breer
Eli Lilly & Company
201 Bountiful Pointe
Wildwood, MO 63040
United States
614-218-3355
breermi@lilly.com
Matt Brewer
University of Wisconsin
107 1/2 Washington St.
Eau Claire, WI 50010
United States
715-460-0258
brewermt@uwec.edu
Holly E. Brianceau
Intervet, Inc.
29160 Intervet Ln. P.O. Box 318
Millsboro, DE 19966
United States
302-934-4232
302-933-4008
holly.brianceau@intervet.com
Kaitlyn Briggs
Cornell University
United States
607-226-3303
krb78@cornell.edu
Dave Bromert
Intervet Schering-Plough Animal
Health
649 Butternut Ct.
Liberty, MO 64068
United States
816-550-3414
dave.bromert@intervet.com
Mary Lou Brongo
617 Kenmore Rd.
Brick, NJ 87230
United States
732-477-9440
mlbrongo@yahoo.com
Holly Brown
310 King Ave.
Athens, GA 30606
United States
706-542-4596
hmbrown@uga.edu
Caroline Brown
University of Tennessee College of
Veterinary Medicine
1324 Sumac Dr.
Knoxville, TN 37919
USA
817-992-0081
cbrow123@utk.edu
carolinebrown@my.unt.edu
William W. Burdett
Intervet, Inc.
8066 N. 130th Rd.
Cairo, NE 68824
United States
308-379-3261
308-485-4321
bill.burdett@intervet.com
Casey Burke
159 Liberty Ln.
Pembroke, VA 24136
United States
540-778-3055
crburke@vt.edu
Ann Busby
310 Kyle Ct.
United States
870-267-2611
anntaylor.busby@okstate.edu
Xin Cao
George Mason University
11102 Cavalier Ct.
Fairfax, VA 22030
United States
732-266-3956
xcao2@gmu.edu
Barbara Butler
Charles River, Ballina
Co Mayo,
Ireland
35 3879919361
barbara.butler@crl.com
Colin Capner
Novartis Animal Health UK
1 Park View
Riverside Way
Watchmoor Park
Camberley, Surrey GU15 3YL
United Kingdom
7557288418
colin.capner@novartis.com
Ronnie Byford
New Mexico State University
P.O. Box MSC 3 BF
Las Cruces, NM 88003
United States
575-571-6980
rbyford@nmsu.edu
Beth Byles
Southern Illinois University
6846 Giant City Rd. Apt. C
Carbondale, IL 62902
United States
618-525-0805
bbyles@siu.edu
T. Michael Burke
10607 Patterson Ave.
Richmond, VA 23238
United States
drb@felinemedicalcenter.comcastbiz John Byrd
.net
907 N. Westbrook Cir.
Mahomet., Il 61853
Erica Burkman
United States
217 586 2004
hlab@horsemenslab.com
340 Timber Creek Drive
Athens, GA 30605
USA
Joseph W. Camp Jr.
770-355-7565
Purdue University College of
eburkman@uga.edu
Veterinary Medicine
725 Harrison St.
West Lafayette, IN 47907
Meghan Burns
United States
Meghan.Burns@virbacus.com
765-496-2463
765-494-9830
Roxanne Burrus
jcamp@purdue.edu
NAMRU-6 Unit 3230 Box 0047
DPO, AA 34031
Bill R. Campbell
United States
Piedmont Pharmaceuticals, LLC
511-614-4133
204 Muirs Chapel Rd. Suite 200
Roxanne.burrus@med.navy.mil
United States
Kevin Burton
336-544-0320, ext. 203
Merial
336-544-0322
285 Quarterhorse Dr
bill.campbell@piedmontpharma.com
Liberty Hill, TX 78642
USA
William C. Campbell
9133758607
wade.burton@merial.com
Drew University
wadeb@austin.rr.com
27 Samuel Way
Madison, NJ 07940
United States
978-668-0489
wcampbel@drew.edu
101
Larry Captini
Ohio State University
captini.1@osu.edu
Doug Carithers
Merial, Ltd.
3239 Satellite Blvd, Building 500
Duluth, GA 30096
United States
678-638-3837, Mobile 770-331-6069
678-638-3817
doug.carithers@merial.com
James Carmichael
3200 Northline Ave.
United States
336-387-1031
james.carmichael@novartis.com
Juliette Carroll
2630 Wapiti
Fort Collins, CO 80525
United States
979-557-3843
Juliecarroll12@gmail.com
Lori Carter
Stillmeadow, Inc.
12852 Park One Dr.
Sugar Land, TX 77478
United States
281-240-8828
281-240-8448
lcarter@stillmeadow.com
Sarah Casey
sjcasey1@vt.edu
Richard J. Cawthorn
Atlantic Veterinary College AVC
Lobster Science Centre
Charlottetown, PE C1A 4P3
Canada
902-566-0584
902-566-0851
cawthorn@upei.ca
H. David Chapman
Department of Poultry Sciences
1260 W. Maple
Fayetteville, AR 72701
United States
479-575-4870
479-575-4202
dchapman@uark.edu
Roxanne Charles
The University of the West Indies
Upper Wharton Street, Success
Village, Laventille
Port of Spain,
Trinidad and Tobago
1-868-355-2372
roxxyanne@hotmail.com
roxanne.charles@sta.uwi.edu
Samuel Charles
Bayer Health Care
P.O. Box 390
United States
913-268-2520
913-268-2541
sam.charles.b@bayer.com
Sam Charles
Bayer Animal Health
12809 Shawnee Mission Pkwy
Shawnee, Kansas 66216
USA
913-268-2520
913-268-2541
sam.charles@bayer.com
Umer Naveed Chaudhry
Calgary, AB T2L 0B6
CN
403-667-4082
unchaudh@ucalgary.ca
Rajendran Chellaiah
USDA
Animal Parasitic Diseases
Laboratory 10300 Edmonston Rd.
Beltsville, MD 20705
United States
301-504-5111
chellairajendran1@gmail.com
Hoyt Cheramie
Merial, Ltd.
859-806-7099
678-638-8631
hoyt.cheramie@merial.com
Jose W. Chernitzky
Facultad de Medicina Veterinaria y
Zoot
UNAM Newton 283 Depto. 702
Colonia Polanco Chapultepec, DF
CP 11560
Mexico
52 55 52030310
jcher13@hotmail.com
M.B. Chhabra
D II / 2518, Vasant Vihar
New Delhi, 110070
India
91-011-26123444
chhabramb@yahoo.com
Bill Chobotar
Andrews University
Department of Biology Price Hall
Room 216
Berrien Springs, MI 49104
United States
269-471-3262
269-471-6911
chob@andrews.edu
Annapoorani Chockalingam
Cornell University
520 The Parkway
Ithaca, NY 14850
United States
607-343-0606
drannss@gmail.com
ac469@cornell.edu
Les Choromanski
Pfizer, Inc.
7000 Poratge Rd. KZO 267-03-303
Kalamazoo, MI 49001
United States
269-833-2503
269-833-4255
leszek.j.choromanski@pfizer.com
Shivani Choudhary
Biomedical Sciences
2051 VETMED
College of Veterinary Medicine
Ames, Iowa 50010
USA
515(450)-4626
shivani@iastate.edu
shivanichoudhary01@gmail.com
Jeffrey N. Clark
JNC Consulting Services, Inc.
38 Wild Rose Ln.
Pittsboro, NC 27712
United States
919-942-9222
depotrd33@yahoo.com
Bill C. Clymer
Consultant to Animal Health
13351 E. FM 1151
Amarillo, TX 79118
United States
806-335-3338
806-335-3531
billclymer@rocketmail.com
Nicole Colapinto
7 Mullford Ave.
Ajax, ON L1Z 1K7
Canada
289-681-6034
nicole.colapinto@merck.com
Rebecca A. Cole
USGS National Wildlife Health
Research Center
6006 Schroeder Rd.
Madison, WI 53711
United States
608-270-2468
608-270-2415
rcole@usgs.gov
Cynthia Cole
cynthia.cole2010@gmail.com
Gerald C. Coles
University of Bristol
Department of Clinical Vet. Science
Langford House
Bristol, BS40 5DU
United Kingdom
44-117-928-9418
44-117-928-9504
gerald.c.coles@bristol.ac.uk
Douglas D. Colwell
Agriculture and Agri-Food Canada
5403 1st Ave. S.
Lethbridge, AB T1J 4B1
Canada
403-317-2254
403-382-3156
doug.colwell@agr.gc.ca
Gary A. Conboy
Atlantic Veterinary College 550
University Ave.
Canada
902-566-0965
902-566-0851
conboy@upei.ca
George A. Conder
Pfizer Animal Health (retired)
9244 Weathervain Tr.
Galesburg, MI 49053
United States
269-665-4948
nannykbc@aol.com
102
Donald P. Conway
Conway Associates
2 Willever Rd.
Asbury, NJ 08802
United States
908-730-8823
conwaydp@earthlink.net
Camille Coomansingh
St. George's University
P.O. Box 7
St. George,
Grenada
473-444-4175
473-439-5068
ccoomansingh@sgu.edu
ccoomansingh@yahoo.com
Helder Cortes
Evora University
Lab of Parasitology Victor Caeiro
Mitra Center
Evora, 7000
Portugal
35198 501-7554
hcec@uevora.pt
Roberto Cortinas
University of Nebraska Lincoln
School of Veterinary Medicine and
Biomedical Sciences
P.O. Box 830905
East Campus Loop and Fair Street
Lincoln, NE 68583-0905
USA
(402) 472-6502
(402) 472-4687
rcortinas@unl.edu
Charles H. Courtney
University of Florida
College of Vet. Medicine P.O. Box
100125
Gainesville, FL 32610-0125
United States
352-392-4700, ext. 5111
352-392-8351
courtneyc@mail.vetmed.ufl.edu
Bobby E. Cowles
Pfizer Animal Health
18503 NW 41st Ave.
Ridgefield, WA 98642
United States
360-606-1443
bobby.cowles@pfizer.com
Carla Craig
858 9th Ave.
Longview, WA 98632
United States
carlaxcj@gmail.com
Thomas M. Craig
College of Vet. Medicine Department
of Vet. Pathobiology
United States
979-845-9191
979-862-2344
tcraig@cvm.tamu.edu
Luiz G. Cramer
Merial, Ltd.
3239 Satellite Blvd.
Duluth, GA 30024
United States
luiz.cramer@merial.com
Larry R. Cruthers
Professional Laboratory &
Research Services, Inc.
1251 NC 32 North
Corapeake, NC 27926
United States
252-465-8686
252-465-4493
cruthers@embarqmail.com
plrsinc@coastalnet.com
Alejandro Cruz-Reyes
Universidad Nacional Autonoma de
Mexico
Moras 556 301
Mexico City, Distrito Federal 30100
Mexico
cruzreyes11@yahoo.com
acr@ib.unam.mx
Cassandra Cullin
110 E. Lakeview Apt. A1
United States
405-808-0172
cassandra.cullin@gmail.com
Rachel Curtis
Veterinary Integrative Biosciences
Department, College of Veterinary
Medicine, Texas A&M University
4458 TAMU
USA
979-458-4924
rcurtis@cvm.tamu.edu
Tracy L. Cyr
Vet. Pathobiology 4467 TAMU
College Station, TX 77843-44
United States
979-458-3276
979-862-2344
tcyr@cvm.tamu.edu
Valdemiro da Silva JUnior
UFRPE
Rua Jorge Couceiro da Costa Eiras
443, Apto 2002, Boa Viagem.
Recife-PE. Brazil
Recife, Pernambuco 51021300
Brazil
5.5813327229e+011
valdemiroamaro@gmail.com
vajunior@dmfa.ufrpe.br
Dean Dailey
Merial
Office 641
Duluth, GA 30096
USA
706-247-5194
678-638-8636
dean.dailey@merial.com
Patricia Daly
Bayer Health Care
39 Waterbury St. Apt. 1
Saratoga Spring, NY 12866
United States
518-581-9617
trish.daly.b@bayer.com
tdalydvm@nycap.rr.com
Gabriel Davila
2949 SW 39th Ave.
Gainesville, FL 32608
United States
352-281-1124
gdavila@ufl.edu
Bradley DeWolf
8066 Heritage Line
Wallaceburg, ON N8A 4L3
China
519-824-4120
bdewolf@uoguelph.ca
Wendell L. Davis
15733 Woodson St.
Overland Park, KS 66223
United States
9138514954
wldavis@swbell.net
wldavis@swbell.net
Danielle Dimon
Infectious Diseases
Athens, GA 30602
United States
404-646-1764
dedimon@gmail.com
Theresa de Blieck
University of Minnesota
3883 County Rd. 74
Saint Cloud, MN 56302
United States
612-868-8643
tessdeblieck@yahoo.com
B. Joe Dedrickson
Merial, Ltd.
207 Ruth's Lane
Wamego, KS 66547-9014
United States
678-428-7405
John B. Dame
678-638-8870
joe.dedrickson@merial.com
College of Vet. Medicine Box 110880 joe.dedrickson@hotmail.com
United States
Nicole DeFraeye
352-392-4700
352-392-9704
5310 Pain Court Line
damej@mail.vetmed.ufl.edu
Pain Court, ON N0P 120
Canada
Sriveny Dangoudoubiyam
519-351-5514
519-355-1247
Gluck Equine Research Center 1435 nicole.defraeye@pfizer.com
Nicholasville Rd.
Lexington, KY 40546
Bart DeLeeuw
United States
859-257-4757, ext. 81167
P.O. Box 37
sriveny.dangoud@uky.edu
Capelle A/D 'jssel, 2900AA
The Netherlands
+31 10 406 4600
Craig Datz
+31 10 406 4293
bart.de.leeuw@pfizer.com
College of Vet. Medicine 900 E.
Campus Dr.
Columbia, MO 65211
Andrew DeRosa
United States
Bayer HealthCare LLC Animal
573-882-7821
Health
573-884-7563
13406 W72nd Street
datzc@missouri.edu
Shawnee, KS 66216
USA
913 200 7090
Amber Davidhizar
andy.derosa@bayer.com
University of Pennsylvania
andy.derosa5@gmail.com
amberdav@vet.upenn.edu
103
Sandra K. Dixon
Sandra Dixon, DVM
P.O. Box 192
Grayson, GA 30017
United States
770-235-5768
jantedixon@yahoo.com
Cynthia Doffitt
Mississippi State University
College of Vet. Medicine 1 Spring St.
Mississippi State, MS 39762
United States
662-325-1154
doffitt@cvm.msstate.edu
Natalia Dogrul
20 LaBonne Vie Dr. Apt. K
Patchogue, NY 11772
United States
613-404-9599
Alektron33@msn.com
Dan T. Domingo
150 E. 42nd St. 150/39/11
New York, NY 10017
United States
212-573-3050
212-309-0428
dan.t.domingo@pfizer.com
William A. Donahue Jr.
Sierra Research Laboratories
5100 Parker Rd.
Modesto, CA 95357
United States
209-521-6380
209-521-6380
srl@clearwire.net
John M. Donecker
Zoetis
707 Parkway Blvd.
Reidsville, NC 27320
United States
336-552-6027
866-590-0323
john.donecker@zoetis.com
Ann R. Donoghue
Donoghue Consulting, LLC
150 N. County Rd. 3
United States
970-482-2044, 970-214-5999
ardonoghue@gmail.com
Mary E. Doscher
147 Gary Dr.
Trenton, NJ 08690
United States
609-586-3185
doschem@msn.com
Jason Drake
7809 Charles Place Dr.
Kernersville, NC 27284
United States
336-387-1027
jason.drake@novartis.com
Michael W. Dryden
Coles Hall 1800 Denison Ave.
Manhattan, KS 66506
United States
785-532-4613
785-532-4039
dryden@vet.ksu.edu
J.P. Dubey
USDA, ARS, ANRI, APDL
Building 1001 BARC-East
Beltsville, MD 20705-2350
United States
301-504-8128
301-504-9222
jitender.dubey@ars.usda.gov
Brock E. Duck
University of Missouri-Columbia
Department of Veterinary
Pathobiology
University of Missouri-Columbia
220 Connaway Hall
Columbia, Missouri 65211
United States
bed89f@mail.missouri.edu
Karen L. Duncan
Intervet, Inc.
29160 Intervet Ln.
Millsboro, DE 19966
United States
302-934-4368
302-934-4203
karen.duncan@sp.intervet.com
Michael T. Dzimianski
Michael Dzimianski, DVM
452 Pony Tr.
Nicholson, GA 30565
United States
706-546-8520
706-546-8520
ponytrail@juno.com
Selene Elizondo
Berengena No.15
Xalapa, 91157
Mexico
2281456950
sln_m85@4hotmail.com
Jenifer Edmonds
Johnson Research, LLC
24007 Highway 20/26
Parma, ID 83660
United States
208-722-5829
208-722-7371
jedmonds@johnsonresearchllc.com
jen_edmonds@frontiernet.net
Matthew Edmonds
24007 Highway 2026
Parma, ID 83660
United States
208-722-5829
208-722-7371
medmonds@johnsonresearchllc.com
Amy Edwards
1818 W. Admiral
United States
amyce@okstate.edu
Jessica Edwards
335 Spring Lake Ct.
Athens, GA 30605
United States
404-558-0438
jessed5@uga.edu
Kristine Edwards
Department of Biochemistry,
Molecular Biology, Entomology &
Plant Pathology
100 Twelve Lane
United States
662-325-2085
662-325-8837
kt20@msstate.edu
ktedwardsdvm@bellsouth.net
Amal K. El-Gayar
200 Charls Haltom #7
United States
979-845-5180
979-845-2344
amalk18@yahoo.com
aelgayar@cvm.tamu.edu
Dave Ellefson
73 Silver Ridge Ln.
Falmouth, VA 22405
United States
605-261-1016
david.ellefson@bimedaus.com
Siobhan Ellison
Pathogenes, Inc.
P.O. Box 970
Fairfield, FL 32634
United States
352-591-3221
352-591-4318
sellison@pathogenes.com
Stacey Elmore
914 Cherry St.
United States
303-579-6667
selmore@colostate.edu
elmore.stacey@gmail.com
David Elsemore
IDEXX Laboratories
1 IDEXX Dr.
Westbrook, ME 04106
United States
207-556-3337
david-elsemore@idexx.com
Richard G. Endris
Intervet Schering-Plough Animal
Health
56 Livingston Ave.
Roseland, NJ 07068
United States
862-245-5133
862-245-3654
richard.endris@sp.intervet.com
Richard Endris
Merck Animal Health
556 Morris Avenue
Summit, NJ 07901
USA
908-473-5618
908-473-5560
apismellifera@verizon.net
richard.endris@merck.com
Michael J. Endrizzi
16036 Eagle River Way
Tampa, FL 33624
United States
336-210-9760
drizzi@verison.com
104
Michael Endrizzi
Zoetis
16036 Eagle River Way
Tampa, FL 33624
USA
336-210-9760
drizz@tamapbay.rr.com
Christian Epe
Petite Glane
St. Aubin FR, CH-1566
Switzerland
+41 26 679 1507
+41 26 679 1228
christian.epe@hotmail.de
Andrew K. Eschner
Merial, Ltd.
4 Pepper Pl.
Gansevoort, NY 12831
United States
518-580-0459
518-580-0460
drandrew.eschner@merial.com
Christopher Evans
Infectious Diseases
Athens, GA 30602
United States
7068704039
ccevans@uga.edu
Kevin Evans
150 E. 42nd St. 150/40/14
New York, NJ 10017
United States
212-733-0910
kevin.evans@pfizer.com
Richard Evans
Pacific Marine Mammal Center
22501 Chase #3102
Aliso Viejo, CA 92656
United States
949-494-3050
949-494-2802
revans@pacificmmc.org
Sidney A. Ewing
Department of Vet. Pathobiology 250
McElroy Hall
Stillwater, OK 74078-2007
United States
405 744-8177
405 744-5275
sidney.ewing@okstate.edu
Maarten Eysker
University of Utrecht
Div. Para. & Tropical Vet.
Medicine Yalelaan 1 P.O. Box
80.165, 3508 TD
Utrecht, 3508TD
The Netherlands
+31 30 253 1223
+31 30 254 0784
m.eysker@vet.uu.nl
Becky Fankhauser
Merial, Ltd.
115 Transtech Dr.
Athens, GA 30601
United States
706-552-2221
becky.fankhauser@merial.com
Silvina Fernandez
Facultad de Ciencias Veterinarias,
UNCPBA
Campus Universitario
B7001BBO, Tandil, Argentina
Tandil, Buenos Aires B7001BBO
Argentina
+54 249 4439852 ext. 257
+54 249 4439852
silvfernandez@gmail.com
sfernand@vet.unicen.edu.ar
Flavia Girao Ferrari
Merck Animal Health
35500 W 91st. street
De Soto, KS 66018
United States
9139918124
flavia.ferrari@merck.com
girao@cvm.msstate.edu
Flavia Ferrari
flaviaferrari@yahoo.com.br
Raymond H. Fetterer
USDA-ARS, APDL, BARC-East
Building 1040
United States
301-504-8762
301-504-5306
raymond.fetterer@ars.usda.gov
Tabitha Finch
tabitha.finch@mail.mizzou.edu
Emily Fitzgerald
1710 e. Northfield blvd
Apt. N3
murfreesboro, tn 37130
united states of america
205-381-3411
eefitz@bellsouth.net
Michael G. Fletcher
Y-Tex Corp.
1825 Big Horn Ave. P.O. Box 1450
Cody, WY 82414
United States
307-587-5515, ext. 115
307-527-6433
mfletcher@ytex.com
Catalina Fung
Peggy Adams Animal Rescue
League
P.O. Box 243094
Boynton Beach, FL 33424
United States
305-213-1393
catifung@yahoo.com
James R. Flowers
College of Vet. Medicine 525 Walnut
Grove Church Rd.
Raleigh, NC 27606
United States
919-513-6404
james_flowers@ncsu.edu
Jennifer Furchak
Hartz Mountain Corp.
31 Chamber Lane
Manalapan, NJ
United States
jfurchak@hartz.com
Laurie Flynn
IDEXX Laboratories
1 IDEXX Dr.
Westbrook, ME 04092
United States
207-556-4226
laurie-flynn@idexx.com
Wayne Forbes
121 Morningtide Dr.
Butler, PA 16002
United States
724-728-4953
Wayne.forbes@sru.edu
Ujvala Deepthi Gadde
7 S. Duncan Ave.
United States
479-966-9834
ujvalagadde@gmail.com
Alvin Gajadhar
Centre for Food-borne and Animal
Parasitology, CFIA
116 Veterinary Rd.
Saskatoon, Saskatchewan S7N 2R3
Canada
306-975-5344
306-975-5711
alvin.gajadhar@inspection.gc.ca
William J. Foreyt
Washington State University
Dept. Vet. Micro. & Path.
Pullman, WA 99164-7040
United States
509-335-6066
509-335-8529
wforeyt@vetmed.wsu.edu
Rajshekhar Gaji
University of Michigan
1737 Cram Cir. Apt. 08
Ann Arbor, MI 48105
United States
734-546-3606
rajgaji@umich.edu
Glenn R. Frank
Heska Corporation
3760 Rocky Mountain Ave.
Loveland, CO 80538-9833
United States
970-493-7272
970-619-3003
gfrank@heska.com
H. Ray Gamble
National Research Council
500 Fifth St. NW Room 557
United States
202-334-2787
202-334-2759
rgamble@nas.edu
Marguerite K. Frongillo
University of South Carolina
3600 Beverly Dr.
Columbia, SC 28204
United States
803-743-0731
mff4@cornell.edu
mff426@gmail.com
Robert D. Garrison
Texas Corners Animal Hospital
7007 W. Q Ave.
Kalamazoo, MI 49009
United States
269-375-3400
bob.garrison@comcast.net
Javier Garza
8001 Jefferson Hwy, Apt. 47
United States
956-451-4552
jgarza7@tigers.lsu.edu
105
Ablesh Gautam
300 Alumni Dr.
Lexington, KY 40503
United States
859-940-3903
ablesh.gautam@uky.edu
Nina Gauthier
Guelph
252 Stone Rd. W. Unit 13
Guelph, ON N1G 2V7
Canada
519-803-5004
gauthien@uoguelph.ca
Carol L. Geake
Hidden Spring Veterinary Clinic
48525 W. Eight Mile Rd.
Northville, MI 48167-9202
United States
248-349-2598
248-349-2517
cgeake29@aol.com
James Geary
Dept. of Pthobiology and Diagnostic
Investugation
United States
269-903-8099
gearyjam@msu.edu
Timothy G. Geary
McGill University
Institute of Parasitology 21,111
Lakeshore Rd.
St. Anne de Bellevue, QC H9X 3V9
Canada
514-398-7612
514-398-7857
timothy.g.geary@mcgill.ca
Claudio Genchi
University of Milan
Fac. of Vet. Medicine Via Celoria 10
Milano, 20133
Italy
+39 02 503 18101
+39 02 503 18095
claudio.genchi@unimi.it
Jinming Geng
IDEXX Laboratories
1 IDEXX Dr.
Westbrook, ME 04092
United States
207-556-6809
jinming-geng@idexx.com
David C. Gerdon
Merial, Ltd.
3239 Satellite Blvd
Duluth, GA 30041
United States
678-638-3865
david.gerdon@merial.com
Richard W. Gerhold
SCWDS College of Vet. Medicine,
501 D.W. Brooks Dr.
Athens, GA 30602-4393
United States
706-542-1741
706-542-5865
rgerhold@vet.uga.edu
Karen Gesy
23-203 Herold Terr.
Saskatoon, S7V 1H4
Canada
306-966-7132
km934@mail.usak.ca
Harold C. Gibbs
588 Kennebec Rd.
Hampton, ME 04444
United States
207-862-3578
kelizabethgibbs@aol.com
John Gilleard
16, Edenstone View
Edgemont, AB
Canada
403-375-0844
jsgillea@ucalgary.ca
Elizabeth Gleim
385 Old Epps Bridge Rd.
Athens, GA 30606
US
404-242-7346
egleim@uga.edu
Daniel Golden
Merial, Ltd.
3239 Satellite Blvd
Duluth, GA 30096
United States
678-638-3703
dan.golden@merial.com
Keith P. Goldman
400 Plaza Dr.
Secaucus, NJ 07094
United States
201-271-4800, ext. 2457
973-429-0699
kgoldman@hartz.com
kgmiddletown@aol.com
Luis A. Gomez-Puerta
Universidad Nacional Mayor de San
Marcos
Facultad de medicine Veterinaria J.
Jose Gabriel Chariarse 296
San Antonio, Miraflores, Lima,
Lima18
Peru
(511)436-8938
(511)436-5780 X 106
lucho92@yahoo.com
David G. Goodwin
College of Vet. Medicine Lab 203
1410 Prices Fork Rd.
United States
540-231-7074
540-231-3426
dagoodwi@vt.edu
Radga Gopal
600 Chatham Park Dr.
Pittsburgh, PA 15220
United States
412 692 9773
radha.gopal@chp.edu
Velmurugan Gopal Viswanathan
USDA, ARS, ANRI
4601 Tonquil St.
United States
301-504-5999
vetvelu@gmail.com
Jonas Goring
IDEXX Laboratories
1 IDEXX Dr.
Westbrooke, ME 04092
United States
613-697-5689
jonas.goring@idexx.com
Michelle Gourley
Western Michigan University
4273 Vine St.
Bridgman, MI 49106
United States
219-393-9655
michelle.l.gourley@wmich.edu
David E. Granstrom
AVMA
1931 N. Meacham Rd. Suite 100
Schaumburg, IL 60173
United States
847-285-6674
847-925-1329
dgranstrom@avma.org
dgranstrom@hotmail.com
Spencer Greenwood
Atlantic Veterinary College Dept. of
Path. & Micro. 550 University
Ave.
Canada
902-566-6002
902-566-0851
sgreenwood@upei.ca
Ellis C. Greiner
Pathobiol Box 110880
Gainesville, FL 32611-0880
United States
352-294-4161
352-392-9704
greinere@ufl.edu
John H. Greve
334 24th St.
Ames, IA 50010
United States
515-232-3520
sdgreve@earthlink.net
Robert B. Grieve
Heska Corporation
3760 Rocky Mountain Ave.
Loveland, CO 80538-9833
United States
970-493-7272
970-619-3003
bob.griever@heska.com
griever@heska.com
Matt Griffin
P.O. Box 197
Stoneville, MS, MS 38776
United States
662-686-3580
griffin@cvm.msstate.edu
Nicole Grosjean
Diagnostic Center for Poplulation
and Animal Health
1720 S Michigan Rd
Eaton Rapids, MI 48827
United States
517-432-5758
517-353-5096
grosjean@dcpah.msu.edu
Caroline Grunenwald
cgrunenw@utk.edu
106
Frank Guerino
Merck Animal Health
556 Morris Ave.
Summit, NJ 07901
United States
908-473-5532
980-473-5560
frank.guerino2@merck.com
fguerino@verizon.net
Jorge Guerrero
10 N. Riding Dr.
Pennington, NJ 08534
United States
609-510-1337
609-737-6793
jorgegu1@hotmail.com
guerrero@vet.upenn.edu
Nicole J. Guselle
Atlantic Veterinary College
Department of Pathobiology &
Micro. 550 University Ave.
Canada
902-566-0595
nguselle@upei.ca
nicoleguselle@gmail.com
Aliessia Guthrie
302-63 Queen St.
Guelph, ON NTE 4RP
Canada
aguthrie@uoguelph.ca
Richard J. Hack
Elanco Animal Health
2001 W. Main St. P.O. Box 708
Greenfield, IN 46140
United States
317-371-8614
317-277-4288
hackri@lilly.com
Sarah Hamer
Veterinary Integrative Biosciences
TAMU 4458
College STation, TX 77843
USA
979-847-5693
shamer@cvm.tamu.edu
Bruce Hammerberg
Hillsborough St.
Raleigh, NC 27606
United States
919-513-6226
919-513 6464
bruce_hammerberg@ncsu.edu
Brian Herrin
203 S. Duncan St. Apt. B
United States
405-756-7629
Brian.h.herrin@okstate.edu
Qian Han
Dept. of Biochemistry 206 Engel Hall
United States
540-231-5779
qianhan@vt.edu
qian_han@yahoo.com
James A. Hawkins
JHawkins Veternary Parasitology
LLC
118 Quail Run Dr.
Madison, MS 39110
United States
601-826 2630
jhawkins4@bellsouth.net
Pete Hann
Bayer Animal Health
3105 Jamin Paul Circle
Edmond, Oklahoma 73013
U.S.
405 471 1597
405 348 2578
pete.hann@bayer.com
Terence J. Hayes
Hoffman-La Roche, Inc.
340 Kingsland St.
Nutley, NJ 7110
United States
973-235-2120
975-235-2981
terence.hayes@roche.com
Terry Hansen
403 Highland Blvd.
Absecon, NJ
United States
203-215-7297
terry.hanson@me.com
Brad Hayes
Lilly House, Priestley Road
Basingstoke, Hampshire RG24 9NL
UK
+44 (0)773 8882007
+44 (0)1256 779515
bradleyha@elanco.com
Aaron Harmon
1447 140th St.
Larchwood, IA 51241
United States
712-477-2811, ext. 2230
aaron.harmon@novartis.com
Elizabeth Harp
Campus Mail 1878 (Biology)
United States
970-203-5462
eharp@lamar.colostate.edu
Theresa A. Hartwell
Stillmeadow, Inc.
12852 Park One Dr.
United States
281-240-8828
281-240-8448
thartwell@stillmeadow.com
au_tig77@hotmail.com
John M. Hawdon
George Washington University
Medical Center
2300 Eye St. NW 519 Ross Hall
United States
202-994-2652
202-994-2913
jhawdon@gwu.edu
Rebekah Heiry
rheiry@yahoo.com
Stephanie Heise
Fahrdorf, 24857
Germany
steph.heise@freenet.de
Klaus Hellmann
KLIFOVET, AG
Geyerspergerstr. 27
Munchen, D-80689
Germany
+49-89-58 00 82 0
+49-89-58 00 82 7777
klaus.hellmann@klifovet.de
klaus.hellmann@klifovet.com
James A. Higgins
USDA-ARS
10300 Baltimore Blvd. Building 173
United States
301-504-6443
301-504-6608
jhiggins@anri.barc.usda.gov
Michael B. Hildreth
South Dakota State University
Dept. Biol. & Micro. College of
Agriculture & Biological
Sciences NPB Room 252
Brookings, SD 57007
United States
605-688-4562
605-688-5624
michael.hildreth@sdstate.edu
Patrick Hilson
Virginia Tech
458 North June Street
Los Angeles, California 90004
USA
2136318890
patrhi13@vt.edu
John Hilson
article9ucc@netzero.net
W. Lance Hemsarth
lhemsarth@hartz.com
Cherly Ann Hirschlein
Merial, Ltd.
5970 Sycamore Rd.
Buford, GA 30518
United States
678-638-3479
678-638-3074
cah4483@aol.com
cheryl.hirschlein1@merial.com
Douglas I. Hepler
Kado Enterprises
815 Cliff Dr.
McLeansville, NC 27310
United States
336-708-2842
336-697-7296
drflea1@bellsouth.net
Sally I. Hirst
Trends in Parasitology
Elsevier 32 Jamestown Rd.
London, WC1X 8RR
United Kingdom
44-20-7611-4128
44-20-7611-4470
s.hirst@elsevier.com
Aaron Herndon
821 E. KC Ct.
United States
405-744-7000
aaron.herndon@okstate.edu
Bryanne Hoar
1617 18 ave NW
Calgary, AB T2M 0X2
Canada
403-220-4224
403-210-3939
bmhoar@ucalgary.ca
107
Eric P. Hoberg
USDA, ARS, US National Parasite
Collection
Animal Parasitic Disease BARC-East
10300 Baltimore Ave. Room 1180
Beltsville, MD 20705-2350
United States
301-504-8588
301-504-8979
eric.hoberg@ars.usda.gov
Tamara Hofstede
Bayer Animal Health
77 Belfield Rd
Toronto, ON M9W 1G6
Canada
519 743 9304
519 743 0703
tamara.hofstede@bayer.com
Johan Hoglund
Swedish Univeristy of Agricultural
Sciences
SWEPAR Ulls Vag 2B
Uppsala, 750 07
Sweden
+46 18 674156
johan.hoglund@bvf.slu.se
Mark Holbert
Stillmeadow, Inc.
Sugar Land, TX
United States
mholbert@stillmeadow.com
Patricia Holman
13009 Tall Timber Dr.
United States
979-845-4202
pholman@cvm.tamu.edu
Rush Holt
724 Louise Ave.
rholt1@mix.wvu.edu
Susan Holzmer
Zoetis
333 Portage Street
Kalamazoo, Michigan 49007
USA
269-833-4977
269-833-9584
susan.holzmer@zoetis.com
sholzmer@gmail.com
Craig Hoover
2775 Mesa Verde Dr. E.
Costa Mesa, CA 92626
United States
714-979-3398
spineheart@live.com
John Hosley
Zoetis
Kalamazoo, MI
269-833-1531
john.d.hosley@zoetis.com
john.h.adds@gmail.com
Joe A. Hostetler
Bayer Health Care
1467 N. 300 Rd.
Baldwin City, KS 66006
United States
913-268-2010
913-268-2878
joe.hostetler@bayer.com
joehostetler@embarqmail.com
Alice Houk
VA/MD Reg. College of Vet.
Medicine
Blacksburg, VA 24061-0442
United States
540-231-7074
aehouk@vt.edu
Vikki Howard
10 Charles Hill Rd.
Kittery Point, ME 39050
United States
603-498-9305
vikki.howard@novartis.com
Daniel K. Howe
Lexington, KY 40546-0099
United States
859-257-4757, ext. 812113
859-257-8542
dkhowe2@uky.edu
Sue Howell
Univ of Georgia
Dept. of Infectious Diseases
501 D.W. Brooks Dr
Athens, Georgia 30602
USA
706 542-0742
jscb@uga.edu
Jeanne Howell
jeannedvm@yahoo.com
Angela Humphreys
2973 Cresco Dr.
New Bloomfield, NJ 08902
US
573-228-1478
angelahumphreys99@gmail.com
Armando Hung
alhungc@upch.edu.pe
Frank Hurtig
Virbac
3200 Meacham Blvd.
Ft. Worth, TX 76137
United States
682-647-3529, cell: 817-360-0418
frank.hurtig@virbacus.com
fhurtig05@msn.com
Emily Jenkins
University of Saskatchewan
Department of Vet. Microbiology
Saskatchewan, SK S7N 5B4
Canada
306-966-2569
emily.jenkins@usask.ca
Douglas E. Hutchens
Bayer Animal Health, GmbH
Am Kleiansacker 3
DÃ¼sseldorf, 40489
Germany
49 1753013742
douglas.hutchens@bayer.com
hutchensde@sbcglobal.net
John Hutcheson
Merck Animal Health
7101 Red Rock Rd.
Amarillo, TX 79118
United States
806-622-1080
806-622-1086
john.hutcheson@merck.com
Marianna Ionita
University of Agronomical Sciences
& Veterinary Medicine
Splaiul Independental, No. 105,
Sector 5
Bucharest, 50097
Romania
40 21 318.04.69
40 21 318.04.98
ionitamary@yahoo.com
Nuzhat Islam
Cornell University
C4114 Veterinary Medical Center
Cornell university
Ithaca, NY 14853
USA
607-253-3394
ni34@cornell.edu
Jesica Jacobs
1339 Riddle Ave
Apt 10
USA
573-645-5900
jjacobs2@mix.wvu.edu
Philippe Jeannin
Merial, Ltd.
Parc Industriel de la Plaine de l'Ain
Allee des Cypres
Saint-Vulbas, 1150
France
+33 04 72 72 33 95
+33 04 72 72 29 57
philippe.jeannin@merial.com
Sandra S. Johnson
7000 Portage Rd.
Kalamazoo, MI 49001
United States
269-833-2660
269-833-2769
sandra.s.johnson@pfizer.com
Mark Jenkins
USDA, ARS, ANRI
BARC-East Building 1040 Room 100
United States
301-504-8054
301-504-6273
mark.jenkins@ars.usda.gov
Colin Johnstone
416 Clinton Rd.
Brookline, MA 02445-4167
United States
508-413-2107
drcjvet@aol.com
Alexandra John
University of Pennsylvania School of
Veterinary Medicine
4002 Pine Street
Philadelphia, PA 19104
USA
alexjohn@vet.upenn.edu
Helen E. Jordan
2923 Fox Ledge Dr.
United States
405-372-6420
wormdoctor@cox.net
Edward G. Johnson
24007 Highway 2026
Parma, ID 83660
United States
208-722-5829
208-722-5119
egjohnson@frontiernet.net
Richard Kabuusus
School of Vet. Medicine
Grenada
473-439-2000
rkabuusu@sgu.edu
Eileen Johnson
College of Vet. Medicine Center for
Vet. Health Science Department of
Patholbiology 250 McElroy Hall
United States
405-744-8549
405-744-5275
eileen.johnson@okstate.edu
Kathy Johnson
Purdue University
4125 Trilithon Ct.
West Lafayatte, IN 47906
United States
774-263-3825
kajohnso@purdue.edu
Mitchell A. Johnson
Intervet
35500 West 91st St.
DeSoto, KS 66018
United States
913-422-6056
mitch.johnson@intervet.com
108
Ray M. Kaplan
Infectious Diseases
Athens, GA 30602
United States
706-542-5670
706-542-0059
rkaplan@uga.edu
ray.m.kaplan@gmail.com
Kevin R. Kazacos
Purdue University
Dept. of Comparative Pathobiology
725 Harrison St.
West Lafayette, IN 47907-2027
United States
765-494-7556
765-494-9830
kkazacos@purdue.edu
Suzanne Keeys
Pac Behavior
333 James Jackson Ave.
Cary, NC 27513
United States
919-468-8301
919-468-8304
sdk@pacbehavior.com
skeeys@campcanine.com
Sarah Keller
Oklahoma State University Center
for Veterinary Health Sciences
205 McElroy Hall
USA
405-744-6633
sarah.keller@okstate.edu
Maxine F. Kellman-Allen
9418 Riverbrink Ct.
Laurel, MD 20723
United States
301-317-9859
301-317-9859
mfkallen@aol.com
Vicky Kelly
Louisiana State University School of
Veterinary Medicine
1909 Skip Bertman Dr.
Baton Rouge, Louisiana 70803
United States
4058021329
vkelly1@tigers.lsu.edu
vickyk@okstate.edu
Thomas J. Kennedy
Consultant
5492 Kennedy Drive.
Waunakee, WI 53597
United States
602 565-9716
tjameskennedy@gmail.com
tkennedy@central.com
Michael Kent
220 Nash Hall
Corvallis, OR 97331
United States
541-737-8652
michael.kent@oregonstate.edu
Stephanie Keroack
370 Missisquol N.
Bromont, QC J2L 2N5
Canada
514-606-6831
stephanie.keroack@pfizer.com
Stephanie Keroack
Zoetis Canada
370 chemin Missisquoi N
Bromont, Quebec J2L 2N5
Canada
514-606-6831
450-534-4943
stephanie.keroack@zoetis.com
stephanie.keroack@yahoo.ca
Jennifer K. Ketzis
Ross University School of Veterinary
Medicine
RUSVM
PO 334
Basseterre,
St. Kitts and Nevis
869 762 2931
jenniferketzis@yahoo.com
April Knudson
Merial, Ltd.
12056 Avenida Sivrita
San Diego, CA 92128
United States
619-417-8787
678-638-8635
april.knudson@merial.com
Muhammad Tanveer Khan
University of Vet. & Animal
Sciences
Lahore, 54000
Pakistan
35 39620856
dr_tanveerkhan@hotmail.com
Jan M. Kivipelto
Dept. of Animal Sciences 459 Shealy
Dr.
United States
352-392-3342
352-392-7652
kivipelt@ufl.edu
Thomas R. Klei
PBS School of Vet. Medicine
United States
225-578-9900
225-578-9916
klei@vetmed.lsu.edu
Ronald D. Klein
Windshadow Farm & Dairy
P.O. Box 249
Bangor, MI 49013
United States
269-599-0467
rdklein@windshadowfarm.com
ron@windshadowfarm.com
Andrew Klingsporn
St Matthews School of Veterinary
Medicine
733 Beaver rd
Glenview, IL 60025
United States
847-571-5318
klingspornandrew@gmail.com
Bruce Klink
Bayer Health Care, LLC
2508 Greenbrook Parkway
Mattews, NC 28104
United States
704-957-5589
704-844-6725
bruce.klink.b@bayer.com
Donald Knowles
USDA, ARS, WSU
3005 ADBF, ADRU
Pullman, WA 99164-6630
United States
509-335-6022
509-335-8328
dknowles@vetmed.wsu.edu
Dennis E. Kobuszewski
Summit VetPharm
8740 Iron Horse Dr.
Irving, TX 75063
United States
804-614-8358
dkobuszewski@summitvepharm.co
m
Denise Kobuszewski
Virbac Corporation
3200 Meacham Blvd.
Ft. Worth, TX 76137
USA
804-614-8358
6826473733
quarterhorsedoc@yahoo.com
denise.kobuszewski@virbacus.com
Katherine M. Kocan
College of Veterinary Health
Sciences Department of Vet.
Pathobiology 250 McElroy Hall
United States
405-744-7271
405-744-5275
kathrine.kocan@okstate.edu
Dorsey L. Kordick
IDEXX Pharmaceuticals, Inc.
4249-105 Piedmont Parkway
United States
336-834-6523
336-834-6525
dorsey.kordick@idexxpharma.com
Sarah Koske
University of Wisconsin
148 E. Gorham Apt. 3R
Madison, WI 53703
United States
413-887-8678
koske@wisc.edu
109
Lisa Kostandoff
19686 Scane Rd.
Ridgetown, ON NOP 2CO
China
lisa.kostandoff@novartis.com
Alexandra Kravitz
Cornell University
ark239@cornell.edu
Rosina C. (Tammi) Krecek
Ross University
School of Vet. Medicine 630 US
Highway 1 Suite 600
Edison, NJ 08837
United States
869-465-2405, ext. 119; mobile: 869665-3196
869-465-1203
tkrecek@cvm.tamu.edu
Timothy Krone
Pima Community College
8181 E. Irvington Rd.
Tucson, AZ 85709
United States
520.206.7414
tkrone@pima.edu
clear_prop33@yahoo.com
Eva Maria Kruedewagen
Opladener Strasse 117
Monheim, 40789
Germany
+49 2173 2036468
eva.kruedewagen@t-online.de
eva.kruedewagen@bayer.com
Ewa Kuczynska
3004 Reed Ct., Apt. 2B
Merrillville, IN 46410
United States
815-295-1509
ewa.ewak@gmail.com
Raymond E. Kuhn
Wake Forest University
Biology Department P.O. Box 7325
Winston-Salem, NC 27109
United States
336-758-5022
336-758-6008
kuhnray@wfu.edu
Daniel Kulke
Heinrich Heine Universitaet
Universitaetstr. 1 Institut fuer
Zoomorphologie und Parasit.
Duesseldorf, NRW 40225
Germany
49 2173 384232
daniel.kulke@uni-duesseldorf.de
Suresh Kumar
Allahabad Agriculture University
50, A-First Floor, Tampale Rd. Jung
Pura Bhogal
New Delhi, 110014
India
011-24374576
011-24374545
sureshkumar84@hotmail.com
nibp@hotmail.com
Bruce Kunkle
Merial, Ltd.
6498 Jade Rd.
Fulton, MO 51112
United States
bruce.kunkle@merial.com
Susan Kutz
University of Calgary Faculty of
Veterinary Medicine
3280 Hospital Dr. NW
Calgary, Alberta T2N 4Z6
Canada
403 210-3824
skutz@ucalgary.ca
William G. Kvasnicka
7131 Meadow View St.
United States
913-441-7946, cell: 775-530-2068
913-441-0937
wkvasnicka@kc.rr.com
Dongmi Kwak
Kyungpook National University
1370 Sankyug 3-dong, Buk-gu
Daegu, 702-701
Republic of South Korea
8715
6876
dmkwak@knu.ac.kr
Elise LaDouceur
Tufts University
1 Arch St.
Westborough, MA 01581
United States
443-629-8883
elise.ladouceur@tufts.edu
Dave Lambillotte
3466 Corte Sonrisa
Carlsbad, CA 92009
United States
dave@safepath.com
Rohollah Lameei
azad eslamic university urmia-iran
vesal 9,vesal avn, farhang
squ,ardabil city,iran
ardabil, ardabil 984451
iran
9.8914453305e+011
roh_vet811@yahoo.com
roh_vet811@yahoo.com
Rolando Landin
P.O. Box 22993
West Palm Beach, FL 33416
United States
roly_landin@hotmail.com
Carlos E. Lanusse
Lab de Farmacologia
Fac de Cien. Veter. Campus
Universitario Universidad Nacional
Tandil, 7000
Argentina
clanusse@vet.unicen.edu.ar
Diane Larsen
Merial, Ltd.
Duluth, GA 30519
United States
678-638-3633
678-638-3636
diane.larsen@merial.com
Mette Larsen
mella@dsc.like.ku.dk
Alice Lee
Cornell University
C4183 VMC, Campus Rd.
Ithaca, NY 14853
United States
607-253-3305
alice.cy.lee@hotmail.com
cl568@cornell.edu
Thomas Letonja
USDA APHIS VS
1210 James Rifle Ct. NE
Leesburg, VA 20176
United States
301-734-0817
301-734-3652
thomas.letonja@aphis.usda.gov
Michael G. Levy
Hillsborough St.
Raleigh, NC 27606
United States
919-513-6293
919-513-6464
mike_levy@ncsu.edu
J. Ralph Lichtenfels
USDA-ARS-ANRI
12311 Whitehall Dr.
Bowie, MD 20715-2350
United States
301-801-2465
2jrcgl@gmail.com
David R. Lincicome
11 Falls Rd.
Roxbury, CT 06783
United States
860-355-1031, 240-286-2438
sheepman@frogmoor.org
wheatens@sbcglobal.net
David S. Lindsay
Center for Molecular Medicine and
Infectious Diseases
Virginia-Maryland Regional College
of Veterinary Medicine
1410 Prices Fork Road
United States
540-231-6302
540-231-3426
lindsayd@vt.edu
Ashley Linton
Fort Collins, CO
United States
ashley.linton@colostate.edu
Janice Liotta
Cornell University
C4-114 VMC Tower Rd.
Ithaca, NY 14853
United States
607-253-3394
jll55@cornell.edu
Annette Litster
Zoetis
817 Elmwood Drive
West Lafayette, IN 47906
USA
+1 765 418 3186
Annette.Litster@zoetis.com
Susan E. Little
Center for Veterinary Health
Sciences Dept. Vet. Pathobiology
250 McElroy Hall
United States
405-744-8523
405-744-5275
susan.little@okstate.edu
Alison Little
OVC
alittl03@uoguelph.ca
110
John E. Lloyd
University of Wyoming
Dept. 3354 Renewable Resources
1000 E. University Ave.
Laramie, WY 82071
United States
307-766-2234
307-766-5025
lloyd@uwyo.edu
Heidi Lobprise
13501 Willow Creek Dr.
Haslet, TX 76052
United States
817-701-8259
Heidi.Lobprise@virbacus.com
J. Mitchell Lockhart
Valdosta State University
Biology Department 1500 N.
Patterson St.
Valdosta, GA 31698
United States
229-333-5767
229-245-6585
jmlockha@valdosta.edu
Michael Loenser
Zoetis
17 Whiteoak Ridge Road
Glen Gardner, NJ 08826
United States
908.500.2787
mike.loenser@zoetis.com
mloenser@yahoo.com
Amanda Loftis
Midwestern University
Dept of Microbiology and
Immunology
19555 N. 59th Avenue
Glendale, AZ 85308
USA
509-590-2166
adloftis@gmail.com
dr_loftis@yahoo.com
Linda L. Logan
College of Veterinary Medicine and
Biomedical Sciences, Mail stop 4467
United States
979 845-5941
llogan@cvm.tamu.edu
Joyce A. Login
Zoetis
164 Fisher Place
Princeton, NJ 08540
United States
609-356-0268
bluesdvm@comcast.net
Lori Lollis
3939 Quail Hollow Rd.
Albany, GA 31721
United States
229-869-7855
vetgal08@gmail.com
Aaron S. Lucas
Dept. of Vet. Medical Science 833
Claytor Square
United States
540-231-7917
aalucas@vt.edu
Charles D. MacKenzie
11649 Jarvis Highway
Dimondale, MI 42881
United States
517-432-3644
mackenz8@msu.edu
tropmed@mac.com
Mary Maclean
490 Barnett Shoals Rd Unit 213
Athens, GA 30605
United States
4108520682
mjmac28@uga.edu
mjmac28@gmail.com
Araceli Lucio-Forster
Cornell University
C4 103 VMC Campus Rd.
Ithaca, NY 14850
United States
607-253-4046
al33@cornell.edu
Ashley Malmlov
USA
970-297-5118
ash.malmlov@colostate.edu
Joan K. Lunney
USDA, ARS, ANRI, APDL
BARC-East, Building. 1040, Room
107
United States
301-504-9368,
301-504-5306
Joan.lunney@ars.usda.gov
John B. Malone Jr.
School of Vet. Medicine
Pathobiological Sciences
United States
225-578-9692
225-578-9701
malone@vetmed.lsu.edu
Elizabeth Lynn
Infectious Diseases
Athens, GA 30602
United States
912-237-0760
blynn210@yahoo.com
Rinosh Mani
76 S. University Pl. Apt 12
United States
405-744-8173
rinosh.mani@okstate.edu
Eugene T. Lyons
Dept. of Vet. Science Gluck Equine
Research Center
United States
859-257-4757, ext. 81115
859-257-8542
elyons1@uky.edu
Alan A. Marchiondo
Zoetis
333 Portage St.
Kalamazoo, MI 49007
United States
269-833-2674, 573-808-0203
269-833-9460
alan.marchiondo@zoetis.com
amkj46@aol.com
Martin Matisoff
Kentucky State University, Honey
Bee Laboratory
8606 Cool Brook Ct.
Louisville, KY 40291
United States
502-243-7637
martin.matisoff@huskers.unl.edu
martinmatisoff@gmail.com
Alan Marchiondo
Zoetis
333 Portage St.
USA
269-833-2674
269-833-9460
amkj46@aol.com
amkj46@aol.com
Mark Mazaleski
Zoetis Inc.
300-306SW
333 Portage St
Kalamazoo, MI 49007-4931
United States
269-833-3203
646-563-1872
mark.e.mazaleski@zoetis.com
seamark@me.com
SE Marley
Ceva Animal Health
8735 Rosehill Road
Suite 160
Lenexa, KS 66215
United States
913.945.4573
s-e.marley@ceva.com
Antoinette Marsh
1950 Westwood Ave.
Columbus, OH 43212
United States
614-282-1154
marsh.2061@osu.edu
Antoinette Marsh
The Ohio State University
1950 Westwood Ave
Columbus, OH 43212
USA
614.282.1154
Antoinette.Marsh@cvm.osu.edu
Claire Mannella
3200 Northline Ave. Suite 300
United States
800-477-2391, ext. 1254
cmannellavet@yahoo.com
Linda S. Mansfield
B43 Food Safety Toxicology Building
A.J. MacInnis
University of California - Los Angeles United States
517-432-3100, ext. 119
Biology Department P.O. Box
517-432-2310
951606
mansfie4@cvm.msu.edu
Los Angeles, CA 90095-1606
United States
310-825-3069
Abdelmoneim Mansour
310-206-3987
TRS Labs, Inc.
mac@biology.ucla.edu
295 Research Dr.
Athens, GA 30605
United States
706-354-1531
abdelmnsr@yahoo.com
Randall J. Martin
Merck Animal Health
10116 Tapestry St.
Forth Worth, TX 76244
United States
800-224-5318
908-337-6417
randy.martin@merck.com
Richard J. Martin
Biomedical Sciences Room 2008
Ames, IA 50011-1250
United States
515-294-2470
515-294-2315
rjmartin@iastate.edu
111
Morgan McArthur
3230 S. 149th St.
New Berlin, WI 53151
United States
414-588-4450
morgan@morganmc.com
William McBeth
812 Springdale Dr.
Exton, PA 19341
United States
610-968-3558
william.mcbeth@pfizer.com
John W. McCall
Infectious Diseases TRS Labs P.O.
Box 5112
Athens, GA 30602
United States
706-542-3945
706-542-3804
jwmccall@uga.edu
Scott McCall
TRS Labs, Inc.
P.O. Box 5112
Athens, GA 30604
United States
706-549-0764
706-353-3590
scmccall@bellsouth.net
trslabs@bellsouth.net
Austin McCall
P.O. Box 5112
Athens, GA 30604
United States
706-769-9519
camfam6@bellsouth.net
Scott McCall
TRS Labs
PO Box 5112
Athens, GA 30604
USA
706-549-0764
706-353-3590
scmccall90@bellsouth.net
James McClanahan
What A Relief
8726 E. Solano Dr.
Scottsdale, AZ 85250-6319
United States
602-317-8443
480-626-7295
mrjimmy2@hotmail.com
Christine M. McCoy
2500 Innovation Way
United States
317-651-4377
mccoycm@elanco.com
ilovefleas@gmail.com
Ashley McGrew
202 E. Elizabeth St.
Ft. Collins, CO 80524
US
303-941-8440
ashlinton98@hotmail.com
Kirsten McGuire
312 Sunset Ave.
Strasburg, PA 17579
US
717-419-4923
keamcguire@yahoo.com
Mary Elizabeth McKenzie
150 E. 42nd St.
New York, NY 10017
United States
212-573-5438
212-573-5348
m.elizabeth.mckenzie@pfizer.com
Michael McMenomy
Kitty Klinic
3447 Lyndale Ave. S.
Minneapolis, MN 55408
United States
612-822-2135
kittyklinic@earthlink.net
Tom McTier
Zoetis Inc
333 Portage Rd
Kalamazoo, MI 49007
United States
269-833-3978
tom.mctier@zoetis.com
Lisa Meader
11705 West Main
Whitmore Lake, MI 48187
United States
llmeader@gmail.com
America Mederos
54 University Ave.
Guelph, ON N1G 1N7
Canada
amederos@ovc.uogurlph.ca
Patrick F.M. Meeus
Zoetis
VMRD, 333 Portage Rd.
Kalamazoo, MI 49007
United States
269-833-2661
patrick.meeus@zoetis.com
pmeeus69@gmail.com
Thomas Miller
VRRC inc
P.O. Box 4142
Sequim, WA 98382
USA
360-582-0985
360-582-985
vrc@olypen.com
Heather Moberly
4462 TAMU
USA
979-847-8624
979-845-7493
hmoberly@tamu.edu
Daniel K. Miller
2209 Mayer Ave.
Lancaster, PA 17603
United States
717-393-7016
danmiller@hotmail.com
milldaniel@gmail.com
Osama Mohammed
King Saud University
Department of Zoology, College of
Science, P.O. Box 2455, Riyadh
11451
Riyadh,
Saudi Arabia
96614675756
obmkkwrc@yahoo.co.uk
omohammed@ksu.edu.sa
Melissa Miller
UGA CVM
501 D.W. Brooks Dr.
Athens, GA 30602
USA
8139282437
Millerm@uga.edu
Shelley Mehlenbacher
Bayer Health Care
P.O. Box 83158
Portland, OR 97283
United States
503-247-9108
Zachary T. Mills
shelley.mehlenbacher.b@bayer.com Merial, Ltd.
3239 Satellite Blvd. Building 500
Room 362
Norbert Mencke
Duluth, GA 30041
Bayer Health Care, AG
United States
Bayer Animal Health
678-638-3834
Kaiser-Wilhelm-Allee 50
678-638-3873
Global Veterinary Services - CAP
zack.mills@merial.com
Leverkusen, 51373
Germany
+49 2173 384921
Pamela S. Mitchell
+49 2173 389694921
norbert.mencke@bayer.com
mencke_norbert@yahoo.com
United States
800-447-2391, ext. 1691
Jeffrey A. Meyer
504-891-8716
Triveritas
pamela.mitchell@novartis.com
P.O. Box 1066
Sheila M. Mitchell
United States
317-318-2083
TyraTech
jeff.meyer@triveritas.com
Durham, NC
United States
919-316-8107
James E. Miller
shmitch2@gmail.com
School of Vet. Medicine Dept. of
Ioan Liviu Mitrea
University of Agronomical Scienses
United States
& Veterinary Medicine
Splaiul Independental, No. 105,
225-578-9652
Sector 5
225-578-9701
jmille1@lsu.edu
Bucharest, 50097
Romania
0040-21-318.04.69
Thomas A. Miller
0040-21-318.04.98
P.O. Box 4142
liviumitrea@yahoo.com
Sequim, WA 98382
United States
360-582-0985
360-582-0986
vrrc@olypen.com
112
Osama Mohammed
King Saud University
Department of Zoology, College of
Science, King Saud University,
POBox 2455, Saudi Arabia
Riyadh, Central Region 11451
USA
9.6650414071e+011
dmccleary@cvm.tamu.edu
omohammed@ksu.edu.sa
Susan Montgomery
CDC
308 Ferguson St. NE
Atlanta, GA 30307
United States
770-488-4520
smontgomery@cdc.gov
Andrew R. Moorhead
College of Vet. Medicine Dept. of Inf.
Diseases 501 D.W. Brooks Dr.
Athens, GA 30602
United States
706-542-8168
amoorhed@uga.edu
Gail Moraru
215 Woodlawn Rd.
Starkville, MS 39759
United States
831-419-0998
moraru@cvm.msstate.edu
Rebecca Morningstar
1781 Nemoke Trl. Apt. 1
Haslett, MI 48840
United States
231-384-5879
rmorning@msu.edu
Ntombi Mudenda
375 W Roosevelt St
Apt 1232
USA
2252215536
nmuden1@tigers.lsu.edu
Jean Mukherjee
Tufts University
200 Westboro Rd.
North Grafton, MA 01536
United States
508-887-4756
508-839-7911
jean.mukherjee@tufts.edu
Jean.Mukherjee@gmail.com
Thomas Murphy
Central Veterinary Research
Laboratory
Backweston Campus Young's
Crossing
Celbridge, Co. Kildare,
Ireland
3531615-7141
tom.murphy@agriculture.gov.ie
Martin Murphy
Charles River, Ballina
Co Mayo,
Ireland
35 3874170323
martin.murphy@crl.com
Tom Nelson
American Heartworm Society
719 Quintard Ave
Anniston, AL 36201
United States
256-236-8387
256-236-5888
drtomnelson@amcvets.com
Eva K. Nace
CDC
4770 Buford Hwy. Suite F-22
Atlanta, GA 30341
United States
770-488-4414
770-488-7761
enace@cdc.gov
ebk5@cdc.gov
Soraya Naem
Urimia University
Fac. Vet. Medicine Nazloo Campus
P.O. Box 1177
Urmia, 37135
Iran
98(441)2776142
98(441)3443442
s.naem@mail.urmia.ac.ir
Yoko Nagamori
130 Apple Pl.
Ames, IA 50010
United States
605-371-6267
yoko@istate.edu
Yoko Nagamori
USA
605-371-6267
yokon@okstate.edu
czesc96@hotmail.com
C. David Nash
Gallatin, TN 37066
K. Darwin Murrell
United States
Department of Veterinary Disease
615-714-7714
Biology, University of Copehagen
Center for Experimental Parasitology dnash@norbrookinc.com
5126 Russett Rd.
Rockville, MD 20853
Sundar Natarajan
United States
10403 46th Ave. Apt. 302
301-460-9307
kdmurrell@comcast.net
United States
301-443-2376
Rainer K. Muser
drnsundar@gmail.com
18 Montgomery Ave.
Rocky Hill, NJ 08553
Muhammad Mudasser Nazir
United States
Virginia Tech.
609-924-1802
1401 Prices Fork Road,
rmuser2391@aol.com
Blacksburg, Virginia 24061
USA
-9431
Gil Myers
mudasser.nazir@uvas.edu.pk
Gil Myers, Ph.D., Inc.
mmnazir@vt.edu
3289 Mt. Sherman Rd.
Magnolia, KY 42757
United States
Mohammed Nazir
270-324-3811
drmudasserch@yahoo.com
270-324-3811
gmyersph@scrtc.com
Harold Newcomb
Intervet
200 Watt St.
Batesville, MS 38606
United States
662-609-6364
harold.newcomb@intervet.com
Bruce Nosky
Merial, Ltd.
P.O. Box 25485
Scottsdale, AZ 85255
United States
480-747-3193
bruce.nosky@merial.com
Larry Nouvel
LNouvel
4657 Courtyard Trail
Plano, Texas 75024
USA
469-223-2854
630-982-6433
lnouvel@covad.net
lnouvel@lnouvel.com
Brain Newman
Medicine
United States
240-893-8473
bmn1127@vt.edu
Samantha O'Brien
2500 Innovation Way
USA
3174989632
samantha.o'brien@elanco.com
Martin K. Nielsen
Lexington, KY
US
859-218-1103
859-257-8542
martin.nielsen@uky.edu
Tom O'Connor
IDEXX Laboratories
1 IDEXX Dr.
Westbrook, ME 04092
United States
207-856-0428
tom-oconnor@idexx.com
Martin Nielsen
319 M.H. Gluck Equine Research
Center,
Department of Veterinary Science
Lexington, Kentucky 40546-0099
USA
859 218 1103
859 257 8542
mknielsen72@gmail.com
Mosun Ogedengbe
Department of Pathobiology, Ontario
Veterinary College.
University of Guelph.50 Stone Road
East.
Guelph, Ontario N1G 2W1
Canada
5198302000
mogedeng@uoguelph.ca
mosun4d@yahoo.com
Robert H.R. Nixon
Pfizer Animal Health Canada
17300 Trans-Canada Hwy.
Kirkland, QC H9J 2M5
Canada
519-432-5502
514-693-4272
rob.nixon@pfizer.com
shannon.topinka@pfizer.com
Thomas J. Nolan
School of Vet. Medicine, 3800
Spruce St.
Philadelphia, PA 19104-6050
United States
215-898-7895
215-573-7023
parasit@vet.upenn.edu
113
Ryan O'Handley
Murdoch University
54 Dotterel Way
Yangebup, WA 6150
Australia
+61 8 9360 2457
+61 8 9310 4144
rohandley@hotmail.com
Pia Untalan Olafson
USDA, ARS
Knipling-Bushland US Livestock
Insects Research Lab 2700
Fredericksburg Rd.
Kerrville, TX 78028
United States
830-792-0322
830-792-0314
pia.olafson@ars.usda.gov
Lauren Olavessen
Merial
8002 Jills Creek Drive
Bartlett, TN 38133
USA
901-356-9965
lauren.olavessen@merial.com
Donya D. Olcott (Dupree)
Sherwood Animal Clinic
101 Hursel Sanner Rd
Hackberry, LA 70645
United States
225-931-8703
ladyvet@gmail.com
Itziar Olea
Oxoid, S.A.
Calle Jose Bergamin, 40, 8-D
Madrid,
Spain
34 913711644
itziar.olea@oxoid.com
Dan A. Ostlind
Merck & Co. (retired)
94 Dearhook Rd
Whitehouse Station, NJ 08889
United States
908-236-9238
908-236-9238
stanton94@embarqmail.com
Nadia Ouellette
Atlantic Veterinary College
550 University Ave.
Canada
902-569-6096
nouellette@upei.ca
nadia-o@hotmail.com
Paul A.M. Overgaauw
Utrecht University Netherlands
Utrecht University
IRAS Div. VPH
PO Box 80175
Utrecht, 3058 TD
The Netherlands
0031-6532-60696
0031-30-253-2365
p.a.m.overgaauw@uu.nl
p.a.m.overgaauw@gmail.com
Brooke Pace
812 Springdale Dr.
Exton, PA 19341
United States
610-968-3369
866-590-1149
brooke.pace@pfizer.com
dalsncows@zoominternet.net
Max Parkanzky
Purdue University College of
Veterinary Medicine
Veterinary Pathology Building,
Room 3
725 Harrison Street
West Lafayette, Indiana 47909-2027
USA
765-494-7558
maxparkanzky@purdue.edu
parkanz2@gmail.com
Azhahianambi Palavesam
UMBI
18763 Nathans Pl.
montgomery vill, MD 20886
United States
202-664-0619
nambibio@gmail.com
palavesa@umbi.umd.edu
Jelena Palic
Dept. Vet. Pathobiology 1789
College Vet. Medicine
Ames, IA 50011
United States
515-294-0959
jelena@iastate.edu
Jelena Palic
Medizinische Kleintierklinik, Faculty
of Veterinary Medicine, Ludwig
Maximilians University
Guttenbrunner Weg 15
Munchen, 81829
Germany
4.9160376892e+011
J.Palic@medizinischekleintierklinik.de
jelena@iastate.edu
James C. Parsons
CSIRO Livestock Industries - AAHL
475 Mickleham Rd.
Attwood, Victoria 3049
Australia
61-3 92174275
61-3 92174161
jim.parsons@dpi.vic.gov.au
Tara Paterson
P.O. Box 7, True Blue
St. George's,
Grenada
473-435-2900
473-435-2997
tpaterson@sgu.edu
John A. Pankavich
American Cyanamid (retired)
513 Thompson Ave.
Lehigh Acres, FL 33972
United States
239-368-3394
pankavichjm@msn.com
Carla C. Panuska
6100
United States
662-325-1198
662-325-1031
panuska@cvm.msstate.edu
Sharon Patton
University of Tennessee
College of Vet. Medicine 2407 River
Dr.
Knoxville, TN 37996-4543
United States
865-974-5645
865-974-5640
spatton@utk.edu
Kelsey Paras
The Ohio State University College of
Veterinary Medicine
paras.4@buckeyemail.osu.edu
Sujatha Parimi
Midwest Veterinary Services, Inc.
1290 County Rd. M Suite A
OAKLAND, NE 68045
United States
402-685-6587
sujatha@mvsinc.net
Sarah Parker
CFIA Saskatoon Lab
116 Veterinary Rd.
Saskatoon, SK S7N 2R3
Canada
306-975-5996
306-975-5711
sarah.parker@usask.ca
Michael Paul
CAPC
1210 Vance Ct.
Bel Air, MD 21014
United States
510-315-0550
410-838-8530
solucky@aol.com
mapaul.dvm@gmail.com
allan paul
univ of illinois college of vet med
2001 s lincoln
urbana, il 61802
usa
217-333-2907
ajpaul@illinois.edu
114
Patricia A. Payne
College of Vet. Medicine, 3005
Payne Dr.
Manhattan, KS 66503
United States
785-532-4604
payne@vet.k-state.edu
Andrew S. Peregrine
Ontario Veterinary College
Department of Pathobiology
Guelph, ON N1G 2W1
Canada
519-824-4120, ext. 54714
519-824-5930
aperegri@ovc.uoguelph.ca
Adalberto A. Perez de Leon
ARS, USDA
2700 Fredericksburg Rd.
Kerrville, TX 78028
United States
830-792-0304
beto.perezdeleon@ars.usda.gov
Raffaele Petragli
Via F.lli Rosselli, 27
Montescudaio PI, PXX 56040
Italy
32 02723102
raffaele@leishmania.it
John E. Phillips
Zoetis
2200 NW 23rd Way
Boca Raton, FL 33431
United States
561-995-1648
561-995-0574
john.phillips@zoetis.com
Celine Picard
Bayer Animal Health
22 Brisson St.Box 235
Limoges, ON K0A 2M0
Canada
613-443-7664
613-443-5364
celine.picard.b@bayer.com
Chris Pinard
Ontario Veterinary College
#310 - 26 Ontario Street
Guelph, Ontario N1E 7K1
Canada
519-546-4463
pinardc@uoguelph.ca
pinardchris@gmail.com
Rhonda D. Pinckney
School of Vet. Medicine Department
of Paraclinical Studies True Blue
St. George's,
Grenada
473-444-4175, ext. 3671
473-439-5068
rpinckney@sgu.edu
pinckney.rhonda2@gmail.com
Jimmy Pitzer
New Mexico State University
P.O. Box MSC 3 BF
Las Cruces, NM 88003
United States
575-571-6980
jpitzer@nmsu.edu
Edward G. Platzer
University of California - Riverside
Dept. of Nematology 4205 Carney
Ct.
Riverside, CA 92521
United States
951-827-4352
edward.platzer@ucr.edu
Kenneth C. Plaxton
Elsevier
Molenwerf 1
Amsterdam, 1014 AG
The Netherlands
31 20 485 3332
31 20 485 3249
k.plaxton@elsevier.com
Matthew Playford
P.O. Box 1118
Camden, 2570
Australia
61 246556464
matt@dawbuts.com
Victoria Polito
16 Parkman St.
Boston, MA 2122
United States
617-543-2552
victoria.polito@okstate.edu
Dana Pollard
dpollard@cvm.tamu.edu
Stephanie Ponder
1707 W. Spring Creek Pkwy.
Plano, TX 75012
United States
sponder@central.com
Fulton, MO 65251
United States
573-642-5977, ext. 1168
573-642-0356
joseph.prullage@merial.com
Linda Marie Pote
6100 Wise Center Spring St.
United States
662-325-1130
662-325-8884
lpote@cvm.msstate.edu
David G. Pugh
Fort Dodge Animal Health
P.O. Box 26
Waverly, AL 36879
United States
334-826-8473
334-826-8473
pughdag@fdah.com
Kerrie Powell
SCYNEXIS, Inc.
P.O. Box 12878
Durham, NC 27713
United States
919-206-7223
919-313-6649
kerrie.powell@scynexis.com
Cassan Pulaski
Louisiana State University School of
Veterinary Medicine Dept of
2512 McGrath Ave
Apt 26
United States
985-590-8416
cpulaski@vetmail.lsu.edu
Kayla Price
5771 Third Line Eramosa RR #33
Rockwood, ON N0B 2K0
Canada
519-824-4120
pricek@uoguelph.ca
Rebecca Price
1168 Agricultural Sciences Building
Morgantown, West Virginia 26505
United States
240-727-1551
rcprice32@aol.com
rcprice@mix.wvu.edu
Roger K. Prichard
McGill University
MacDonald Campus 21111
Lakeshore Rd.
Ste-Anne-de-Bel, QC H9X 3V9
Canada
514-398-7729
514-398-7594
roger.prichard@mcgill.ca
Helen Profous-Juchelka
Merck Sharp & Dohme Corp.
126 East Lincoln Ave.
Rahway, NJ 07065
United States
732-594-5569
732-594-3624
helen_profous@merck.com
Frank L. Prouty
9225 Indian Creek Pky Suite 400
United States
913-664-7041
913-217-6872
frank.prouty@pfizer.com
Shon Pulley
2500 Innovation Way
Greenfield, Indiana 46140
USA
317-277-2187
317-655-0394
pulley_shon_x1@elanco.com
Trisha Putro
Bristol-Myers Squibb
Route 206 & Provinceline Rd.
Princeton, NJ 08543
United States
609-252-5557
609-252-6896
trisha.putro@bms.com
Tariq Qureshi
Bayer Animal Health
490 Franklin Cir.
Yardley, PA 19067
United States
913-268-2070
913-268-2541
tariq.qureshi@bayer.com
t7qureshi@comcast.net
Tariq Qureshi
t7qureshi@gmail.com
Alex Raeber
Prionics AG
Wagistrasse 27A
Schlieren, Zurich, 8952
Switzerland
4144 200-2000
alex.raeber@prionics.com
Joseph B. Prullage
Merial, Ltd.
6498 Jade Rd.
115
Manish Rai
c/o Nitin Raj, 128/13-C 3rd Fl.
New Delhi, 110014
IN
11 26348328
nibp@hotmail.com
Robert H. Rainier
10720 Club Chase
Fishers, IN 46038
United States
317-598-9120
317-598-9120
bnrain@sbcglobal.net
Brenda J. Ralston
Alberta Agriculture Food & Rural
Development
97 East Lake Ramp NE
Airdrie, AB T4A 0C3
Canada
403-948-8545
403-948-2069
brenda.ralston@gov.ab.ca
Siva Ranjan
8 Wheatston Ct.
Princeton, NJ 08550
United States
609-799-7746
sivaja.ranjan@gmail.com
Vijayaraghara Rao
McGill University
2111 Lakeshore Ste. Anne de Bell.
Montreal, QC H9X 349
Canada
vijayaraghara.rao@mail.mcgill.ca
Lisa Rascoe
Auburn University
College of Vet. Medicine 151 Greene
Hall
Auburn, AL 36849
United States
251-554-8948
334-844-2652
rascoln@aubirn.edu
Thilakar Rathinam
1260 W Maple St
Department of Poultry Science
Fayetteville, Arkansas 72701
USA
thilak@uark.edu
Mason Reichard
of Pathobiology 250 McElroy Hall
United States
405-744-8159
405-744-5275
mason.reichard@okstate.edu
Craig R. Reinemeyer
East Tennessee Clinical Research,
Inc.
80 Copper Ridge Farm Rd.
Rockwood, TN 37854
United States
865-354-8420
865-354-8421
creinemeyer@easttenncr.com
Renne Rember
P.O. Box 83
Ester, AK 99725
United States
907-978-7453
docney@gmail.com
Robert S. Rew
Research Consulting
400 N. Wawaset Rd.
West Chester, PA 19382
United States
610-918-1248
484-356-0452
rewnr@yahoo.com
Heather Rhoden
1114 S. Duck St.
US
402-366-1369
hrtracy@okstate.edu
Elizabeth Rich
Lincoln Memorial University
6965 Cumberland Gap Pkwy
Harrogate, TN 37752
USA
(423) 869-3611
Elizabeth.c.rich@gmail.com
Robert K. Ridley
College of Vet. Medicine Coles Hall
Manhattan, KS 66506
United States
785-532-4615
785-532 4851
ridley@vet.ksu.edu
Sherri Rigby
Bayer Healthcare
804 Greentree Arch
Virginia Beach, Virginia 233451
United States of America
757-575-6711
sherri.rigby@bayer.com
G.C. Ritchie
6741 Layton Rd.
Liberty, NC 27298
United States
336-402-4831
gc.richie@novartis.com
William Robbins
550 University Ave.
Charlottetown, Prince Edward Island
C1A 4P3
Canada
902-566-0787
wtrobbins@upei.ca
Douglas Ross
Bayer Animal Health
P.O. Box 390
Shawnee Mission, KS 66201-0390
USA
913/991-8268
913/268-2541
douglas.ross@bayer.com
Tiffany Rushin
Virginia-Maryland Regional College
of Veterinary Medicine
Center for Molecular Medicine and
Infectious Disease
trushin@vt.edu
Alan P. Robertson
2058 College of Vet. Medicine
Ames, IA 50011
United States
515-294-1212
515-294-2315
alanr@iastate.edu
Mary G. Rossano
255 Handy's Bend Rd.
Wilmore, KY 40390
United States
859-257-7552
859-323-1027
mary.rossano@uky.edu
Bill Ryan
Ryan Mitchell Associates
16 Stoneleigh Park
Westfield, NJ 07090-3306
United States
908-789-2095
wgryan@yahoo.com
Jessica Rodriguez
4467 TAMU
US
713-594-2914
jyrodriguez@cvm.tamu.edu
Alexa Rosypal
University of North Carolina at
Chapel Hill
Department of Path./Lab. Med. CB
#7525, 805 Brinkhous-Bullitt
Chapel Hill, NC 27599-7525
United States
919-966-4294
919-966-0704
arosypal@email.unc.edu
R. Rodriquez
2106 Sugar Hill Dr.
Deer Park, TX 77536
United States
713-594-2914
jrod.dvm@gmail.com
Erin Rogers
Oklahoma State University Center
for Veterinary Medicine
3398E 6th AVe Apt K
Stillwater, oklahoma 74074
United States
2147739049
erin.r.rogers@okstate.edu
Raffaele A. Roncalli
29 Louise Rd.
Milltown, NJ 08850
United States
732-940-4070
732-940-7881
r.roncalli@gmail.com
Douglas Ross
Bayer Health Care
Animal Health Division 12809
Shawnee Mission Parkway
United States
913-268-2828
913-268-2541
douglas.ross.b@bayer.com
Alexa Rosypal
Johnson C. Smith University
100 Beatties Ford Rd
Charlotte, NC 28216
USA
704-378-1206
acrosypal@jcsu.edu
arosypal@gmail.com
Emily Rubinson
Maxwell H.Gluck Equine Research
Center
Department of Veterinary Science
United States
(859) 257-4757
(859) 257-8542
efru222@uky.edu
Tony Rumschlag
614 White Pine Dr.
Noblesville, IN 46062-8533
United States
317-773-5181
317-773-1224
tony.rumschlag@merial.com
Tony Rumschlag
2500 Innovation Way
USA
317-997-2386
rumschlag.a@elanco.com
116
Meriam Saleh
US
msaleh1@vt.edu
Nina Samoylovskaya
All -Russian institute helminthology
named K.I.Skryabin
117418 Tsurupi 17-4 Moscow
Russia
Moscow, 117418
Russia
79262250063
74991245655
ninasamoylovskaya@gmail.com
rhodiola_rosea@mail.ru
Justin Sanders
220 Nash Hall
Corvallis, OR 97331
USA
541-737-1859
Justin.Sanders@oregonstate.edu
Tejbir Sandhu
1618 3rd St Cir E
Palmetto, FL 34221
United States
951-708-1456
tejbirsandhu@yahoo.com
Gursimrat Sandu
afh10gsa@student.lu.se
Nicholas C. Sangster
Charles Sturt University
School of Agricultural and Veterinary
Sciences Locked Bag 588
Wagga Wagga, NSW 2678
Australia
02 6933 4107
02 6933 2812
nsangster@csu.edu.au
Monica Santin-Duran
USDA, ARS, ANRI, EMSL
BARC-East 10300 Baltimore Ave.
Building 173
United States
301-504-6774
301-504-6608
msantin@anri.barc.usda.gov
monica.santin-duran@ars.usda.gov
Raj Saran
DuPont
5 Tether Vourt
Wilmington, DE 19808
United States
302-366-5357
raj.k.saran@usa.dupont.com
Mohamed Z. Satti
St. Matthew's University
School of Vet. Medicine 6504
Kenneaw Rd.
Canton, MI 48187
United States
345-927-5752
mohd_satti@hotmail.com
Rebecca Sayre
P.O. Box 104
Porta Costa, CA 94569
United States
209-603-0270
US
rssayre@cvm.tamu.edu
Rudolf Schenker
Lanzenbergstrasse 4
Magden,
Switzerland
41 61 843 01 97
ruedischenker@sunrise.ch
Philip J. Scholl
5138 Neptune Cir.
Oxford, FL 34484
US
352-399-5249
pjhvs@aol.com
Jennifer Schori
Educational Concepts
2021 S. Lewis Ave., Ste. 760
Tulsa, OK 74104
USA
856-935-1156
918-749-1987
dr.jen@cliniciansbrief.com
jlschori@msn.com
Bettina Schunack
Schwarzwaidalle 215
Basel, 4058
Switzerland
4179 8260307
bettina.schunack@novartis.com
George C. Scott
800 Hessian Cir.
United States
Alireza Sazmand
610-793-1852
No. 512, Atlasi Apartment, Atlasi Sq., gcsascott@aol.com
Safaiyeh
Yazd, Yazd 8915865359
Jeffrey Scott
Iran
Cornell University
Alireza_sazmand@yahoo.com
4 Ayla Way
Ithaca, NY 14850
John Schaefer
United States
Cornell University
607-255-7723
258 Bundy Rd.
jgs5@cornell.edu
Ithaca, NY 14850
United States
R. Lee Seward
607-273-2939
Seward Farms, LLC
js967@cornell.edu
P.O. Box 6
Eaton, CO 80615-0006
James H. Schafer
United States
Schafer Veterinary Consultants
970-834-0216
800 Helena Ct.
970-834-0216
sewardfarm@aol.com
United States
970-224-5103
Jennifer Sexsmith
970-224-5882
jschafer@schaferveterinary.com
1700 Allard Ave.
Grosse Pointe Woods, MI 48236
Peter M. Schantz
United States
2856 Woodland Park
313-401-3992
Atlanta, GA 30345
jennifer.sexsmith@novartis.com
United States
404-982-0591
pschantz@comcast.net
Karen Shearer
Zoetis
R.R.#1
Buckhorn, ON K0L 1J0
Canada
705-657-3043
705-657-3043
karen.shearer@zoetis.com
Greg L. Simons
11205 W. 140th Pl.
United States
800-447-2391
913-851-0472
greg.simons@novartis.com
SungShik Shin
Chonnam National University
Yongbong-Dong
Gwangju, 500-757
Republic of Korea
+82 62 530 2860
+82 62 530 2809
sungshik@jnu.ac.kr
vetpia@hanmail.net
Hal R. Sinclair
Teva Animal Health, Inc.
8609 N.W. Shannon Ave.
Kansas City, MO 64153
United States
816-676-6108
816-676-6877
hal.sinclair@tevausa.com
Barbara Shock
313 Oak Grove Dr.
Grantsville, WV 26147
United States
706-542-1741
bshock@uga.edu
Jeffrey Shryock
Merial
6498 Jade Rd.
Fultom, MO 65251
USA
573-642-5977
jeffrey.shryock@merial.com
Rick Sibbel
Intervet/Schering-Plough AH
1401 NW Campus Dr.
Ankeny, IA 50023
United States
515-249-2111
rick.sibbel@sp.intervet.com
Gary Sides
13355 CR 37
Sterling, CO 80751
United States
970-520-5953
gary.sides@pfizer.com
Everett E. Simmons
Burnet Road Animal Hospital
8511 Burnet Rd.
Austin, TX 78757
United States
512-452-7606
rtoth9196@aol.com
Anthony Simon
150 East 42nd St.
New York, NY 10017
United States
646-509-9140
tony.simon@pfizer.com
117
Andrew Skidmore
Merck Animal Health
32 Bountiful Dr
Hackettstown, NJ 07840
USA
7164742715
andrew.skidmore@merck.com
andrewskidmore57@gmail.com
Terry Skogerboe
81 Rivercrest Dr.
Pawcatuck, CT 06379
USA
tskogerboe@comcast.net
Frank Slansky
Department of Entomolgy &
Nematology Building 970 P.O. Box
110620
United States
352-332-2001
352-392-0190
fslansky@ufl.edu
Owen Slocombe
Ontario Vet. College 29 Oak St.
Guelph, ON N1G 2N1
Canada
519-821-7222
519-824-5930
oslocomb@uoguelph.ca
Robyn L. Slone
PLRS
1251 NC 32 North
Corapeake, NC 27926
United States
252-465-8686
252-465-4493
rslone@embarqmail.com
Larry L. Smith
Larry L. Smith DVM Research &
Development, Inc.
108 Davis St.
Lodi, WI 53555
United States
608-592-3275
608-592-5118
smithoffice@charter.net
Joe Smith
University of California Davis
joesmith@ucdavis.edu
Karen Snowden
College of Vet. Medicine, Dept. of
Vet. Pathobiology, Mail Stop 4467
US
979-862-4999
979-862-2344
ksnowden@cvm.tamu.edu
Daniel E. Snyder
2500 Innovation Way
United States
317-277-4439
312-277-4167
snyder_daniel_e@elanco.com
drdan@iquest.net
Mark D. Soll
Merial, Ltd.
Duluth, GA 30096-4640
United States
678-638-3605
678-638-3668
mark.soll@merial.com
Karen Sommers
G050 Agricultural Sciences Building
PO Box 6108
United States
740-310-6998
ksommer2@mix.wvu.edu
Karine Sonzogni-Desautels
McGill University
3730 Boul. Laurier Est
Saint-Hyacinthe, J2R 2B7
CN
514-398-7608
karine.sonzognidesautels@mail.mcgill.ca
Julia Steinke
1509 Stover St.
Fort Collins, CO
United States
jrsteink@gmail.com
Smaragda Sotiraki
NAGREF-VRI
NAGREF Campus Thermi
Thessaloniki, 57001
Greece
30231 936-5373
smaro_sotiraki@yahoo.gr
Maci Stephens
m.stephens@uky.edu
Jennifer Spencer
Auburn University
of Pathobiology 166 Greene Hall
Auburn, AL 36849
United States
334-844-2701
334-844-2652
spencja@vetmed.auburn.edu
Madeleine S. Stahl
Intervet, Inc.
29160 Intervet Ln. P.O. Box 318
Millsboro, DE 19966
United States
302-934-4362
302-934-4281
madeleine.stahl@sp.intervet.com
David G. Stansfield
United States
336-255-0201
336-387-1030
drdavid_s@yahoo.com
david.stansfield@pfizer.com
Lindsay Starkey
250 McElroy Hall
United States
405-744-3236
lindsay.starkey@okstate.edu
Michael Stegemann
Pfizer, Ltd.
Animal Health Group IPC 896,
VMR&D Ramsgate Rd.
Sandwich, CT13 9NJ
United Kingdom
+44 1304 646121
+44 1304 656257
michael.stegemann@pfizer.com
Joyce Steinbock
BioMed Diagnostics, Inc.
1388 Antelope Rd.
White City, OR 97503
United States
541-830-3000
541-830-3001
joyces@biomeddiagnostics.com
Ashley Steuer
University of Tennessee College of
Veterinary Medicine
6315 Kingston Pike Apt. 1406
Knoxville, Tennessee 37919
USA
(269) 352-5670
asteuer@utk.edu
Jay Stewart
CAPC
P.O. Box 230
aumsville, OR 97325
United States
503 448 1855
cjstewar@hotmail.com
T. Bonner Stewart
5932 Forsythia Ave.
United States
225-766 8327
jstewa11@bellsouth.net
Roger Stich
Dept. of Pathobiology 209
Connaway Hall
Columbia, MO 65211
United States
573-882-3148
stichrw@missouri.edu
Heather D. Stockdale
College of Vet. Medicine 2484 Royal
Point Dr.
Green Cve, FL 32043
United States
352-294-4142
hdstockdale@vetmed.ufl.edu
Heather Stockdale Walden
2484 Royal Pointe Dr.
Green Cove Springs, FL 32043
United States
904-282-2591
hdstockdale@ufl.edu
118
Bob Storey
College of Vet. Medicine
501 D.W. Brooks Drive
Athens, GA 30602
United States
706-542-0195
bstorey@uga.edu
Bert E. Stromberg
University of Minnesota
1971 Commonwealth Ave.
St. Paul, MN 55108
United States
612-625-7008
612-625-5203
b-stro@umn.edu
Chunlei Su
University of Tennessee - Knoxville
701 Dawson Creek Ln.
Knoxville, TN 37922-9007
United States
612-625-7008
612-625-5203
csu1@utk.edu
Miguel T. Suderman
Cell Systems 3-D, LLC
351 Columbia Memorial Pkwy. Suite
C-2
Kemah, TX 77565
United States
281-389-1861
msuderman@cellsystems3d.com
Woraporn Sukhurnavasi
Chulalongkorn University, Faculty of
Veterinary Science, Parasitolog
Henri-Dunant Rd., Pathumwan
Bangkok, Thailand 10330
Thailand
662-218-9664
668-5-351-1080
vetkwan@hotmail.com
Sivapong Sungpradit
Department of Molecular
Microbiology and Immunology
615 N.Wolfe
Baltimore, MD 21205
United States
410-955-3442
sungpradit@yahoo.com
sungpradit@gmail.com
Xun Suo
China Agricultural University
College of Vet. Medicine
Yuanmingyan West Rd. 2# Haidian
Disrict
Bejing, 100193
China
861061 273-4325
suixun@cau.edu.cn
Md. Hasanuzzaman Talukder
777 Chestnut Ridge Rd., Apt. 203
US
304-680-4500
mhtalukder03@yahoo.com
Patrick Tanner
Merial, Ltd.
P.O. Box 661
Crawford, GA 30630
United States
706-759-3859
patrick.tanner@merial.com
Tiffany Taylor
Tuskegee University
90 Neal St.
Crawford, GA 30602
United States
1tmtaylor@gmail.com
Stephan Tegarden
Intervet/Schering-Plough
22694 W. 267th St.
Paola, KS 66071
United States
913-422-6828
stephan.tegarden@sp.intervet.com
Kevin B. Temeyer
USDA, ARS
Knipling-Bushland US Livestock
Insect Research Lab. 2700
Fredericksburg Rd.
Kerrville, TX 78028
United States
830-792-0330
830-792-0314
kevin.temeyer@ars.usda.gov
Astrid M. Tenter
Hermann-Hesse-Str. 15
Hannover, 30539
Germany
astrid.tenter@zoonosesinternational.com
Jerold H. Theis
University of California - Davis
School of Medicine Dept. of Med.
Micro. 1 Shields Ave.
Davis, CA 95616
United States
530-752-3427
530-752-8692
jhtheis@ucdavis.edu
V.J. Theodorides
1632 Herron Ln.
United States
610-696-5493
610-701-6395
vjtheodorides@axs2000.net
Gabriela Perez Tort
Jefe de Clinic: Hosp. Vet de Virreyes
Acceso Norte 2502 San Fernando
Pcia de Buenos Aires
Buenos Aires,
Argentina
00 54 11 4745-8612, 00 54 11 45802820
gabrielapt@gmail.com
Jennifer Thomas
Center for Veterinary Health
Sciences
Veterinary Pathobiology
250 McElroy Hall
Stillwater, Oklahoma 74078
United States
405-744-4484
jennifer.e.thomas@okstate.edu
Alex Thomasson
Merial, Ltd. St. Matthews University
SVM, University of Florida CVM
2600 Haven St
Inverness, FL 34452
US
352-422-6527
jatvet@hotmail.com
alex.thomasson@merial.com
Patricia Thomblison
2711 SW Westport Dr.
Topeka, KS 66614
United States
785-271-6005
785-273-9505
dr.pat@cliniciansbrief.com
patthom@cox.net
Patricia Thomblison
2711 SW Westport Drive
Topeka, KS 66614
USA
785-271-6025
patthomb@gmail.com
Adam Thompson
Guelph
778 Stonepath Cir.
Pickering, L1V3T
Canada
athomps2@uoguelph.ca
Marilyn Thompson
Companion Animal Hospital
7297 Commerce St
Springfield, VA 22150
USA
703-866-4100
703-866-4926
Pupdoc@aol.com
Philip R. Timmons
SCYNEXIS, Inc.
3501C Tricenter Blvd.
Durham, NC 27713
United States
919-549-5233
919-549-5240
phil.timmons@scynexis.com
Thomas Tobin
ttobin@aol.com
Michael Ulrich
Cheri-Hill Kennel & Supply, Inc.
17190 Polk Rd.
Stanwood, MI 49346
United States
231-823-2392
231-823-2925
misiu@frontier.com
Jennifer Towner
190 Round Table Ct.
Athens, GA 30606
United States
912-441-9921
jtowner1@uga.edu
Govind G. Untawale
281 Faemindale Rd.
Wayne, NJ 07470
United States
201-271-4800, ext. 7760
201-271-0357
guntawale@hartz.com
Abhishek Tripathi
Loyola University of Chicago
5920 South Cass Ave
Westmont, IL 60559
United States
484-620-9629
h.m.t.abhi@gmail.com
atripathi@lumc.edu
Joseph F. Urban Jr.
USDA, ARS, BHNRC, NRFL
BARC-East Building 307 C
United States
301-504-5528
301-504-9062
urbanj@ba.ars.usda.gov
Joseph P. Tritschler
Virginia State University
Agriculture Research P.O. Box 9061
Petersburg, VA 23842
United States
804-524-5957
804-524-5186
jtritsch@vsu.edu
Jennifer Uribe
University of Illinois
1506 S. Jefferson St.
Lockport, IL 60441
United States
630-220-8824
uribe@illinois.edu
Ba Cuong Tu
Modern Veterinary Therapeutics,
LLC
18301 SW 86 Ave.
Miami, FL 33157
United States
305-232-6645, cell: 786-200-2495
bacuong@yahoo.com
cuong.tuba@modernveterinarythera
peutics.com
Leah Tucker
NCSU College of Veterinary
Medicine
608 Oleander Road
Raleigh, North Carolina 27603
USA
3369062958
lktucker@ncsu.edu
lktucker89@gmail.com
Wendy Vaala
Schering Plough - Intervet
51476 Pleasant View Rd.
Alma, WI 54610
United States
608-685-4560
wendy.vaala@sp.intervet.com
Frans Van Knapen
Veterinary Public Health Division
Dept. IRAS P.O. Box 80175, 3508
TD
Utrecht, 3508TD
The Netherlands
+31 30 253 5367
+31 30 253 2365
d.vanknapen@uu.nl
Bradley van Paridon
511 9A Street NE
Calgary, AB T2E 4L3
Shannon Tyler
CN
Ross University School of Veterinary 403-305-2670
Medicine
bradley.vanparidon@uleth.ca
P.O. Box 334
Basseterre, St Kitts
St Kitts
18697603006
shannontyler@students.rossu.edu
119
Andrea Varela-Stokes
Basic Sciences Wise Center Spring
St.
Mississippi State University, MS
39762
United States
662-325-1345
stokes@cvm.msstate.edu
Julie Vargas
233 Sleepy Creek Dr.
Athens, GA 30606
United States
478-461-3185
jewells@uga.edu
Marie Varloud
marie.varloud@ceva.com
Adriano Vatta
Zoetis Inc.
333 Portage Street
USA
269-833-3083
adriano.vatta@zoetis.com
adrianovatta@gmail.com
Luisa Velasquez
Luisav@okstate.edu
Jozef Pieter Vercruysse
Ghent University
UGent
Faculty of Veterinary Medicine
Lab Parasitology
Salisburylaan 133
Merelbeke, OVL B-9820
Belgium
+32 9 2647390
+32 9 2647496
jozef.vercruysse@ugent.be
Saurabh Verma
2008 BMS,
College of Vet Med
ISU
Ames, Iowa 50011
United States
5152942547
sverma@iastate.edu
drsverma@gmail.com
Guilherme Verocai
3330 Hospital Drive NW
Calgary, AB T2N 4N1
Canada
4036043896
4032106693
gverocai@gmail.com
gverocai@gmail.com
Nathan Voris
Zoetis
10610 E. David Allen Rd
Columbia, MO 65201
USA
573-441-9714
nathan.voris@zoetis.com
nathanvoris@gmail.com
Isabelle Verzberger
Atlantic Vet. College
550 University Ave.
Canada
902-566-0843
isabelle.ve1@gmail.com
John M. Vetterling
Parasitologic Services
P.O. Box 475
Fort Collins, CO 80522-0475
United States
970-484-6040
970-221-0746
sprague@frii.com
Susan E. Wade
Cornell University
2250 N. Triphammer Rd, Unit 5B
Ithaca, NY 14850
United States
607-253-6182
sew9@cornell.edu
Stan Veytsman
St. Georgia's University
2725 E. 63 St.
Brooklyn, NY 11234
US
718-975-7542
sveytsma@sgu.edu
Bradley J. Waffa
University of Tennessee - Sewanee
735 University Ave.
Sewanee, TN 37383
United States
630-484-3435
bwaffa@sewanee.edu
Thomas Vihtelic
MPI Research
54943 N. Main St.
Mattawan, MI 49071
United States
269-668-3336
thomas.vihtelic@mpiresearch.com
Brent Wagner
University of Saskatchewan
52 Campus Dr.
Saskatoon, SK S7N 5B4
Canada
306-966-7217
brent.wagner@usask.ca
Eric Villegas
Environmental Protection Agency
26 W. Martin Luther King Dr. MS:320
Cincinnati, OH 45268
United States
513-569-7017
513-569-7117
villegas.eric@epa.gov
Cecelia Waugh-Hall
University of Technology
237 Old Hope Road,
Kingston 06,
Jamaica
(876)536-0018
celiawaugh@yahoo.com
Nadine Vogt
nvogt@uoguelph.ca
Janis Weeks
NemaMetrix LLC and University of
Oregon
1724 Sylvan St.
Eugene, OR 97403
USA
541-543-9984
janis.weeks@nemametrix.com
jweeks@uoregon.edu
Marcela von Reitzenstein
Zoetis Inc.
333 Portage street
Kalamazoo, MI 49007
USA
269-833-2410
marcela.vonreitzenstein@zoetis.com Andre Weil
Avogadro
marvonrei@inbox.com
185 Alewife Brook Parkway Suite
410
Cristiano Von Simson
Cambridge, MA 02138
Bayer Animal Health
United States
P.O. Box 390
403-201-4712
403-201-3643
United States
andre.weil@avogadro-lab.com
913-268-2867
cvsimson@gmail.com
cristiano.vonsimson@bayer.com
Emily Wessling
Cornell University
egw28@cornell.edu
120
Stephen K. Wikel
University of Connecticut Health
Center
School of Medicine 263 Farmington
Ave. MC3710
Farmington, CT 06030
United States
860-679-3369
860-679-8130
swikel@up.uchc.edu
Heike Williams
Intervet Innovation GmbH
Zur Propstei
Schwabenheim, 55270
Germany
+49 6130 948 240
+49 6130 948 513
heike.williams@intervet.com
James C. Williams
5214 N. Chalet Ct.
Baton Rouge, LA 70808-4843
United States
225-766-4728
jascwill@agctr.lsu.edu
Jeffrey F. Williams
HaloSource, Inc.
1631 220th St. Suite 100
Bothell, WA 98021
United States
425-974-1949, 208-390-7178
425-882-2476
jwilliams@halosource.com
Sherry Wilson
181 Sulphur Springs Rd.
Warrenton, NC 27589
United States
swilson@summitvetpharm.com
John B. Winters
Beverly Hills Small Animal Hospital
353 N. Foothill Rd.
Beverly Hills, CA 90210
United States
310-276-7113
310-859-0411
bhsah@adelpia.net
Adrian Wolstenholme
Dept. of Infectious Diseases College
of Vet. Medicine
Athens, GA 30602
United States
adrianw@uga.edu
Ming M. Wong
University of California - Davis
Vet. Path. Micro. & Immun.
Davis, CA 95616
United States
916-756-0460
916-756-0460
rfzwong@ucdavis.edu
Rob Woodgate
4 Atkins Pl.
Estella, NSW
Australia
rwoodgate@csu.edu.au
Jerry Woodruff
Fort Dodge Animal Health
8182 Moran Canyon Rd.
North Platte, NE 69101
United States
308-539-0475
jwoodruf@fdah.com
Debra Woods
VMRDCIPCD883 Ramsgate Rd.
Sandwich, Kent, CT139NJ
United Kingdom
44 13046473
debra.j.woods@pfizer.com
Heidi Wyrosdick
University of Tennessee
2407 River Dr.
Knoxville, TN
USA
865-974-5645
hwrosdi@utk.edu
Dehai Xu
USDA Agricultural Research
Services
P.O. Box 952 Aquatic Animal Health
Research
Auburn, AL 36831-0952
United States
334-887-3741
334-887-2983
dxu@ars.usda.gov
Michael Yabsley
SCWDS College of Vet. Medicine
Athens, GA 30602
United States
706-542-1741
myabsley@uga.edu
Tom A. Yazwinski
Dept. of Animal Science 1120 W.
Maple
United States
479-575-4398
479-575-5756
yazwinsk@uark.edu
Timothy Yoshino
Dept. Pathobiological Sciences,
University of Wisconsin, School of
Veterinary Medicine
2115 Observatory Drive
Madison, WI 53706
United States
608-263-6002
yoshinot@svm.vetmed.wisc.edu
Daniel Zarate Rendon
160 Dudley Dr., Apt. 505
Athens, GA 30606
United States
706-255-1544
dazre02@gmail.com
Hee-Jeong Youn
Seoul National University
College of Veterinary Medicine, 599
Gwanak-ro, Gwanak-gu
Seoul, 151-742
Korea
-2067
-1467
younhj@snu.ac.kr
younhj@snu.ac.kr
Dante Zarlenga
USDA, ARS, ANRI
Bovine Func. Genomics Lab 9555
Michaels Way
Ellicott City, MD 21042
United States
310-504-8754
dante.zarlenga@ars.usda.gov
Guan Zhu
Veterinary Pathobiology 4467 TAMU
David Young
United States
Young Veterinary Research Services 979-218-2391
7243 East Ave.
979-845-9972
Turlock, CA 95380
gzhu@cvm.tamu.edu
United States
209-632-1919
Dan Zhu
209-632-1944
Cornell University
youngdvm@yvrs.com
203 Williams Street
Ithaca, NY 14850
Silene Young
USA
Bayer Health Care
607-379-0845
P.O. Box 390
dz254@cornell.edu
Shawnee Mission, KS 66201-0390
zhudan105@hotmail.com
United States
913-268-2867
Zia-ur-Rehman
913-268-2878
Massey university
Skidogs1@gmail.com
Hopkirk research Institute, IVABS,
Massey university
Lisa Young
Palmerston North, 4442
Elanco
New Zealand
2500 Innovation Way
6.4210816714e+011
z.u.rehman@massey.ac.nz
USA
ziyakh@gmail.com
3172920361
youngli@elanco.com
Gary L. Zimmerman
Zimmerman Research
Cole Younger
1106 W. Park PMB 424
Stillmeadow, Inc.
Livingston, MT 59047
12852 Park One Dr.
United States
406-223-3704
United States
zresearch@msn.com
281-240-8828
cyounger@stillmeadow.com
Erich Zinser
Anne M. Zajac
7000 Portage Rd.
Kalamazoo, MI 49001
United States
Medicine
269-833-2401
erich.w.zinser@pfizer.com
United States
540-231-7017
540-231-6033
azajac@vt.edu
121

Proceedings AAVP Membership Directory

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