Transmissible Diseases Handbook

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  Transmissible Diseases Handbook  

TABLE OF CONTENTS I.

INTRODUCTION

J. Kaandorp

II.

GENERAL CONSIDERATIONS

J. Kaandorp

III.

EUROPEAN UNION

J. Kaandorp

1. EUROPEAN UNION 2. DATA ON EU MEMBER STATES 3. EU ANIMAL HEALTH STRATEGY AND ANIMAL HEALTH ADVISORY COMMITTEE 4. EU ANIMAL HEALTH LAW 5. EAZA POSITION STATEMENT ON THE ANIMAL HEALTH STRATEGY FOR THE EUROPEAN UNION 6. OIE IV.

LINKS

J.Kaandorp

1. EU VETERINARY FACULTIES 2. USEFUL (VETERINARY) WEBLINKS V.

ANIMAL HEALTH LEGISLATION IN EUROPE

P. Dollinger

VI.

RECOMMENDATIONS FOR THE APPLICATION OF EAZWV – IDWG – DEFRA – ANNEX C TO THE BALAI - DIRECTIVE BMVEL – RVV – DG SANCO

VII.

POST-MORTEM PROCEDURES

G. Dorrestein et al.

VIII. GUIDELINES FOR CLEANING AND DISINFECTION IN M. Kiupel et al. ZOOLOGICAL GARDENS IX.

BLUETONGUE IN NON-DOMESTIC RUMINANTS: S. Sanderson EXPERIENCES GAINED IN EAZA ZOOS DURING THE

2007 & 2008 BTV8 AND BTV1 EPIZOOTICS X.

TUBERCULOSIS IN ZOO SPECIES: DIAGNOSTIC EAZWV Tuberculosis Working UPDATE AND MANAGEMENT ISSUES Group 1. ELEPHANTS (ELEPHAS MAXIMUS)

XI.

W. Shaftenaar

AVIAN INFLUENZA 1. VACCINATION OF NON-DOMESTIC AVIAN J. Philippa SPECIES AGAINST HIGHLY PATHOGENIC AVIAN INFLUENZA (HPAI) VIRUSES 2. WAZA/EAZA/EAZWV AVIAN INFLUENZA

XII.

RECOMMENDATIONS

ON WAZA – EAZA – EAZWV

VACCINATION OF NON-DOMESTIC CARNIVORES: A J. Philippa REVIEW

XIII. LIST OF LABORATORIES XIV. TEMPLATE FACT SHEET XV. INSTRUCTIONS FOR AUTHORS XVI. FACT SHEETS

Transmissible Diseases Handbook

I.

INTRODUCTION

Jacques Kaandorp Past - President EAZWV, IDWG chairman, DFZVO chairman, EAZA veterinary committee chairman, member EAZA legislation committee

At the 2000 EAZWV congress in Paris there was a proposal to establish an Infectious Diseases Working Group (IDWG) within the EAZWV. This idea emerged when the problems of diagnosing and handling paratuberculosis were discussed in a presentation. It was obvious that commonly agreed recommendations were urgently needed for improved control of the infectious diseases that threaten our collections. Also, as Europe becomes more and more united it is necessary to deal with European politics and legislation, and intensify international collaboration. The Foot and Mouth Disease (FMD) outbreak in 2001 clearly underlined the importance of this initiative and made agreement on European standards a matter of even more urgency. During this outbreak we saw how difficult it was to control a disease in domesticated animals because of public opinion, politics, money and difficulties in understanding the disease and agreeing on legislation. With this in mind it is obvious that in lesser-studied animals, like our exotics, even more misunderstandings may arise when dealing with such diseases. In 2006 Highly Pathogenic Avian Influenza spread across Europe and threatened zoo collections. Outbreaks of Classical Swine Fever now and then occur. Blue Tongue has made serious problems in the ability of transporting our hoof stock and thus jeopardizing our endangered species programs. African Swine Fever, and maybe in the near future diseases like West Nile Virus, African Horse Sickness, and Monkey pox may also threaten our collections. In recent history even the SARS epidemic (related to an animal Corona virus) indeed confirmed the above and made the concept and ideas of The Transmissible Diseases Handbook even more important. European laws (e.g. the Balai Directive) are still under discussion and in development. Different attitudes in Europe (e.g. vaccination vs. non-vaccination against FMD and Avian Influenza), different levels of veterinary faculties and laboratories, and thus differences in the possibility of diagnosing diseases, pose further problems for European zoo veterinarians. As a consequence, we have to deal with different approaches to infectious diseases. Standardising our work is therefore in the interests of us all. To achieve this goal we need to work together to define procedures for transmissible diseases and propose recommendations for people dealing with exotic animals. Only in this way will it be possible to make comparisons, and to statistically analyse our similarities and differences. The IDWG brings together experienced zoo veterinarians and specialists in infectious diseases from several European countries and even from countries on the other side of the Atlantic. The idea of the Group is to combine our knowledge in order to help our animals by dealing more efficiently with future disease outbreaks which may threaten our collections. Putting efforts into the creation of a reference manual published under the umbrella of our Association is an important step in the process of European standardisation, and should provide a useful tool for zoo practitioners, zoo managers and European legislative authorities dealing with wildlife and zoo animals.

Transmissible Diseases Handbook

II.

GENERAL CONSIDERATIONS

Jacques Kaandorp Past-President EAZWV, IDWG chairman, DFZVO chairman, EAZA veterinary committee chairman, member EAZA legislation committee

This handbook on transmissible diseases, created by the IDWG, is not intended to provide extensive descriptions of all transmissible diseases which may occur in a zoo collection, but we hope that it will be a useful tool for every European zoo and/or wildlife veterinarian encountering a problem related to such diseases. The handbook should help to find answers to the question “what should I do” when an infectious disease is suspected. Furthermore, in regard to inter-zoo exchanges, it is very important to standardise surveillance programs for infectious diseases in European zoos. This handbook should help to overcome the differences present within Europe and to attain the standardisation required by the Balai Directive (92/65/EC). EAZWV recommendations for application of the directive have therefore been included. These recommendations are accepted by SANCO in Brussels as a guide for the authorities to audit the approvals of zoos under the Balai Directive. (SANCO/10479/2005) As mentioned above, the handbook is not supposed to replace good scientific books. On the contrary, such books will still be necessary for finding detailed information. The handbook is basically a review. It summarises information on various diseases: susceptible animals, zoonotic potential, clinical symptoms, pathology, diagnostic methods, qualified laboratories, treatment, prevention, experts who may be consulted, legislation (esp. European laws) and relevant literature. The detailed table of contents and the standardised format of the fact sheets will help the reader to find information quickly. Almost all the fact sheets were reviewed by two experts. For a few fact sheets only one reviewer could be found. The search engine on this CD-ROM will help the reader to find his way through the book. The handbook should be a “living” book. Our European world is changing fast and the information will soon be outdated if we don’t revise it regularly. A continuous updating process is a prerequisite for a reliable and useful tool! Ideally, the handbook should include all diseases mentioned in the Balai Directive (92/65/EC) as well as other diseases relevant for exchanges of animals between zoos (OIE list A and B and certain zoonoses). Again it looks like we have succeeded in this! In addition to the fact sheets, there are general chapters about European legislation, the integral texts of the relevant European regulations, the EAZWV recommendations on the Balai Directive, post-mortem procedures, guidelines for cleaning and disinfection, vaccination of non-domestic carnivores, Tuberculosis, Blue Tongue and lists of reference laboratories. The authors of the various fact sheets have done their best to meet the requirements. Almost all fact sheets from the second edition have been updated! Our living book needs to be updated regularly. We therefore urge all readers to send us their comments for incorporation in future editions.

II.

General Considerations

The chapters on legislation, the list of reference laboratories as well as OIE list A and B have been included to compensate for the empty sections in many fact sheets. Many European legislation relevant for exotic animals are added in order to have a more complete dossier. A list of IDWG members is not included in the handbook, because readers can address questions directly to them if necessary, since addresses from the authors (read IDWG members), can be found in the fact sheets themselves. The legislation texts can be easily downloaded and printed as PDF-files in the various European languages at http://eur-lex.europa.eu/ This time the IDWG has again chosen to bring this edition digitally on a CD-ROM or downloadable at www.eazwv.org and www.eaza.net and not in print. This has reduced the costs of producing the book and will also reduce the costs of spreading it enormously. Another advantage was the easy expansion of the book, since the second edition already comprised of 662 pages. This book contains more legislation and new chapters, thus a lot more pages, and a printed version would become too massive to distribute and would also be a lot more expensive. We also hope that the book in this format will find its way globally more easily. This fourth edition, like the previous editions, will be spread to all EAZWV members, all EAZA zoos, Central Veterinary Officers of the 27 EU Member States, the O.I.E., and SANCO in Brussels. Another advantage of this version is the search engine which is included on this CD-ROM. At this point I would like to thank the authors of all the fact sheets for their tremendous work. Norin Chai did the editing of the fact sheets of this fourth edition. For the final editing Ayla Bayens was of great help. I would also like to thank EAZA and EAZWV for their financial support. All people involved understood the need to standardise some aspects of our work in the context of a rapidly-changing Europe. Without their involvement it wouldn’t have been possible to create this handbook. Let’s hope that all EAZWV members as well as the EAZA zoo community will appreciate this work, make use of it, and help us to improve it! For comments you are kindly requested to contact the chairman of the IDWG: Jacques Kaandorp, e-mail: [email protected]

Transmissible Diseases Handbook

III. EUROPEAN UNION Jacques Kaandorp Past-President EAZWV, IDWG chairman, DFZVO chairman, EAZA veterinary committee chairman, member EAZA legislation committee

1. EUROPEAN UNION 2. DATA ON EU MEMBER STATES 3. EU ANIMAL HEALTH STRATEGY AND ANIMAL HEALTH ADVISORY COMMITTEE 4. EU ANIMAL HEALTH LAW 5. EAZA POSITION STATEMENT ON THE ANIMAL HEALTH STRATEGY FOR THE EUROPEAN UNION 6. OIE

1. European Union The European Union is a unique economic and political partnership between 27 democratic European countries aiming at peace, prosperity and freedom for its 498 million citizens. The EU is run through bodies such as  the European Parliament representing the people of Europe  the Council of the European Union representing national governments  the European Commission representing the common EU interest  the Court of Justice making sure that EU law is interpreted and applied in the same way in all EU countries  the Court of Auditors checking that the EU’s funds are spent legally, economically and for the intended purpose

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European Union

2. Data on the 27 EU Member States EU Member State Austria Belgium Bulgaria Cyprus Czech Republic Denmark Estonia Finland France Germany Greece Hungary Ireland Italy Latvia Lithuania Luxembourg Malta Netherlands Poland Portugal Romania Slovenia Slovakia Spain Sweden United Kingdom

Year of EU entry 1995 Founding member 2007 2004 2004 1973 2004 1995 Founding member Founding member 1981 2004 1973 Founding member 2004 2004 Founding member 2004 Founding member 2004 1986 2007 2004 2004 1986 1995 1973

Political system Federal republic Constitutional Monarchy Republic Republic Republic Constitutional Monarchy Republic Republic Republic Federal republic Republic Republic Republic Republic Republic Republic Constitutional Monarchy Republic Constitutional Monarchy Republic Republic Republic Republic Republic Constitutional Monarchy Constitutional Monarchy Constitutional Monarchy

Capital city Vienna Brussels Sofia Nicosia Prague Copenhagen Talinn Helsinki Paris Berlin Athens Budapest Dublin Rome Riga Vilnius Luxembourg Valetta Amsterdam Warsaw Lisbon Bucharest Ljubljana Bratislava Madrid Stockholm London

Population 8.3 million 10.7 million 7.6 million 0.8 million 10.4 million 5.5 million 1.3 million 5.3 million 63.8 million 82.2 million 11.2 million 10.1 million 4.4 million 59.6 million 2.3 million 3.4 million 0.5 million 0.4 million 16.4 million 38.1 million 10.6 million 21.5 million 2.0 million 5.4 million 45.3 million 9.2 million 61.2 million

Veterinarians 2704 13709 3422 271 6655 2330 1081 2124 27338 35098 3547 3500 3887 22604 2256 1923 129 No data 5978 9321 3665 9562 940 No data 9600 2553 29971

Currency Euro Euro Lev Euro Czech koruna Danish krone Estonian kroon Euro Euro Euro Euro Forint Euro Euro Lats Litas Euro Euro Euro Zloty Euro Romanian leu Euro Euro Euro Krona Pound sterling

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European Union

3. The EU Animal Health Strategy (2007-2013) where “prevention is better than cure” and the Animal Health Advisory Committee The EU’s Animal Health Strategy provides the framework for EU animal health and welfare measures until 2013. Give the devastating impact that serious disease outbreaks can have on farmers, society and the economy, the strategy is based on the principle that “prevention is better than cure”. The aim is to put greater focus on precautionary measures, disease surveillance, controls and research, in order to reduce the incidence of animal disease and minimise the impact of outbreaks when they do occur. However, the new strategy encompasses much more than just the control of animal diseases. It also focuses on issues which are inextricable linked to animal health, such as public health, food safety, animal welfare, sustainable development and research. The “Animal Health Advisory Committee” includes representatives from non governmental organisations (including EAZA) across the animal health sector, consumers and governments. It is aimed at gathering specialised inputs. It provides strategic guidance on the appropriate/acceptable level of animal or public health protection, and on priorities for action and communication. It also follows the Animal Health Strategy’s progress. The Committee is consulted on all impact assessments and advises the European Commission on the best means of delivering agreed outcomes. www.one-health.eu

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European Union

4. The EU’s Animal Health Law Since its creation, the European Community has had as goal the creation of a single market to allow the free circulation of different commodities, including live animals and products of animal origin. To ensure the safe trade of live animals and animal products, harmonised health requirements for all EU Member States had to be established. The process of establishing the EU health requirements requires continual updates because these requirements need to consider a number of evolving factors, such as new scientific knowledge, and emerging animal diseases. This process of continual updating of the legislation means that a number of different regulatory measures are now in place. Currently, EU veterinary legislation includes more than 400 different legislative acts. This large body of legislation has created difficulties for different actors, such as the Member States’ veterinary authorities, commercial operators, farmers, and breeders to understand the law. This is also true for the zoo community. To tackle this problem, the European Commission decided to review the current EU animal health legislation when it published its Animal Health Strategy in 2007. As part of its Strategy, the European Commission decided that it should create a new Animal Health Law, which would provide a single and clearer regulatory framework for all EU animal health regulation. The aims of this legal framework would be to simplify, clarify and provide greater flexibility for EU animal health legislation, and also to introduce important new elements, such as for example a preventive approach to disease control. An extensive consultation process for the EU Animal Health Law is launched end September 2009 and will end in December 2009. The aim of this consultation is to gather the views of all concerned parties and to take these comments into consideration when drafting the final legislative text. The target date for the formal adoption of the proposal by the commission is the end of 2010.

III.

European Union

5. EAZA Position Statement on the Animal Health Strategy for the European Union

Introduction This statement presents the position of the European Association of Zoos and Aquaria (EAZA) on the Animal Health Strategy for the European Union (2007-2013) “Prevention is better than cure” and its resulting Action Plan. It is also endorsed by the European Association of Zoo and Wildlife Veterinarians (EAZWV) with approximately 700 individual members from zoos, zoo related research institutions and universities. In general EAZA is pleased with the strategy and would like to congratulate the European Commission on its publication. The slogan “Prevention is better than cure” is much appreciated and reflects the approach of EAZA and its member institutions. The flexible approach to vaccination is a highly important aspect for the European zoo and aquarium community as is the link between animal welfare and animal health. A clearer and simplified regulatory framework is necessary, especially as EU animal health regulations are mostly directed at agricultural animals and the impact of such legislation for zoo and aquarium animals is often unclear or open to multiple interpretations. The development of one general EU Animal Health Law is therefore strongly supported by EAZA. The remainder of this statement will provide further detail on EAZA’s position. EAZA’s current status and general position;  As laid down in EAZA’s constitution the objects of the association are: a. to promote co-operation for the furtherance of wildlife conservation, through internationally coordinated breeding programmes of wild animals and in situ conservation; b. to promote education, in particular environmental education; c. to promote scientific study; d. to represent the interests of its members;  EAZA represents 324 members from 35 countries, 300 of which maintain public collections of animals. More than 280 institutions of the total EAZA membership are located within the European Union. EAZA member institutions receive approximately 140 million visitors a year and house more than 250,000 animals, excluding fish and invertebrates. EAZA member institutions employ 20,000 staff members, 5000 of which are seasonal;  The ‘EAZA Minimum Standards for the Accommodation and Care of Animals in Zoos and Aquaria’ include extensive paragraphs on animal welfare, health, hygiene, surveillance and veterinary aspects.  EAZA members are often important economic drivers and cultural centres in their local communities;  EAZA has a significant social role in educating European citizens about animals, their conservation, and overarching threat processes such as climate change. Zoos and aquaria have been demonstrated to host a far more representative visitor social spectrum than either museums or science centres;

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European Union

EAZA has adopted the World Zoo and Aquarium Conservation Strategy (2005) which articulates the modern role of zoos and aquaria and their commitment to conservation; EAZA institutions in the European Union comply with Council Directive 1999/22/EC relating to the keeping of wild animals in zoos; EAZA encourages its member institutions to apply for approval under article 13 (2) of Council Directive 92/65/EEC; EAZA has a Memorandum of Understanding with the European Association of Zoo and Wildlife Veterinarians (EAZWV) and supported the publication of the Transmissible Disease Handbook (2007). This is recognised as a key publication of high relevance across all sectors concerned with public health. EAZA will also be supporting the publication of the 2009 edition of this important document; All EAZA members must join the International Species Information System and use the Animal Record Keeping System (ARKS) software to keep up to date records of their animal collections; EAZA collections exchange approximately 25,000 animals annually; Emerging infectious disease outbreaks generally do not originate from zoo and aquarium collections. EAZA zoos have highly trained specialised veterinarians and therefore can quickly recognise newly arriving emerging infectious diseases, and thus can serve as emergent disease sentinels. Zoos and aquariums can suffer from outbreaks of diseases, through potential loss of stock, obstruction of transfers of animals between collections as part of vital conservation breeding programmes and loss of visitor revenue if the institution is within geographical areas of disease outbreak and human movement restrictions occur; Populations of endangered species kept in EAZA collections are often irreplaceable and some held in EAZA institutions are extinct in the wild.

We believe;  Keeping and displaying healthy animals under good welfare conditions in EAZA collections is of crucial importance to reach EAZA’s main objectives;  Animal exchanges between EAZA member institutions (largely in the framework of EAZA’s breeding programmes) are imperative to ensure healthy and sustainable populations of wild animals in human care into the future;  EAZA’s European Endangered species Programmes (EEPs) and European studbooks (ESBs) should be managed independently, where appropriate, from ex situ populations in other regions and from wild populations, unless specifically part of global endangered species programmes. Nevertheless, occasional imports of unrelated stock are important to ensure long-term genetic variability;  Contamination risks in zoos and aquaria are significantly lower than in the agricultural industry, e.g. because of surveillance, housing conditions, reduced numbers of animals and individual care for most species;  Compared to 4.3 million cattle, 21.7 million pigs and 794 million poultry traded between EU member states in 2006 the number of animals exchanged between EAZA members pose an extremely low health risk;  Breeding programmes for endangered species are jeopardized by unclear legislation, lack of uniform implementation of EU legislation by member states and slow decision making processes making animal exchanges difficult or even impossible. This obstruction can lead to compromised welfare conditions and obstruction of conservation initiatives;  EAZA’s breeding programmes and the Animal Record Keeping System (in the process of being replaced by the Zoological Management Information System – ZIMS) are suitable means for identification and traceability of zoo and aquarium species.

III.

European Union

What we would like to improve in the future;  EAZA to be consulted at an early stage when the EU is formulating animal health and welfare legislation to ensure that wild animal species as held in zoos and aquaria are clearly and appropriately included in the regulatory definitions where relevant and excluded where irrelevant;  The position of EAZA zoos and aquaria to be fully considered when designing EU animal health regulations;  The health risks posed by animal collections of zoos and aquaria better evaluated in light of the existing EU animal health regulations;  Exchanges of animals in the framework of recognised EAZA breeding programmes simplified and prioritised in relation to EU legislation pertaining to animal health;  Implementation of the BALAI Directive 92/65 to be harmonized across EU Member States;  EAZA breeding programmes and ISIS ARKS registration formally recognised as suitable means for identifying and tracing of zoo and aquarium animals;  In relation to Council Regulation 1/2005 of 23 December 2004 animal transports between zoos carried out only by the institutions themselves should not be considered “commercial transports”. In a future review of Council Regulation 1/2005 the wide diversity of species to be transported between zoos and aquaria and thereafter their differing and specific welfare needs should be recognised;,  To continue to be able to vaccinate where appropriate in the case of emerging disease outbreaks (refer to the ‘EAZA Minimum Standards for the Accommodation and Care of Animals in Zoos and Aquaria’) ;  To have, through approval under 13(2) of Council Directive 92/65, the possibility to import wild animals from third countries where there is an agreed and demonstrable programme need, particularly in relation to recognised and accountable EAZA conservation breeding programmes, and where the import can be demonstrated to not be of any detriment, either of welfare or persistence, to wild populations.

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European Union

6. The O.I.E. (World Organisation for Animal Health) (Organisation International des Epizooties) The OIE is an intergovernmental organisation based in Paris, with a mandate from its 174 Member Countries and Territories to improve world animal health. In this capacity, the OIE is responsible for ensuring transparency of the animal disease situation worldwide, including diseases transmissible to humans, and the sanitary safety of world trade in animals and animal products. The OIE publishes international standards in all fields covered by its mandate, including animal welfare and consumer protection. At the global level, the OIE has modernised its worldwide information system on animal diseases (including zoonoses) with the creation of WAHIS, a mechanism whereby all countries are linked on-line to a central server that collects all the compulsory notifications sent to the OIE, covering 100 priority terrestrial and aquatic animal diseases, as well as any emerging disease. Together with the WHO (World Health Organisation) and the FAO (Food and Agricultural Organisation of the United Nations), the OIE has created GLEWS, the Global Early Warning System, a platform shared by the three organisations to improve early warning on animal diseases and zoonoses worldwide. The OIE, WHO and FAO (with the support of UNICEF, the United Nations System Influenza Coordinator and the Worldbank) have prepared a consensus document on global measures needed to coordinate medical and veterinary health policies more effectively, taking into account new requirements to prevent and control zoonoses. In this perspective veterinarians play a crucial role in protecting animals and society as a whole from the negative effects of diseases such as Avian Influenza, Foot and Mouth Disease and Bluetongue. Veterinarians play a crucial role at each stage of the food chain, from “stable to table”. For example, by checking that only healthy animals are slaughtered for human consumption, or by alerting the authorities at the first signs of disease on the farm or in our zoos. Veterinarians across the European Union, through their work, help to ensure that the goal of “One World, One Health” for all can be reached, encompassing both animals and people in good health. The concept “One World, One Health” urges the international community to consider the link between animal diseases and public health. It is well known that 60% of known human infectious diseases have their source in animals, as do 75% of emerging human diseases and 80% of the pathogens that could potentially be used in bio terrorism. The unprecedented flow of commodities and people gives pathogens of all kinds the opportunity to spread and multiply around the world to find genetic reassortment opportunities, and climate change can enable them to expand in their range. www.one-health.eu

Transmissible Diseases Handbook

IV. LINKS Jacques Kaandorp Past-President EAZWV, IDWG chairman, DFZVO chairman, EAZA veterinary committee chairman, member EAZA legislation committee

1. EU VETERINARY FACULTIES 2. USEFUL (VETERINARY) WEBLINKS

1. EU Veterinary Faculties Austria Veterinärmedizinische Universität Wien www.vu-wien.ac.at

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Belgium Université de Liège Faculté de Médecine Vétérinaire www.ulg.ac.be/fmv University of Ghent Faculty of Veterinary Medicine www.ugent.be/di/en Bulgaria Trakia University. Faculty of Veterinary Medicine, Student’s Campus, Stara Zagora www.uni-sz.bg University of Forestry, Sofia. Faculty of Veterinary Medicine www.edu.ltu.bg/typo3 Czech Republic Veterinárni a Farmaceutická Universita www.vfu.cz Denmark Københavns Universitet, Det Biovidenskablige Fakultet www.life.ku.dk Estonia Estonian Agricultural University Faculty of Veterinary Medicine Institute of Veterinary Medicine and Animal Sciences www.eau.ee Finland University of Helsinki Faculty of Veterinary Medicine www.vetmed.helsinki.fi

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France École Nationale Vétérinaire d’Alfort (ENVA) www.vet-alfort.fr École Nationale Vétérinaire de Lyon (ENVL) www.vet-lyon.fr École Nationale Vétérinaire de Nantes (ENVN) www.vet-nantes.fr École Nationale Vétérinaire de Toulouse (ENVT) www.envt.fr Germany Veterinärmedizinische Fakultät der Freien Universität Berlin www.fu-berlin.de Fachbereich Veterinärmedizin der Universität Giessen www.uni-giessen.de Tierärztliche Hochschule Hannover www.tiho-hannover.de Veterinärmedizinische Fakultät der Universität Leipzig www.uni-leipzig.de Tierärztliche Fakultät der Ludwig-Maximillians Universität München www.vetmed.uni-muenchen.de Greece Aristoteles University – Thessaloniki. School of Veterinary Medicine University of Thessaly – Faculty of Veterinary Science www.vet.uth.gr Hungary University of Veterinary Science in Budapest www.univet.hu Ireland University College Dublin Faculty of Veterinary Medicine www.ucd.ie Italy Facoltá di Medicina Veterinaria www.uniba.it/ Universitá di Bologna Facoltá di Medicina Veterinaria www.vet.unibo.it Universitá di Camerino – Facoltá di Medicina Veterinaria www.unicam.it Universitá di Messina Facoltá di Medicina Veterinaria www.unime.it Universitá di Milano Facoltá di Medicina Veterinaria www.veterinaria.unimi.it Universitá di Napoli Federico Il Facoltá di Medicina Veterinaria www.unina.it Facoltá di Medicina Veterinaria Universitá degli studi di Padova www.veterinaria.unipd.it Universitá di Parma Facoltá di Medicina Veterinaria www.unipr.it Universitá di Perugia Facoltá di Medicina Veterinaria www.unipg.it/-facvet

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Universitá di Pisa Facoltá di Medicina Veterinaria www.unipi.it Universitá di Sassari Facoltá Medicina Veterinaria www.uniss.it Universitá degli Studi di Teramo www.unite.it/ Universitá di Torino Facoltá Medicina Veterinaria www.veter.unito.it/ Latvia Latvia University of Agriculture Veterinârmedicìnas fakultâte www.llu.lv/?mi=152 Lithuania Lithuanian Veterinary Academy www.lva.lt/en/ Netherlands Universiteit Utrecht Faculty of Veterinary Medicine www.vet.uu.nl Poland Uniwersytet Przyrodniczy w Lublinie – Wydzial Medycyny Weterynaryjnej www.ar.lublin.pl Uniwersytet Warmińsko – Mazurski w Olsztynie –Wydzial Medycyny Weterynaryjnej www.uwm.edu.pl/wmw Szkola Glówna Gospodarstwa Wiejskiego w Warszawie – Wydzial Medycyny Weterynaryjnej www.sggw.pl Uniwersytet Przyrodniczy we Wroclawiu - Wydzial Medycyny Weterynaryjnej www.ar.wroc.pl Portugal Escola Universitária Vasco da Gama – Coimbra www.euvg.net Universidade de Évora www.uevora.pt Universidade Técnica de Lisboa – Faculdade de Medicina Veterinária www.fmv.utl.pt Universidade Lusófona de Humanidades e Tecnologias – Lisboa Universidade do Porto – Instituto de Cièncias Biomédicas de Abel Salazar www.up.pt Universidade de Trás-os-Montes e Alto Douro – Vila Real www.utad.pt Romania University of Agricultural Sciences and Veterinary Medicine Bucharest www.fmvb.ro Faculty of Veterinary Medicine, Cluj-Napoca University of Agronomy and Veterinary Medicine – lasi Faculty of Veterinary Medicine, Timisoara Slovenia University of Ljubljana www.vf.uni-lj.si

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Slovak Republic University of Veterinary Medicine in Košice www.uvm.sk/ Spain Universitat Autònoma de Barcelona Facultat de Veterinària www.uab.es/fac-veterinaria Universidad de Córdoba Facultad de Veterinaria www.uco.es/organiza/centros/veterinaria Facultad de Veterinaria de Cáceres www.veterinaria.unex.es Universidad de León – Facultad de Veterinaria www.unileon.es Universidad de Murcia Facultad de Veterinaria www.um.es/veterina/ Facultad de Veterinaria de las Palmas de Gran Canaria www.vet.ulpgc.es Universidad Complutense de Madrid, Facultad de Veterinaria www.ucm.es/info/webvet Universidade de Santiago de Compostella – Licenciado en Veterinaria (LUGO) www.facveterinarialugo.org/ Universidad Politécnica de Valencia – Departamento de Ciencia Animal www.www.dcam.upv.es/dcia/ Universidad de Zaragoza Facultad de Veterinaria www.wzar.unizar.es/acad/fac/vete/unizar.html Universidad Alfonso X (Madrid) www.uax.es/oferta_docente/titulaciones/vet Sweden University of Agricultural Sciences Faculty of Veterinary Medicine www.slu.se United Kingdom University of Bristol www.bristol.ac.uk/fmus/ University of Cambridge Veterinary School www.vet.cam.ac.uk University of Edinburgh Royal (Dick) School of Veterinary Studies www.vet.ed.ac.uk/ University of Glasgow Faculty of Veterinary Medicine www.gla.ac.uk/faculties/vet/ University of Liverpool Faculty of Veterinary Science www.liverpool.ac.uk/vets/ Royal Veterinary College www.rvc.ac.uk University of Nottingham www.nottingham.ac.uk

Links

IV.

2. Useful (Veterinary) Weblinks Veterinary links The World Organisation for Animal Health www.oie.int OIE Animal Disease Information Summaries www.oie.int/eng/ressources/en_diseasecards.htm OIE Terrestrial Animal Health Code www.oie.int/eng/normes/mcode/en_index.htm Food and Agricultural Organisation of the United Nations www.fao.org The World Health Organisation www.who.int WHO mediacentre www.who.int/mediacentre/factsheets/en Federation of Veterinarians of Europe www.fve.org A guide to the use of the Internet for Vets www.vts.intute.ac.uk/he/tutorial/vet/ Veterinary Information Network www.vin.com International Veterinary Information Service www.ivis.com Veterinary Sciences Tomorrow www.vetscite.org World Veterinary Association www.worldvet.org Veterinary Events Worldwide www.vetagenda.com The World Small Animals Veterinary Association www.wsava.org The Federation of European Companion Animal Veterinary Associations www.fecava.org European Association for the Evaluation of Veterinary Establishments www.eaeve.org Centre for Disease Control and Prevention USA www.cdc.gov/nczved/dfbmd/disease_listing.html The Merck Veterinary Manual www.merckvetmanual.com/mvm/index.jsp EAZWV European Association of Zoo and Wildlife Veterinarians www.eazwv.org AAZV American Association of Zoo Veterinarians www.aazv.org EAZA European Association of Zoos and Aquaria www.eaza.net WAZA World Association of Zoos and Aquaria www.waza.org European communication platform for veterinary students www.vetstart.org International Veterinary Student Association www.ivsa.org

Links

IV.

European (Union) links Gateway to the European Union http://europa.eu European Parliament www.europarl.europa.eu Council of the European Union www.consilium.europa.eu European Commission http://ec.europa.eu Directorate-General for Health and Consumers http://ec.europa.eu/dgs/health_consumer/index_en.htm Directorate-General for Agriculture and Rural Development http://ec.europa.eu/dgs/agriculture/index_en.htm Directorate-General for Maritime Affairs and Fisheries http://ec.europa.eu/dgs/fisheries/index_en.htm Directorate-General for Environment http://ec.europa.eu/dgs/environment/index_en.htm European Food Safety Authority www.efsa.europa.eu European Centre for Disease Prevention and Control http://ecdc.europa.eu European Medicines Agency www.emea.europa.eu Access to European law http://eur-lex.europa.eu

Links

Transmissible Diseases Handbook

VI. RECOMMENDATIONS FOR THE APPLICATION OF ANNEX C TO COUNCIL DIRECTIVE 92/65 (“BALAI”) AS AMENDED BY COUNCIL REGULATION (EC) NO 1282/2002 OF 15 JULY 2002 (OJ L 187/3) IN APPROVED ZOOS 1. Introduction The amendment of 15 July 2002 to the Council Directive 92/65/EEC laying down animal health requirements governing trade in and imports into the Community of animals, semen, ova and embryos not subject to animal health requirements laid down in specific Community rules referred to in Annex A (1) to Directive 90/425/EEC (“BALAI”) was made in the legal form of a regulation. As a consequence, the Annexes A, C and E are directly applicable in Member States. ANNEX C contains, however, a few points which leave considerable room for interpretation. The present recommendations aim at contributing to a uniform interpretation of these points, and thus at achieving the ultimate goal of this annex, namely to facilitate the exchange of animals between approved zoos easily and without major health risks. These recommendations are the result of two meetings held on 15/16 September 2003 and 5 February 2004 at Cologne Zoo with the participation of representatives of the European Commission (DG SANCO - Health and Consumer Protection), the Department for Environment, Food & Rural Affairs (DEFRA), the Rijksdienst voor de Keuring van Vee en Vlees (RVV), the Bundesministerium für Verbraucherschutz, Ernährung und Landwirtschaft (BMVEL), the European Association of Zoo and Wildlife Veterinarians (EAZWV), the EAZWV Infectious Diseases Working Group (IDWG) and zoo veterinarians from France, Germany, Italy, The Netherlands and the United Kingdom representing also their respective professional organisations at the national level. The recommendations are meant to provide some practical guidance to both, Veterinary Authorities and zoo veterinarians, throughout the European Union. EU veterinary legislation applies also to the British Crown Dependencies (Channel Islands, Isle of Man), Andorra, Monaco and San Marino. As Council Directive 92/65/EEC and Commission Regulation (EC) No. 1282/2002 are texts with EEA relevance (Regulation 1282/2002 was incorporated into the EEA Agreement by Decision of the EEA Joint Committee of 14 March 2003), and are also part of the equivalent sector of the Bilateral Agreement on Agriculture with Switzerland (Regulation 1282/2002 is included in the 2003 update), they apply in Liechtenstein, Norway, and Switzerland too. Note: To make reading of this document easier, the term “Bodies, institutes and centres” is usually replaced by the word “zoo”

2. Contents A. The term ‘animals’ B. The approved veterinarian C. The annual disease surveillance plan

VI.

D. The added animals procedure E. Quarantine / isolation requirements F.

The certificates

Recommendations

VI.

Recommendations

A. The term ‘animals’ 1.

‘Animals’ means specimens of animal species other than those referred to in Directives 64/432/EEC, 90/426/EEC, 90/539/EEC, 91/67/EEC, 91/68/EEC, 91/492/EEC and 91/493. (Directive 92/65/EEC Article 2 (b))

2.

The following animal species are referred to in the Directives mentioned above: Directive

Article

90/426/EE Art. 2 C (b)

3.

Species

Comments

Equidae means wild or domesticated animals of the equine (including zebras) or asinine species

i.e. all Equidae species

64/432/EE Art. 2 (b) and C (c)

Bovines including Bison bison (American bison) and Bubalus bubalus (Domestic buffalo) and swine for slaughter, breeding or production Art. 2 (1 Ovine and caprine animals for slaughter, breeding and fattening and 2)

Wild cattle, except American bison, and wild Suidae, including ‘feral pigs’, do not fall under 64/432/EEC

91/68/EE C

Wild species do not fall under 91/68/EEC

90/539/EE Art. 2 C (1)

Fowl, turkeys, guinea fowl, ducks geese, quails, pigeons, pheasants, partridges and ratites reared or kept in captivity for breeding, the production of meat and eggs for consumption, or for re-stocking supplies of game

‘breeding’ in this context means production of hatching eggs for the production of animals for breeding, the production of meat and eggs for consumption, or for restocking supplies of game

91/67/EE C

‘aquaculture animals’ i.e. live fishes, crustaceans or molluscs coming from a farm, including those from the wild intended for a farm

i.e. animals in a zoo or aquarium do not fall under 91/67/EEC if animals are transferred from a zoo to another zoo, this transfer is covered by 91/67.

Art. 2 (1)

Although the animals listed in the above tabulation do, in principle, not fall under 92/65/EEC, it would make no sense to exclude them from the health surveillance plan (see Section C paragraph 3 of these recommendations).

VI.

Recommendations

B. The approved veterinarian 1.

In order to be granted official approval under Article 13 of the Directive, a zoo must secure by contract or legal instrument the services of a veterinarian approved by and under the control of the competent authority (ANNEX C paragraph 1.(g)).

2.

The role of the approved veterinarian is to ensure that the requirements of the present directive and other related legislation are complied with on a day to day basis.

3.

Where this veterinarian is a member of a practice, other members of the same practice may be included provided that they are also approved by the competent authority and individually nominated in writing.

4.

The approved veterinarians shall comply mutatis mutandis with the requirements referred to in Article 14(3)(B) of Directive 64/432/EEC (ANNEX C paragraph 1.(g)(i)).

5.

Taken directly from Directive 64/432- Article 14 (3)(b) “According to this article, the approved veterinarians must: i. meet the conditions for pursuing the veterinary profession; ii. have no financial interest or family links with the owner of or person responsible for the holding (but see point 6 below); iii. possess particular knowledge in the field of animal health as it applies to animals of the species concerned. This means that they must:  regularly update their knowledge, especially as regards the relevant health regulations,  meet the requirements laid down by the competent authority to ensure the proper functioning of the network,  provide the owner of or person responsible for the holding with information and assistance in order that all steps are taken to ensure that the holding's status is maintained, particularly on the basis of programmes agreed with the competent authority,  ensure compliance with the requirements concerning: (i) the identification and health certification of the animals of the collection, the animals introduced and those traded; (ii) compulsory reporting of infectious animal diseases and any other risk factor for animal health or welfare, and for human health; (iii) establishing as far as possible the cause of death of animals and where they are to be consigned; (iv) the hygiene conditions of the herd and of the livestock production units.”

6.

For the purposes of the implementation of the present Directive point ii. above is of less importance than it is in Directive 64/432/EEC as zoo animals have a conservation value rather than an economic value and because, for the purposes of the present Directive, the approved veterinarian is working under the supervision of the official veterinarian. It is thus up to the official veterinarian to assess whether there could be a conflict of interest, and whether the approved vet appointed by the zoo fulfils the requirements above, and in particular has appropriate specialist knowledge in relation to zoo animals.

7.

The competent authority shall draw up lists of approved veterinarians and of the approved holdings participating in the network. If the competent authority finds that a participant in the network no longer fulfils the conditions set out above, it shall suspend or withdraw approval, without prejudice to any penalties that may be applied.

VI.

Recommendations

C. The Annual Disease Surveillance Plan 1.

The approved veterinarian has to draw up and implement an annual disease surveillance plan (ANNEX C paragraph 1 (g) (ii) 1st indent). This plan is subject to annual audits by an official veterinarian from the competent authority (ANNEX C paragraph 2 (a) (ii).

2.

For the purposes of approval under the present directive the surveillance plan must cover those diseases listed in Annex A (and B if relevant).

3.

The surveillance plan may also include other general measures as may be required under Council Directive 1999/22/EC of 29 March 1999 relating to the keeping of wild animals in zoos (“Zoo Directive”, O.J. L94/24 0f 09.04.1999), and specific measures for individual taxonomic groups as may be agreed by the relevant Taxonomic Advisory Group from the European Endangered Species Program (EEP) of the European Association of Zoos and Aquaria (EAZA). As a general rule such specific measures would be elaborated by the EAZWV Infectious Diseases Working Group and subsequently be integrated into the Husbandry Guidelines for the taxon concerned.

4.

Animals for the purpose of this surveillance programme mean those species that are covered by article 13 of Directive 92/65/EEC; namely any species susceptible to the diseases listed in Annex A or Annex B. In practice this means all mammals, all birds, fishes of the salmonid group, and honey bees but not other invertebrates. Where animals of the domestic species are kept within a zoo premises, for example in a children's zoo, they will be regarded as part of the zoo collection and subject to all the same conditions as the rest of the collection as far as approval is concerned, including the surveillance programme. Note in this context that there may be specific requirements for domestic animal species that are covered by other Directives. Reptiles and amphibians and fishes other than salmonids are not included.

5.

The Annual Disease Surveillance Plan and the measures based thereon must include a.

Immediate notification to the competent authority if there is any cause for suspicion that animals may be affected by any disease, including zoonoses, that is notifiable under Community legislation (including 92/65 and Directive 82/894 and other relevant Community legislation) or national legislation.

b.

Close observation of each animal at least once per day by suitably qualified staff, under the direction of the approved veterinarian (in the case of large group species, such as fish in an aquarium, the veterinarian may decide that observation of the group is sufficient).

c.

Immediate notification of the approved veterinarian by zoo staff if any animals appear unwell or die (in the case of large group species, notification may be triggered by mortality above an agreed, expected level).

d.

Laboratory examination to establish the infective agent in any live animals that appear to be affected by an infectious disease (in the case of large group species, such as fish in an aquarium, the veterinarian may decide that a representative sample is sufficient). In the case of suspicion of a disease that is listed on Annex A and B and/or is notifiable under national legislation, the official veterinarian must be informed immediately. The official veterinarian will be responsible for arranging disease control precautions and further investigation for diseases notifiable under national legislation, and may direct that samples should be taken and submitted to a designated laboratory.

VI.

Recommendations

e.

Procedures for newly arrived and diseased animals, taking into account the relevant risk factors and, therefore, including handling practices, clinical examination and specific tests as appropriate.

f.

Regular parasitological examination of faecal samples (individual or group samples, depending on the housing system) in particular with regard to zoonotic parasites. It is recommended by EAZWV that all relevant groups should be checked at least once a year; the frequency of examination should be related to the prevalence of parasites.

g.

Opportunistic examination and taking of appropriate samples from immobilised or otherwise restrained animals. EAZWV recommends all serum samples to be retained and stored at –180 C or below.

h.

Specific guidelines for the systematic testing of specific animal species may be developed and recommended by the Infectious Diseases Working Group of EAZWV.

i.

Post mortem examination without unnecessary delay to check for significant pathology, and as far as possible to establish the cause of death in every animal that dies or foetus that is aborted (but the approved veterinarian may exercise discretion where there is clearly no suspicion of infectious disease, such as obvious trauma, or euthanasia of a healthy animal; and where it has been established that an infectious disease is affecting a group, the veterinarian may decide, in consultation with the competent authority if necessary, that a representative sample is sufficient).

j.

The vaccination programme should be based on the availability of safe vaccines. It should take into account the species involved and the risk of diseases likely to occur in the zoo, and may cover zoonotic diseases other than those mentioned in Annex A or B, but these vaccinations must be in compliance with the applicable legislation.

k.

Records must be kept in an easily accessible form, to be available as necessary for audit purposes, and retained for at least 10 years, to show at least the following information:  All cases of disease, and treatment if applicable.  Preventive actions such as vaccinations.  Results of blood tests and other diagnostic procedures.  Results of post mortem examinations including records of stillbirths.  Observations during any periods of precautionary isolation.  Reports to the veterinary authority of any suspicion of Annex A diseases or diseases notifiable under national law.

l.

Zoo vets should be aware that they will be asked for specific information on diseases under the zoonosis directive and should therefore be able to extract this information easily.

VI.

Recommendations

D. The Added Animals Procedure 1.

2.

3.

1

General a.

Only animals coming from another approved body, institute or centre may be introduced into an approved zoo (ANNEX C paragraph 2 (b). There is, however a derogation from this requirement provided that certain conditions are respected (ANNEX C paragraph 3).

b.

By way of derogation from Article 5(1) of the Directive, ANNEX C paragraph 3 allows also for the introduction of “apes” (i.e. non-human primates)1 from non-approved sources.

c.

Animals having an origin other than an approved body, institute or centre may be introduced in an approved zoo provided they undergo a quarantine under official control before being added to the collection (ANNEX C paragraph 3).

d.

Animals from approved zoos and from non-approved sources should not be transported together in the same container or vehicle.

e.

The animals must be accompanied by transport permits or any other documentation as may be required by national legislation. ANIMO requirements apply.

Animals from another approved establishment a.

Animals coming from another approved establishment in the same member state fall outside the scope of Directive 92/65, and hence under Community legislation there is no requirement for the animal to be accompanied by the model health certificate in Annex E. However, national rules governing certification may apply. For the same reason, there is no official requirement for post-arrival isolation, although the establishment may choose to carry out isolation and/or testing for its own private purposes.

b.

If the animals are coming from an approved establishment in another Member State they must be accompanied by the model health certificate in Annex E type 3. Depending of the health situation there may be additional requirements imposed by EU or national legislation.

Animals from a non-approved establishment in the same Member State a.

Animals coming from a non-approved establishment in the same member state fall outside the scope of Directive 92/65, and hence under Community legislation there is no requirement for the animal to be accompanied by the model health certificate in Annex E. However, national rules governing certification may apply. However, in accordance with Annex C of Directive 92/65, the animals must undergo post-arrival isolation in the isolation area, designated in the terms of approval, for at least 30 days or such longer period as may be required by the approved veterinarian and/or the competent authority to be satisfied that the health status of the animals is not inferior to the health status of the other animals in the collection.

b.

During isolation the animals may be required to undergo testing for any disease on Annex A that the approved veterinarian and the competent authority consider appropriate. They should have particular regard to diseases for which national programmes are in place, such as tuberculosis, brucellosis, Aujeszky's disease. The approved veterinarian may also wish to carry out specific testing for any diseases for

“apes” is zoologically not correct but it is the term used in the core text of the directive

VI.

Recommendations

which the premises runs its own surveillance, control and eradication programme, covering diseases other than those listed in Annex A. c.

4.

The arrival of the new animals must be recorded by the receiving establishment, as laid down in Annex C. There is no official requirement for any other health certification.

Animals from a non-approved establishment in another Member State a. Member states may, by way of derogation, allow the movement of animals from non approved establishments in another member state. The Veterinary Service of the receiving country has to be informed under paragraph 3 of ANNEX C and may lay down specific conditions under which transfer must take place. In addition, in accordance with Annex C of Directive 92/65, the animals must undergo post-arrival isolation in the isolation area, designated in the terms of approval, for at least 30 days or such longer period as may be required by the approved veterinarian and/or the competent authority.

5.

6.

Animals from non-approved establishment to a non-approved establishment a.

Between establishments in the same Member State: National rules apply

b.

Between establishments in different Member States: Not covered by Annex C of Directive 92/65. This means that the rules laid down in the body of the Directive apply in full. Therefore, where no provisions are laid down, the Member State of destination can request specific requirements for introduction ( e.g. requests for additional guarantees such as the status, negative tests or treatments see Article 6 of Directive 92/65 for details )

Animals from third countries a.

Animals being imported into the Community must fulfil the animal health conditions as laid down in Directive 92/65. However where harmonised rules for a particular species have not been laid down in the Directive, then national rules shall apply, although these also should be based on the animal health principles laid down in the Directive. EU rule, or if not applicable national rules, for licensing, health certification, post-import quarantine and testing will apply, as appropriate for the species. The importing zoo must apply for a specific import licence, which will contain the conditions relevant to the species and place of origin.

(Please note that new legislation (amending Decision 79/542/EEC)is due to come into force in the near future that will lay down a list of third countries and harmonised rules (certification and animal health requirements) for the importation of all ungulates (including all Artiodactyla). Because of these rules, imports of such animals will only be allowed exclusively from a small number of third countries authorised for each species. However a draft Decision is under discussion which foresees a particular regime for importation of live animals originating in any third country but imported after a residency period in St.Pierre and Miquelon (a little island in the Atlantic ocean close to Canada) where they will spend a period in a quarantine station. During this period, specific testing will be carried out on the animals. For the moment, these special conditions are limited to the import of live Camelidae, but the intention is to extend this possibility to other species. Following introduction of this new legislation national rules will therefore no longer apply to imports belonging to the order Artiodactyla.

VI.

Recommendations

E. Quarantine / Isolation requirements 1.

2.

Definitions a.

‘Isolation’ and 'quarantine' are not precisely defined in European Union legislation, and one word is usually described by reference to the other. For example in the poultry trade directive 90/539/EEC: ‘Quarantine station shall mean facilities where the poultry is kept in complete isolation and away from direct or indirect contact with other poultry, so as to permit long-term observation and testing’ (Council Directive 90/539/EC, Article 2).

b.

The Office International des Epizooties (OIE) Terrestrial Animal Health Code defines a quarantine station as 'a facility under the control of the veterinary authority where a group of animals is maintained in isolation., with no direct or indirect contact with other animals, in order to undergo observation for a specified length of time and, if appropriate, testing and treatment.

c.

In order to emphasize the difference between quarantine of the above types and quarantine required for added animals under the BALAI Directive, the latter is referred to as 'isolation' throughout this document.

d.

The conditions below refer to isolation for added animals entering a BALAI approved zoo from a non-approved source within the EU (or in Norway, Liechtenstein, and Switzerland).

Principles a.

In order to be granted approval, zoos must have adequate means for isolating animals, and have available adequate quarantine facilities and approved procedures for animals from non-approved sources. (ANNEX C paragraph 1. (b).

b.

Incoming animals must be isolated as necessary (ANNEX C paragraph 1 (g) (iv).

c.

Animals having an origin other than an approved body, institute or centre may be introduced in an approved zoo provided they undergo quarantine under official control before being added to the collection (ANNEX C paragraph 3).

d.

For the purposes of the approval under the Directive only the diseases listed in Annex A have to be taken into account. In addition, it should be considered that the introduction of new animals susceptible to these diseases is only possible either from within the EU or from listed third countries.

e.

A risk analysis has to be made and the quarantine / isolation requirements must cope with the risk. Quarantine requirements for comparable livestock could provide some guidance. In this context it is noted that management procedures could be adjusted easily to each individual case, but that the availability of suitable facilities is a prerogative for approval and has to be seen without a specific case in mind.

f.

For the purposes of the approval , i.e. for the introduction of animals from nonapproved sources within the European Union or from listed Third Countries where such lists exist, the following information may be useful when considering, which general requirements to apply: There are three risk groups:  Primates: can be imported from anywhere (no Third Countries List), they may be carriers of zoonoses.

VI.

Recommendations

 Birds: the introduction from areas where OIE list A diseases exist can not be excluded (occurrence of NCD in wild birds), and the relevant diseases, NCD, AI and Psittacosis are easily transmitted via the air or, in the case of West Nile Virus, by mosquitoes.  Mammals other than primates: introduction only from areas free from highly contagious diseases, all relevant diseases not transmittable by air over a longer distance, in most cases direct contact required. g.

Consequently, the following general requirements apply:  Primates: The quarantine requirements laid down in the OIE Terrestrial Animal Health Code (Chapter 2.11 and Appendix 3.5.1) shall be respected (ANNEX C paragraph 3).  Birds must be isolated in buildings and the possibility of disease transmission by air or insects has to be taken into account. Windows should be kept closed. EAZWV strongly recommends that the isolation rooms should be ventilated, and the exhausted air should pass through a dust filter.  Mammals should, as a general rule, be isolated indoors, but no special precautions have to be taken regarding the exhausted air to cope with the relevant diseases listed in Annex A of the Directive. If, for specific reasons, mammals have to be isolated outdoors, the ground should be solid and easy to disinfect. If this is not possible, the isolation enclosure should be relatively small to allow for other treatment of the soil, e.g. removal of top soiling. No zoo will be able to have specific isolation facilities for all mammalian taxa, which may include a diverse range of species, including e.g. big cats, dolphins, elephants, hippopotamuses. In such cases it should be possible to use the standard facility for isolation purposes.

h.

3.

In order to be granted approval, zoos must have available adequate quarantine / isolation facilities. This wording does not imply that the facilities are on the ground or owned by the zoo concerned. In addition the option exists for several zoos to jointly operate a facility, or have contracts among themselves. In this last case, the option should be specified in the annual plan.

General conditions - Structural requirements a.

Location

The isolation quarters must be physically separated from other animal accommodation by a reasonable distance, taking into account the species concerned and the ability of the relevant viruses to spread on the air. This distance can be much reduced if the exhausted air is filtered (for animals originating from within the EU or from listed Third Countries the use of dust filters is sufficient, otherwise High Efficiency Particulate Extraction (HEPA) may be required). b.

Demarcation

The limits of the isolation area must be clearly demarcated by walls or fences as appropriate. This does not preclude the possibility that specific areas or pens within the premises may be designated as isolation areas for a limited time and a particular purpose, provided that they meet the general requirements. c.

Access

There must be a double door system to prevent escape at the entry/exit with sufficient space between the doors to allow one to be closed before the other is opened. Entry/exit

VI.

Recommendations

doors must be lockable and must display a notice stating: 'QUARANTINE: No Admission to Unauthorised Persons'. d.

Hygiene barrier

Facilities must be available at the entry/exit point for attendants to change overalls, to change and disinfect boots, to wash hands, and if appropriate to shower. e.

Loading/Unloading.

Suitable facilities must be available to load or unload animals between transport crates and isolation pens without the risk of escape. f.

Restraint

Suitable crush or penning facilities should be available within reasonable access of the isolation area, so that animals may be safely restrained for clinical and diagnostic procedures such as blood sampling. The route from isolation to restraint must not put other animals at risk of infection from the introduced animals. g.

Inspection

The design of the pens or cages within the isolation area must be such that the animals may be visually inspected at any time, with adequate light and ease of access. h.

Disinfection

The physical structure and all equipment must be made of such materials that they can be effectively cleansed and disinfected, or destroyed after use. i.

Vermin

The design must be suitable to minimise access by rodents, wild birds and insects, as appropriate for the species in question. Where drains are present, they must be fitted with rodent proof covers. j.

Feed Store

The feed store must be suitably protected from vermin. k.

Waste Disposal

Adequate storage facilities must be available to contain the litter and animal waste produced during the isolation period, and the storage facility must be bird and vermin proof. There must be facilities to dispose of the waste either during or after the isolation period in a way which will ensure that there is no risk of the spread of disease. l.

Post Mortem

Refrigeration facilities or equivalent must be available within the isolation area, or in a suitably disease-protected location nearby, to hold carcases of animals that die until they can be subject to post mortem examination. Procedures for conveying carcases safely to the storage facility must be laid down in writing by the approved veterinarian. 4.

General conditions - Management procedures a.

Surveillance

Every animal in isolation must be visually inspected at least once a day by suitably competent staff. Any signs of illness must be recorded and reported immediately to the responsible veterinarian, who should make a further examination of the affected animals without any unreasonable delay. b.

Staff

VI.

Recommendations

The premises must have designated staff who are present on a sufficiently regular schedule to ensure surveillance of the animals on a daily basis, and more frequently if appropriate. c.

Hygiene

Staff entering the isolation premises must always change into protective clothing and footwear. On leaving, the overalls and footwear must be removed and left within the isolation area, and the footwear must be disinfected. Hands must be washed, or otherwise disinfected, on entering and leaving. d.

Equipment

None of the moveable items used in the isolation unit should be taken outside the unit, or used with other stock outside the unit, for the entire duration of the isolation period. e.

Waste

Litter and waste material must be collected regularly, stored in the containers provided, and disposed of either during or after the isolation period in such a way that disease agents will not be spread. f.

Disinfection

Premises must have an effective programme, laid down in writing by the approved veterinarian, for cleansing and disinfection after each isolation session; approved disinfectants must be specified and used in the programme; and an appropriate resting period (usually 7 days) must be specified after each cleansing and disinfection operation. g.

Transport Crates

Crates or cages used for transport, if to be re-used, must be made of materials which allow effective cleaning and disinfection, and this should be carried out within the isolation unit. If not re-used, the crates and cages must be destroyed in such a way that disease agents cannot be spread. h.

All-in, All-out

An ‘all-in, all-out’ policy should be followed in the isolation unit. If it is necessary to add animals whilst others are already present in the unit, the isolation period of all of them must be extended until the latest completion date of any of the animals. i.

Illness

If any animals become ill during isolation and the approved veterinarian considers that they need to be moved to a specialised hospital facility for diagnosis or treatment, he/she must ensure that this is done under his/her personal supervision in such a way as to ensure no possible risk of disease spread. In particular the approved veterinarian must personally supervise the arrangements for maintaining isolation throughout the movement, and for disinfecting any vehicles, rooms and equipment with which the animal has had contact. j.

Disease and Death

Any sign of any disease or death during isolation must be reported immediately to the approved veterinarian. All suspicions of any infectious disease on Annex A and any deaths in isolation must be reported immediately to the competent authority. Carcases of animals, which die during isolation, and if necessary those that are dead on arrival, must be submitted to a post mortem examination without unreasonable delay. k.

Designated Attendants

VI.

Recommendations

The establishment must designate suitable staff to attend to the animals in isolation, taking appropriate precautions to ensure that there is no risk of transferring infection from the isolation unit to any other animals, and the arrangements must be agreed in writing by the approved veterinarian. l.

Visitors

Visitors must not be allowed to enter the isolation unit. If personnel apart from the designated attendants need to enter for essential maintenance etc., they must be required to wash thoroughly on entering and leaving, and wear protective clothing which shall be put on prior to entering and removed prior to leaving. There must be a visitors' book to record the dates, names and addresses of all visitors. m. Records The person in charge of the isolation unit must keep the following records, which should be retained for at least ten years 

the date, number and identification of animals entering and leaving the isolation facility.



copies of the export health certificates and border crossing certificates accompanying imported animals.



significant health observations, cases of illness and deaths on a daily basis.



dates and results of testing



dates and types of treatment



dates and names and addresses of persons entering the isolation unit.

n.

Duration

Isolation should normally last for at least 30 days, unless a longer period is required to exclude specific risks such as rabies. 5.

Additional requirements specifically for birds a.

Ventilation

Birds must always be isolated in buildings. There must be no possibility of access by wild birds or by mosquitoes. As a general rule, windows should be kept closed, the isolation rooms should be ventilated, and the exhausted air should pass through a dust filter. If the isolation facility is more than 250m away from other bird enclosures, this requirement can be waived and ventilation through open windows can be permitted. In such case all ventilation openings must be covered with a double layer of wire mesh. b.

Air Space

If there are separate units within the isolation facility, each unit must occupy a separate airspace so as to be an isolated epidemiological unit. If this cannot be achieved, all the birds in isolation must remain until the completion date of the last birds to enter. 6.

Additional requirements specifically for ungulates a.

Fencing

If the isolation area includes open paddocks, situations where there may be stock in adjacent paddocks must be avoided. The isolation paddocks must be surrounded by

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Recommendations

double fences allowing a suitable sanitary gap between the fences. A minimum gap of at least 3 meters should normally be satisfactory, but taking account of the species concerned, the competent authority may require a different standard. Both fences must be escape-proof. b.

Herds in Isolation

If the isolation facility is intended to contain large groups of animals, there must be additional provision so that any individual that appears to be unwell can be separated and kept apart from the rest of the group, with facilities for testing and treatment as appropriate. 7.

Additional requirements specifically for primates (based on the OIE Terrestrial Health Code, chapter 2.10.1. and Appendix 3.5.1. (2002 edition) a.

Zoonoses

Any biting or scratching incidents involving humans, or other events in which humans are exposed to primate blood or saliva, are to be reported immediately to the responsible zoo veterinarian, who should consult with medical authorities as appropriate. b.

Protection of Attendants

The overalls and boots provided for entry to the isolation facility should completely cover the attendant's body, and suitable masks, visors, goggles and gloves should also be provided (where this raises issues such as welfare and socialisation, the approved veterinarian should consult with the competent authority who may agree to alternative methods providing equivalent security). c.

Staff Training

The responsible veterinarian or physician should ensure that all attendants are fully instructed in the procedures necessary to protect their own health, as well as the health and welfare of the animals in isolation. Personnel must not eat, drink, smoke or store food for human use within the isolation rooms. d.

Staff Health

Personnel working within the isolation area should be encouraged to provide baseline serum samples, which would be stored for study and comparison if appropriate. Additional serum samples may be collected periodically as an aid to epidemiological investigations. Staff should be encouraged to report any signs of illness immediately to their medical adviser. e.

Ventilation

If natural ventilation is used, the openings must be covered with a double layer of mesh, each of which is individually strong and secure enough to prevent the escape of the animals. Ventilation intakes and outlets must not be so close to any other animal holding area as to present a disease risk. If forced ventilation with HEPA filtration is used, there should be provision to maintain adequate ventilation in the event of a technical failure. Separate units must be ventilated separately. f.

Washing facilities

Washing facilities with hot and cold running water should be available for personnel to wash hands within each animal holding room. Personnel should wash or otherwise disinfect hands at frequent intervals whilst working within the isolation premises.

VI.

g.

Recommendations

Footbaths

Footbaths should be available not only at the entrance/exit of the isolation premises, but also between individual holding rooms within the premises. The footbaths should contain an approved disinfectant agreed by the approved veterinarian. Personnel should use the footbaths as they pass from one room to another. h.

Equipment

Each holding room should have its own complete range of dedicated equipment, and equipment should not be transferred from one holding room to another. After use all equipment including work surfaces should be effectively cleaned and disinfected. Because of the aerosol risk power hoses should not be used, except with the agreement of the approved veterinarian. i.

Group Separation

Separate groups entering the isolation premises from different sources or on different occasions must remain physically and epidemiologically isolated from each other. Separate groups must be accommodated in separate, isolated units. Animals may not be transferred between groups. However where this raises issues such as welfare and socialisation, the approved veterinarian in consultation with the competent authority may agree to mixing animals, provided that isolation conditions then apply to all those in contact with the introduced animals. j.

Cage Discipline

No animals may be removed from their cages, albeit within the self-contained isolation premises, without the specific authority and supervision of the responsible zoo veterinarian. k.

Duration

OIE recommends a quarantine duration of at least 30 days when the primate is sent from another premises under veterinary supervision, and at least 12 weeks if it is coming from circumstances without veterinary supervision or from the wild. In Great Britain under rabies regulations the duration of quarantine for primates must be at least 6 months.

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Recommendations

F. The certificates Where certificates are required 1.

‘Animals’ in the terms of article 2(b) of Directive 92/65/EEC must be accompanied by a certificate corresponding to the specimen in Annex E (Directive 92/65, Article 5).

2.

Bovines and swine falling under the scope of Directive 64/432/EEC must be accompanied by a certificate conforming to the model set out in Annex F of that Directive.

3.

Ovines and caprines falling under the scope of directive 91/68/EEC must be accompanied by a certificate conforming to the model set out in Annex E of that Directive.

4.

Equines (including zebras) must be accompanied by a certificate complying with Annex C of Directive 90/426/EEC.

5.

Poultry and ratites falling under the scope of Directive 90/539/EEC must be accompanied by a certificate complying with Annex IV of that Directive.

Transmissible Diseases Handbook

V.

ANIMAL HEALTH LEGISLATION IN EUROPE

Peter Dollinger VDZ / zooschweiz / EAZWV Office Berne, Switzerland (Chapter updated in 2009)

1. Introduction As a result of the Uruguay round, an Agreement on the Application of Sanitary and Phytosanitary Measures (the "SPS Agreement") was developed and adopted. It entered into force with the establishment of the World Trade Organization on 1 January 1995. It concerns the application of food safety and animal and plant health regulations. The Agreement allows countries to set their own standards. But it also says regulations must be based on science. They should be applied only to the extent necessary to protect human, animal or plant life or health. And they should not arbitrarily or unjustifiably discriminate between countries where identical or similar conditions prevail. Member countries are encouraged to use international standards, guidelines and recommendations where they exist. However, members may use measures which result in higher standards if there is scientific justification. They can also set higher standards based on appropriate assessment of risks so long as the approach is consistent, not arbitrary. All European countries have issued legislation to control the spread of transmissible diseases within their territory and to protect livestock and the human population from the introduction of exotic diseases. As a function of diseases occurring in a given country, of the trade patterns prevailing, and of the desired level of protection, legal provisions may vary from one country to another. There are, however, two mechanisms leading in many respects to a standardisation of the legal requirements throughout the whole of Europe: a. all countries, except the Holy See (where hardly any animals are kept), Monaco and San Marino (which have to apply EC legislation) are members of the Office International des Epizooties, and b. 27 countries (Austria, Belgium, Bulgaria, Cyprus, Czech Republic, Denmark, Estonia, Finland, France, Germany, Greece, Hungary, Ireland, Italy, Latvia, Lithuania, Luxemburg, Malta, Poland, Portugal, Romania, Slovakia, Slovenia, Spain, Sweden, The Netherlands and the United Kingdom) are members of the European Union. EU veterinary legislation applies also in the British Crown Dependencies (the Channel Islands and the Isle of Man), the Faeroe Islands, Andorra (Protocol on veterinary matters supplementary to the agreement in the form of an exchange of letters between the European Economic Community and the Principality of Andorra - OJ L 148 06.06.1997 p.16) and San Marino (Decision No 1/94 of the EC-San Marino Cooperation Committee of 28 June 1994 on Community veterinary regulations to be adopted by the Republic of San Marino - OJ L 238 13.09.1994 p.25). Norway has adopted Community legislation in the framework of the EEA Agreement (Iceland and Liechtenstein are also members of the EEA Agreement, but have been exempted

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Animal Health Legislation in Europe

from its veterinary provisions). Switzerland and the EU have concluded a bilateral agreement by which the equivalence of the respective animal health legislation has been mutually recognised (Agreement between the European Community and the Swiss Confederation on Trade in Agricultural Products). This agreement entered into force in spring 2002. In the meantime, Switzerland has been almost fully integrated into the EU veterinary area, and Annex 11 of the agreement, dealing with veterinary matters, has been made applicable also to Liechtenstein. As a consequence, veterinary checks on shipments between Switzerland, Liechtenstein and the EU have been waived, and Switzerland carries out EU Third Country checks at the two intercontinental airports of Zurich and Geneva. Finally, there is a REGULATION (EC) No 998/2003 on the animal health requirements applicable to the non-commercial movement of pet animals and which addresses also the movement of pet animals between the EU and the Holy See and Iceland The present chapter is, therefore, limited to an introduction to OIE mechanisms and standards and to the EU legislation on animal health.

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Animal Health Legislation in Europe

2. The Office International des Epizooties (OIE) The OIE is an independent world wide organisation whose membership is made up from 172 sovereign states. The headquarters are located at Paris. The organisation maintains an internet site at the URL http://www.oie.int which provides up-to-date information on the animal health situation in member countries and gives online access to some relevant OIE publications.

OIE’s objectives are to 

Ensure transparency in the global animal disease situation



Collect, analyse and disseminate veterinary scientific information



Encourage international solidarity in the control of animal diseases



Safeguard world trade by publishing health standards for international trade in animals and animal products



Improve the legal framework and resources of national Veterinary Services



Provide a better guarantee of food of animal origin and to promote animal welfare through a science-based approach

The OIE was originally founded to combat highly contagious cattle diseases, like rinderpest and foot-and-mouth disease, and focuses today primarily on diseases affecting agricultural livestock and which have either severe economic implications or a zoonotic potential. In the OIE Terrestrial Animal Health Code, it makes recommendations to the veterinary administrations of importing and exporting countries on how to regulate trade in animals in order to prevent the introduction of transmissible diseases. The complete text of the Terrestrial Animal Health Code 2008 can be found at the URL http://www.oie.int/eng/normes/mcode/en_sommaire.htm. The Code contains a list of diseases for which recommendations are made: 

Multiple species diseases: This section contains 26 diseases including anthropozoonoses (e.g. brucellosis), parasitoses (e.g. echinococcosis) and viral diseases affecting a wider range of taxa (e.g. foot-and-mouth disease).



Cattle diseases contain 14 diseases including e.g. BVD and IBR/IPV, but also BSE and bovine tuberculosis, which, interestingly, are not rated as multiple species diseases.



Sheep and goat diseases include a list of eleven, like e.g. Maedi-Visna, scrapie, or sheep and goat pox.



There are also 11 equine diseases, including e.g. African horse sickness, equine infectious anemia, or glanders.



The seven swine diseases include traditionally well-known, highly contagious diseases, such as African and classical swine fever, but also diseases, which were discovered, or were considered of economic relevance, only recently such as Nipah virus or the porcine reproductive and respiratory syndrome.



Avian diseases include 14 traditionally well-known diseases. Avian flu is listed as “Highly pathogenic avian influenza in birds and low pathogenicity notifiable avian influenza in poultry”, meaning that low pathogenic avian influenza in wild birds does not fall under the scope of the Code.

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

Animal Health Legislation in Europe

The remaining three categories are made up by two Lagomorph diseases (myxomatosis and rabbit haemorrhagic disease), seven diseases of bees, and two other diseases, namely camel pox and leishmaniosis.

The Code recommends measures mainly with regard to diseases in domestic animals. Wildlife is rarely explicitly mentioned, but the same criteria may be applied mutatis mutandis. A separate standard which is also regularly updated is the International Aquatic Animal Health Code. It currently refers to 10 fish, 7 crustaceans, and 7 mollusc diseases. To improve the knowledge about the presence of infectious diseases in wildlife, however, and to create awareness, an OIE Working Group on Wildlife Diseases has produced annual reports since 1992. In 1996, an OIE-EAZWV Working Group began with the drafting of a recommendation on zoonoses transmissible from non-human primates. In 1998, the draft was adopted and included as Chapter 6.9 in the OIE Terrestrial Animal Health Code. The chapter focuses on defining the health of non-human primates and on the practice of protective measures against disease transmission. It emphasises the process of quarantining after international transportation, and stresses that some degree of risk for zoonotic disease transmission should always be recognised. In 1999, an annex on quarantine requirements was adopted. It is now Chapter 5.9 of the Code. For standards for testing non-human primates, the OIE Manual of Standards for Diagnostic Tests and Vaccines (the full text of the Standards Manual is found at http://www.oie.int/eng/normes/mmanual/a_summry.htm) recommends consulting the following document: Health monitoring of non-human primate Colonies. Recommendations of the Federation of European Laboratory Animal Science Associations (FELASA) Working Group on Non-Human Primate Health accepted by the FELASA Board of Management, 21 November 1998 (http://www.lal.org.uk/pdffiles/LAfel5.pdf) .

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Animal Health Legislation in Europe

3. The Veterinary Legislation of the European Union

In 1964, the European Union – at that time still as the European Economic Community – started to issue legislation in the veterinary field. The first of these legal acts has been amended many times but is still in effect. Originally, the EU has produced only Directives and Decisions in the veterinary field, as of 1990, also Regulations were issued, e.g. in the context of BSE, animal by-products, pet animals, or circus animals.

As at March 1, 2009, there were 938 valid acts in the field of animal health and zootechnics. Many of these have been repeatedly amended, and in a number of cases it had become necessary to amalgamate the original version and all amendments to a consolidated version serving as a documentation tool. 

Directives are addressed to the member states and must be implemented through national legislation. This leaves the national authorities with some degree of freedom regarding the ways by which they want to achieve the goals and policies set by the EU.



Decisions must be implemented to the letter (examples: certificates, lists of approved establishments).



Regulations are addressed directly to EU citizens and companies, i.e. the authorities have to follow them very closely.

If the EU has decided to control or eradicate a disease, the measures agreed upon are binding for all member states. This means e.g. that an individual member state is not allowed to vaccinate against a given disease if a non-vaccination policy is applied by the Community. In areas not regulated by the EU, Member States are more or less free to set their own rules. These must not lead to a distortion of intra-Community Trade. The countries may, however, apply for additional guarantees if they have been able to eliminate a disease from their territory or parts thereof. The following are some EU acts of relevance either for the monitoring and control of transmissible diseases or laying down conditions for the intra-community trade and import from third countries. The text of these acts or other legislation can be found at the URL http://eur-lex.europa.eu/. Most of the texts referred to are relevant also in the EEA context (Norway, Liechtenstein) or under the bilateral agreement with Switzerland. A. Trade and placing on the market The legislation referred to in this section defines the conditions under which animals may be moved between EU Member States. As a general rule, it is required that the animals come from an area and from a holding which is free from certain diseases. The animals themselves must be identified in agreement with prescribed marking systems (e.g. cattle, sheep and goats must be double ear-tagged), or must be otherwise identifiable (e.g. horse passport), and they must be healthy, in particular free from specified diseases, and fit for transport. Most requirements apply to all Member States. As a result of a particularly favourable health situation, certain member States are, however, allowed to request additional guarantees (e.g. Denmark, Austria and others regarding IBR/IPV, or the majority of Member States regarding

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Animal Health Legislation in Europe

Aujeszky’s Disease) and/or to take specific precautions (e.g. United Kingdom regarding rabies). 

Council Directive 64/432/EEC of 26 June 1964 on animal health problems affecting intraCommunity trade in bovine animals and swine - OJ P 121 29.07.1964, p.1977.



Council Directive 91/68/EEC of 28 January 1991 on animal health conditions governing intraCommunity trade in ovine and caprine animals - OJ L 46, 19. 2. 1991, p. 19.



Council Directive 90/426/EEC of 26 June 1990 on animal health conditions governing the movement and import from third countries of equidae - OJ L 224, 18. 8. 1990, p. 42.



Commission Decision 93/623/EEC of 20 October 1993 establishing the identification document (passport) accompanying registered equidae - OJ L 298, 3.12.1993, p. 45–55.



Commission Regulation (EC) No 504/2008 of 6 June 2008 implementing Council Directives 90/426/EEC and 90/427/EEC as regards methods for the identification of equidae - OJ L 149, 7.6.2008, p. 3–32.



Council Directive 90/539/EEC of 15 October 1990 on animal health conditions governing intraCommunity trade in, and imports from third countries of, poultry and hatching eggs - OJ L 303 31.10.1990, p.6.



Commission Decision 2006/605/EC of 6 September 2006 on certain protection measures in relation to intra-Community trade in poultry intended for restocking of wild game supplies - OJ L 246, 8.9.2006, p. 12–14.



Council Directive 91/67/EEC of 28 January 1991 concerning the animal health conditions governing the placing on the market of aquaculture animals and products - OJ L 046 19.02.1991, p. 1.



Council Directive 92/65/EEC of 13 July 1992 laying down animal health requirements governing trade in and imports into the Community of animals, semen, ova and embryos not subject to animal health requirements laid down in specific Community rules referred to in Annex A (I) to Directive 90/425/EEC -OJ L 268, 14. 9. 1992, p. 54 – BALAI Directive.



Regulation (EC) No 998/2003 of the European Parliament and of the Council of 26 May 2003 on the animal health requirements applicable to the non-commercial movement of pet animals and amending Council Directive 92/65/EEC – OJ L 146 13.06.2003, p.1.



2003/803/EC: Commission Decision of 26 November 2003 establishing a model passport for the intra-Community movements of dogs, cats and ferrets - OJ L 312, 27.11.2003, p. 1–13.



Commission Regulation (EC) No 1739/2005 of 21 October 2005 laying down animal health requirements for the movement of circus animals between Member States - OJ L 279 22.10.2005, p. 47.



Regulation (EC) No 1760/2000 of the European Parliament and of the Council of 17 July 2000 establishing a system for the identification and registration of bovine animals and regarding the labelling of beef and beef products and repealing Council Regulation (EC) No 820/97 - OJ L 204, 11.8.2000, p. 1–10.



Commission Regulation (EC) No 644/2005 of 27 April 2005 authorising a special identification system for bovine animals kept for cultural and historical purposes on approved premises as provided for in Regulation (EC) No 1760/2000 of the European Parliament and of the Council OJ L 107 28.04.2005, p. 18.



Commission Regulation (EC) No 509/1999 of 8 March 1999 concerning an extension of the maximum period laid down for the application of ear-tags to bison (Bison bison spp.) - OJ L 60, 9.3.1999, p. 53.

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Animal Health Legislation in Europe



Council Regulation (EC) No 21/2004 of 17 December 2003 establishing a system for the identification and registration of ovine and caprine animals and amending Regulation (EC) No 1782/2003 and Directives 92/102/EEC and 64/432/EEC - OJ L 005 09.01.2004, p. 8.



Council Directive 2008/71/EC of 15 July 2008 on the identification and registration of pigs (Codified version) - OJ L 213, 8.8.2008, p. 31–36.

After many years of lobbying by the zoo community and negotiations between EAZWV and the EU Commission, the BALAI Directive was amended by Commission Regulation (EC) No 1282/2002 of 15 July 2002 (OJ L 187/3). As a consequence, the Directive became more acceptable to the zoo community and its Annexes A, C and E are now directly applicable in Member States. As ANNEX C contains a few points which leave considerable room for interpretation. EAZWV, in cooperation with the EU Commission and some national veterinary services, developed recommendations aiming at contributing to a uniform application of the Directive, and thus at achieving the ultimate goal of this annex, namely to facilitate the exchange of animals between approved zoos easily and without major health risks. In spite of this effort, differences between Members States in implementing the Directive remained, and, in 2009, e.g. France still does not apply the Directive with regard to zoos. The core body of Directive 92/65/EEC, the Regulation 1282/2002 and the recommendations are annexed to this chapter. B. Importation from third countries A common import regime was established in 1972. On the basis of an assessment of the health situation in Third Countries and of the reliability of their veterinary services, the EU Commission has – for certain species – drawn up lists of countries eligible to export animals to the Community starting. As far as harmonised import conditions exist, the animals must be accompanied by an official veterinary health certificate which follows the models given in the specific directives or decisions, they must undergo a border veterinary check at an approved checkpoint on arrival, and they must undergo a quarantine period under supervision by the official veterinarian. The original Council Directive 72/462/EEC applied to cattle, sheep, goats and swine only. Successively other Directives regulating the import of other species were adopted, and in 2004 the old 72/462/EEC was replaced by a new Directive, which adds all wild even-toed ungulates, rhinos, tapirs and elephants to the list of regulated species. 

Council Directive 2004/68/EC of 26 April 2004 laying down animal health rules for the importation into and transit through the Community of certain live ungulate animals, amending Directives 90/426/EEC and 92/65/EEC and repealing Directive 72/462/EEC - OJ L 139 30.04.2004 p. 320.

Directive 2004/68/EC addresses the import and transit of even-toed ungulates of wild species, and of rhinos, tapirs and elephants. Wild equids still fall under the scope of Directive 90/426/EEC. In Annex I, a list of the relevant taxa is given. Annex II defines the basic conditions for a country being considered free from certain diseases. Annex III establishes the basic requirements for veterinary certificates. Annex IV replaces Annex F of Directive 90/426/EEC, and Annex V contains a list of implementing rules for the import of live animals, meat and meat products, which shall remain in force until replaced by measures adopted under the new regulatory framework. 

Council Decision 79/542/EEC of 21 December 1979 drawing up a list of third countries from which the Member States authorize imports of bovine animals, swine and fresh meat - OJ L 146 14.06.1979 p. 15.

Decision 79/542/EEC underwent frequent amendments – in 2008 alone it was amended six times, and it is strongly recommended to consult the consolidated text, which now contains also model certificates.

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Animal Health Legislation in Europe



Commission Decision 2004/212/EC of 6 January 2004 on Community health conditions on imports of animals and fresh meat including minced meat from third countries and amending Decisions 79/542/EEC, 2000/572/EC and 2000/585/ - OJ L 073 11.03.2004, p. 11.



Commission Decision 2004/211/EC of 6 January 2004 establishing the list of third countries and parts of territory thereof from which Member States authorise imports of live equidae and semen, ova and embryos of the equine species, and amending Decisions 93/195/EEC and 94/63/EC - OJ L 073 11.03.2004, p. 1.



2007/240/EC: Commission Decision of 16 April 2007 laying down new veterinary certificates for importing live animals, semen, embryos, ova and products of animal origin into the Community pursuant to Decisions 79/542/EEC, 92/260/EEC, 93/195/EEC, 93/196/EEC, 93/197/EEC, 95/328/EC, 96/333/EC, 96/539/EC, 96/540/EC, 2000/572/EC, 2000/585/EC, 2000/666/EC, 2002/613/EC, 2003/56/EC, 2003/779/EC, 2003/804/EC, 2003/858/EC, 2003/863/EC, 2003/881/EC, 2004/407/EC, 2004/438/EC, 2004/595/EC, 2004/639/EC and 2006/168/EC - OJ L 104, 21.4.2007, p. 37–50.



Commission Regulation (EC) No 798/2008 of 8 August 2008 laying down a list of third countries, territories, zones or compartments from which poultry and poultry products may be imported into and transit through the Community and the veterinary certification requirements OJ L 226, 23.8.2008, p. 1–94.



Council Directive 72/462/EEC of 12 December 1972 on health and veterinary inspection problems upon importation of bovine animals and swine and fresh meat from third countries OJ L 302 31.12.1972, p.28.



Commission Decision 93/198/EEC of 17 February 1993 laying down a model for the animal health conditions and veterinary certification for the importation of domestic ovine and caprine animals from third countries - OJ L 086 06.04.1993, p.34.



Council Directive 90/426/EEC of 26 June 1990 on animal health conditions governing the movement and import from third countries of equidae - OJ L 224 18.08.1990, p.42.



Commission Decision 93/197/EEC of 5 February 1993 on animal health conditions and veterinary certification for imports of registered equidae and equidae for breeding and production - OJ L 086 06.04.1993, p.1.



Commission Decision 2003/459/EC of 20 June 2003 on certain protection measures with regard to monkey pox virus - OJ L 154, 21.6.2003, p. 112–113).

Decision 2003/459/EC aims at preventing the introduction of monkey pox by the importation of Prairie dogs (Cynomys spp.) from the United States. 

Council Directive 90/539/EEC of 15 October 1990 on animal health conditions governing intraCommunity trade in, and imports from third countries of, poultry and hatching eggs - OJ L 303 31.10.1990, p.6.



Commission Decision 96/482/EC of 12 July 1996 laying down animal health conditions and veterinary certificates for the importation of poultry and hatching eggs other than ratites and eggs thereof from third countries including animal health measures to be applied after such importation - OJ L 196 07.08.1996, p.13.



Commission Regulation (EC) No 318/2007 of 23 March 2007 laying down animal health conditions for imports of certain birds into the Community and the quarantine conditions thereof - OJ L 84, 24.3.2007, p. 7–29.



Commission Decision 2006/696/EC of 28 August 2006 laying down a list of third countries from which poultry, hatching eggs, day-old chicks, meat of poultry, ratites and wild game-birds, eggs and egg products and specified pathogen-free eggs may be imported into and transit through the Community and the applicable veterinary certification conditions, and amending Decisions 93/342/EEC, 2000/585/EC and 2003/812/EC - OJ L 295, 25.10.2006, p. 1–76.



2003/881/EC: Commission Decision of 11 December 2003 concerning the animal health and certification conditions for imports of bees (Apis mellifera and Bombus spp.) from certain third countries and repealing Decision 2000/462/ - OJ L 328, 17.12.2003, p. 26–31.



Commission Decision 2004/595/EC of 29 July 2004 establishing a model health certificate for the importation into the Community for trade of dogs, cats and ferrets - OJ L 266 13.08.2004, p. 11.

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Animal Health Legislation in Europe



Commission Decision 2004/824/EC of 1 December 2004 establishing a model health certificate for non-commercial movements of dogs, cats and ferrets from third countries into the Community OJ L 358 03.12.2004, p. 12.



Commission Decision 2005/64/EC of 26 January 2005 implementing Council Directive 92/65/EEC as regards import conditions for cats, dogs and ferrets for approved bodies, institutes or centres OJ L 027 29.01.2005, p. 48.

In the case of taxa not regulated by Community legislation, national rules apply. Please note that new legislation is likely to come into force in the near future that will further harmonise rules (certification and animal health requirements) for the importation of other wild animals. Because of these rules, imports of such animals will only be allowed from a small number of third countries authorised for each species. However a draft Decision has been discussed which foresaw a particular regime for importation of live animals originating in any third country but imported after a residency period in St. Pierre and Miquelon (a little island in the Atlantic Ocean close to Canada) where they will spend a period in a quarantine station. During this period, specific testing would be carried out on the animals. For the moment, these special conditions are limited to the import of live Camelidae, but the intention has been expressed to extend this possibility to other species. In the event of a disease appearing in a country from which imports normally are permitted, the Commission may decide on specific protective measures, including a temporary import ban, e.g.: 

Commission Decision 2006/563/EC of 11 August 2006 concerning certain protection measures in relation to highly pathogenic avian influenza of subtype H5N1 in wild birds in the Community and repealing Decision 2006/115/EC - OJ L 222, 15.8.2006, p. 11–19.



Commission Decision 2005/759/EC of 27 October 2005 concerning certain protection measures in relation to highly pathogenic avian influenza in certain third countries and the movement from third countries of birds accompanying their owners - OJ L 285 28.10.2005, p. 52.

C. Biosecurity measures The following legislation contains specific rules for controlling and eradicating certain diseases. In the case of the outbreak of a highly contagious or of an emerging disease, the Commission will take specific Decisions ad hoc, e.g. defining the applicable infection and surveillance zones and the specific trade restrictions to be observed. 

Council Directive 77/391/EEC of 17 May 1977 introducing Community measures for the eradication of brucellosis, tuberculosis and leucosis in cattle - OJ L 145 13.06.1977, p.44.



2004/226/EC: Commission Decision of 4 March 2004 approving tests for the detection of antibodies against bovine brucellosis within the framework of Council Directive 64/432/EEC OJ L 68, 6.3.2004, p. 36–37.



Council Directive 85/511/EEC of 18 November 1985 73/53/EEC: introducing Community measures for the control of foot-and-mouth disease - OJ L 315, 26. 11. 1985, p. 11.



Council Directive 90/423/EEC of 26 June 1990 amending Directive 85/511/EEC introducing Community measures for the control of foot-and-mouth disease, Directive 64/432/EEC on animal health problems affecting intra-Community trade in bovine animals and swine and Directive 72/462/EEC on health and veterinary inspection problems upon importation of bovine, ovine and caprine animals and swine, fresh meat or meat products from third countries - OJ L 224, 18. 8. 1990, p. 13.



Commission Decision 2001/303/EC of 11 April 2001 laying down the conditions for the control and eradication of foot-and-mouth disease in endangered species in application of Article 13 of Directive 85/511/EEC - OJ L 104 , 13/04/2001, p.3.



Council Directive 2000/75/EC of 20 November 2000 laying down specific provisions for the control and eradication of bluetongue – OJ L327 22.12.2000, p.74.

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Animal Health Legislation in Europe



Commission Decision 2004/558/EC of 15 July 2004 implementing Council Directive 64/432/EEC as regards additional guarantees for intra-Community trade in bovine animals relating to infectious bovine rhinotracheitis and the approval of the eradication programmes presented by certain Member States - OJ L 249, 23.7.2004, p. 20–25.



Regulation (EC) No 999/2001 of the European Parliament and of the Council of 22 May 2001 laying down rules for the prevention, control and eradication of certain transmissible spongiform encephalopathies – OJ L147 31.05.2001, p.1. This regulation has been amended very frequently, for the last time by Commission Regulation (EC) No 163/2009 of 26 February 2009.



Commission Decision 2007/453/EC of 29 June 2007 establishing the BSE status of Member States or third countries or regions thereof according to their BSE risk - OJ L 172, 30.6.2007, p. 84–86.



Commission Regulation (EC) No 546/2006 of 31 March 2006 implementing Regulation (EC) No 999/2001 of the European Parliament and of the Council as regards national scrapie control programmes and additional guarantees and derogating from certain requirements of Decision 2003/100/EC and repealing Regulation (EC) No 1874/2003 - OJ L 94, 1.4.2006, p. 28–31.



Council Directive 92/119/EEC of 17 December 1992 introducing general Community measures for the control of certain animal diseases and specific measures relating to swine vesicular disease - OJ L 62, 15. 3. 1993, p. 69.



Council Directive 2001/89/EC of 23 October 2001 on Community measures for the control of classical swine fever – OJ L316 01.12.2001, p.5 (note: this directive repeals 80/217/EEC).



Council Directive 2002/60/EC of 27 June 2002 laying down specific provisions for the control of African swine fever and amending Directive 92/119/EEC as regards Teschen disease and African swine fever - OJ L 192, 20.7.2002, p. 27–46.



Commission Decision 2008/185/EC of 21 February 2008 on additional guarantees in intraCommunity trade of pigs relating to Aujeszky’s disease and criteria to provide information on this disease (notified under document number C(2008) 669) (Codified version) (Text with EEA relevance) - OJ L 59, 4.3.2008, p. 19–30.



Council Directive 92/35/EEC of 29 April 1992 laying down control rules and measures to combat African horse sickness - OJ L 157, 10. 6. 1992, p. 19.



Council Directive 2005/94/EC of 20 December 2005 on Community measures for the control of avian influenza and repealing Directive 92/40/EEC - OJ L 10, 14. 1. 2006, p. 16.



Commission Decision 2005/464/EC of 21 June 2005 on the implementation of survey programmes for avian influenza in poultry and wild birds to be carried out in the Member States OJ L 164 24.06.2005, p. 52.



Commission Decision 2005/731/EC of 17 October 2005 laying down additional requirements for the surveillance of avian influenza in wild birds - OJ L 274 20.10.2005, p. 93. Validity extended until 31 December 2009 by Commission Decision 2009/6/EC of 17 December 2008.



Commission Decision 2005/734/EC of 19 October 2005 laying down biosecurity measures to reduce the risk of transmission of highly pathogenic avian influenza caused by Influenza virus A subtype H5N1 from birds living in the wild to poultry and other captive birds and providing for an early detection system in areas at particular risk - OJ L 274 20.10.2005, p. 105.



2006/415/EC: Commission Decision of 14 June 2006 concerning certain protection measures in relation to highly pathogenic avian influenza of the subtype H5N1 in poultry in the Community and repealing Decision 2006/135/EC - OJ L 164, 16.6.2006, p. 51–60.



2006/563/EC: Commission Decision of 11 August 2006 concerning certain protection measures in relation to highly pathogenic avian influenza of subtype H5N1 in wild birds in the Community and repealing Decision 2006/115/EC - OJ L 222, 15.8.2006, p. 11–19.



2007/598/EC: Commission Decision of 28 August 2007 concerning measures to prevent the spread of highly pathogenic avian influenza to other captive birds kept in zoos and approved bodies, institutes or centres in the Member States - OJ L 230, 1.9.2007, p. 20–26.

V.

Animal Health Legislation in Europe



Council Directive 92/66/EEC of 14 July 1992 introducing Community measures for the control of Newcastle disease - OJ L 260, 5. 9. 1992, p. 1.



Council Directive 2006/88/EC of 24 October 2006 on animal health requirements for aquaculture animals and products thereof, and on the prevention and control of certain diseases in aquatic animals - OJ L 328, 24.11.2006, p. 14–56.

D. Notification of diseases Disease notification is the basis for disease control. Any veterinarian suspecting the presence of a disease which is notifiable under EU or national legislation is under an obligation to immediately contact the competent official veterinarian or the competent veterinary office/service and to communicate their suspicion. 

Council Directive 82/894/EEC of 21 December 1982 on the notification of animal diseases within the Community - OJ L 378, 31. 12. 1982, p. 58.



Commission Decision 2005/176/EC of 1 March 2005 laying down the codified form and the codes for the notification of animal diseases pursuant to Council Directive 82/894/EEC - OJ L 059 05.03.2005, p. 40. Amended by Decisions 924/2006/EC and 2008/755/EC (no consolidated text available).

Zoo veterinarians must be particularly aware of the list of notifiable diseases contained in Annex a of Directive 92/65/EEC (BALAI). E. Mixed texts 

Directive 2003/99/EC of the European Parliament and of the Council of 17 November 2003 on the monitoring of zoonoses and zoonotic agents, amending Council Decision 90/424/EEC and repealing Council Directive 92/117/ - OJ L 325 12.12.2003, p.31.



Council Directive 89/608/EEC of 21 November 1989 on mutual assistance between the administrative authorities of the Member States and cooperation between the latter and the Commission to ensure the correct application of legislation on veterinary and zootechnical matters - OJ L 351, 2. 12. 1989, p. 34.



Regulation (EC) No 1774/2002 of the European Parliament and of the Council of 3 October 2002 laying down health rules concerning animal by-products not intended for human consumption - OJ L 273 10.10.2002, p.1.



Commission Decision 2003/322/EC of 12 May 2003 implementing Regulation (EC) No 1774/2002 of the European Parliament and of the Council as regards the feeding of certain necrophagous birds with certain category 1 materials - OJ L 117, 13.5.2003, p. 32–34.

Regulation 1774/2002 is extremely complex, sometimes contradicting itself, and the public interest in some of the provisions is not evident. Within six years, there have been 49 amendments, 33 derogations, two corrections and nine consolidated versions. In addition, the Regulation was affected by four court cases. From a zoo perspective, in particular the definition of Category 1 material contained in Article 4 (1) a) is not acceptable and cannot be implemented. Dead animals other than farmed animals and wild animals, including in particular pet animals, zoo animals and circus animals, and experimental animals are considered to fall under this category and are assumed to be directly disposed of as waste by incineration in an incineration plant. Therefore, it is theoretically not permitted to feed guinea pigs, laboratory rats or laboratory mice to reptiles, owls, raptors or small carnivores, which may create a conflict with national animal welfare requirements. While the regulation permits a zoo to feed to its carnivores or raptors a sick or wounded animal that perished in the wild, it prevents the feeding of healthy surplus animals shot or euthanised for

V.

Animal Health Legislation in Europe

management reasons at a zoo. If a zoo, however, were to declare itself being a game farm, the same deer could be killed for human consumption. Under Commission Decision 2003/322/EC the feeding of free-living vultures in France, Greece, Italy, Portugal and Spain with certain category 1-materials is allowed. It could be argued that the same should be allowable in the case of vultures kept by zoos. F. Veterinary checks The following Directives and Decisions describe the veterinary checks applicable in intracommunity trade and on importation, and they define the list of approved border checkpoints. The most important recent development in this field is the introduction of the TRACES System on April 1, 2004. This TRAde Control and Expert System combines the functions of the previous ANIMO and SHIFT systems by creating a single central database to track the movement of animals and certain types of products both within the EU and from outside the EU. Consequently the duplication of data is avoided. TRACES is designed to be used directly by economic operators under the control of the competent veterinary authorities, so relevant information can easily be shared with customs authorities. 

Council Directive 89/662/EEC of 11 December 1989 concerning veterinary checks in intraCommunity trade with a view to the completion of the internal market OJ L 395, 30.12.1989, p. 13–22.



Council Directive 90/425/EEC of 26 June 1990 concerning veterinary and zootechnical checks applicable in intra-Community trade in certain live animals and products with a view to the completion of the internal market - OJ L 224, 18. 8. 1990, p. 29.



Commission Decision 94/339/EC of 25 May 1994 laying down detailed rules for the application of Article 9.1 of Council Directive 90/425/EEC concerning veterinary and zootechnical checks applicable in intra-Community trade in certain live animals and products with a view to the completion of the internal market - OJ L 151 17.06.1994, p.38.



Commission Regulation (EC) No 599/2004 of 30 March 2004 concerning the adoption of a harmonised model certificate and inspection report linked to intra-Community trade in animals and products of animal origin - OJ L 094 31.03.2004, p. 44.



Council Directive 91/496/EEC of 15 July 1991 laying down the principles governing the organization of veterinary checks on animals entering the Community from third countries and amending Directives 89/662/EEC, 90/425/EEC and 90/675/EEC - OJ L 268, 24. 9. 1991, p. 56.



Commission Decision 2001/812/EC of 21 November 2001 laying down the requirements for the approval of border inspection posts responsible for veterinary checks on products introduced into the Community from third countries – OJ L306 23.11.2001, p.28.



Commission Decision 2001/881/EC of 7 December 2001 drawing up a list of border inspection posts agreed for veterinary checks on animals and animal products from third countries and updating the detailed rules concerning the checks to be carried out by the experts of the Commission – OJ L326 11.12.2001, p.44. Amended by 2003/506/EC - OJ L 172 10.07.2003, p.16.



Commission Regulation (EC) No 136/2004 of 22 January 2004 laying down procedures for veterinary checks at Community border inspection posts on products imported from third countries - OJ L 021 28.01.2004, p. 11.



2007/275/EC: Commission Decision of 17 April 2007 concerning lists of animals and products to be subject to controls at border inspection posts under Council Directives 91/496/EEC and 97/78/EC - OJ L 116, 4.5.2007, p. 9–33.



Commission Decision 97/794/EC of 12 November 1997 laying down certain detailed rules for the application of Council Directive 91/496/EEC as regards veterinary checks on live animals to be imported from third countries - OJ L 323, 26.11.1997, p. 31–36.

V.

Animal Health Legislation in Europe



Commission Regulation (EC) No 282/2004 of 18 February 2004 introducing a document for the declaration of, and veterinary checks on, animals from third countries entering the Community OJ L 049 19.02.2004, p. 11.



Regulation (EC) No 882/2004 of the European Parliament and of the Council of 29 April 2004 on official controls performed to ensure the verification of compliance with feed and food law, animal health and animal welfare rules - OJ L 165, 30.4.2004, p. 1–141.



Commission Decision 2003/623/EC of 19 August 2003 concerning the development of an integrated computerised veterinary system known as Traces - OJ L 216 28.08.2003, p. 58.



Commission Decision 2004/292/EC of 30 March 2004 on the introduction of the Traces system and amending Decision 92/486/EEC (OJ L 094 31.03.2004, p. 63), amended by Commission Decision 2005/515/EC of 14 July 2005 - OJ L 187 19.07.2005, p. 29.

4. Zoonoses – a special problem Only a small proportion of the more than 200 communicable diseases known to be common to man and animals are contained in the lists of the OIE Terrestrial Animal Health Code. With a focus mainly on food-borne diseases, the European Union obliges Members States to ensure that data on the occurrence of zoonoses and zoonotic agents and antimicrobial resistance related thereto are collected, analysed and published without delay. Annex I of Directive 2003/99/EC contains a list of 8 diseases or agents, which should be included in monitoring, and another 16 diseases which should be monitored according to the epidemiological situation. The zoonoses covered by the second list may not always be regulated by national legislations. The consequence of this situation is that zoonoses often are only detected after an animal has been introduced into the collection and has either fallen sick and died, or other animals or humans have been infected. This situation is largely due to veterinary administrations primarily addressing diseases of agricultural livestock. Apart from a few zoonoses addressed by veterinary services in all countries, like rabies, brucellosis, bovine tuberculosis etc., there is no official network of officially approved diagnostic and reference laboratories for the bulk of zoonotic diseases. When confronted with an import application for zoo animals, import conditions are often established on an ad hoc basis which may not necessarily be scientifically sound. To reduce the risk of introducing zoonoses by international trade and of their spreading in zoo collections and to zoo staff, measures have to be taken at several levels.

Measures by veterinary administrations Import requirements for zoo animals should be defined in compliance with the OIE Code. Where no such standards exist, a sound risk assessment has to be made, or quarantine procedures of national zoo organisations may be followed if these are available. Certification requirements should not be overemphasised, but proper quarantine should be ensured at the importing zoos. Veterinary supervision of zoos should be mandatory, and this could be best achieved by subjecting the operation of a zoo to licensing and to approval under the BALAI Directive (92/65/EEC). Measures by the zoological gardens Zoos should keep high hygienic standards for animals, keepers and food, implement veterinary controlled quarantine for all incoming animals (if there are no other requirements usually 30 days, unless the judgement of the veterinarian allows for shortening this period of time), avoid contact to neighbouring farms, implement a control programme for rats and mice, and attempt to exclude other local free-ranging wild mammals from the zoo, as these

V.

Animal Health Legislation in Europe

may be potential carriers of zoonoses. There should be close clinical, parasitological and post-mortem surveillance of the collections and of free roaming wild animals, treatment or vaccination of susceptible animals for relevant zoonoses, collection of blood samples for direct diagnosis and for establishing serum banks. Exposed staff should be included in the surveillance and prophylactic measures. In Children's Zoos, there should be educational signage about how to behave for minimising the risk of disease transmission, hand-washing stations should be available and eating and drinking should not be allowed in the contact area. Measures by the zoo and wildlife veterinarians’ organisations Zoonoses should be a prominent feature in scientific venues. Regional organisations should co-operate with the OIE Working Group on Wildlife Diseases, and should follow the example of the European Wildlife Disease Association (EWDA) in producing regional reports. As with the AAZV (American Association of Zoo Veterinarians) in North America, they should co-operate with the zoo organisations of their region in establishing procedures for minimising the zoonosis risk. EAZWV should continue to co-operate with OIE in improving international standards, and should involve other groups from within the WAWV (World Association of Wildlife Veterinarians) family.

5. Certification procedures International movements of many animal species are only possible if the animal is accompanied by a veterinary certificate. Unless otherwise defined, such certificates have to be issued by an official veterinarian who, however, would often have to base his statement on the findings of another veterinarian. Hence it follows that zoo or institute veterinarians may be required to certify certain facts to the official veterinarian, who in turn will use the information received for issuing an official certificate. Under the revised BALAI Directive, the veterinarian of an approved zoo or institute is authorised to issue certificates for certain species when moved between EU Members States or between the EU and a Third Country under a bilateral agreement. In order to maintain confidence in the certification process, it is necessary that certification is based on the highest possible ethical standards, the most important of which is that the professional integrity of the certifying veterinarian must be respected and safeguarded. It is essential not to include in the requirements additional specific matters which cannot be accurately and honestly signed by a veterinarian. For example these requirements should not include certification of an area as being free from non-notifiable diseases, the occurrence of which the signing veterinarian is not necessarily informed about. Equally, to ask certification for events which will take place after the document is signed is unacceptable when these events are not under the direct control and supervision of the signing veterinarian. Guidelines for certifying veterinarians have been drawn up by professional organisations, the certification process is described in the OIE Terrestrial Animal Health Code, and the EU also has defined requirements. All these texts are very similar to each other. The following is the text of the articles 5.2.2 and 5.2.3 of the OIE Terrestrial Animal Health Code. Preparation of international veterinary certificates Certificates should be drawn up in accordance with the following principles:

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Animal Health Legislation in Europe

1.

Certificates should be designed so as to minimize the potential for fraud including use of a unique identification number, or other appropriate means to ensure security. Paper certificates should bear the official identifier of the issuing Veterinary Authority. Each page of a multiple page certificate should bear the unique certificate number and a number indicating the number of the page out of the total number of pages. Electronic certification procedures should include equivalent safeguards.

2.

They should be written in terms that are as simple, unambiguous and easy to understand as possible, without losing their legal meaning.

3.

If so required, they should be written in the language of the importing country. In such circumstances, they should also be written in a language understood by the certifying veterinarian.

4.

They should require appropriate identification of animals and animal products except where this is impractical (e.g. day-old birds).

5.

They should not require a veterinarian to certify matters that are outside his/her knowledge or which he/she cannot ascertain and verify.

6.

Where appropriate, they should be accompanied, when presented to the certifying veterinarian, by notes of guidance indicating the extent of enquiries, tests or examinations expected to be carried out before the certificate is signed.

7.

Their text should not be amended except by deletions which must be signed and stamped by the certifying veterinarian. The signature and stamp must be in a colour different to that of the printing of the certificate.

8.

Replacement certificates may be issued by a Veterinary Authority to replace certificates that have been, for example, lost, damaged, contain errors, or where the original information is no longer correct. These must be clearly marked to indicate that they are replacing the original certificate. A replacement certificate should reference the number and the issue date of the certificate that it supersedes. The superseded certificate should be cancelled and where possible, returned to the issuing authority.

9.

Only original certificates are acceptable.

Certifying veterinarians Certifying veterinarians should: 1.

be authorised by the Veterinary Administration of the exporting country to sign international veterinary certificates;

2.

only certify matters that are within their own knowledge at the time of signing the certificate, or that have been separately attested by another competent party;

3.

sign only at the appropriate time certificates that have been completed fully and correctly; where a certificate is signed on the basis of supporting documentation, the certifying veterinarian should be in possession of that documentation before signing;

4.

have no conflict of interest in the commercial aspects of the animals or animal products being certified and be independent from the commercial parties.

V.

Annex:

Animal Health Legislation in Europe

Diseases listed in the OIE Terrestrial Animal Health Code and applicable EU legislation

Disease

OIE List

Notifiable under 82/894/EEC (yes/no) - Zoonosis under 2003/99/EC (A/B)* - other relevant EU law**

Anthrax

Multiple species

no - 64/432/EEC (E-I) - 91/68/EEC (BI) - 92/65/EEC (A)

Aujeszky's disease

Multiple species

no - 64/432/EEC (E-II)- 2008/185/EC

Bluetongue

Multiple species

yes - 92/65/EEC (A) - 2000/75/EC 2004/68/EC

(Bovine) brucellosis (Brucella abortus)

Multiple species

no – A - 64/432/EEC (E-I) 77/391/EEC - 92/65/EEC (A) 2004/226/EC

(Caprine/ovine) brucellosis (Brucella melitensis)

Multiple species

no - A -91/68/EEC (B-I) - 92/65/EEC (A)

(Porcine) brucellosis (Brucella suis)

Multiple species

no - A - 64/432/EEC (E-I) - 92/65/EEC (A)

Crimean Congo haemorrhagic fever

Multiple species

no - 2006/696/EC -

Echinococcosis/hydatidosis

Multiple species

no - A

Epizootic haemorrhagic disease

Multiple species

no - 92/119/EEC

Equine encephalomyelitis (Eastern)

Multiple species

yes - 90/426/EEC

Foot and mouth disease

Multiple species

yes - 64/432/EEC (E-I) - 2003/85/EC (repealing 85/511/EEC) - 90/423/EEC 91/68/EEC (B-I) - 92/65/EEC (A) 2001/303/EC - 2004/68/EC

Heartwater

Multiple species

no

Japanese encephalitis

Multiple species

yes - 90/426/EEC

Leptospirosis

Multiple species

no

New world screwworm (Cochliomyia hominivorax)

Multiple species

no

Old world screwworm (Chrysomya bezziana)

Multiple species

no

Paratuberculosis

Multiple species

no - 91/68/EEC (B-III)

Q fever

Multiple species

No

Rabies

Multiple species

no - B - 64/432/EEC (E-I) - 91/68/EEC (B-I) - 92/65/EC

Rift Valley fever

Multiple species

yes - 92/65/EEC (A) - 2004/68/EC

Rinderpest (cattle plague)

Multiple species

yes - 92/65/EEC (A) - 2004/68/EC

Surra (Trypanosoma evansi)

Multiple species

no -

Trichinellosis

Multiple species

no - A - 377/96/EEC

Tularemia

Multiple species

no - 92/65/EEC

Vesicular stomatitis

Multiple species

yes - 90/426/EEC - 92/65/EEC (A) 2004/68/EC

West Nile fever

Multiple species

no

Bovine anaplasmosis

Cattle

no

Bovine babesiosis

Cattle

no

Bovine genital campylobacteriosis

Cattle

no - A

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Animal Health Legislation in Europe

Bovine spongiform encephalopathy

Cattle

yes -92/65/EEC (A) (all TSE) 2001/9/EC - R(EC)999/2001

Bovine tuberculosis (Mycobacterium bovis)

Cattle

no - A - 64/432/EEC (E-I) - 77/391/EEC - 92/65/EEC (A) - 2000/504/EC

Bovine viral diarrhea

Cattle

no

Contagious bovine pleuropneumonia

Cattle

yes - 64/432/EEC (E-I) - 92/65/EEC (A) - 2004/68/EC

Enzootic bovine leukosis

Cattle

no - 64/432/EEC (E-I) - 77/391/EEC

Haemorrhagic septicaemia

Cattle

no

Infectious bovine rhinotracheitis/infectious pustular Cattle vulvovaginitis

no - 64/432/EEC (E-II) - 2000/502/EC

Lumpy skin disease

Cattle

yes 64/432/EEC - 92/65/EEC (A) 2004/68/EC

Theileriosis

Cattle

no

Trichomonosis

Cattle

no

Trypanosomosis (tsetse-transmitted)

Cattle

no

Caprine viral arthritis/encephalitis

Sheep and goat

no - 91/68/EEC (B-III)

Contagious agalactia

Sheep and goat

no - 91/68/EEC (B-III)

Contagious caprine pleuropneumonia

Sheep and goat

no

Enzootic abortion of ewes (ovine chlamydiosis)

Sheep and goat

no

Maedi-visna

Sheep and goat

no - 91/68/EEC (B-III)

Nairobi sheep disease

Sheep and goat

no

Ovine epididymitis (Brucella ovis)

Sheep and goat

no - A - 91/68/EEC (B-I) - 92/65/EEC (A)

Peste des petits ruminants

Sheep and goat

yes - 92/65/EEC (A) (concerns also Suidae) - 2004/68/EC

Salmonellosis (S. abortusovis)

Sheep and goat

no - A

Scrapie

Sheep and goat

no - 91/68/EEC - 92/65/EEC (A) 2001/9/EC - R(EC)999/2001

Sheep pox and goat pox

Sheep and goat

yes - 92/65/EEC (A) - 2004/68/EC

African horse sickness

Equine

yes - 92/35/EEC, 92/36/EEC, 92/65/EEC (A)

Contagious equine metritis

Equine

no -

Dourine

Equine

yes - 90/426/EEC

Equine encephalomyelitis (Western)

Equine

yes - 90/426/EEC

Equine infectious anaemia

Equine

yes - 90/426/EEC

Equine influenza

Equine

no -

Equine piroplasmosis

Equine

no -

Equine rhinopneumonitis

Equine

no -

Equine viral arteritis

Equine

no -

Glanders

Equine

yes - 90/426/EEC

Venezuelan equine encephalomyelitis

Equine

yes - 90/426/EEC

African swine fever

Swine

yes - 64/432/EEC (E-I) - 92/65/EEC (A) - 2002/60/EC - 2004/68/EC

Classical swine fever

Swine

yes - 64/432/EEC (E-I) - 92/65/EEC (A) - 2001/89/EC - 2004/68/EC

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Animal Health Legislation in Europe

Nipah virus encephalitis

Swine

no

Porcine cysticercosis

Swine

no

Porcine reproductive and respiratory syndrome

Swine

no

Swine vesicular disease

Swine

yes - 64/432/EEC (E-I) - 92/65/EEC (A) - 92/119/EEC - 2004/68/EC

Transmissible gastroenteritis

Swine

no - 64/432/EEC (E-II)

Avian chlamydiosis

Avian

no - B - 92/65/EEC (A) (in Psittaciformes)

Avian infectious bronchitis

Avian

no -

Avian infectious laryngotracheitis

Avian

no -

Avian mycoplasmosis (M. gallisepticum)

Avian

no - 90/539/EEC

Avian mycoplasmosis (M. synoviae)

Avian

no

Duck virus hepatitis

Avian

no

Fowl cholera (Avian pasteurellosis)

Avian

no

Fowl typhoid (Salmonella gallinarum)

Avian

no - A - 90/539/EEC

Highly pathogenic avian influenza in birds and low Avian pathogenic notifiable influenza in poultry as defined in Chapter 10.4

yes - 92/65/EEC (A) - 2005/94/EC 2005/464/EC - 2005/731/EC 2005/734/EC - 2005/759/EC 2006/415/EC - 2006/563/EC 2006/598/EC

Infectious bursal disease (Gumboro disease)

Avian

no

Marek's disease

Avian

no

Newcastle disease

Avian

yes - 92/65/EEC (A) - 92/66/EEC

Pullorum disease

Avian

no - A - 90/539/EEC

Turkey rhinotracheitis

Avian

no

Myxomatosis

Lagomorph

no - 92/65/EEC

Rabbit haemorrhagic disease

Lagomorph

no - 92/65/EEC

Acarapisosis of bees

Bee

no - 92/65/EEC

American foulbrood of honey bees

Bee

no - 92/65/EEC (A)

European foulbrood of honey bees

Bee

no

Small hive beetle infestation (Aethina tumida)

Bee

yes - 92/65/EEC (A)

Tropilaelaps infestation of honey bees

Bee

yes - 92/65/EEC (A)

Varroosis of honey bees

Bee

no - 92/65/EEC

Camel pox

Other

no

Leishmaniosis

Other

no

Ovine pulmonary adenomatosis

--

no - 91/68/EEC (B-III)

Caseous lymphadenitis

--

no - 91/68/EEC (B-III)

Ebola in non-human primates

--

92/65/EEC (A)

Monkey pox in rodents and non-human primates

--

92/65/EEC (A) - 2003/459/EC

Porcine enterovirus encephalomyelitis (Teschen)

--

92/65/EEC (A)

Avian tuberculosis

--

no - A

Listeriosis

--

no - A

Verotoxigenic Escherichia coli

--

no – A

Calicivirus

--

no – B

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Animal Health Legislation in Europe

Hepatitis A virus

--

no – B

Influenza virus

--

no – B

Viruses transmitted by arthropodes

--

no – B

Borreliosis and agents thereof

--

no – B

Botulism and agents thereof

--

no – B

Leptospirosis and agents thereof

--

no – B

Tuberculosis other than mentioned above

--

no – B

Vibriosis and agents thereof

--

no – B

Yersiniosis

--

no – B

Anisakiasis and agents thereof

--

no – B

Cryptosporidiosis and agents thereof

--

no – B

Cysticercosis and agents thereof

--

no – B

Toxoplasmosis and agents thereof

--

no – B

Other zoonoses an zoonotic agents

--

no – B

* A = Monitoring is mandatory, B = To be monitored according to the epidemiological situation: ** 64/432/EEC: (E-I) = Compulsorily notifiable, (E-II) = Member States may have national control or monitoring programmes 91/68/EEC: (B-I) = Compulsorily notifiable, (B-III) = Member States may have national control or monitoring programmes 92/65/EEC (A) = Notifiable in the context of the BALAI Directive

Transmissible Diseases Handbook

VII. POST-MORTEM PROCEDURES Gerry M. Dorrestein and Marein M. van der Hage Diagnostic Pathology Laboratory Dutch Research Institute for Avian and Exotic Animals (NOIVBD) Veldhoven,The Netherlands

1. Introduction Pathology is not a science for pathologists alone. The findings of the pathologist should be one of the bases of the clinician’s understanding of disease and the pathophysiology of disease. Changes in anatomic pathology are the foundation of disease, and understanding these changes will give the clinician an advantage in selecting the right diagnostic tools and therapeutic approach. Determination of the cause of death in zoo animals is often difficult and may require the close co-operation of a number of disciplines. Some zoo veterinarians perform a post-mortem examination in the zoo. But in doing the post-mortem, careless observation and sampling can result in useless, and sometimes even harmful, information. Those who perform postmortem examinations in this manner are running a risk, as overlooking fundamental changes in tissue can be costly to both pocketbook and intellectual development (Cheville, N.F., 1999, In: Introduction to Veterinary Pathology, Iowa State University Press/Ames). When a post-mortem is performed in the zoo itself, or samples are collected for diagnostic purposes, the results and information deriving from this activity are highly protocol sensitive. For a diagnostic laboratory to contribute fully to the final diagnosis, the specimen(s) collected must be selected carefully and preserved in suitable conditions. A thorough post-mortem examination of animals that die or are euthanised is a necessary adjunct to any good clinical practice. The purpose of this chapter is to assist the zoo veterinarian in the performance of a thorough post-mortem examination and in the correct selection, preservation and transportation of pathological and biological specimens. This chapter will not be a complete protocol, but more of a guide. As with any activity, it is essential to prepare yourself before you begin. This includes reading and preparing your protocols and setting up an area or room, with the proper equipment, where you can perform the post-mortem. Keep in mind that the facilities need to be “infectious disease proof”; every post-mortem should be treated as an infectious problem until proven otherwise. As the basis for this article we have used the excellent booklet, “Post-mortem procedures for wildlife veterinarians and field biologists” written by M.H. Woodford, D.F. Keet and R.G. Bengis (2000) and published jointly by the Office International des Epizooties (O.I.E.), Care for the Wild International and the Veterinary Specialist Group/Species Survival Commission of the World Conservation Union (IUCN). This little 55 page booklet is very comprehensive and should be present in every zoo where post-mortems are performed. It is available through the OIE, 12 Rue de Prony, 75017, Paris France (ISBN 92-9044-419-6).

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We will not try to rewrite this booklet here, but several parts will be completely reproduced in this chapter, as is allowed by the OIE. There are several reasons for performing a post-mortem or having one done. These can include: finding the cause of death, confirming a diagnosis, investigating unsuccessful therapy, increasing knowledge, or simply satisfying curiosity. In the zoo. Dead animals can also function as sentinels, indicating the presence of subclinical infectious diseases which may not be immediately related to the cause of death. Diagnostic pathology is not limited to a post-mortem. The pathologist uses the clinical history (including haematology, blood chemistry and therapeutic measurements), the gross description, culture results and other data, as well as the cytological and histological appearance of the lesions, to make a diagnosis. Absence of any of these, or incorrect submission of tissues, will hamper this process. And remember: it is better to store and preserve too much material than to realise after some time that the proper material required for a diagnosis has already been discarded. We also consider it one of the functions of a “scientifically-minded” zoo to make optimal use of their animals and associated data. This includes gathering scientific information about the live animals regarding housing, feeding, behaviour, breeding, etc. Ideally, this would include maintaining a data bank of sera and organs or tissues, for future retrospective studies of diseases and their agents.

2. Submission of carcasses or specimens When a zoo veterinarian has access to the services of a pathological institute that will perform the post-mortem, procedures should be established for the submission of the intact carcass. These specialists have the necessary experience and training, and their work will yield the best results. In some cases the zoo veterinarian or technician will perform the postmortem examination and submit the appropriate tissue to a diagnostic laboratory. Based on the gross post-mortem findings, material will be collected and sampled for follow-up investigation. The quality of information received from such an examination is directly proportionate to the quality and choice of the specimens submitted and the information that accompanies them. When in doubt about “what” and “how” samples should be collected and packed, the laboratory should always be contacted before sending in any materials. In many situations it is not possible to start a post-mortem immediately after death. To promote the rapid cooling of a small carcass, the fur or plumage should be thoroughly soaked with cold water to which a small amount of soap or detergent has been added to aid complete wetting of the coat or plumage and skin. The carcass should be placed in a plastic bag, all excess air squeezed out, the bag sealed or tied, and then refrigerated. Larger animals should be stored in a cool environment as soon as possible. Big animals will not cool down quickly enough to prevent extensive autolytic post-mortem changes and should be necropsied as soon as possible. When this is not possible, then opening the abdomen can be helpful in lowering the core body temperature. The animal should be kept refrigerated until the post-mortem is performed or the carcass has been shipped to the laboratory. In general, providing the carcass has been cooled immediately upon death and can be delivered to the laboratory within 72 to 96 hours of the time of death, it should be refrigerated (not frozen). Small animals can be packed with sufficient ice or cool packs to keep the carcass cold until arrival at the laboratory. If delivery to a laboratory is expected to be delayed beyond 96 hours post-mortem the carcass should be frozen immediately rather than simply refrigerated. Frozen tissue specimens or carcasses must be packed with sufficient ice to keep them frozen until arrival at the laboratory.

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Refrigerated or frozen specimens should be packed in a sturdy, insulated (Styrofoam®) box, preferably in a leak-proof sealbag, and shipped to the laboratory by a private courier service, which guarantees same- or next-day delivery to the laboratory. Most laboratories cannot receive specimens over the weekend; it is thus advisable not to ship refrigerated or frozen specimens on Fridays or weekends. Remember, it is crucial that sufficient refrigerant be packed with the specimen and that it be adequately insulated to insure that it will remain cold (or frozen) until it is received by the laboratory personnel. In instances where the carcass is extremely small, such as embryos, nestlings or very small adult animals, the entire carcass may be submitted for histological examination. This is best accomplished by opening the body cavity, gently separating the viscera and fixing the entire carcass in formalin solution. When you perform a post-mortem yourself or collect diagnostic material it must be done systematically. The correct selection of material for further examination, and the correct sampling, storage and shipping of material, will increase the quality of results tremendously. A written report of the post-mortem findings will help the zoo veterinarian to keep track of the disease status of the zoo collection. EVEN A NEGATIVE FINDING IS A FINDING, SINCE IT MEANS THAT THE LESIONS/CHANGES YOU WERE LOOKING FOR ARE NOT PRESENT.

3. Post-mortem site, protective clothing and equipment For details see Woodford et al. (2000), Section I: Preparing for a post-mortem examination. It is helpful to have a set of instruments designated for post-mortem examinations. These should be thoroughly cleaned and sterilised after use. A separate room to perform the postmortem is also advisable. Instruments that are used for post mortem examinations should not be used for living animals. It is important to wear adequate protective clothing. The instrument pack should include post-mortem knives, forceps, two scalpel handles (one for cutting, one for burning organ surfaces before taking a microbiology sample), stout scissors and/or poultry shears (for cutting bones), and fine scissors for dissection. Tiny animals such as finches, lizards or rodents require fine instruments such as iris scissors. For large animal post-mortem examinations special instruments such as a vibrating (cast-cutting) saw may be used. Other useful equipment includes a (gram) scale, a hand-lens or dissecting microscope, and paper tissues. In addition to instruments, one should have at hand: - 10% neutral buffered formalin (= 4% formaldehyde), - 70% alcohol for wetting and disinfecting the skin, - 96-100% ethyl alcohol (for fixing specimens suspected of having gout, and 100mg/1ml for PCR testing), - a bottle with normal saline (0.9% NaCl) with a pipette (for parasitological examination), and - appropriate containers. Other material for ancillary diagnostic procedures include: - syringes and needles to obtain samples for serology, haematology, or cytology, - clean glass slides for impression smears,

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a stain set for cytology (e.g. "Diff-QuickR, Hemacolor, Stamp or Machiavelli), clean glass slides and coverslips for wet mounts (parasitology), burner for heating and sterilising a scalpel blade before taking a sample for microbiology, sterile swabs or culture tubes with appropriate transport media for bacterial, or fungal culture, - transport media (96% ethanol or buffered 4 M guanidine isothiocyanate) for PCR testing (viruses, mycobacteria and chlamydia) - petri dishes or freezer-proof tubes for submission of tissues for viral isolation.

-

It may also be helpful to have a camera available for documentation of gross lesions. A standard checklist and post-mortem report form will assist in recording observations.

4. Euthanasia The method of euthanasia may affect specimens submitted to the pathologist. High doses of barbiturates are caustic to tissues and cause crystallisation in and on organs. Such changes may be mistaken for early gout, but will also change and mask macroscopic and microscopic lesions. When euthanasia solutions are used at an appropriate dosage, e.g. pentobarbital 200 mg/kg bw intraperitoneally or T61 0.5 ml/kg intramuscular, few alterations are seen The euthanasia agent can also be administered intravenously, or into the spinal cord at the base of the skull with the head flexed (especially in larger birds). Administering such agents slowly to effect is helpful to prevent undesired artificial changes. Serum or heparinised haematological samples should be collected prior to euthanasia. The blood may be centrifuged and serum or plasma submitted or saved and frozen pending postmortem results. This may be helpful in diagnosis of endocrine disorders or viral infections. Routine haematological tests may also be performed on these samples.

5. Impression Smears Impression smears of fresh cut organs or altered surfaces are not common practice at postmortems. They are a useful and often underestimated adjunct to a complete post-mortem examination. In our protocol, two sets of impression smears are made at every post-mortem from liver, spleen, lung and rectum. Organs with pathological changes are automatically added to this list. For a first impression of the presence of bacteria, yeasts or protozoa, this technique is very useful. Tissue phases of parasites such as Atoxoplasma spp, Toxoplasma spp., Plasmodium spp, Hemoproteus spp., Leucocytozoon spp, and Trypanosoma spp. are mostly readily identified in impression smears of liver, spleen and lungs. Immunofluorescent staining for Chlamydia spp. can be carried out in specific laboratories on the impression smears of these organs in all post mortem examinations of suspected cases in reptiles and birds especially within the families Columbiformes and Psittaciformes. Immunohistochemistry staining on fixed paraffin-embedded histological sections is a good alternative when available. The now-days confirmation of Chlamydiosis is by PCR. Also, the cell-type of lymphoreticular and haematopoietic neoplasms is easier to diagnose from impressions of liver, spleen and bone marrow than from histology. To make a good impression smear (actually a touch preparation), it may be easier to hold the slide when touching with the tissue. Grasp a small piece of the tissue with forceps so that a fresh cut, well-blotted surface faces downward. Lower the tissue to the clean slide touching it lightly. Retract quickly without dragging the tissue across the slide. Make several "touch preps" on each slide. Impressions are generally more useful when air-dried. If other fixation is necessary (e.g. heat fixation for acid-fast stains), it can be done after air-drying . Exudate or any other fluids may be prepared for cytological evaluation by having a thick drop air-dried.

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6. Fixation for Histopathology Several philosophies may determine the choice of tissues for histopathological examination: 1. Economic reasons; these are poor grounds for decision-making, but in this case it is better to collect the tissues and, after consulting the pathologist, the selected tissues should be sent in but additional tissue samples should be retained "just in case." 2. Completeness; this is especially valid for a scientific, research situation. Collect all tissues listed in table 1. 3. A standard selection completed with a choice based on the post-mortem findings. This list is practical and will in most cases lead to sufficient diagnostic support. In table 1 the standard selection is marked with an asterisk (*). Normally, selected tissues are fixed in neutral-buffered formalin for histopathological examination. If the formalin solution is not freshly prepared on a frequent basis, formic acid will be formed. A layer of pieces of marble at the bottom of the container or bottle will bind the formic acid to a precipitate, keeping the formalin neutral. This prevents the formation of “formalin pigment” in histological specimens; a confusing, annoying and unnecessary artefact that can be present in improperly fixed tissue samples. Ten percent buffered formalin penetrates only about 2-5 mm in 24 hours, so specimens must be less than 10 mm thick. Penetration is slower in very bloody, dense tissues (e.g. congested spleen or liver) and more rapid in relatively porous tissue (e.g. lung). Formalin will not penetrate well into the brain through the unopened calvarium or into bone marrow unless the bone has been cracked. The biggest problem seen with submission of fixed tissues is inadequate fixation due to prior severe autolysis or an inadequate volume of fixative allowing continuing decomposition. Proper initial fixation is achieved if at least ten times the volume of formalin as volume of tissue is used. When preparing specimens for mailing the amount of formalin may be reduced after tissues have been fixed for 12-24 hours. Wet formalin-fixed tissue may be conveniently stored and shipped in heat-sealed plastic bags. Other fixatives, such as those required for electron microscopy (EM), are not usually necessary, since formalin fixed tissue is easily refixed with glutaraldehyde and the main structures (including viruses) are preserved. For EM fixation very fresh tissue in tiny parts (12 mm3) is essential. The number of tissue specimens submitted to the histopathology laboratory may depend on the cost per sample. If you do not send the complete set of specimens, it is prudent to save the rest in formalin while awaiting a diagnosis. If only grossly visible lesions or limited tissue specimens are submitted, a diagnosis may not be possible. When specific lesions are observed at post-mortem, the tissue specimens collected should include a small margin of normal tissue adjacent to the lesion. Too often, the limited tissue specimens submitted suggest a diagnosis, which cannot be confirmed because other tissues have already been discarded. Tissue specimens for histopathology should not be frozen. Freezing creates crystals and ruptures cells, making histopathology virtually useless. Tissues for toxicological analysis should be frozen. They may be frozen at -20oC after being wrapped in aluminium foil. The optimum temperature for freezing tissues for virus isolation is -70oC. If this cannot be accomplished, the tissues for viral isolation should be sent (by rapid mail) in sterile containers on wet ice to the laboratory. For detailed information on sampling etc. see Woodford et al. (2000), Section III: “The collection and field preservation of biological and pathological specimens” and the Appendices.

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Table 1: Tissues routinely collected for histopathology. skin

thyroid glands

rectum

ovary & oviduct

(feather follicles)

parathyroid glands

caeca

testes (male)

trachea

oesophagus

(cloaca)

pectoral muscle

lung*

(crop)

spleen*

bone marrow

(air sac) heart

*

kidneys* adrenal glands

*

(proventriculus )

liver *

*

stomach/ventriculus

gall bladder

duodenum*

pancreas*

small intestine

thymus brain spinal cord *

(cloacal bursa )

(ischiatic nerve)

*

Standard selection of tissues for routine histopathological examination. (..) Tissue specimens from birds. Selection of additional tissue specimens will depend upon gross lesions observed at post-mortem.

7. Autopsy protocol There are probably as many ways to dissect an animal as there are pathologists. One should choose a procedure with which one is familiar and feels comfortable, and then use it consistently. No matter what procedure is used, each post-mortem should be performed in as regular and thorough a manner as can be accomplished by the prosector and a "complete" set of tissues and specimens be collected for subsequent histopathological, parasitological, toxicological, serological, and biochemical examination. The veterinarian should review the appended detailed checklist of organs to be examined, observations to be made, ancillary tests to be performed and specimens to be collected, prior to disposal of the remains. The following procedure and checklist is used for avian species at the Diagnostic Pathology Laboratory of the Dutch Research Institute of Avian and Exotic Animals (NOIVBD) in Veldhoven, The Netherlands (www.noivbd.nl). For a protocol for mammals see Woodford et al. (2000), Section II: “Post-mortem procedures”.

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Avian Necropsy Protocol Gerry M. Dorrestein1, Marein M. van der Hage, and Marja J. L. Kik2 1

Diagnostic Laboratory of the NOIVBD, Veldhoven, The Netherlands and 2

Department of Veterinary Pathobiology, Utrecht University, The Netherlands

Before starting the necropsy procedure (see also reference list), the packing material is to be inspected for the presence of mites or lice!

I. History Read the history - including identification, physical findings, medical history and pertinent laboratory data - and summarise the most relevant data on your work sheet. Make a note of leg band numbers, transponders or other identifying marks.

II. External examination of the carcass First make a carcass identification based upon identification: species, age, and colour pattern as well as leg band, tattoo or microchip implant data. Record information about general bodily condition, weight, muscle mass, joints, integument (incl. beak and nails), plumage, (for defects, ectoparasites, faeces), body orifices (eyes, ears, nostrils and vent), uropygial gland, traumata, and abnormalities. Palpate the skeleton. The feeding status can be judged based upon the muscles on the keel and the filling of the crop and intestines. When heavy metals are suspected (e.g. rifle bullets or ingested lead) survey radiographs may be taken. Examples (of alterations found at external examination): Broken feathers due to feather picking; diagnosis: normal feathers on the head. Altered feathers with constrictions at the base caused by PBFD; diagnosis: histology of skin with feather-follicles; PCR test. thickened dry skin caused by Malassezia sp; diagnosis: cytology skin scraping. Look for feather and skin parasites. Swelling above the eye or dilated nostrils with a plug in parrots due to vitamin A deficiency; diagnose: histological examination with metaplastic changes in salivary glands. Conjunctivitis and sinusitis related to ornithosis, chlamydiosis or psittacosis Conjunctivitis with pox-lesions; diagnosis: cytology, histology and culture. Abdominal or other swellings, tumours, egg-related peritonitis: diagnosis: histology Cloacal mucosal prolapse, papilloma; diagnosis: histology.

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III. Preparation of the bird Small birds are wetted and plucked, all other birds should be wetted with alcohol 70% before the necropsy. This is done to allow better visualisation of the skin, to part the feathers to permit incision of the skin and to prevent loose feathers from irritating or harming the prosector (zoonosis) or contaminating the viscera. The bird is positioned on its back, in small birds the wings and legs are pinned to a dissecting board with nails or needles, large birds are fixed on a metal tray with pieces of rope.

IV. Post mortem examination General remarks: -

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1.

use a gram-scale for body weight and measuring the size of organs, open all tube-like structures, cut all parenchymatous organs in slices to find small focal lesions, tissue for optimal formalin fixation should preferably not exceed 3 to 4 mm in thickness (5 mm maximum), ratio of tissue to formalin required for adequate fixation is 1:10, collect tissue samples during the necropsy to prevent desiccation. Do not wait till the gross examination is finished, remember to collect and submit specimens from a broad spectrum of organs and systems, collect at least heart, lung, liver, spleen, kidney, gonad, and adrenal, and a piece of intestine (duodenum and pancreas) for histopathology, when suspecting a viral problem, collect 100mg tissue in 1ml ethanol 96% for a PCR, freeze tissue as soon as possible at -20oC, or collect tissue on wet ice till shipment. when you suspect a bacteriological problem, make a impression smear and look before you select you culture media or send an organ or swab to a laboratory for culturing.

An incision is made in the skin along the ventral midline from the mandible over the sternum to the cloaca. The skin is reflected to expose the subcutis, crop, pectoral muscles, keel, abdominal wall, leg muscles and fat. Watch for colour of the muscles, parasites, haemorrhages, and oedema. Judge the amount of food in the crop. In pigeons a vascular plexus in the deep layers of the cutis of the cervical region can be seen, the plexus venosus intracutaneous collaris. This plexus can be mistaken for an extensive haemorrhage. Examples Stripes in leg- or breast-muscle; sarcosporidiosis; diagnosis: cytology of such a stripe reveals the bradyzoites. A large dark spot distal to the keel; swollen liver: diagnosis: see under 3. Changes of the skin; cnemidocoptes, yeast-infection; diagnosis: wet mount and cytology smear.

2.

Make an incision through the pectoral muscle along the sides and around the posterior border of the sternum through the abdominal muscles; cut with heavy rongeurs, scissors or poultry shears through the ribs, coracoid bones, and clavicle to remove the sternum.

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Examine the inside of the sernum, the air sacs and pericardial sac, and make impression smears (when abnormalities or inflammations are seen). During dissection of the keel the air sacs are easily seen. Normal air sacs appear as glistening transparent membranes. Examples Opaque air sacs or (fibrinous) inflammation: chlamydiosis; diagnosis: cytology with special staining, PCR Opaque air sacs or obvious inflammation: bacterial infection; diagnosis: rods or cocci in cytology smear; culture and sensitivity test. Air sacs covered with white/yellow plaques: fungal infection; diagnosis: wet mount (heated with chlorallactophenol), showing hyphae, culture. Air sacs solid with white/yellow material; chronic fungal infection, mostly aspergillosis; diagnosis: wet mount showing hypha, culture. Air sacs, esp. cervical and prescapular, with small black dots in passerines and small psittacines; Sternostoma tracheocolum infestation; diagnosis: magnifying-glass and wet mount. Air sacs filled with food: forced feeding; diagnosis: wet mount and histology. Pericardial sac filled with fluid: inanition, cachexia; diagnosis: muscle wasting, oedema and gelatinous fat-tissue. Pericardium covered with white chalky deposits: visceral gout; diagnosis: wet mount with crystals; often in combination with nephritis.

3.

Identify the (para)thyroids cranial and lateral to the syrinx along the carotid arteries. Remove the thyroids when required. Look for the thymus along the neck in juvenile birds. The liver is examined in situ (examples see 5). Examples In budgerigars enlarged thyroid glands; diagnosis: histology. Parrots (especially African greys) hyperparathyroidism; diagnosis: histology. "Abscesses"; Salmonella or E.coli infections; diagnosis: rod shaped bacteria in cytology, culture.

4.

Remove the heart with the carotids and thyroids attached and cut across the apex to check for an "open" lumen and to assess the thickness of the ventricle walls. Open the heart and large vessels and examine the valves and endocardial surface. Keep in mind that the right atrioventricular valve in birds is a muscular structure. Examples Yellow plaques on the wall inside the large vessels; the vessels are stiff: atherosclerosis: diagnosis: macroscopic (gross) examination, histology. Epi- or endocardial haemorrhages: septicaemia or agonal event; diagnosis: continue post mortem. Gelatinous, serous pericardial fat; starvation, chronical illness: diagnosis; continue post mortem. Changes (inflammation, necrosis) in the myocardium: myocarditis; diagnosis: cytology, histology, microbiological isolation, continue post mortem. Cardiomyopathy with muscle cysts: sarcocystis; diagnosis: cytology, histology. An enlarged lumen of the left ventricle and only slight difference in thickness of the ventricle walls: heart failure; diagnosis: congestion of the lungs and/or liver.

5.

Examine and measure the liver. Take a sample for cytology and histology. Decide if you want to do a bacteriological culturing or freeze piece of liver-tissue for virological or toxicological testing.

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Separate the rest of the liver from the viscera by holding the ligaments in the forceps and cutting them with scissors, examine the gallbladder (if present). For a thorough examination, slice the liver at regular intervals. Examples Enlarged red variegated liver with pale areas: hepatitis; diagnosis: cytology with many inflammatory cells; histology. Enlarged liver with necrotic areas: hepatitis by chlamydiosis, herpesvirusinfection; diagnosis: imprints, culture, histology, PCR. Very extensive acute liver necrosis: suspected for peracute or acute hepatitis by bacterial septicaemia, polyoma-, herpes- adeno- or reovirus; in juvenile African grey parrots acute circovirus infection diagnosis: macroscopic (gross) examination, cytology, histology, virology (PCR), culture. Focal yellow proliferation with often central necrosis; tuberculosis; diagnosis: see above. Small round yellow necrotic foci: salmonellosis or yersiniosis; diagnosis: imprints with rod-shaped bacteria; culture. Evenly enlarged, often variegated, pale liver: leucosis; diagnosis: macroscopic (gross) examination; included often other organs; cytology and histology. Evenly enlarged, often variegated, pale soft liver: degeneration; diagnosis: cytology hepatocytes with vacuoles; histology. Enlarged orange, yellow liver: fatty liver; diagnosis: macroscopic (gross) examination, cytology, histology with Sudan III stain. Liver with necrotic ulcer: histomoniasis (black head); diagnosis: histology.

6.

The spleen can be found by cutting the oesophagus in the bifurcation of the trachea and with combined blunt and sharp dissection remove the viscera leaving the lungs and kidneys. Do not cut the cloaca, but bend the viscera caudally. This exposes the spleen in the angle between the proventriculus, gizzard (and liver). Examine, remove and measure the spleen; make impression smears from a fresh cut surface after blotting to remove excess blood. Examples Spleen-swelling together with air sac opacity: chlamydiosis; diagnosis: see above. - Very large swollen and cherry red spleen in parrots watch for herpesvirus infection (= Pacheco's) or sarcocystis; diagnosis: liver necrosis with intranuclear inclusion bodies or protozoa, cytology, histology, IFT, PCR, virus isolation. - Very large swollen and cherry red spleen in penguins and some other species; Plasmodium infection; diagnosis: cytology for parasites in macrophages and severe pneumonia, histology Swollen and pale: (bacterial) septicaemia; diagnosis: cytology with bacteria, culture. Multiple irregular yellow foci in the spleen; tuberculosis; diagnosis: the same foci in other organs, in imprint non-staining rods, acid-fast. Differentiation avian/bovine strains by culture or PCR. Large firm spleen: tumour; diagnosis: histology. Enlarged friable spleen with multiple, milliary necrotic foci: salmonellosis, yersiniosis; diagnosis: the same foci in liver and caeca; imprint with rod shaped bacteria; culture. Homogeneous red enlarged spleen in canaries and finches: atoxoplasmosis; diagnosis: cytology.

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Small, grey spleen: lymphoid depletion; stress, viral infection; diagnosis: cytology, histology, virus isolation.

Examine the adrenals, gonads (determine sex) and genital tract, and the kidney with ureters in situ. Remove the kidneys by applying gentle traction to the cranial vessels. Notice the adrenals and look for the sciatic nerve in the middle division of the kidneys. In our laboratory the kidneys are not routinely screened in cytology, but only when pathological changes are seen. Examples A swelling inside the oviduct: egg-binding, egg concrements; diagnosis: open the oviduct. Irregular swellings related to kidney or gonads: tumour; diagnosis: macroscopic (gross) examination and histology. Pale swollen kidneys with white striation: urate congestion; diagnosis: dehydration; histology (fixation 100% alcohol!!). Irregular pale swollen kidney with white foci: nephitis with "renal gout"; diagnosis: histology (fixation 100% alcohol). Irregular swollen kidney with multifocal abscessation: bacterial infection; diagnosis: histology, cytology, culture. Enlarged red kidneys: acute nephritis; diagnosis: histology. Pale swollen friable kidneys: kidney degeneration; diagnosis: histology. White, firm small kidneys: chronic kidney fibrosis; diagnosis: macroscopic (gross) examination NB the adrenals are important for the histological diagnosis of proventricular dilatation disease (PDD, avian bornavirus infection) in psittacines.

8.

Free the lungs by applying gentle traction to the trachea and oesophagus and cut the attachment to the ventral ribs and backbone at the thoracic inlet. This may be difficult as there is no pleural space in birds. Using blunt and sharp dissection will free the lungs. Inspect the lungs. Open the oesophagus. To open the syrinx, trachea and main bronchi a strip has to be cut out; cut through the lungs at intervals; make an impression smear from the lungs. Examples Dark coloured grey lungs: lung oedema; diagnosis: on cut surface clear serosal fluid. Dark coloured wet red lungs: lung congestion; diagnosis: from a cut surface only blood; the lungs are supple and evenly bright red; watch for congestion in other organs and alterations of the heart. Think also of polytetrafluoroethylene (TeflonR) toxicosis, acute mycotic infection, Plasmodium and sarcocystis. Dark firm lungs often variegated and focal changes: pneumonic foci; diagnosis: cut surface; cytology (inflammation cells); histology. Dark, supple, dry lungs: atelectasis; diagnosis: on cut surface only a dark colour of the surface of the lung and dried up. Scattered through the lungs white/yellow foci: aspergillosis, tuberculosis; diagnosis: wet mount with hyphae (aspergillosis), acid fast rods (in routine quick staining, non-stained rods) (tuberculosis); culture and histology. Irregular scattered pneumonic foci: bacterial pneumonia; eg. Salmonella spp. or Yersinia spp.; diagnosis: cytology and culture. In the syrinx of parrots white material: syringeal mycosis; based on a metaplasia due to vitamin A deficiency; diagnosis: see aspergillosis. In the trachea red worms: Syngamus spp, black dots: Sternostoma mites; mucous and fibrin: avipoxvirus, cytomegalovirus

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9.

Post-Mortem Procedures

Examine the "abdominal" viscera. Open and inspect the proventriculus and gizzard (with koilin layer). Examine the contents for foreign bodies and heavy metals. The bowel should be opened by making cuts at intervals to examine the contents and the wall for changes and parasites. Open and inspect the caeca when present. Look for the pancreas, bursa and umbilical sac. Take the following samples: ---> contents from the duodenum and rectum for parasitology (including direct examination on very fresh specimens for flagellates as well as other techniques) ---> contents from the rectum for a smear for staining (Diff Quick®) ---> contents from the rectum for microbiology Examples Crop Thickened wall with white material: yeast infection; diagnosis: smear of the material; culture. Thickened wall with mucous material: capillaria infection; diagnosis: smear of scraping of the epithelium; histology. Thickened wall with grey/yellow material, sometimes with trapped air bubbles; trichomoniasis; diagnosis: wet mount; cytology; histology. Local yellow necrotic ulceration: pox-lesions; diagnosis: macroscopic (gross) examination; histology; virusculture. Local red mucosal thickening: papillomas: diagnosis: histology. Stomach (proventriculus and ventriculus) Dilated proventriculus and gizzard, often stuffed with seeds (sunflower): gastric dilatation syndrome; diagnosis: histology; (ganglio)neuritis, lymfoid infiltrates in the adrenals.. An empty proventriculus with excess of mucous: Macrorhabdes ornithogaster (formally "megabacteria"); diagnosis: wet mount and cytology. Swollen red glands in proventriculus: Tetrameres spp; diagnosis: parasitologic examination. Intestines Haemorrhagic contents duodenum: coccidiosis; diagnosis: wet mount, cytology. Haemorrhagic, black contents in the entire small intestine: haemorrhagic diathesis; diagnosis: history (fasting during high energy need for over 24 hours), macroscopic (gross) examination. Pseudomembraneous covering of the duodenal wall: hexamitiasis; in cranes; diagnosis: wet mounts, cytology and histology. Thickened wall with or without blood in the lumen: enteritis; diagnosis: wet mount and cytology; parasitology; microbiology. Beware: in psittacines very rarely coccidia, often ascaridia; in small passerines rarely worms, often coccidia spp. Haemorrhagic contents: lead intoxication, clostridium infection, pseudomonas infection, Giardia spp.; diagnosis: lead in gizzard; lead analysis liver and kidneys; cytology, culture. Clear watery contents in small intestine with flabby wall: hexamitiasis; diagnosis: fresh wet mount, cytology, histology. Yellow non-digested starch and broken seeds in small passerines: Cochlosoma or Campylobacter spp.: diagnosis: fresh wet mount, cytology, selective culture. Enlarged caeca with pseudomembraneous to necropurulent content; typhlitis; diagnosis: galliformes: histomoniasis ("blackhead"); diagnosis: cytology, histology (often with liver lesions)

VII.

-

Post-Mortem Procedures

Caeca with nodular lesions: parasitic typhlitis; pheasants: Heterakis isolonga; diagnosis: worms and ova; histology

Cloaca Congested, swollen red mucosa: papilloma: diagnosis: histology. Bursa Especially in young birds for detecting virus infections e.g. circovirus: diagnosis: histology, PCR.

BEWARE: TRY TO ESTABLISH A RELATIONSHIP BETWEEN THE CLINICAL HISTORY AND THE POST-MORTEM FINDINGS

10.

Open the anterior part of the oesophagus from the beak, make a wet mount. Remove the tongue and cut the salivary glands. Inspect the beak, choanae and oesophagus. Examples Tongue with yellow "abscesses" at the location of the salivary glands in psittacines: metaplasia, due to vitamin A deficiency; diagnosis: wet mount, diet history, histology. see also crop/intestines (eg. trichomoniasis, avipox, candidiasis). Chronic, necrotic lesions especially in commisures: tuberculosis: diagnosis: cytology (acid fast stain), histology, culture.

11.

Cut across the beak through the nostrils and sinuses. Examples The presence of turbid mucus: sinusitis; diagnosis: wet mount, cytology, culture.

12.

13.

Inspect the joints, bones, bone marrow, brains. Joints of the wings, legs and feet should be opened and examined. If exudate is present, cytology should be done as well as a microbiological examination. White, chalky deposits may represent urate deposition. Bone marrow is most easily collected from the tibiotarsus for both cytology and histology. In bone marrow tubercular lesions can be found; often visible on X-rays. Ecchymoses within the calvarium are a common agonal change and do not imply head trauma. Examination of the nervous system and associated tissues is governed by the presence or absence of neurological or ocular disease. The brain may be removed by deskinning the head, making a sagittal incision through the calvarium and removing the bony calvarium to expose the brain. When sampling for histology, it is often better to leave the brain inside the skull, after opening it, and immerse the whole head in formalin. The muscles of the legs and the sciatic nerve running on the posterior surface of the femur should be examined.

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Post-Mortem Procedures

V. Final activities 1.

Bacterial cultures are done from the liver and the rectum and all abnormal organs, especially when bacteria are seen in the imprints! The following media are selected: blood agar, selective Enterobacteriaceae agar (brilliant-green-agar) and serum broth. The intestinal contents are collected in tetrathionate broth as an enrichment medium for Salmonella spp. When special microorganisms are expected (e.g. anaerobes, Campylobacter spp.) contact the laboratory.

2.

When a mycotic problem is suspected a Malt-agar, or other selective culture medium, is selected as well.

3.

The impression smears are allowed to dry, stained with 'HemacolorR' or "Diff QuickR" and Stamp or Macchiavello (for Chlamydia), and examined by microscope with objective 100x in immersion oil. The slide for an IFT for Chlamydia is fixed in cold acetone (freezer -20°C) and sent to the laboratory.

4.

POSITIVE cytology for Chlamydia requires sampling for IFT or PCR.

5.

Examine the wet mounts of gut contents.

6.

Samples collected for ancillary diagnostics should be packed, labelled and stored properly, until shipment. See that each sample is provided with the essential documentation .

7.

Make a detailed report and use this to document the samples.

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Post-Mortem Procedures

References 1.

2.

3.

4.

5.

6.

Graham, D. L. 1992. Checklist for necropsy of the pet bird and preparation and submission of necropsy specimens - A mnemonic aid for the busy avian practitioner. AAV Introduction to Avian Medicine and Surgery, New Orleans, LA, USA, pp. 1-4. Lowenstine, L. J. 1986. Necropsy procedures. In: Harrison, G. J., and L. R. Harrison (eds). Clinical Avian Medicine and Surgery, P.A. Saunders, Philadelphia, Pennsylvania. Pp. 298 - 309. Latimer, S. L., and P. M. Rakich. 1994. Necropsy examination. In: Ritchie, B. W., G. J. Harrison, and L. R. Harrison (eds). Avian Medicine: Principles and application. Wingers Publishing, Inc. Lake Worth, FL, USA. pp. 355-379. Dorrestein, G. M. 1996. Cytology. In: Beyon, P. H., N. A. Forbes, and N. H. HarcourtBrown (eds), Manual of Raptors, Pigeons and Waterfowl, BSAVA, London, UK. Pp. 5562 Dorrestein Gerry M. and Wit, Martine de (2005) Chapter 7. Clinical pathology and necropsy. In: BSAVA Manual of Psittacine Birds. 2nd ed. N. Harcourt-Brown and Chitty J. (eds). pp. 60-86. Dorrestein, G. M. 1997. Diagnostic Necropsy and Pathology and Avian Cytology. In: Altman, R. B., S. L. Clubb, G. M. Dorrestein, and K. Quesenberry (eds). Avian Medicine and Surgery, WB Saunders, Philadelphia, Pennsylvania, USA. Pp. 158-169 and pp. 211222.

VII.

Post-Mortem Procedures

Necropsy report form and check list The following checklist can be use during the post-mortem examination as well as writing the necropsy report:

1. Bird species, weight, age/leg band number, sex, summarized history. 2. Date of necropsy, your name. 3. Macroscopic (gross) examination: External examination - general bodily condition: muscle mass: robust, well muscled, moderately muscled, thin, emaciated, depot fat) - feathers/integument/ectoparasites - palpate skeleton - body openings/oral cavity Internal examination - fat/subcutis/body wall - body cavities (air sacs/pleura/peritoneum) - (para)thyroids, thymus - spleen (size) - heart, aorta, other vessels - liver, gall bladder, bile ducts - reproductive system (gonads, repr. tract) - urinary tract (kidneys, ureters) and adrenal glands. - respiratory tract (nasal/sinus, choanal, larynx, trachea, syrinx, air sacs, lungs) - digestive tract (beak, tongue, oropharynx, oesophagus, crop, proventriculus, gizzard, duodenum and pancreas, small intestine, yolk sac, caeca, rectum (colorectum), cloaca, bursa of Fabricius, vent) - special senses (eyes, ears, nares) - musculoskeletal system: muscles, skeleton (sternum, ribs, vertebrae, long bones), bone marrow, joints - brain, pituitary, spinal cord, meninges, peripheral nerves Wet mounts (crop, rectum, etc.) Cytology (liver, spleen, lung, rectum) Chlamydiosis TENTATIVE (DIFFERENTIAL) DIAGNOSIS Ancillary diagnostics: bacteriology, mycology, virology, parasitology, toxicology, others Tissue saved: Tissues submitted for histopathology:

Transmissible Diseases Handbook

VIII. GUIDELINES FOR CLEANING AND DISINFECTION IN ZOOLOGICAL GARDENS Matti Kiupel, Robin Mecklem, Birgit Hunsinger and Rachel E. Marschang Diagnostic Center for Population and Animal Health (MK) and Office of Radiological, Chemical and Biological Safety (RM), Michigan State University, Michigan, USA Institut für Umwelt- und Tierhygiene (REM, BH), Hohenheim University, Garbenstr. 30, 70599 Stuttgart, Germany Introduction Hygienic measures are a major component for prevention of infectious diseases in zoological exhibitions. A clean environment is not only a key feature in disease control, but also attractive to the public. Preventive health care consists of multiple components of equal importance including food storage, preparation and handling, insect and vermin control and cleaning and disinfection (Fowler, 1978). The purpose of this chapter is to provide guidelines for cleaning and disinfection to the range of people who would be involved in managing a disease emergency involving a zoo, zoological garden, game park, circus or any other facility keeping exotic animals. For the information in this chapter to be effective, it is important that the described cleaning and disinfection methods are incorporated into the routine husbandry and emergency procedures. The goal is to provide individual zoos with information that can be used to develop their own specific cleaning and disinfection protocols. Each of the fact sheets in the “Transmissible Disease Handbook“ contains disease specific information about prevention and control of diseases in zoos and suggested disinfectants for animal housing. Therefore, the purpose of this chapter is not to provide detailed information on specific disinfectants for the diseases listed in this handbook, but rather to provide an overview of the guiding principles of cleaning and disinfection in zoological gardens and to review the various classes of chemical disinfectants. Defining Disinfection The term “disinfectant” is defined as “an agent that frees from infection, usually a chemical agent but sometimes a physical one, such as x-rays or ultraviolet light, that destroys diseases or other harmful microorganisms but may not kill bacterial spores” (Block, 2000). Disinfectants are used on inanimate surfaces and are assumed to act rapidly and efficiently to kill or inhibit growth of microorganisms. In the veterinary care environment, disinfection is most effective for diseases that are not vector-borne, but are acquired by direct contact with contaminated fluids or animal products (Quinn, 2000). Therefore it is important to recognize that disinfectants play a critical, yet limited role in infection control. Insect and vermin control are equally important in preventing infectious diseases, especially for the control of insect-

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Guidelines for Cleaning and Disinfection in Zoological Gardens

borne diseases that present particular challenges in zoos. Techniques available to minimize insect-borne disease spread include the use of anthelmintics and insecticides through direct application on at-risk species or through the spraying of housing of susceptible animals. The assistance of epidemiologists and entomologists should be sought to establish insect traps within the zoo for insect identification and to identify potential areas of a high-risk exposure. For situations where resistant microorganisms are present, or animals may be susceptible to infection due to their health status or the invasiveness of the procedure to be performed, a sterilization method should be used for treatment of contaminated items or surfaces. Sterilization is achieved when all living microorganisms and bacterial endospores have been destroyed. Common methods of sterilization include, dry heat, moist heat, and exposure to specific chemical compounds. For dry heat sterilization, surfaces must be exposed to 160 C to 170 C for periods of 2 to 4 hours (Heinsohn et al., 1995). Sterilization by steam may be accomplished by exposing contaminated surfaces to moist heat at 121 C for at least 15 minutes (Quinn, 2000) This will not be sufficient for certain heat resistant organisms such as some thermophilic spores and prions. Chemical sterilization may be suitable in situations where items to be sterilized cannot withstand the temperatures and physical conditions required for dry heat or steam sterilization. Chemical sterilants may be used in the form of a vapor or gas, such as formaldehyde or ethylene oxide, or as an immersion liquid such as glutaraldehyde. These chemical compounds and the requirements for their proper use will be discussed in a later section of this chapter. The efficacy of a certain disinfection or sterilization protocol will depend on a number of factors. The nature of the surface that is to be disinfected plays an important role. An uneven surface will be more difficult to disinfect than an even one. Dirt will interfere with the disinfection. Another important factor are the pathogens present. Some pathogens are more difficult to inactivate than others. The most difficult to inactivate are bacterial spores and prions. For disinfection, non-enveloped viruses can also be a challenge. Role of Cleaning and Disinfection in Infection Control Having an effective cleaning and disinfection plan is a crucial step in every biosecurity program. Such a program should be instituted for every new building and facility and should be revised after the occurrence of an infectious disease and prior to the introduction of new animals. The main purpose of a cleaning and disinfection program is to reduce the number of pathogens (disease-causing agents) in the environment and thereby to reduce the potential for diseases to occur. The first step in an effective disease control plan is an exact identification of the routes how infectious agents may enter zoological gardens (inputs), and how they may spread throughout the zoo to other facilities and outside the zoo to become a threat to farming operations or possibly humans. Inputs into and outputs from zoological institutions may vary depending on the type of the facility. Animal inputs may include: animals introduced from other facilities within the same institution, animals from institutions, either from within Europe, or imported from another continent, animals confiscated by customs/quarantine officers, sick or injured animals brought in by members of the public, free-ranging animals, which may be either native (rats, mice, birds), or feral (including cats and dogs), and animals imported from farming

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Guidelines for Cleaning and Disinfection in Zoological Gardens

operations. Feed inputs include dry processed preparation (concentrates, hay, pellets, seed etc.) and wet feed, including fresh fruit, fish, meat, vegetables and pasture silage. Biological specimens confiscated by customs and quarantine officers are sometimes brought to zoos for identification. Semen and embryos may be imported for breeding purposes. Various vehicles move into the facility and may be contaminated. Other materials entering the facility include materials used during the importation of the animals (hay, sawdust, crates etc.). Personnel entering the premises for normal work purposes may have contact with other animals (pets/farm animals) outside of work hours. Local and international visitors pass through the premises on a daily basis and may introduce disease. Animals may leave the facilities for a number of reasons including: exchange and sale of animals, relocation of animals to other facilities within and outside the institution, use of animals in public relations exercises, i.e. animals taken to shows, shopping centers, schools and television stations, animals taken home by staff for hand rearing, free-ranging of native and feral animals, including cats and dogs that may have had contact with animals/animal waste. Waste materials that have to be removed from the premises may include hay and composted feces, effluent, some of which has secondary treatment, and a small portion with tertiary treatment, waste-water. Biological specimens and feces are sent to laboratories for testing and biological specimens are sometimes also sent to museums, veterinary schools etc. In some institutions, offal and carcasses are taken off the property for disposal or may be sent to veterinary schools for necropsy. Vehicles, crates and packing material used in the transportation of animals are all regularly moved off the premises. Possible outputs related to people include personnel who have been in contact with animals and waste products (contaminated clothing and footwear), and visitors who may have been in contact with animals. Due to the public role of zoological gardens, it will be impossible to clean and disinfect all inputs and outputs. In particular, visitors and the media would not except the imposition of cleaning and disinfection programs onto them. Therefore a strict cleaning and disinfection program of the animal premises incorporated into daily routines becomes even more important. In addition, it is useful to identify premises according to their exposure to infectious agents to select the proper cleaning and disinfection regimes and to possibly limit the access to such premises. Areas (could be all or part of a facility) in which an infectious disease exists, is believed to exist, or which may harbor the infective disease agent should be classified as an infected premise (IP). Dangerous contact premises (DCP) are premises containing animals with no clinical signs that were exposed to an infectious disease and therefore will be subjected to disease control measures. Suspect premises (SP) are areas containing animals that show no clinical signs, but may have been exposed to an infectious disease through possible contact with infected animals or facilities, people, equipment, semen or embryos, or animals with evident disease symptoms, but no confirmed diagnosis. Using this classification in combination with specific cleaning and disinfection programs and other measures, such as insect and vermin control, as well as treatment, vaccination and quarantine of infected and exposed animals or animals at high-risk for infections, will help to control and to eliminate an infectious disease after an outbreak has occurred. In some cases, it may also be necessary to consider culling certain animals that have been exposed to disease. Disposal of animal carcasses in the case of a disease outbreak should also be incorporated into these plans.

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Guidelines for Cleaning and Disinfection in Zoological Gardens

Cleaning before Disinfection The first step in any cleaning and disinfection program is cleaning. Cleaning is the removal of organic material (i.e., feces, urine, blood, bedding, food, dust etc.). Disease agents are often protected by such materials and can survive the disinfection process. Therefore, thorough cleaning of a building prior to the disinfection is required. The cleaning process can include a dry cleaning and a wet cleaning step. Dry cleaning physically removes the organic material before the actual wet cleaning. Wet cleaning, as the name implies, involves the use of water. There are 4 basic steps in the wet cleaning process: soaking, washing, rinsing, and drying. The use of detergents will often benefit the wet cleaning process. However, it is more important to have pressure washers with the proper pressure (500-800 psi) to ensure that all organic materials are removed from the facilities. The final step for ensuring proper cleaning is to dry the wet areas of the building quickly. If the building is not dried properly, the excess moisture can result in bacteria multiplying to higher levels than prior to cleaning. It is vital to make sure the cleaning procedure is done properly, as an improper cleaning can actually increase the load of infectious agents! If done properly, a good cleaning can remove 90% of all pathogens (Fotheringham, 1995). Special care must be taken when cleaning facilities that are contaminated with suspected zoonotic or highly contagious animal pathogens. All personnel should wear protective clothing, footwear and if necessary face masks, goggles, and headwear. After the cleaning of facilities, proper care must be taken of the protective clothing and equipment that has been used for cleaning and disposable products should be disposed as biohazards. Organic material removed during cleaning may also contain pathogens and should be treated accordingly. The last step in a cleaning and disinfection program is the actual disinfection process that will further reduce pathogens in the facilities. Disinfection is especially useful in reducing infection risks in young animal nursery facilities, and in routine cleaning operations of animal quarters and feeding utensils. To maximize the effectiveness of a cleaning and disinfection program, it is crucial to modify such a program based on the suspected pathogens that should be eliminated or reduced. In addition, specific disinfectants may be selected for certain known microbial contaminants following an infectious disease outbreak. It is also important to remember that many disinfectants can be toxic to the animals, or may be caustic or corrosive. Animals must usually be excluded from facilities being disinfected. After a suitable environmental exposure time to such disinfectants premises should be rinsed thoroughly, before animals are allowed to return. Levels of Disinfection Disinfectants are tested for their bacteriocidal, tuberculocidal, sporicidal, fungicidal, virucidal (against enveloped and non-enveloped viruses), and antiparasitic (against eggs and coccidia) effects. They can generally be categorized into 3 groups based on their ability to inactivate certain microorganisms. High-level disinfectants are effective against bacterial endospores under specific conditions. Intermediate-level disinfectants include those products that can inactive tubercle bacilli, but do not kill bacterial spores. Such products are commonly marketed by manufacturers as “tuberculocidal”. Finally, normal (“low-level”) disinfectants include products that kill vegetative bacteria and fungi but are not reliable for the destruction of bacterial endospores, tubercle bacilli or small non-enveloped viruses within a practical period of time (Boothe, 2000).

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Guidelines for Cleaning and Disinfection in Zoological Gardens

When selecting a disinfectant for use on items that will come in contact with patients during animal care procedures, the invasiveness of the procedure and composition of the medical device must be considered. A system for classifying such devices was developed by Spaulding in the 1970s. This system has been recognized and referenced since that time by many human healthcare organizations like the Association for Professional in Infection Control and Epidemiology (Rutala, 1996). Although this system was developed for human healthcare, there are notable similarities to procedures and devices used in veterinary care environments. These principles may be adapted for disinfectant selection purposes in zoological gardens and other exotic animal containments. The Spaulding system categorizes devices into 3 groups based on the risk of infection involved with their use: critical, semicritical and noncritical. Critical devices are those that bear a substantial risk of acquiring infection if contaminated with microorganisms at the time of use. Devices that are introduced directly into the body such as needles, scalpels, catheters and implants fall into this category. These items must be sterilized in between animal use (Favero and Bond, 2000). When choosing a sterilization method, consideration must be given to the impact of the sterilization process on the integrity of the device. Semicritical devices are those items that come in contact with mucous membranes but do not normally penetrate the skin. Examples of these devices include endoscopes, bronchoscopes, and urinary catheters. Sterilization of semicritical devices is desirable, but high-level disinfection is a minimum requirement (Favero and Bond, 2000). High-level disinfectants may also act as cold liquid sterilants under specific conditions of use. Therefore, some products may be appropriate for both critical and semicritical devices. Product versatility should be one of the considerations for disinfectant selection. Non-critical devices are items that only come in contact with an animal’s intact skin, and therefore bear the lowest relative risk of disease transmission. Stethoscopes, otoscopes, electrodes, and common animal restraining tools are a few examples of non-critical devices. Proper treatment of this category of devices in between animals will depend on the nature and degree of contamination sustained during use. Where contamination is minimal, simple scrubbing with detergent and warm water may be acceptable to assure safety. However, the use of low- or intermediate-level disinfectants may be appropriate to assure that disinfection has been accomplished (Favero and Bond, 2000). Factors that Impact Efficacy of Disinfection Effective disinfection can only be achieved if the following interrelated factors are taken into account: Nature of the item or surface to be disinfected Items with smooth, non-porous surfaces are the easiest to disinfect because they can be effectively cleaned of organic debris and this will allow for ample surface contact with the disinfectant applied. As a rule of thumb, the “rougher” the surface (i.e. crevices, damaged finish), the more difficult cleaning and subsequent disinfection will be. To account for this, increased contact time of the disinfectant or a higher disinfectant concentration on these surfaces may be beneficial (careful with increased concentrations: toxicity and corrosive or caustic effects may also be increased).

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Guidelines for Cleaning and Disinfection in Zoological Gardens

Contact time and other factors The surface contact time is the minimum required duration of exposure of the contaminated surface to the disinfecting agent in order to achieve the level of disinfection that is claimed by the manufacturer. Therefore, careful consideration should be given to the manufacturer’s recommendations and conditions of use. These recommendations will also address other factors that may impact efficacy such as pH, water hardness, performance in the presence of organic debris and optimal temperatures for product use. Presence of organic material The presence of organic material on surfaces to be disinfected will generally compromise disinfection. Any type of contamination will reduce the surface contact time of the applied disinfectant and therefore its effectiveness. Blood, blood products, body fluids and feces contain proteins that will bind and inactivate some disinfections or slow their action. This reinforces the importance of cleaning surfaces prior to disinfection whenever possible. If organic debris cannot be removed, an increased concentration of the disinfectant or contact time may be warranted. Furthermore there are also products available that will be effective in the presence of organic debris. Number and type of microorganisms present In general, items that have a higher level of microbial contamination will require a longer exposure to chemical products to achieve disinfection. Level and type of microbial contamination may be presumed by the nature of a procedure that a device was used for, the health status of the animal, or other epidemiological factors associated with the veterinary care environment. Resistance of microorganisms Microorganisms vary in their susceptibility to disinfectants. In general, obligate intracellular bacteria, such as Mycoplasma, are the most susceptible to disinfection. Gram-positive and Gram-negative bacteria, enveloped viruses and fungal spores are less susceptible, but are not considered to be resistant to disinfection. Highly resistant microorganisms include nonenveloped viruses and mycobacteria. Although these organisms are hearty, they are not as resistant as bacterial endospores or protozoal oocysts and even more prions, which are resistant to most disinfectants (Quinn and Markey, 2000). The level of disinfection should be based on the most resistant microorganism posing a risk in a given situation. Type and concentration of disinfectant As previously noted, microorganisms vary in their overall resistance to disinfection. However, resistance will also depend on the type of disinfectant used. Each class of chemical products acts differently on the cells. It is important to assure that the mode of action for the disinfectant is compatible for the type of cells to be destroyed. General modes of action for commonly used disinfectant classes are addressed in a later section of this chapter. Regarding concentration, with all other variables constant, the higher the concentration, the greater the effectiveness of the disinfection and the shorter the contact time (Favero and Bond, 2000). However, there are exceptions to this rule including alcohol and iodophors that have an optimal range of concentration. While the concept of using concentrated

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Guidelines for Cleaning and Disinfection in Zoological Gardens

disinfectants to achieve faster, more effective disinfection may seem like an attractive timesaver, caution should be exercised. Chemical disinfectants are often hazardous to human and animal health and an increased concentration is likely to equate to increased chemical exposure hazard to personnel, animals and the visiting puplic. Factors for Selecting a Disinfectant Because all disinfectants are unique in their chemical action and properties, it is unlikely that one disinfectant will fit all needs. However, for veterinary care purposes, there are a number of desirable qualities that have been identified and should be considered for product selection. These include (Quinn and Markey, 2000): o o o o o o o o o o

Wide antimicrobial range Absence of chemical hazards (i.e. toxicity, teratogenicity, carcinogenicity) Compatible with a wide range of chemicals Non-corrosive Active in the presence of organic debris Stable at ambient temperatures Long shelf-life Effective over wide range of temperatures Inexpensive and readily available Nonpolluting and biodegradable

In Europe, there are a number of organizations that test disinfectants for use in animal husbandry. In Germany, the German Veterinary Medical Society (DVG) provides guidelines for testing disinfectants and publishes a list of disinfectants that have been tested for use in animal husbandry. The list also contains information on effectivity of the products against different categories of pathogens, and suggested concentrations and incubation times. Products in this list have also been tested under conditions meant to simulate “real life” outside the laboratory, and include effectivity in a protein-contaminated environment and on “rougher” porous surfaces. The European Committee for Standardization (CEN) is currently establishing European guidelines for disinfectant testing in the veterinary field. Overview of Categories of Chemical Disinfectants There are a number of compounds that have specific use as disinfectants in the veterinary care environment. An overview of the main classes of chemical disinfectants is given in this section but periodic consultation to current literature is required as new products are constantly under investigation and emerging infectious diseases continue to rise. For lists of tested disinfectants see e.g. the DVG (German Veterinary Medical Society) for products used for veterinary medicine or in the food industry or the DGHM (German Society for Hygiene and Microbiology) for disinfection in human medicine and hospitals. At this time, no general European lists are available. Alcohols Alcohols are inexpensive, relatively nontoxic, and colorless, and ethyl and isopropyl alcohol are most widely used. They are considered to be intermediate level disinfectants that inactivate organisms by denaturing proteins. While they are effective for destruction of enveloped viruses and tubercle bacilli, they are less effective against non-enveloped viruses

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Guidelines for Cleaning and Disinfection in Zoological Gardens

(particularly isopropyl alcohol) and are not sporocidal (Maillard and Russell, 1997). A concentration of 70% of ethyl alcohol is most effective. When used as a surface disinfectant, rapid evaporation of these compounds makes sufficient contact time difficult to achieve. Additionally, alcohols tend to harden and swell plastic tubing when used over time, and may be absorbed by rubber products that can lead to irritation of the skin and mucous membranes (Favero and Bond, 2000). Aldehydes Formaldehyde (available in an aqueous 37% solution) and glutaraldehyde are two common examples of this class. Formalin in solutions of 3% - 8% is effective for intermediate to high level disinfection. It is less effective than glutaraldehyde in the presence of organic matter. Despite discussions on potential carcinogenicity, skin irritation, and irritating fumes, aldehydes are common compounds in disinfectant preparations used in the veterinary field (i.e. breeding, husbandry, transport and disposal of all animals). Glutaraldehyde, which inactivates microorganisms by alkylation, is also an intermediate to high level disinfectant. At 2% active ingredient, glutaraldehyde solutions are effective against bacterial endospores. Although minimally affected by organic matter, product effectiveness is influenced by a number of factors such as pH and temperature. Glutaraldehyde solutions are most effective at alkaline pH and effectiveness increases with temperature (Quinn and Markey, 2000). Glutaraldehyde solutions are noncorrosive and are typically used in immersion baths for high level disinfection of semi-critical devices (i.e. endoscopes) that cannot withstand steam or heat sterilization. Due to the toxic nature of this compound, special considerations must be made for proper handling and storage of solutions. Immersion baths should have tight-fitting lids and be placed in an exhaust hood to minimize personnel exposure to vapors. Additionally, to eliminate skin and mucous membrane exposure, splash goggles and appropriate chemical resistant gloves should be worn. Aldehyds should not be used at temperatures below 10 °C. Alkalis Sodium hydroxide (NaOH), or lye is a caustic alkali that has a wide virucidal spectrum when used at a 2 percent concentration (prepare by mixing 1/3 cup of NaOH pellets per gallon of water – careful! hot). This is effective against most bacteria, and enveloped and nonenveloped viruses, although somewhat higher concentrations may be necessary for some viruses. This concentration is effective against virusesincluding the causes of Avian Influenza, Rinderpest and Pest of Small Ruminants, Newcastle Disease, Malignant Catarrhal Fever, Lumpy Skin Disease and Sheep and Goat Pox, African Horsesickness and Bluetongue, Foot-and-Mouth Disease, and Swine Vesicular Disease. Sodium hydroxide should not be used on wood. Sodium hydroxide is also one of the few compounds that can be used for the inactivation of prions (in much higher concentrations) (Taylor, 2000). An overview on deactivation of prions is given below. A pH above 12 is required to for the inactivation of bacilli such as Mycobacterium bovis. Chemical exposure risks to personnel and animals as well as surface incompatibility are factors for the limited use of such products. Ammonium hydroxide, a weak base, has a strong activity against coccidial oocytes (Williams, 1997). The high pH of even low aqueous solutions and intense fumes require protective clothing when using this alkali.

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Guidelines for Cleaning and Disinfection in Zoological Gardens

Sodium carbonate, containing 0.1% sodium silicate, has significant virucidal activity and is commonly used for the decontamination of aircrafts. Biguanides Biguanides are a group of cationic compounds commonly used for hand washing and skin preparation. Chlorhexidine, a low-level disinfectant that is incompatible with anionic compounds, is the most commonly used compound in this group. Its activity is pH dependent and it is inactivated by organic debris (McDonnell and Russell, 1999). It is more active against grampositive than gram-negative bacteria and Pseudomonas spec. and Proteus spec. are resistant against chlorhexidine (Widmer and Frei, 1999). Although it is active against enveloped viruses, it can not be used against non-enveloped viruses, mycobacteria and most fungal species, such as dermatophytes, i.e. Microsporum canis, are resistant (DeBoer et al., 1995) Because chlorhexidine has a longer residual activity on teat skin than most other disinfectants, it is widely used for mastitis control (Quinn and Markey, 2000). Halogens Chlorine Compounds Chlorine and chlorine releasing compounds, such as hypochlorite, chlorine dioxide, sodium dichloroisocyanurate, and chloramine-T, are used in some countries for disinfection of surfaces, equipment, buildings and vehicles as well as disinfection of milking installations and in the food industry. Sodium hypochlorite (NaOCL or household bleach) solution is an intermediate-level disinfectant when properly diluted. The mode of action for this solution is dependent on the formation of undissociated hypochlorous acid that will oxidize peptide links and denature proteins (Maris, 1995). Solutions diuted to final concentration must be prepared directly before use and should not be stored for more than 1 day. At a concentration of 0.1%, this compound is effective against microbial agents of diseases, including enveloped viruses and Foot-and-Mouth Disease virus. It can be prepared at the time of use by adding approximately 30 cc (ml) of household bleach to a gallon of water (or 1 gallon of bleach plus 50 gallons of water). As pH decreases microbicidal activity (as well as corrosivity) increases. In concentrated form chlorine-based disinfectants are usually unstable, and affected by light and heat. They can lose 50% of their concentration in a month when left in an open container (Widmer and Frei, 1999). Additionally, the corrosivity of active solutions can be damaging to surfaces and equipment and irritating to the skin and eyes. Bleach solutions are readily inactivated by organic debris, limiting their effectiveness to situations where devices or surfaces have been thoroughly cleaned prior to disinfection. In areas heavily contaminated with secretions, excretions, and soil, there is a considerable organic demand for available chlorine and disinfection should be repeated at least once. Under such circumstances a 3% solution of sodium hypochlorite should probably be used (3 liters of bleach to 2 liters of water). This concentration is effective against a variety of agents of viral diseases including Avian Influenza, Rinderpest and Pest of Small Ruminants, Malignant Catarrhal Fever, Lumpy Skin Disease and Sheep and Goat Pox, and African Horsesickness and Bluetongue as well as other non-enveloped viruses. At concentrations used for water

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Guidelines for Cleaning and Disinfection in Zoological Gardens

treatment it is not effective against Cryptosporidium parvum (Fayer, 1995). Halogens rapidly lose efficacy on rough surfaces such as concrete. Iodine Compounds Iodine compound disinfectants are more effective in the presence of organic matter than chlorine compounds, and less chemically reactive. Inorganic iodine and iodophors, compounds in which iodine has been complexed with polymers to sustain the release of free iodine and to increase water solubility, have both been used for disinfection. However, the instability of inorganic iodine in the environment, skin irritations, hypersensitivity reactions, and intense staining of contaminated surface limit its use. At appropriate dilutions, iodophors are bactericidal, mycobactericidal, sporicidal, fungicidal, and virucidal. They are also widely used as teat dips (Saran, 1995). Because the activity of iodophors is greater at a low pH, the compound should not be used in alkaline conditions or mixed with other compounds. Certain acidified iodophor compounds that contain a generic formulation of polyethoxysubstituted polypropoxy-ethane complex are extremely effective disinfectant compounds with strong virucidal activity (Maillard and Russell, 1997). The active ingredient must provide a minimum use concentration of 0.02% titratable iodine. Organic and Anorganic Acids Acids inhibit the growth of microorganisms and a number of compounds are commonly used as preservatives in the food industry. Acetic acid has been used for the treatment of wounds infected with Pseudomonas spec. (Lemarie and hosgood, 1995). The use of acids may be warranted under certain circumstances in the animal care environment. Formic acid is the most effective of these substances and should be used at a concentration of 4%. Citric acid, formic acid, and strong mineral acids, such as phosphoric acid, are effective for inactivating the Foot-and-Mouth Disease virus (Quinn and Markey, 2000). Two mineral acids, hydrochloric acid and sulfuric acid, are sometimes used for cleaning and disinfection of animal housing. A 2.5% concentration of hydrochloric acid can be used to inactivate endospores of Bacillus anthracis on the skin and is also effective against Rotaviruses and Vesicular Stomatitis virus (Maillard and Russell, 1997). However, the activity of these compounds is highly pH dependent and these compounds are corrosive and hazardous to workers and animals. To minimize chemical exposure risk to personnel, citric acid should be used when possible. Peroxygen Compounds Strong oxidizing agents such as hydrogen peroxide and peracetic acid, have a broad antimicrobial spectrum. Hydrogen peroxide is fast acting, nonpolluting, and decomposes to oxygen and water. Because it is unstable in solution, stabilizers are usually added. It is bactericidal, fungicidal and virucidal, but should not be used against Mycobacteria (Maillard and Russel, 1997). It is quickly inactivated by protein contamination of treated surfaces. Peracetic acid is a strong oxidizing agent and an even stronger disinfectant that has algicidal, bactericidal, fungicidal, virucidal and sporicidal activity. Unfortunately it can corrode most metals, rubber and has been suspected to act as a cocarcinogen (Sattar and Springthorpe, 1999). Oxidising agents rapidly lose efficacy on rough surfaces such as concrete.

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Guidelines for Cleaning and Disinfection in Zoological Gardens

Phenolic Compounds Phenol is cited in the literature as being the original standard against which other disinfectants were compared (Quinn and Markey, 2000). Due to its toxicity and unpleasant odor, the use of phenol for disinfection purposes has been largely replaced by the use of synthesized phenolic compounds. Examples include ortho-phenylphenol and ortho-benzylpara-chlorophenol. These compounds are a complex group of chemicals often found in various combinations and concentrations in disinfectant products. The effectivity of phenolic compounds increases with the length of the alkyl chains and number of halogen atoms in the molecule, but this also increases the sensitivity of the compounds to organic material, lowers their water solubility and increases their toxicity, so that these compounds are not frequently found in disinfectants for veterinary medical use. These products are considered to be intermediate to low level disinfectants. For veterinary care purposes, it is important to note that these compounds generally leave a residue on surfaces that may increase chemical exposure for personnel and animals. Additionally, pigs and cats are susceptible to the toxic effects of these compounds (Quinn and Markey, 2000). Also available are substituted phenolic compound, that have potent virucidal activity when prepared as a 1% solution of stock disinfectant. These types of compounds are effective against Eimeria tenella (Williams, 1997) and the viruses of African Swine Fever, Avian Influenza, Hog Cholera, and Velogenic Newcastle Disease, but not effective against the viruses of Foot-and-Mouth Disease and Swine Vesicular Disease (Widmer and Frei, 1999). Quaternary Ammonium Compounds (QACs) Quaternary ammonium compounds (QACs) are cationic detergents that may be used for low level disinfection, especially surface disinfection. Benzalkonium chloride and dodecyl dimethyl ammonium chloride are common examples of this group. QACs are relatively nontoxic making them appropriate for general cleaning purposes. However, QACs are inactivated by organic debris, metal salts in water (i.e. hard water) and anionic detergents (Heinsohn et al., 1995). Even some gram-negative bacteria, such as Pseudomonas spec., can survive disinfection with QAC and may grow in QAC solutions. Such limitations must be considered when developing procedures for the use of the products in the animal care environment. QACS are found in many disinfectant formulations used in the food industry. Inactivation of prions Because of the political impact transmissible spongiform encephalopathies have had in the past few years and the public perception of these diseases, this paragraph has been included. Prions are characterized by extreme resistance to conventional inactivation procedures including irradiation, boiling, dry heat, and chemicals (formalin, betapropiolactone, alcohols). While prion infectivity in purified samples is diminished by prolonged digestion with proteases, results from boiling in sodium dodecyl sulfate and urea are variable. Sterilization of rodent brain extracts with high titers of prions requires autoclaving at 134 C at 3 bar for 1 h . Denaturing organic solvents such as phenol or chaotropic reagents such as guanidine isothiocyanate or alkali such as NaOH can also be used for sterilization. Prions are inactivated by 1 N NaOH (>1 h), 4.0 M guanidinium hydrochloride or isocyanate, 2,5 % sodium hypochlorite (2% free chlorine concentration; >1 h), followed by heating in an autoclave at 134 C at 3 bar for >1 h (Anonymus, 2003). It is recommended that dry waste be autoclaved at 134 C at 3 bar for at least 1 hour (depends on the thickness of the material) or incinerated. Large volumes of infectious liquid waste

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Guidelines for Cleaning and Disinfection in Zoological Gardens

containing high titers of prions can be treated with 1 N NaOH (final concentration) before autoclaving at 134 C at 3 bar for 1 hour. Disposable plasticware, which can be discarded as a dry waste, is highly recommended. Because the paraformaldehyde vaporization procedure does not diminish prion titers, biosafety cabinets must be decontaminated with 1 N NaOH, followed by 1 N HCl, and rinsed with water. Although there is no evidence to suggest that aerosol transmission occurs in the natural disease, it is prudent to avoid the generation of aerosols or droplets during the manipulation of tissues or fluids and during the necropsy of experimental animals. It is further strongly recommended that gloves be worn for activities that provide the opportunity for skin contact with infectious tissues and fluids. Formaldehydefixed and paraffin-embedded tissues, especially of the brain, remain infectious. Considerations for Disinfectant Use Disinfectant products, as well as range of applications vary widely. However, careful consideration of manufacturer’s product information in conjunction with a basic understanding of limitations to the disinfection process will go a long way in assuring that disinfection is an effective part of an infection control program. As previously addressed, selecting the proper disinfectant for specific needs is essential. Before purchasing a product, request and review information from the manufacturer. Many manufacturers have product sheets that will go into more detail than the information on the product label and may even include efficacy studies that could impact the decision-making process. Review the material safety data sheet (MSDS) for the product to determine the nature of hazards associated with product use and specific product disposal requirements. Remember that disinfectants are designed to destroy living cells. Therefore, all disinfectants are likely to be hazardous in one way or another to personnel and to client animals if the product has residual properties or is improperly used. Before disinfecting surfaces or devices, it is essential to clean the surface first if at all possible. Organic debris can inactivate many disinfectants. Additionally, feces and other body excretions can shield pathogens from contact with the disinfectant thus hindering disinfection. When using a disinfectant, it is essential that the product be used in accordance with the manufacturer’s instructions. Note that the manufacturer’s efficacy claims are based on its prescribed dilution ratio, method of preparation, method of application, surface contact time and shelf-life. To clarify, contact time is the minimal amount of time that a product must be in contact with the surface to be disinfected in order to achieve the level of disinfection claimed by the manufacturer. Shelf-life is the amount of time that a diluted product is actively effective for use once prepared from a concentrated product. There are a number of organizations in Europe that provide lists of disinfectants for use in animal husbandry, human medicine, and the food industry. Examples are the German Veterinary Medical Society (DVG), and the Germany Society for Hygiene and Microbioloy (DGHM). These lists can be consulted for commercial disinfectant preparations that can be used in certain situations and against different types of pathogens. Under circumstances where it is critical to assure that disinfection practices are efficacious, (i.e. infectious disease outbreaks, immunosuppressed animal housing), it may be beneficial to perform a surface contamination test. There are several techniques described in the

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Guidelines for Cleaning and Disinfection in Zoological Gardens

literature that may be used for surface testing (Tamasi, 1995). The floor may be the surface of choice for such a test because it is likely to receive the most contamination of the environmental surfaces, and this surface often has irregularities that will make disinfection a challenge. In cases of disease outbreaks, disinfection procedures may be dictated by law and close cooperation with veterinary authorities may be necessary. Summary Cleaning and disinfection are a major component of disease prevention and eradication in every zoological exhibition. Despite the development of antimicrobial drugs and effective vaccines, infectious diseases remain a major threat to animals kept in captivity as well as our wildlife population. The proper selection and proper use of disinfectants requires extensive knowledge not only about the effectiveness of specific disinfectant compounds, but also about the infectious agent, the conditions under which the disinfectant will be used and the side effects of each compound, including toxic, caustic and corrosive properties. However, disinfection is only one aspect of an effective infectious disease control program in any animal or human health environment. As described in this chapter proper sanitation consists of multiple components in addition to cleaning and disinfection. Insect and vermin control, and proper food storage, handling, preparation and distribution are equally important for a healthy zoo environment. Furthermore, sanitation should be part of a preventive veterinary medicine program that would further include training of personnel, proper nutrition and feeding, routine surveillance and prophylactic medicine including vaccination, quarantine of new and sick animals, proper holding facilities, correct disposal of waste products and dead animals. This chapter can only provide a brief overview of the guidelines of cleaning and disinfection. Before using a disinfectant, the manufacturer’s label should always be studied. When dealing with a specific disease problem, further information regarding disinfection can be found in the disease specific fact sheets. Additionally, the listed references will provide more detailed information.

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Guidelines for Cleaning and Disinfection in Zoological Gardens

References Anonymus 2003. Beschluss 603 des Ausschusses für Biologische Arbeitsstoffe: Schutzmaßnahmen bei Tätigkeiten mit Transmissibler Spongiformer Enzephalopathie (TSE) assoziierten Agenzien in TSE Laboratorien. BarbBl. 2003-3. Pp. 55 Deutsche Veterinärmedizinische Gesellschaft (DVG). Frankfurter Str. 89, 35392 Giessen, Germany. www.dvg.net. Deutsche Gesellschaft für Hygiene und Mikrobiologie (DGHM). Institut für Hygiene und Mikrobiologie, Univeristät Würzburg, Josef-Schneider-Str. 2, 97080 Würzburg, Germany. www.dghm.org. Block, S. 2000. Definition of terms. In: Block, S. (ed). Disinfection, sterilization and preservation. 5th ed. Lippincott Williams & Wilkins, Philadelphia. Pp. 19 - 28. Boothe, H. W. 1998. Antiseptics and disinfectants. Vet. Clin. North Amer.: Small Anim. Pract. 28: 233 - 248. DeBoer, D. J., K. A. Moriello, and R. Cairns. 1995. Clinical update on feline dermatophytosis: part II. Comp. Cont. Edu. Prac. Vet. 17: 1471 – 1480. Bruins, G., and J. A. Dyer. 1995. Environmental considerations of disinfectants used in agriculture. Rev. Sci. Tech. 14: 81-94. Favero, M., and W. Bond. 2000. Chemical disinfection of medical and surgical materials. In: Block, S. (ed). Disinfection, sterilization and preservation. 5th ed. Lippincott Williams & Wilkins, Philadelphia. Pp. 881 - 917. Fayer, R. 1995. Effect of sodium hypochlorite exposure on infectivity of Cryptosporidium parvum oocysts for neonatal BALB/c mice. Appl. Environ. Microbiol. 61: 844 – 846. Fotheringham, V. J. C. 1995. Disinfection of livestock production premises. Rev. Sci. Tech. 14: 191 – 205. Fowler, M. E. 1978. Sanitation and Disinfection. In: Fowler, M. E. (ed). Zoo and Wild Animal Medicine. W. B. Saundres, Philadelphia. Pp. 21 - 30. Heinsohn, P., R. Jacobs, and B. Concoby. 1995. (eds) Biosafety Reference Manual, 2nd ed. Fairfax: American Industrial Hygiene Association. 101 - 110. Heuschele WP. 1995. Use of disinfectants in zoos and game parks. Rev. Sci. Tech. 14: 447 - 454. Jarroll, E. L. 1999. Sensitivity of protozoa to disinfectants: intestinal protozoa. In: Russell, A. D., W. B. Hugo, G. A. J. Ayliffe (eds). Principles and practice of disinfection, preservation, and sterilization, 3rd ed. Blackwell Scientific Publisher, Oxford. Pp. 251 - 257. Jeffrey, D. J. 1995. Chemicals used as disinfectants: active ingredients and enhancing additives. Rev. Sci. Tech. 14: 57 - 74.

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Guidelines for Cleaning and Disinfection in Zoological Gardens

Lemarie, R. J., and G. Hosgood. 1995. Antiseptics and disinfectants in small animal practice. Comp. Cont. Edu. Prac. Vet. 17: 1339 – 1350. Maillard, J. Y., and A. D. Russell. 1997. Viricidal action and mechanisms of action of biocides. Sci. Prog. 80: 287 – 315. Maris, P. 1995. Modes of action of disinfectants. Rev. Sci. Tech. 14: 47 - 55. McDonnell, G., and A. D. Russell. 1999. Antiseptics and disinfectants: activity action and resistance. Clin. Microbiol. Rev. 12: 147 – 179. Quinn, P., and B. Markey. 1999. Viricidal activity of biocides. activity against veterinary viruses. In: Russell, A. D., W. B. Hugo, G. A. J. Ayliffe (eds). Principles and practice of disinfection, preservation, and sterilization, 3rd ed. Blackwell Scientific Publisher, Oxford. Pp. 187 –196. Quinn, P., and B. Markey. 2000. Disinfection and disease prevention in veterinary medicine. In: Block, S. (ed). Disinfection, sterilization and preservation. 5th ed. Lippincott Williams & Wilkins, Philadelphia. Pp.:1069 - 1103. Ruano, M. 2001. Efficacy comparisons of disinfectants used by the commercial poultry industry. Avian Dis. 45:972-977. Russell, A. D., 1996. Activity of biocides against mycobacteria. J. Appl. Bact. 81: 87S - 101S. Russell, A. D., 1998. Microbial susceptibility and resistance to chemical and physical agents. In: Collier, L., A. Balow, M. Sussman (eds). Topley and Wilson’s microbiology and microbial diseases, 9th ed. Arnold, London. Pp. 149 -184. Russell, A. D., 1999. Antifungal activity of biocides. In: Russell, A. D., W. B. Hugo, G. A. J. Ayliffe (eds). Principles and practice of disinfection, preservation, and sterilization, 3rd ed. Blackwell Scientific Publisher, Oxford. Pp. 149 -167. Russell, A. D., V. S. Yarnych, and A. V. Koulikovskii (eds). 1984. Guidelines on Disinfection in Animal Husbandry for Prevention and Control of Zoonotic Diseases. World Health Organization (Veterinary Public Health Unit), Geneva, Switzerland. Rutala, W. 1996. APIC guidelines for selection and use of disinfectants. Am. J. Inf. Contr. 24: 313 - 342. Saran, A. 1995. Disinfection in the dairy parlour. Re. Sci. Tech. 14: 207 – 224. Sattar, S. A., and S. Springthorpe. 1999. Viricidal activity of biocides: activity against human viruses. In: Russell, A. D., W. B. Hugo, G. A. J. Ayliffe (eds). Principles and practice of disinfection, preservation, and sterilization, 3rd ed. Blackwell Scientific Publisher, Oxford. Pp. 168 –186. Seymore S. Block. 2001. Disinfection, Sterilization, and Preservation. 5th Ed. Lippincott, Williams, & Williams, Philadelphia, Pa.

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Springthorpe, S., and S. A. Sattar. 1990. Chemical disinfection of virus-contaminated surfaces. In: Straub, C. P. (ed). Critical reviews in environmental control. Vol. 20. CRC Press, Boca Raton, Florida. Pp. 169 - 229. Stellmacher, W. 1966. The Testing of Heavy-Duty Disinfectants Against Bacteria. National Veterinary Medicine Testing Institute, Berlin, Pp. 547 - 575. Stone, S.S., and W.R. Hess. 1972. Effects of some disinfectants on African swine fever virus. Appl. Microbiol., 24: 115 - 122. Tamasi, G. 1995. Testing disinfectants for efficacy. Rev. Sci. Tech. 14: 75 - 79. Taylor, D. M. 2000. Inactivation of transmissible degenerative encephalopathy agents: a review. Vet. J. 159: 10 - 17. WHO/CDS/CSR/APH/2000.3, 1999. Report of a WHO consultation: WHO Infection Control Guidelines for Transmissible Spongiform Encephalopathies, Geneva, Switzerland, March 23-26, Annex III. Widmer, A. F. and R. Frei. 1999. Decontamination, disinfection and sterilization. In: Murray, P. R., E. J. Barron, and M. A. Pfaller (eds). Manual of clinical microbiology. ASM Press, Washington, DC. Pp. 138 – 164. Williams, R. B. 1997. Laboratory tests of phenolic disinfectants as oocysticides against the chicken coccidium Eimeria tenella. Vet. Rec. 141: 447 – 448.

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Table 1: Effectiveness of classes of disinfectants against various pathogens Class of Disinfectant Alcohols Aldehydes Alkalis Biguanides

Halogens

Organic and Anorganic Acids

Peroxygen Compounds

Phenols

Chlorine Iodine Compounds Compounds

Infectious Agent

obligat intracellular gram-positive Bacteria

gram-negative bacilli endospores

Fungi

including spores

enveloped Viruses non-enveloped

Protozoa including oozysts

Prions

highly effective

effective

limited effectiveness

not effective

This table provides only a general overview of the effectiveness of various classes of disinfectants. The effectiveness of a specific compound in each of these classes may vary. For more detailed information please review the information provided in the text. Always consult the manufacturers label for correct use and concentration of each compound. The table is based on the following sources: Boothe, 1998; Bruins and Dyer, 1995; Jaroll, 1999; Jeffrey, 1995, Lemarie and Hosgood, 1995; Maillard and Russell, 1997; McDonnell and Russell, 1999; Quinn and Markey, 1999 and 2000; Russell, 1996, 1998 and 1999; Sattar and Springthorpe, 1999; Sringthorpe and Sattar, 1990; Taylor, 2000; Widmer and Frei, 1999; Williams, 1997.

QAC

Transmissible Diseases Handbook

IX. BLUETONGUE IN NON-DOMESTIC RUMINANTS: EXPERIENCES GAINED IN EAZA ZOOS DURING THE 2007 & 2008 BTV8 AND BTV1 EPIZOOTICS. Stephanie Sanderson Chester Zoo Introduction Until the introduction of Bluetongue virus serotype 8 (BTV8) in 2006, Bluetongue had not been a significant disease problem in Europe. The ensuing epizootic and subsequent incursion of BTV1 have caused widespread mortality and morbidity in livestock and been a major focus for international disease control in the EU. This chapter provides information pertaining to these European epizootics – focusing primarily on the clinical species susceptibilities of non-domestic ungulates and their response to vaccination. The data presented has been derived from the literature and two EAZWV endorsed Bluetongue web surveys of EAZA zoos covering the 2007 and 2008 disease seasons. General disease information on Bluetongue can be found in disease fact sheet 7. BTV8 in Europe: an atypical virus strain Bluetongue is an insect borne disease caused by an orbivirus and affecting mainly domestic sheep breeds and occasionally cattle. It is transmitted by midges of a few select Culicoides species and its global distribution is largely defined by suitable climatological factors for these species. Bluetongue viruses have been found on all continents excepting Antarctica but the disease is generally only endemic in the tropics and subtropics (34° S to 53°N OIE terrestrial animal health code). (Hately 2009, MacLachlan 2009). Until 2006, BTV was not considered to be a significant threat to Central and Northern European livestock as the resident Palaearctic midge species were not competent vectors. However the unexpected occurrence of an atypical BTV virus in Maastricht in 2006 has challenged established thinking on the behaviour of this disease. Key differences from other BTV serotypes and strains include:  Ability to use Palaearctic midges as vectors (C.obsoletus and C.pulicaris)  Significant morbidity and mortality in cattle as well as sheep  Fairly frequent vertical transmission in pregnant ruminants – a rare event in other BTV strains. Other features of BTV epidemiology in temperate climates include a marked seasonality with clinical cases occurring almost exclusively between July and December and with recrudescence occurring the following summer. It is still unclear how the virus is maintained from year to year. The source of this BTV8 strain is still unknown. Phylogenetic studies indicate the likely origin to be sub-Saharan Africa and it has been postulated that it could have been introduced either by an infected animal originating from Africa, imported infected vectors or as an illegally imported vaccine strain. (EFSA 2007)

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Bluetongue in non-domestic ruminants

Other BTV subtypes in Europe A number of BT serotypes have been present in Southern Europe from 1900’s onwards. The spread of different BT serotypes since 1998 is shown in figure 1. The distribution of serotypes in May 09 is shown in figure 2. BTV1 has been seen on numerous occasions in Southern Europe however it was not until 2008 that it spread northwards and is also now capable of spread via Palaearctic midges. Cases of BTV6 and BTV11 were also reported in the Netherlands and Belgium in 2008. These are postulated to have been introduced via illegal imports of modified live vaccine containing these serotypes from Southern Africa. At the time of writing it appears that these serotypes have not become established unlike BTV8 and BTV1. Increased surveillance has also uncovered a previously unknown orbivirus in Toggenberg goats which may indeed be a new serotype of BTV. Its clinical significance is currently unknown. (Chaignat et al. 2009)

IX.

Bluetongue in non-domestic ruminants

Fig 1. Spread of Bluetongue throughout Europe since 1998. Picture, Institute of Animal Health, Pirbright

Fig 2.

IX.

Bluetongue in non-domestic ruminants

Species susceptibility to clinical infection It is generally accepted that all ruminant species and some camelids are likely to be capable of supporting BTV infection. Clinical expression of the disease is however highly variable between viral serotypes, species, breeds and indeed environmental conditions and individual health status. Animals indigenous to endemic areas appear to be clinically resistant. (Verwoerd & Erasmus, 2004). The pathology and pathogenesis of the disease is reviewed in detail by MacLachlan et al. and the clinical picture seen in domestic species is well documented. (MacLachlan et al. 2009). The classical clinical signs of fever, nasal discharge, dypsnea, cyanosis of the tongue, oral lesions and ulcers, oedema of the head and neck, lameness and hyperaemia of the coronary band are due to virus mediated vascular injury. These signs are most frequently seen in sheep where mortality rates can be up to 30% or higher. Cattle rarely show clinical disease (with the exception of the European BTV8 strain).(MacLachlan et al. 2009) Little is published about clinical susceptibility of non-domestic species. In North America, white-tailed deer (Odocoileus virginianus), prong-horn antelope (Antilocapra americana) and desert bighorn sheep (Ovis canadensis) are known to develop severe disease similar to that described in domestic sheep. (Verwoerd & Erasmus, 2004). Abortion and death has also been seen in dogs injected with BT contaminated vaccine and in a European lynx fed infected meat (EFSA 2007; Jauniaux et al. 2008, MacLachlan 2009). Natural, asymptomatic infection of African carnivores has also been reported. With the incursion of BTV8 into Europe, European Zoos were in a unique position to contribute to knowledge on species susceptibility to disease as they hold a naïve individuals representing a wide taxonomic and geographic spectrum. A survey of all 313 EAZA zoos was undertaken in January 2008 to collate data on clinical disease seen during the 2007 BTV season. 49 zoos had confirmed BTV8 cases within 20km and could be classified as at risk of infection. These 49 zoos held over 1000 susceptible individuals of 53 different species and 7 ruminant families indigenous to Europe, North and South American, Africa and Asia. Clinical disease was seen in 62 individuals (6% of the at risk population), spread between 13 zoos (27% of at risk collections). (Sanderson et al. 2008). Mortality and morbidity rates and the clinical picture seen in each affected species is summarised inTable 1 (Sanderson et al. 2008). Bovidae are the most susceptible family of ruminants to clinical disease, with four species showing morbidity rates of greater than 20% and mortality rates of greater than 10%. The average case fatality rate for the affected Bovidae species was 69%. All the affected ruminant species in this study were indigenous to Europe, Asia or South America. Clinical signs in these species and are consistent with those recorded for BTV8 infection domestic livestock. (Elbers et al., 2007) and with those reported in yak (Mauroy et al. 2008;). It is noteworthy that despite over 200 African ruminants of 20 species being held by zoos in at risk areas, none of these were reported to have shown clinical signs of infection. This is consistent with observations in Africa that indigenous antelope do not develop clinical disease (Verwoerd & Erasmus, 2004). Data was also gathered by European Zoos on species susceptibility to BTV1. There is yet insufficient data on BTV1 to draw any firm conclusions however experiences so far suggest a similar clinical picture and species susceptibility to BTV8.

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Bluetongue in non-domestic ruminants

Table 1: Morbidity, mortality, case fatality and clinical signs reported in ruminant species by zoos situated within 20km of confirmed BTV8 outbreaks during the period August 2006 - December 2007 (Sanderson et. al. 2008) AFFECTED SPECIES

Abs. No.

Clinically affected Abs. No.

Morbidity * Rate %

Laboratory confirmation Abs. No.

Abs. No.

Mortality ** Rate %

Case Fatality *** Rate (%sick that died)

519

55

10.60

25

38

7.32

69.09

30

10

33.33

3

5

16.67

50.00

European wisent (Bison bonasus)

20

8

40.00

4

4

20.00

50.00

Yak (Bos grunniens)

35

6

17.14

6

6

17.14

100.00

17

2

11.76

2

2

11.76

100.00

Sudden death

101

6

5.94

5

2

1.98

33.33

Lethargy, fever, swelling head and neck, mouth ulcers, drooling, difficulty eating, conjunctivitis, respiratory difficulty, lameness, inflammation coronary band, sudden death.

208

2

0.96

2

2

0.96

100.00

Lameness

34

2

5.88

2

2

5.88

100.00

Nasal discharge, sudden death.

Siberian ibex (Capra sibirica)

4

1

25.00

0

0

0.00

0.00

Swelling of head and neck.

Muskox (Ovibos moschatus)

5

1

20.00

1

0

0.00

0.00

Lethargy, fever, conjunctivitis, abortion.

AFFECTED CERVIDAE

83

5

6.02

5

2

2.41

40.00

Fallow deer (Dama dama)

43

2

4.65

2

2

4.65

100.00

AFFECTED CAMELIDAE

40

2

5.00

2

2

5.00

100.00

Bactrian camel (Camelus bactrianus)

8

1

12.50

1

1

12.50

100.00

AFFECTED BOVIDAE American bison (Bison bison)

Blackbuck (Antilope cervicapra) Sheep/mouflon (Ovis aries) Goat (Capra hircus) Alpine ibex/Tur (Capra ibex)

At risk individuals (BTV8 within 20km)

Deaths

Clinical Signs Reported in Affected Animals.

Lethargy, fever, mouth ulcers, drooling, difficulty eating, conjunctivitis, corneal oedema, lameness, inflammation coronary band, sudden death. Lethargy, fever, mouth ulcers, drooling, difficulty eating, conjunctivitis, corneal oedema, respiratory difficulty, lameness, inflammation coronary band, sudden death. Nasal discharge, conjunctivitis, corneal oedema, drooling, difficulty eating, respiratory difficulty, lameness, inflammation of coronary band, sudden death.

Mouth ulcers, difficulty eating, drooling, lameness, sudden death

Sudden death

Alpaca 32 1 3.13 1 1 3.13 100.00 Sudden death (Lama pacos) *Morbidity rate= number clinically affected / number at risk; **Mortality rate = number that die/number at risk; ***Case fatality = number that die / number clinically affected. Note: Morbidity and mortality rates for the 2006 BTV8 epidemic in domestic livestock were 20% and 5% for domestic sheep and 7% and 3% in domestic cattle (Elbers et al., 2006)

IX.

Bluetongue in non-domestic ruminants

Control Strategies for BTV: legislative framework Bluetongue is a disease of global importance. Its ability to cause death and debilitating disease across international borders has led to its inclusion in the World Organization for Animal Health (Office International des Epizooties) Terrestrial Animal Health Code which in turn has implications for trade. In EU Member States, control and eradication provisions are laid out in Council Directive 2000/75/EC. Measures include vector control, restriction to movements of live ruminants from affected areas to non-infected regions where the vector is present and the use of vaccines. Of these, vaccination is the mainstay of control in areas where BTV has become established. The OIE guidelines on movement controls, diagnostic methods and vaccine production can be found at: http://oiebtnet.izs.it/btlabnet/. Further information on EU control measures are laid out http://ec.europa.eu/food/animal/diseases/controlmeasures/bluetongue_en.htm

in

In addition, individual European Countries have their own legislation and detailed disease control plans (e.g. http://www.defra.gov.uk/animalh/diseases/notifiable/bluetongue/about/index.htm ). Vaccination Mass vaccination has been identified by the European Commission as the most efficient veterinary measure in combating bluetongue. Mass emergency vaccination campaigns can be used to achieve the following objectives (European Commission, 2008/655/EC, Savini et al. 2007): 1. 2. 3. 4.

prevention of clinical disease limiting regional spread of BT allowing regional/country eradication safe movement of animals between affected and free zones.

A variety of vaccines have been developed of three different types: modified live vaccines (MLV), inactivated vaccines (either whole killed virus preparations or virus like particle produced from recombinant bacluovirus) and recombinant vaccinia, capripoxvirus or canarypox virus vectored vaccines. (EFSA 2007). Only vaccine types currently approved under EC approved national disease programmes (MLV and killed whole virus preparations) will be discussed further here. MLV have been used for over 40 years in endemic bluetongue areas (Verwoerd D. W. and Erasmus, B. J. 2004).They are quick to produce (8-10wks), highly immunogenic and can confer long lasting protection after a single dose. Using live virus has significant disadvantages as there is potential for under attenuation causing symptomatic disease, milk drop and foetal pathology, and for infection of the vector population leading to local spread and potential for reassortment with field strains leading to new serotypes. For these reasons, inactivated vaccines are preferred even though they take longer (6-8mths) to develop, are more costly and require regular boosters in order to maintain efficacy. (EFSA 2007; OIE 2008) There is little or no cross protection between different serotypes of Bluetongue, hence vaccines are produced specifically in response to circulating BTV serotypes and strains. All of the vaccines currently available in the EU for the control of BTV1 and BTV8 are inactivated vaccines using saponin and aluminium hydroxide as adjuvants. (Table 2)

IX.

Bluetongue in non-domestic ruminants

Table 2: BTV 1 & 8 Vaccines licensed for use in 2008. Manufacturer

BTV8 Vaccine Trade Names

BTV1 Vaccine Trade Names

Intervet

Bovilis BTV8

CZ Veterinaria

Bluevac 8

Bluevac 1

Fort Dodge

Zulvac8 Bovis Zulvac8 Ovis

Zulvac1 Bovis Zulvac1 Ovis

Merial

BTVPUR AlSap 8

Virbac

SYVAZUL 1

Safety Trial and field experience has found these vaccines to be safe in domestic species (EMEA 2009, Gethman et al. 2009, Eschbaumer et al 2009). An overview of field experience following administration of over 60million doses of BTV8 vaccine in 12 countries was undertaken by the European Medicines Agency (EMEA 2009). Mass vaccination campaigns often necessitate deviations from normal procedure. Large groups of animals are brought together, less attention is paid to their individual health status, needle hygiene is less good and government instructions may deviate from those of the manufacturers (eg minimum age of vaccination, target species, duration of immunity). In addition, compensation schemes in some countries may lead to over reporting of certain adverse reactions. Despite these factors, adverse reactions were seen in less that 1 in 10,000 animal. Those recorded are typical for other inactivated vaccines and include local reactions and non-severe general reactions such as pyrexia (fever) and lethargy. A survey of all 313 EAZA zoos was undertaken in February 2009 to collate data on vaccination in non-domestic species. Over 2000 individuals of 57 species in 47 institutions in 9 European countries were vaccinated for BTV8 using 5 of the products on the market during 2008. Adverse reactions occurred at a rate of 0.5% with half of these being local reactions and 40% being abortions. The slightly higher rate could well be due to the relatively small sample size and also because the species studied are not used to handling and are likely to have been more stressed than their domesticated counterparts. Nonetheless, the abortion rate is still well below that considered acceptable for vaccines. (EAMEA 2009) Efficacy Vaccine efficacy can be assed both by response to virus challenge (both clinical and levels of viraemia) and serological response induced by immunisation. (Savini et al. 2008). Whilst experimental virus challenge under laboratory conditions provides the most accurate measures of efficacy, are the mainstay of vaccine testing and are required for vaccine licensing (European Parliament 2001), field experiences are also provide a useful data. The licensed vaccines have been shown to be efficacious in domestic animals (Eschbauer et al. 2009, Gethmann et al. 2009 ) The 2008 European Zoo Survey (Sanderson et al. 2009) found that of the 37 bovidae (cattle, sheep, goat and antelope sp) and girrafidae tested post vaccination, 100% seroconverted post vaccination as did 87% of the 40 South American camelids tested. Of the 9 cervidae (deer sp) represented only 50% seroconverted. No vaccinated animals succumbed to clinical disease post vaccination, despite virus circulating in the area. These data suggest that the inactivated BTV8 vaccines are efficacious in bovidae, girrafidae and to a lesser extent camelids. The sample size in the cervidae is too small to draw any firm conclusions and further work needs doing to evaluate efficacy in these species.

IX.

Bluetongue in non-domestic ruminants

Conclusions BTV poses a significant risk of mortality and morbidity in naïve non-domestic ruminants. The clinical picture is similar to that seen in domestic livestock with species indigenous to temperate areas of Europe, Asia and the Americas being most severely affected. Species indigenous to Africa, the putative source of BTV8, were clinically unaffected. This suggests that there is a genetic resistance to particular BTV serotypes. Inactivated BTV8 and BTV1 vaccines have been used in many European zoos both on a voluntary basis as part of national control measures. Adverse reactions were rare and in line with those seen in the domestic species for which they are licensed. Vaccination produced a reliable immune response and no animals showed clinical evidence of infection post immunisation despite the presence of circulating virus in the region. These vaccines would appear to be safe in non-domestic ruminants and efficacious in the bovidae and camelidae. Further work is required to evaluate their efficacy in cervidae. Further work is underway within the zoo community to expand our knowledge on vaccine efficacy and duration of immunity in non-domestic species. Data is also being collected on species susceptibilities to other BTV serotypes as they appear.

IX.

Bluetongue in non-domestic ruminants

References Chaignat, V., Worwa, G., Scherrer, N., Hilbe, M., Ehrensperger, F., Batten, C., Cortyen, M., Hofmann, M., Thuer, B., (2009). Toggenburg Orbivirus, a new bluetongue virus: Initial detection, first observations in field and experimental infection of goats and sheep. Veterinary Microbiology 138, 11-19. Elbers, A.R.W , K. Mintiens , C. Staubach , G. Gerbier , E. Meroc , H.M. Ekker , F.J. Conraths , A.N. van der Spek , A. Backx (2007) Epidemiological analysis of the 2006 bluetongue virus serotype 8 epidemic in north-western Europe; Nature and severity of disease in sheep and cattle. In EFSA scientific report: Epidemiological analysis of the 2006 bluetongue virus serotype 8 epidemic in north-western Europe http://www.efsa.europa.eu/EFSA/1178620925100/efsa_locale1178620753812_Bluetongue.htm EMEA CVMP.(2009)An overview of field safety data from the EU for bluetongue virus vaccines serotype 8 emerging from the 2008 national vaccination campaigns. London: EMEA/CVMP/652019/2008. Eschbaumer, M., Hoffmann, B., Konig, P., Teifke, J.P., Gethmann, J.M., Conraths, F.J., Probst, C., Mettenleiter, T.C., Beer, M., (2009). Efficacy of three inactivated vaccines against bluetongue virus serotype 8 in sheep. Vaccine 27, 4169-4175. European Commission, 2008/655/EC: Commission Decision of 24 July 2008 approving the emergency vaccination plans against bluetongue of certain Member States and fixing the level of the Community's financial contribution for 2007 and 2008, Off J Eur Union L214/66 (August). The European Parliament and the Council of the European Union (2001). Directive 2001/82/EC of the European Parliament and of the Council of 6 November 2001 on the Community code relating to veterinary medicinal products. Off J Eur Commun;L311/1(November). European Food Standards Agency (EFSA) (2007) Report of the Scientific Panel on Animal Health an Welfare (AHAW) on the EFSA Selfmandate on bluetongue origin and occurrence. EFSA Journal 479,1-29; 480, 1-20 Gethmann, J., Huttner, K., Heyne, H., Probst, C., Ziller, M., Beer, M., Hoffmann, B., Mettenleiter, T.C., Conraths, F.J., (2009). Comparative safety study of three inactivated BTV8 vaccines in sheep and cattle under field conditions. Vaccine 27, 4118-4126. Hateley, G., 2009. Bluetongue in northern Europe: the story so far. In Practice 31, 202-+. Jauniaux, T.P., De Clercq, K.E., Cassart, D.E., Kennedy, S., Vandenbussche, F.E., Vandemeulebroucke, E.L., Vanbinst, T.M., Verheyden, B.I., Goris, N.E., Coignoul, F.L., (2008). Bluetongue in Eurasian lynx. Emerging Infectious Diseases 14, 1496-1498. Mauroy, A., Guyot, H., De Clercq, K., Cassart, D., Thiry, E., Saegerman, C., (2008). Bluetongue in captive yaks. Emerging Infectious Diseases 14, 675-676. Maclachlan, N.J., Drew, C.P., Darpel, K.E., Worwa, G., (2009). The Pathology and Pathogenesis of Bluetongue. Journal of Comparative Pathology 141, 1-16. Office International des Epizooties (OIE). Bluetongue. In: Manual of diagnostic tests and vaccines for terrestrial animals. Paris: OIE; 2008. Sanderson, S., Garn, K., Kaandorp, J. (2008) Species Susceptibility to Bluetongue in European Zoos during the Bluetongue Virus Subtype 8 (BTV 8) Epizootic Aug 2006-Dec 2007. Proc. European Association of Zoo and Wildlife Veterinarians.

IX.

Bluetongue in non-domestic ruminants

Sanderson, S., Edwards, S.J., Setzkorn, C and Baylis M. (2009) Survey of Species Susceptibility to Bleutongue virus and Bluetongue Vaccine Usage in European Zoos During 2008. Proc. International Conference on Diseases of Zoo and Wild Animals. 1. 2. Savini, G., MacLaclalan, N.J., Sanchez-Vinaino, J.M., Zientara, S., (2008). Vaccines against bluetongue in Europe. Comparative Immunology Microbiology and Infectious Diseases 31, 101-120. Verwoerd D. W. and Erasmus, B. J. (2004) in: Infectious Diseases of Livestock. Eds Coetzer J.A.W & Tustin R.C. Oxford University Press pp 1201-1220

Transmissible Diseases Handbook

X.

TUBERCULOSIS IN ZOO SPECIES: DIAGNOSTIC UPDATE AND MANAGEMENT ISSUES.

EAZWV Tuberculosis Working Group

1. Introduction One person out of three is currently infected by tuberculosis over the world. More than 2 millions of people die from it every year. Recent raising of Multi Drug Resistance or even Extreme Drug resistance, co-infection with HIV and incidence of non tuberculous mycobacteria (NTM) makes the battle against tuberculosis more difficult: the disease is now placed in the top 3 list by WHO, together with AIDS and Malaria. Most of the mycobacteria from the ‘tuberculosis complex’ have the ability to infect wild species, in whom the pathogenesis, receptivity and immune responses vary widely. Genetic key factors (e.g. Interferon Gamma receptor or vitamin D genotypes) are obviously acting towards these differences (SCHLUGER, 2005). Course of disease and occurence of latent infection vs. open disease are variable among species, from extremely sensitive old world monkeys to apparently resistant equids. Table 1: Tuberculosis complex mycobacteria and their reported hosts. Mycobacteria of the Major historical known Reported Wild and Zoo Host Tuberculosis complex host or burden Human, Non Human Elephant, NHP, Beisa Oryx, M.tuberculosis Primates (NHP) Addax, Goats, Birds, Lowland Tapir, Giraffes, Springboks, mongoose, Rhinoceros Cattle (+buffalo, Bison) All ruminants, Badgers, M.bovis Possums, Meerkats, Big Cats, Canids, Rodents, NHP, Wild boars Elephants, Camelids, Rhnoceros, Onager, Horse, Birds Human Cattle, Swine, NHP M.africanum Vole, Camelids NW Monkeys, Big Cats M.microti Pinnipeds Camel, Tapir, Big Cats M.pinnipedii Goat, Sheep, Swine Swine, Cattle Wild Boars, Red M.caprae & WT deer, Camel, Bison Human ? M.canetii Hyraxes Meerkats « Dassie bacillus »

X.

Tuberculosis in Zoo Species

2. European regulation National programs Domestic species of cattle in zoos and safari parks should normally be subjected to routine tuberculin testing as often as the indicated testing interval for the area in which the zoo is located. Zoo species are generally exempted from statutory TB testing and, in any case, there is no recognized, approved screening test for TB in species other than bovines and deer. Several texts are defining sanitary policy for tuberculosis within country members. The EU policy mainly focuses on the eradication of bovine tuberculosis and is based on two fundamental principles: 1/ The Member States are primarily responsible for the eradication of bovine tuberculosis and may receive community financial support for the eradication program 2/ Eradication of bovine tuberculosis in the EU must be the final target and the Member States must consider eradication as the defined aim. Hence, most of the EU regulations apply only to M.bovis- sometimes M.tuberculosis screening. Table 2: European legislation concerning animal tuberculosis. LEGISLATION RELATED TO THE ERADICATION OF BOVINE TUBERCULOSIS

Council Directive 64/432/EEC of 26 June 1964 on animal health problems affecting intra-Community trade in bovine animals and swine Council Directive 77/391/EEC of 17 May 1977 introducing Community measures for the eradication of brucellosis, tuberculosis and leucosis in cattle Council Directive 78/52/EEC of 13 December 1977 establishing the Community criteria for national plans for the accelerated eradication of brucellosis, tuberculosis and enzootic leukosis in cattle Commission Decision 2003/467/EC of 23 June 2003 establishing the official tuberculosis, brucellosis, and enzootic-leukosis-free status of certain Member States as regards bovine herds Commission Decision 2003/849/EC of 28 November 2003 approving the programmes for the eradication and monitoring of animal diseases and for the prevention of zoonoses presented by the Member States for the year 2004 Council Decision 90/424/EEC of 26 June 1990 on expenditure in the veterinary field Council Decision 90/638/EEC of 27 November 1990 laying down Community criteria for the eradication and monitoring of certain animal diseases Based on the Commission Decision 1999/467/EC of 15 July 1999 seven states of the European Union were classified as free of bovine tuberculosis: Denmark, Germany, Luxembourg, the Netherlands, Austria, Finland, and Sweden. In 2004 Belgium, the Czech Republic, and France were added to this group and on 4th March 2005 an eleventh country, Slovakia was added. Switzerland also has the status “free of bovine tuberculosis”.

X.

Tuberculosis in Zoo Species

BALAI application within Europe The BALAI directive 92/65 requires that all ruminants traded between institutions must come from officially free herds, as it’s mentioned in TRACES health certificates. Moreover, to be BALAI approved, an institution must be free of “bovine tuberculosis”, as listed into the Annex A of the directive, for at least three years, or “tuberculosis” (this term includes all mycobacteria of the TB complex) for primates, felidae and ruminants, if the member state has a control monitoring progam. During a transfer between two approved bodies, Tb testing is not required by the BALAI directive, but many member states (UK, Sweden,…) added TB test as an additional requirement when importing ungulate, whatever institutions it is coming from. Given these requirements, a lot of zoo mammals are still being exchanged in national and international transfers without any serious individual testing.

3. Quick overview of current diagnostic methods Standard screening tools like skin testing and sputum smears have limited application in wildlife species, especially when prevalence is low. Inversely, investigations of cell-mediated immunity through an vitro assay of gamma interferon have numerous advantages (good sensitivity), as long as technical limits are known and can be improved. Furthermore, new tools based on the investigation of humoral immunity seem very promising for the detection of antibody directed against certain immunogenic mycobacterial antigens in a wide range of species. All these methods are currently evaluated in field studies, despite difficulties to ensure rigorous validation. Thus, diagnostic methods are hardly homogenic and never validated for any zoo species. Bearing in mind that validation will not easily be possible, zoo actors must be aware of actual tests available. Limits of unspecific diagnosis Clinical signs of TB are rarely seen prior to death in zoo animals (LYASCHENKO & AL, 2006, MONTALI & AL, 2001). If cough and dyspnea are noticed, this is always in a very late and irreversible stage of the pulmonary form of disease. The most frequent sign noticed among all mammals is chronic weight loss. Imagery techniques can help, especially laparoscopy followed by biopsy of suspect granulomas or tissue. A CT scan is useful to detect lesions in non-palpable lymph nodes. On the other hand, though X-rays are a regular step in human diagnostics, it is not practically useful in zoo veterinary medicine: the size of animals may prevent X-R use, references images are often missing, skills required to interpret pictures are often out of “average zoo vet” range and, on top of that, a lot of species do not show any calcified lesions. Limits of direct examination Culture stands as the “gold standard” method, but requires at least 2 to 6 weeks delay before a test result can be obtained, even with recent fast techniques (VARGAS & al, 2005) which are still not validated in veterinary medicine. Hence, faster testing still relies on microscopic examination and mycobacterium DNA amplification methods.

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Tuberculosis in Zoo Species

The threshold number of bacilli needed to obtain a positive microscopic exam (Ziehl Neelsen and other stains) is around 104 bacilli / ml, which is a rather important and infectious value. In a culture, it is possible to detect bacilli loads superior to 101-102 / ml. As a comparison, less than 10 colony forming units (CFU) of M.tuberculosis per ml is enough to infect a cynomolgus macaque (LIN & al, 2006). Many biological samples contain very few bacilli (highly calcified lesions, intermittent shedding in biological fluids like bronchial secretions or milk). As an example, 15 to 20% of active pulmonary forms are not even confirmed by any culture in human being (FRIEDEN & al, 2003). Therefore, only positive culture findings provide evidence of disease, whereas negative culture results may not rule out infection in exposed/suspected animals. Detection of DNA material in biologic samples have been used already in various zoo species: elephant trunk wash (MIKOTA & AL), broncho-alveolar lavage (FLYN & al, 2003), gastric lavage,.. Amplification methods (PCR) are now well developed and help specify the mycobacteria with the use of selected probes. However, some of these biological samples are likely to host many other bacteria that are impairing PCR efficacy. Within the last years, very sensitive and specific PCR became available, but it still can’t be used as a broad screening tool as described in table 3. Table 3: Example of PCR sensitivity in human sputum with 5% prevalence. From (VEZIRIS, pers. com.). With those given sensitivity and specificity, a positive PCR result is meaning an even probability (49%)of TB infected or free animal, mainly because of the low prevalence. The lower the prevalence is, the lower the PPV turns out. Disease No Disease PCR Positive predictive value (Active TB) (Latent or no TB) Se=72% PPV=3.6/(3.6+3.8) Sp= 96% =49% 5 95 Prevalence 5% PCR + 3.6 3.8 Negative predictive value NPV=91.2/(91.2+1.4) PCR 1.4 91.2 =98% When there is enough DNA (rich sample or culture), molecular typing should be performed. Spoligotyping or other methods (VNTR,..) allow to identify strains of mycobacteria and these techniques can be a very useful tool to track the epidemiological circuit of the TB. In recent years, several tuberculosis strains were spoligotyped and their fingerprint can be compared to other circulating strains. Whenever a tuberculosis mycobacterium is discovered, this kind of typing should always be performed, or at least samples must be kept frozen (-25°C or 80°C) until further typing is available. Cell mediated immunity (CMI) exploration Memory of T cells regarding previous contact with Mycobacterium can be assessed by presenting selected antigen(s) to them, either in vivo (skin test or “Mantoux” test) or in vitro (Lymphoblastic Transformation Test –LTT- or Interferon Gamma tests). Skin testing is far from being validated for zoo species, as there are great variations between tegument and dermal structure within species. The test relies on local inflammatory cell recruitment, which can be low or absent for many reasons: tegument cellular organization, immunosuppressive status, superficial temperature, … Skin test sensibility is often poor: e.g 70 to 90% in human, 50 to 90% in zoo hooftstock (COUSINS AND FLORISSON, 2005). Specificity depends on antigen (tuberculin) injected intradermally but could also be low because of species-specific features (e.g. orangutans) or co-infection by other non tuberculous mycobacteria (e.g. avium complex), leading to false positive results.

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Tuberculosis in Zoo Species

Materials and methods of application of the skin test vary between countries and vets: standardized quality of tuberculin is still missing within EEC, reading methods also vary between vets with a lot of “distant” appreciations of TST local reaction. Whenever possible, a close exam, palpation and caliper measurement of tegument is strongly recommended in order to avoid false negative results. In vitro tests of the cellular immunity rely on (re)stimulation of T lymphocytes memory. Thus, first compulsory step is to keep cells alive until they reach the lab. Blood sample should reach lab for stimulation no later then 8-10h after collection and should be kept at ambient temperature until there. Particular care must be paid to homogenic and full mixing of blood and anticoagluant (heparine) at collection. Sometimes part of experimental study, LTT is not a current option to a zoo vet because of its limited reproducibility and the use of radio-elements. Gamma interferon (IFNg) tests stand as a good option as they are already available and marketed for cattle, non human primates, deer and humans. These tests are performed in two steps, for a total run of 48h minimum. The first step is to incubate T cells with selected mycobacterial antigen (usually bovine and avium ppd), leading them to produce IFNg when they were previously in contact with the same antigen.. The second step is to reveal the amount of IFNg produced through an ELISA or an ELISPOT (more sensible). This step can be delayed as soon as stimulation wells have been frozen. Commercial kits are available, designed for cattle (BOVIGAM®), primates (PRIMAGAM®), deer (CERVIGAM®, but production is discontinued at this time) and human (QUANTIFERON GOLD®). Recent studies show that these tests can be used in some exotic species : for example a cattle-designed test will detect INFg of a large range of exotic Bovidae, but also some animals standing outside of this family (e.g. Girafidae). On the other hand, other artiodacyl species just don’t do well with the usual positive control antigens, or are not detected by ELISA (RIQUELME, 2009). This also counts for the primate-referred test: although a list of “validated” species is provided on its leaflet, field studies showed contrasted results (LÉCU, 2008, RIQUELME, 2009). In order to overcome the problem of specificity in ELISA detection of INFg, in-house modified tests can be created. One solution is the detection of mRNA coding for INFg, which seems to have broad nucleotid sequences, shared by a lot of different mammal species. This can only be achieved by experienced laboratories. Another solution is to design a specific interferon antibody, as is currently being developed in elephants and rhinoceros. Whatever kind of cellular exploration is chosen, the duration of the cell-mediated immunity is rather unknown. All studies concerning longest periods of time (VERVENNE & AL, 2004) and work on BCG vaccination protection length suggest that stimulation tends to go back to a baseline level, which remains unknown beyond a year. Humoral immunity exploration Serodiagniosis of tuberculosis suffered from “bad reputation” because early trials were assessing antibody against broad mycobacterial antigens, showing a very low specificity in these tests. During the course of an infection, Th1 activity (CMI pathway) is thought to be initially greater than Th2 (humoral route) in order to control and confine infection. An inversion of this Th1/Th2 balance control is often associated with a relapse or with active disease (DOHERTY & ROOK, 2006). Thus, seeking for antibodies maybe of little help to screen for latent infected animals, but is becoming more relevant to detect and monitor more active, “sick” and shedding individuals. It has been clearly noticed also in human (ABEBE & al, 2007) that some antibody rise occurs during the shift from latent to active disease, so that a prognostic value may be added to certain serological results

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Tuberculosis in Zoo Species

Some mycobacterial antigens elicit humoral response in a wide range of species. Most relevant antigens have been selected to design ELISA and Rapid Lateral Flow Technology test, with good results in a lot of various species (elephants, primates, tapirs, camels, deer,…). However, the panels of immunostimulant antigen will remain directly linked to the type of mycobacterium involved, host species, timeframe of disease and also individual condition. Some commercial tests validated for Asian and African elephants (ElephantTB STATPAK®, DPP® VetTB) or primates (PrimaTB STATPAK®) also look promising in non target species (GREENWALD ET AL. 2009; LYASHCHENKO ET AL. 2008). Sensibility (Se) and specificity (Sp) of these serological assays are encouraging. However, Se and Sp values are issued from descriptive studies performed on either very sensitive species (i.e prone to start disease soon after infection) or under very clear enzootic conditions, maybe jeopardizing the real Se and Sp. Moreover, serological profiles of animals in long-term quiescent infection status are still remaining poorly known. Repeated testing seems to be the only way to adapt with these actual limitations, as titers is thought to arise when relapse is about to happen. As zoonotic risk increases a lot within this shifting time, detection of early serological change may be relevant to trigger a deeper screening (direct exam) and preventive measures towards staff and public.

4. Current recommendations Within some species, current captive population shows more than 10% of infected herds. To draw a parallel, for a country to keep the TB-free status regarding bovine tuberculosis, the OIE requires that the percentage of herds confirmed infected with M. bovis has not exceeded 0.1% per year for 3 consecutive years. All the zoo community should make an effort, even if available tools are still – and will likely remain - unvalidated. Crossing the three exploration paths (direct, CMI and serology) is a good way to partially get rid of some of their respective limitations. The aim of detecting TB in zoo species has two levels: the first one is to protect staff (keepers, vet) and public from zoonotic contamination. Few actual tools (serology, PCR) are aimed at this predictive purpose, focusing on the active (=excretion) phase of the disease. The second level is the census of latently infected individuals and their monitoring. For this purpose, CMI testing can then help a lot, as long as their sensibility / specificity limits are taken into account. Table 4: Testing categories available. A positive test from one of these boxes should always initiate a test from another box.

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Tuberculosis in Zoo Species

Any repeated positive outcome in the immunological investigationary tests should trigger exploration of the direct exam category, in order to determine the shedding status of the animal. If direct exam is positive, measures should be taken to avoid contamination of staff, surrounding animals and premises. “Euthanasia or treatment” is a choice that should be considered first with veterinary officials, occupational physician, all related actors and then according to global captive population status and dynamics (EEP, TAG, …). Depending on TB extension analysis, treatment may be an alternative to euthanasia, to prevent infected animals from shedding mycobacteria. Treating an animal for TB implies following strict rules of drug administration, pharmacokinetic check, observance and excretion follow-up, that are very hard to deal with. Zoos who engage in treatment must stick to these rules and require permission for treatment from the official authorities. It must be borne in mind that failure in the treatment could lead to emergence of resistant strain as it was already noticed (LYASHCHENKO ET AL, 2006). The effect of a successful treatment is to bring the animal back to a latent stage, which means that reactivation will always be possible in any stages of its life when treatment is discontinued. Thus, treated animals should be monitored closely for the rest of their life. It could be recommended that treated animals remain in the premises where they have been treated or are transferred to an institution with at least the same level of monitoring abilities. An updated database –working group leaded- is under progress, in order to gather and share information on tests, categorized by zoo species. The feedback of test results is of primary concern, as well as follow-up of suspect populations and fingerprinting of circulating strains. Zoo vets, TAG & EEP advisor must arouse this interest within their relative animal & human population. It’s not acceptable anymore to exchange animals without any medical history including TB. Table 5: Current TB risk and monitoring levels, and methods of diagnostic available for relevant selected species in zoo. Refer to Annex 3 for more specific recommendations when existing. Species Risk Recommended methods (G): see TB Recommended monitoring guidelines level in Annex 3 Depends of status of  Skin test: 0.1ml of bovine PPD, 1mg/ml or Bovidae surrounding animals, and at least 2000 UI/test injected intradermally above all, contact with feral into a shaved cervical area. Reading at 24, animals. 48 and 72h includes palpation and Maintenance monitoring = measuring the skin thickness with an testing on occasions: appropriate caliper. Caudal tail fold less exchange, anesthesia – if appropriate to wild species. purpose doesn’t interfere  Bovigam® : blood should be tested within with TST (infection, use of AI 8h after sampling. Refer to literature for drugs,…) adjusting TST and elicit booster effect.  Serological test Old world monkeys are  Skin test with 0.1ml bovine ppd (at least Primates highly susceptible to 2000 IU/test, 1000 to 10000 times greater infection, new world (G) than human) into eyelid or abdomen skin. monkeys are fairly resistant Reading at 24, 48 and 72h. Old but able to develop TB. mammalian tuberculin may be used to OIE recommends quarantine increase sensibility but it decreases on recently imported specificity

X.

primates with 2 to 3 skin test  at a 2-week intervals.  Carnivores

Not a standard test to  perform except when there is a risk of exposure (Tb cases  in zoo or fed with suspect carcass) 

Sealions

Some South American sealions (O.byronia) infected with M.pinnipedii. Likely to come from imported animals. Transmission to other species of pinnipeds (seals, Californian sealion) and terrestrial mammals occurred. Test should be required for any trade or as soon as animals are trained enough. As bovids

(G)

Camelids

   



 Tapirs (G)

Elephant (G) Deers

Malayan tapirs are most at  risk with a not yet defined proportion of captive population infected.  Infected Asian elephants are  still detected. Clusters of infected feral  deer remain in Europe. 

Rhinoceros M.tuberculosis infection from  infected human reported in  black rhino  M.bovis infected white and black rhino reported in zoos.

Tuberculosis in Zoo Species

Primagam® : whole blood or white cells should be stimulated 8h max. after sampling. Refer to literature for validated or successful species. RapidTest (PrimaTB STATPAK®) on serum Comparative cervical test(CCT) maybe used Serological assays looks promising (ELISA and RapidTest) In-house interferon test could be used (badgers), see references. Skin test with 0.1ml bovine ppd in cervical area, up the shoulder. RapidTest (ElephantTB STATPAK®) : prefer serum to whole blood ELISA on serum Stains, PCR and culture of any nasal discharge

Skint test as indicated for bovids but one has to be aware that the percentage of false positive and false negative is much higher than in bovids Serologic tests (ELISA, ElephantTB STAT PAK) TST or Comparative skin test can be performed with 5000 IU=0.1ml bovine ppd tuberculine and 2000 IU avian ppd tuberculine injected in the inguinal area. Non specific reactions are not uncommon Serological test showed good results Serological tests show the best sensibility and specificity : ELISA, ElephantTB STATPAK® and DPP®VetTB Comparative Cervical Test with avian and bovine ppd (M.avium infection is not rare in cervids) Serologic tests (ELISA, CerivdTB STAT PAK®, DPP®VetTB) Caudal Fold skin test Gastric lavage with culture and PCR Serologic tests (ELISA, CerivdTB STAT PAK®, DPP®VetTB)

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Annex 1 References ABEBE, F., HOLM-HANSEN, C., WIKER, H.G., BJUNE, G. 2007. Progress of Serodiagnosis of Mycobacterium tuberculosis infection. Scandinavian Journal of Immunology 66: 176191 ARTOIS, M., LOUKIADIS, E., GARIN-BASTUJI, B., THOREL, M.F., HARS, J. 2004. Infection des mammifères sauvages par Mycobacterium bovis, risques de transmission aux bovines domestiques. Bulletin Epidémiologique de l’AFFSA 13: 1-3 CHAPEL, H., HAENEY, M., MISBAH, S., SNOWDEN, N. 2006. Essentials of Clinical Immunology, 5th Edition. Blackwell Publishing, Oxford. 368pp. CLIFTON-HADLEY, R.S.,SUATER-LOUIS, C.M., LUGTON, I.W., JACKSON, R., DURR, P.A., WILESMITH, J.W. 2001. Mycobacterium bovis infections. In Infectious diseases of wild mammals (ed E.S. Williams & I.K.Barker). pp 340-361. Iowa State Unisversity Press. COOK RA (1993): Review and update on USDA Cervid TB regulations and their effects on AAZPA accredited zoos. Proc.Am.Assoc.Zoo.Vet., 205-207. COUSINS, N.V. & FLORISSON N. 2005. A review of tests available for use diagnosis of tuberculosis in non-bovine species. Rev sci tech Off int Epiz. 24 (3): 1039-1059. CRANFIELD, M.R., THOEN, C.O., KEMPSKE, S. 1990. An outbreak of Mycobacterium bovis infection in hoofstock at the Baltimore Zoo. Proceedings of the Annual Meeting of the American Association of Zoo Veterinarians, Denver, Colorado, October 21-26 1990 : 128-137. DOHERTY, T.M. & ROOK, G. 2006. Progress and hindrances in tuberculosis vaccine developpement. Lancet 367: 947-949. FITZGERALD, S.D., KANEENE, J.B., BUTLER, K.L. & al. 2000. Comparison of post mortem techniques for the detection of Mycobacterium bovis in white tailed deer (Odocoileus vigrinianus). J Vet DIagn Invest 12(4): 322-327. FLYNN, J.L., CAPUANOA, S.V., CROIXA, D., PAWARA, S., MYERSA, A., ZINOVIKA, A., KLEINB, E. 2003. Non-human primates: a model for tuberculosis research. Proceedings from the 4th World Congress on Tuberculosis. Tuberculosis 83(1-3): 116-118. FRIEDEN, T.R., STERLING, T.R., MUNSIFF, S.S., WATT, C.J., DYE, C. 2003. Tuberculosis. Lancet 362: 887-899. GREENWALD, R., LYASHCHENKO, 0., ESFANDIARI, J., MILLER, M. MIKOTA, S., OLSEN, J.H, BALL, R., DUMONCEAUX, G., SCHMITT, D. MOLLER, T., PAYEUR, J.B., HARRIS, B., SOFRANKO, D., WATERS, R. AND LYASHCHENKO, K.P. 2009. Highly Accurate Antibody Assays for Early and Rapid Detection of Tuberculosis in African and Asian Elephants. Clin. Vaccine Immunol.; 16(5) : 605-612. GROBLER D.G., MICHEL A.L., DE KLERK L.M., BENGIS R.G. 2002. The gamma-interferon test: its usefulness in a bovine tuberculosis survey in African buffaloes (Syncerus caffer) in the Kruger National Park. Onderstepoort J Vet Res. 69(3):221-227. HAAGSMA, J. & EGER, A. 1990. ELISA for diagnosis of Tuberculosis and Chemotherapy in zoo and wildlife animals. Proceedings of the Annual Meeting of the American Association of Zoo Veterinarians, Denver, Colorado, October 21-26 1990: 107-110. JURCYNSKI, K., LYASHCHENKO, K., GOMIS, D., LÉCU, A., TORSTCHANOFF, S., KLARENBEEK, A., MOSER, I. 2007. Mycobacterium pinnipedii infection is South American sea lions (Otaria byronia) in Europe. Proceeding 43rd Internationalen

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Symposiums über die Erkankungen des Zoo un Wildtiere, Edinburgh 2007 : 180. KIERS A, KLARENBEEK A, MENDELTS B, VAN SOOLINGEN D, KOËTER G. 2008. Transmission of Mycobacterium pinnipedii to humans in a zoo with marine mammals. Int J Tuberc Lung Dis. 12(12): 1469-73. LECU, A. & RIQUELME, L. 2008. Evolutions des outils diagnostiques de la tuberculose des éspèces sauvages. Bulletin de l’académie vétérinaire de France, 161 (2) : 151-158. LEWERIN,S.S., OLSSON,S.L., ELD,K., ROKEN,B., GHEBREMICHAEL,S., KOIVULA,T., KALLENIUS,G., BOLSKE,G., 2005. Outbreak of Mycobacterium tuberculosis infection among captive Asian elephants in a Swedish zoo. Vet Rec. 156:171-175. LIN, P.L., PAWAR, S., MYERS, A., PEGU, A., FUHRMAN, C., REINHART, T.A., CAPUANO, S.V., KLEIN, E., FLYNN, J.L. 2006. Early Events in Mycobacterium tuberculosis Infection in Cynomolgus Macaques. Infection and Immunity 74 (7): 3790–3803. LYASHCHENKO KP, GREENWALD R, ESFANDIARI J, OLSEN JH, BALL R, DUMONCEAUX G, DUNKER F, BUCKLEY C, RICHARD M, MURRAY S, PAYEUR JB, ANDERSEN P, POLLOCK JM, MIKOTA S, MILLER M, SOFRANKO D, WATERS WR. 2006. Tuberculosis in Elephants: Antibody Responses to Defined Antigens of Mycobacterium tuberculosis, Potential for Early Diagnosis, and Monitoring of Treatment. Clin. Vaccine Immunol. 13, 722-732. LYASHCHENKO, K.P., GREENWALD, R., ESFANDIARI, J., GREENWALD, D., NACY, C.A., GIBSON, S., DIDIER, P.J., WASHINGTON, M., SZCZERBA, P., MOTZEL, S. & AL. 2007. PrimaTB STAT-PAK Assay, a Novel, Rapid Lateral-Flow Test for Tuberculosis in Nonhuman Primates. Clin. Vaccine Immunol 14(9): 1158-1164. LYASHCHENKO, K.P., SINGH, M. , COLANGELI, R, GENNARO, M.L. 2000. A multi-antigen print immunoassay for the development of serological diagnosis of infectious diseases. Journal of Immunological Methods. 242(1-2): 91-100. LYASHCHENKO K.P., GREENWALD, R., ESFANDIARI, J., CHAMBERS, M.A., VICENTE, J., GORTAZAR, C., SANTOS, N., CORREIA-NEVES, M., BUDDLE, B.M., JACKSON, R., O'BRIEN, D.J., SCHMITT, S., PALMER M.V., DELAHAY, R.J., WATERS, W.R. 2008. Animal-side serologic assay for rapid detection of Mycobacterium bovis infection in multiple species of free-ranging wildlife. Vet Microbiol.;132(3-4): 283-92. MIKOTA SK,LARSEN RS, MONTALI RJ. 2000. Tuberculosis in Elephants in North America. Zoo Biology 19, 393-403. MILLER, M. 2008. Current diagnostic methods for tuberculosis in Zoo Animals. In Fowler, M.E & MILLER, E.R. 2008. Zoo and Wildlife Medicine, current therapy, volume 6. Saunders Elsevier Eds. Pp 10-19. MONTALI, R.J., MIKOTA, S.K., CHENG, L.I. 2001. Mycobacterium tuberculosis in zoo and wildlife species. Rev sci tech Off int Epiz. 20(1): 291-303. MORAR, D., TIJHAARB, E., NEGREA, A., HENDRIKS, J., VAN HAARLEM, D., GODFROID, J., MICHEL, A.L., RUTTEN V.P.M.G. 2007. Cloning, sequencing and expression of white rhinoceros (Ceratotherium simum) interferon-gamma (IFN-γ) and the production of rhinoceros IFN-γ specific antibodies. Veterinary Immunology and Immunopathology. 115 (1-2): 146-154. MOSTOWY, S., INWALD, J., GORDON, S., MARTIN, C., WARREN, R., KREMER, K., COUSINS, D., BEHR, M.A. 2005. Revisiting the Evolution of Mycobacterium bovis. J Bacteriol. 187(18): 6386–6395. OIE. 2007.: Terrestrial Animal health Code. Chapter 3.2.2. Bovine Tuberculosis. RIQUELME, L. 2009. La tuberculose chez les espèces sauvages captives : possibilité d’utilisation du test interferon gamma pour son diagnostic ante-mortem. Thèse Med. Vet.

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Alfort; n°35, 239p. ROBBE-AUSTERMAN, S., KRULL, A.C., STABEL, J.R. 2006. Time delay, temperature effects and assessment of positive controls on whole blood for the gamma interferon ELISA to detect paratuberculosis. J.Vet.Med. B 53: 213-217. SCHLUGER, N.W. 2005. The pathogenesis of tuberculosis: the first one hundred (and twenty-three) years. Am J Respir Cell Mol Biol. 32(4): 251-256. SOCKETT DC (1993): Mycobacterium bovis infection in U.S. Deer and Elk Farms. Proc.Am.Assoc.Zoo.Vet., 198-201. THOEN CO, STEELE JH, GILSDORF MJ (2006): Mycobacterium bovis infection in animals and humans. 2nd Edition. Blackwell Publishing. 329pp. VARGAS D, GARCÍA L, GILMAN RH, EVANS C, TICONA E, ÑAVINCOPA M, LUO RF, CAVIEDES L, HONG C, ESCOMBE R, MOORE DAJ. 2005. Diagnosis of sputum-scarce HIV-associated pulmonary tuberculosis in Lima, Peru. Lancet 365, 150–52 VERVENNE RAW, JONES SL, VAN SOOLINGEN D, VAN DER LAAN T, ANDERSEN P, HEIDT PJ, THOMAS AW, LANGERMANS JAM. 2004. TB diagnosis in non-human primates: comparison of two interferon-g assays and the skin test for identification of Mycobacterium tuberculosis infection. Veterinary Immunology and Immunopathology 100, 61–71 WOOD, P. R. & JONES, S. L. 2001. BOVIGAM® : an in vitro cellular diagnostic test for bovine tuberculosis. Tuberculosis 81(1-2): 147-155.

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Annex 2 List of National Laboratory regarding Mycobacteria diagnostic. VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 34.11.11 Fax: (44.1932) 34.70.46 Glyn Hewinson Email: [email protected] AFSSA Alfort Unité Zoonoses Bactériennes 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 María Laura Boschiroli-Cara Email: [email protected] Veterinary Research Institute Hudcova 70, 62132 Brno CZECH (Rep.) Tel: (420.5) 33.33.16.01 Fax: (420.5) 33.33.12.29 I. Pavlik Email: [email protected] VISAVET Laboratorio de vigilancia veterinaria, Facultad de Veterinaria, Universidad Complutense de Madrid Avda. Puerta de Hierro, s/n. Ciudad Universitaria 28040. Madrid SPAIN J.M. Vizcaino Email: [email protected] Community reference laboratory for bovine tuberculosis from July 2008 until July 2013. Friedrich-Loeffler-Institute Federal Research Institute for Animal Health Suedufer 10 D-17493 Greifswald - Insel Riems GERMANY Phone +49 38351 7-0 Fax +49 38351 7-219 Dr Irmgard Moser Email : [email protected] Central Veterinary Institute - Lelystad Edelhertweg 15 8203 AA Lelystad THE NETHERLANDS Phone: +31 320 238159 Fax: +31 320 238050 Douwe Bakker

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Email : [email protected] National Veterinary Institute (SVA) Department of Bacteriology Section of Mycoplasma and Mycobacteria Diagnosis S-Uppsala SWEDEN Göran BÖLSKE E-mail: [email protected] Institut National des Recherches Veterinaires (INRV) Laboratory of Tuberculosis B-Brussel BELGIUM Jacques Godfroid E-mail: [email protected] Laboratorio Nacional de Investigaçao Veterinaria Ministerio de Agricultura, do Desenvolvimento Rural e das Pescas Estrada de Benfica 701 1500 Lisboa PORTUGAL Tel. 351 1 7115200 Fax. 351 1 7115383 Alicia Amado Email : [email protected] Department of Bacteriology National Veterinary and Food Research Institute Hämeentie 57 Helsinki FINLAND Tel. 358 9 393 18 26 Fax. 358 9 393 18 11 Sinikka Pelkonnen Email : [email protected] Sections of Bacteriology, Virology and Serology, and Pathology National Veterinary Institute 8156 Dep N-0033, Oslo NORWAY Tel. 47 22 96 46 75 Fax. 47 22 60 09 81 Oeivind Oedegaard Email : [email protected] Institute of Veterinary Research of Thesaloniki 80, 26th October Street Thessaloniki GREECE Tel. 30 31 55 20 28 Fax. 30 31 55 40 22 Zoi Dimareli-Malli

Tuberculosis in Zoo Species

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Email : [email protected] EAZWV Working Group members

Tuberculosis in Zoo Species

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Tuberculosis in Zoo Species

Annex 3 Tuberculosis guidelines for some species Malayan Tapirs (Tapirus indicus), from M.Hoyer 2009.

South American Sea Lions (Otaria byronia) issued from the 1st Sealion TB Meeting, Duisburg, Sept. 2009. Contact

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Elephants (Elephas maximus) W.Shaftenaar, vet advisor of Elephant EEP 26 June 2009 These recommendations for the control of tuberculosis in captive Asian en African elephants are aimed to prevent the spread of Mycobacterium spp. that can cause tuberculosis in mammals. The recommendations are based on the document “TB testing in captive elephants in the EEP, 23 July 2008”, (see annex to this document and are reflecting the current possibilities for testing within Europe. The document will be updated when new relevant developments become available. The interpretation of the available diagnostic tests is under constant evaluation and the panel of experts involved in TB-testing in elephants in recent years will be consulted when questions arise. Glossery Antibody test (serum or plasma): ELISA. At present, the Central Veterinary Institute Lelystad is the only institute in Europe running this ELISA on a routine base. Antigens used: M.bovis, MPB70 and M.paratuberculosis. Address: Central Veterinary Institute, DSU Edelhertweg 15, 8219 PH Lelystad, the Netherlands Elephant TB STAT-PAK Assay: Also known as ERT. Test to be performed by a qualified zoo veterinarian or veterinary institute. The test is available through the following website: www.zootest.com Culture of suspected material to be sent to: National Veterinary Laboratory Tuberculin to be obtained from: National Veterinary Institute Trunk wash for culture and/or PCR See definition in the annex How to act when an elephant has to be transported from or to a zoo that participates in the elephant EEP? Before the transport of an elephant to or from an institute that participates in the elephantEEP, the veterinary advisor to the elephant TAG must be notified. Together with the institute that is sending the animal as well as the receiving institute, the veterinary advisor will propose the measures to be taken in order to reduce the risk of transmission of tuberculosis. In case of disagreement, the studbook coordinator will be informed and the EAZA veterinary committee will be consulted if the parties involved do not resolve these issues. The advice of the veterinary advisor is always subject to the (inter-)national veterinary legislation applicable in the countries involved at the time of shipment. The veterinary advisor will base his advice on the pre-transport screening and specific test results as described below. Pre-transport screening A report about the history of the animal should be made, including the following data: • Species / ID / date of birth / location of birth • Locations where the animal has been kept during its entire life, including dates of entry in any new location

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• Has the animal previously been suspected of tuberculosis or treated for this disease? • Has there been any known direct or indirect contact with confirmed or suspected TBcases in herd mates or other mammalian species, including humans? • List of data when blood was sampled, tested and stored at below -20°C • Test results of any TB-test performed in the animal in the past • Does the animal show clinical signs that are suggestive for tuberculosis? Specific tests The animal(s) that will be transported must be tested 4-6 weeks prior to transport for the presence of specific antibodies using the following tests: ELISA’s and Elephant TB STATPAK Assay (ERT). Serum or heparin plasma of the elephant(s) must be shipped on dry ice to the Central Veterinary Institute (CVI) in Lelystad to be tested in the ELISA’s that are routinely used for elephants. The Elephant TB STAT-PAK Assay will also be performed at the CVILelystad. Costs for these tests will be covered by the sender of the samples. On the same day when the blood sample is taken, a comparative skin test will be done: 2000 IU of ppd avian tuberculin* will be injected intracutaneously in a thin part of the skin on the base of the backside of the ear. At the same time 3000-5000 IU of ppd bovine tuberculin* will be injected intracutaneously on the contra-lateral ear at a similar location. If only one ear can be used, both injections should be given at a minimum distance of 10 cm from each other. * Standardization of the quality of the tuberculin is of outmost importance. In case of any doubt, the tuberculin can be provided by the veterinary advisor for the elephant TAG. The results of all tests and the anamnesis data will be evaluated by the zoo vet and the veterinary advisor of the elephant-TAG. The latter will write his advice to the EEP studbookcoordinator. What to do when an elephant is suspected of tuberculosis? An elephant can become suspected of tuberculosis for different reasons. The anamnesis may contain certain data that raise suspicion or one or more tests may give a positive result. When this is the case, the veterinary advisor will consult international experts in the field of tuberculosis and may propose additional tests, which may include: repeated antibody tests (ELISA, ERT), additional intradermal tuberculin test, trunk wash and other tests available at that particular moment). Attempts will also be made to send a serum sample to the USA to perform a Multi Antigen Print Immuno Assay. It is the responsibility of the zoo veterinarian to consult the local veterinary authorities about the situation. Willem Schaftenaar, DVM Veterinary advisor to the elephant TAG Rotterdam zoo PO box 532 3000 AM Rotterdam, the Netherlands Tel: +31 10 4431 485 Fax: +31 10 4431 414 E-mail: [email protected] TB testing in captive elephants in the EEP 23 July 2008 TB testing in elephants is a concern for all elephant keeping institutions. In Europe, no validated tests for elephants, approved by EU-authorities or any other European country, are available at this time. However, veterinary authorities often request elephants to be tested on TB when they are moved to other countries. Zoos that receive elephants must be well aware

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of the risk of the import of TB in their collection. This makes it very important to build up a well documented history of elephant herds, including regular testing and monitoring other animals and personnel. In the EAZWV-recommendations for approval of zoos according to the EU Directive 92/65, the trunk wash procedure for regular testing on tuberculosis (TB) has been discussed. In the final recommendations, a more general text was chosen (see point C.5.h.): “Specific guidelines for the systematic testing of specific animal species may be developed and recommended by the Infectious Diseases Working Group of EAZWV”. This means, that at this time, no official guidelines for TB-testing of elephants are described under the BALAIdirective. However, this does not mean that zoos keeping elephants should not prepare themselves to establish a routine testing protocol. The risk of introducing TB in a zoo through elephants is quite realistic as we can see from cases in the past decade both in the USA as well as in Europe. This document does not touch the issue of introduction of TB through other animal species and human contact, but should leave no doubts that a complete TB-surveillance is the only way to reduce the risks of TB in a zoo. Every national or regional government may have its own interpretation about how to deal with suspected TB-cases. Much depends on the relationship between the zoo veterinarian and the local veterinary authorities. Generally it helps if the zoo has a clear policy regarding its health surveillance system. There is one approach that should not be practiced: the “sleeping dog policy”. Zoos that make all efforts to be certified according to the EU Directive 92/65 should have the intrinsic desire to stay free of infectious diseases like TB. This document describes methods that can be used to collect information about the TB-status of individual elephants and an elephant herd. One should be aware that the European situation can not be compared to the situation in the USA, where one federal government determines guidelines for the entire country. Legislation in the EU and other non-EU-member states is far from uniform. Quality of PPD’s even within the EU differs significantly between the member states. This makes it impossible to write a guideline with an official status. This document can only help to convince institutions that keep elephants to use any means available to minimize the risk of contracting and spreading tuberculosis in their animals and personnel. Finally it may help decision making in case of a planned elephant transfer; the reliability of a “TB- history report” depends a priori of the amount of data collected over the years, not only of a single test taken “on the day of transport”. What are the diagnostic possibilities on live elephants? As there are no tests that provide 100% accuracy, it is of outmost importance that each serum or plasma sample obtained from elephants should always be banked at below -20°C. This can be of great value for the interpretation of future test results and the evaluation of the TB-status in an individual elephant as well as the herd status. To test elephants for carrying TB-organisms, a very limited range of tests can be used, of which none has been validated for elephants: 1. Immunologic tests None of these tests are 100% conclusive, but positive results should be followed by further investigations : a. (Comparative) intradermal tuberculin test. In most countries, this test is only validated for cattle, but it is used in a large range of other species. In elephants it may have some diagnostic value, but results should be interpreted with great care. Bovine and avian tuberculins are injected intradermally in an area where the skin is thin, like the ear base area. The skin reaction is measured after 3, 4 and 5 days (swelling,

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b.

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Tuberculosis in Zoo Species

temperature, sensibility). This test may provide information about the cellular immune response of the animal as a reaction on previous contact of the animal with Mycobacterium sp. A positive response on bovine tuberculin should be followed by further investigations. Humoral antibodies: ELISA: some laboratories run an ELISA for the detection of antibodies. There is no official ELISA recognized in Europe for use in exotic mammals. The Central Veterinary Institute in Lelystad (the Netherlands) is the only official institute that runs these ELISA’s on a routine base for zoos. The antigens used at this CVI are M.bovis, MPB70 and M.paratuberculosis. Positive results should be followed by further investigations. The Elephant TB STAT-PAK Assay is officially recognized in the USA and can be obtained in Europe too (www.zootest.com) However, the test has no official status in Europe at this moment. Gamma interferon test: not yet available for elephants. This test is currently under development at the Department of Infectious diseases and Immunology of the Veterinary Faculty in Utrecht, the Netherlands. A positive test result should be followed by further investigations. 2. Culture of Mycobacterium sp.

For several years the method of choice for sample collection has been the trunk wash*. If positive, this is a conclusive test: you know that you have an animal that is excreting the Mycobacterium sp. Action should be undertaken immediately, regardless the consequences it may have for the status of the zoo. No zoo can allow its personnel to work with an animal which is a serious risk for its employees without taking protective measures nor to expose other animals to this disease. If the culture is negative, the animal may still be a carrier of Mycobacterium sp. and may excrete the pathogens in low quantities. It may take several months before you get the result back from the lab; in the Netherlands the Central Veterinary Institute in Lelystad declares a culture to be negative for M. bovis-complex only after 4 months. A two phase TB diagnostic method combining the culture method and the PCR technique may shorten this period. However the PCR is especially useful, when the samples are ZN positive, otherwise many cycles are needed to get a signal and this will result in a certain numbers of false positive reactions in negative samples. NB: In recent years the sensitivity of the trunk wash method has been proven to be extremely low. If you are dealing with a TB-positive animal, the risk of spreading the Mycobacteria by using this method is a great concern! In the view of the authors this method is not recommended unless performed by qualified persons that are aware of the risks and low sensitivity of the test. * Trunk wash is an active manipulation at the elephant trunk which can be performed in free and protected contact systems in non immobilized elephants after they are conditioned for this procedure. The principle is that a sterile saline solution (approx. 100 ml) is injected in each nostril of the trunk. The nostrils must be kept closed by an elephant keeper and the trunk has to be lifted actively by the elephant or passively by the keeper so that the solution is running up to the basis of the trunk. The mixture of the solution and trunk mucus is collected in sterile plastic bags by active blowing of the elephant through its trunk or by lowering the level of the nostrils. The sample must be sealed and stored at 4°C for immediate culture or stored at –20°C if culture is to be performed at a later stage. Trunk wash under non-contact situation requires a full anesthesia of the elephant and a portable fluid pump and sucking system which allows the operation under sterile condition. The external pump and sucking system will be connected to a sterile PVC tube (1 cm diameter) with a length of approx. 2 meter. The amount of sterile solution and the collection bag are like described before.

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Every zoo should work on a TB-free status, also in elephants. This can be accomplished by regular testing, especially when new animals are being brought in frequently. We propose to test: a.

b. c.

Every elephants at least opportunistically whenever blood can be collected, by ELISA’s (Central Veterinary Laboratory-Lelystad, using antigens derived from M.bovis, MPB70 and M. paratuberculosis) and the Elephant TB STAT-PAK Assay. Training of the elephants should include blood collection; New animals by ELISA’s (Central Veterinary Laboratory-Lelystad, using antigens derived from M.bovis, MPB70 and M. paratuberculosis) and Elephant TB STAT-PAK Assay) and skin test; All suspect animals and contact animals by all immunological tests available. It is advisable to repeat the antibody tests 6 weeks after the intradermal tests. Potential carriers of TB may possibly react by an increase in the production of antibodies as measured by ELISA’s or (if the first sample was negative) by the Elephant TB STATPAK Assay. One has to be aware that this phenomenon may not occur in elephants, though it has been seen in a few individual cases. As in other tests described here, it may contribute to obtain more information about an unknown TB/health status.

What to do when an elephant has died or will be euthanized? An intense search for lesions of tuberculosis is encouraged in all elephant necropsies. This should include all elephants that die or are euthanized for other reasons even though TB is not suspected. Be advised that elephant TB is likely to be caused by Mycobacterium tuberculosis and M. bovis which are contagious to humans. Therefore be prepared with proper protective apparel, and contain any suspicious organs or lesions as soon as possible. Ideally, elephants should be tuberculin tested 3 days prior to euthanasia and bled for serology (ELISA and Elephant TB STAT-PAK) at the time of euthanasia. Serum from elephants that die naturally should be harvested from post mortem blood for serological assays. All elephants undergoing necropsies should have a careful examination of the tonsillar regions and submandibular lymph nodes for tuberculous appearing lesions. Take any nodes that appear caseous or granulomatous for culture (freeze or ultrafreeze), and fixation (in buffered 10% formalin). In addition, search thoracic organs carefully for early stages of TB as follows: after removal of the lungs and trachea, locate the bronchial nodes at the junction of the bronchi from the trachea. Use clean or sterile instruments to section the nodes. Freeze half of the lymph node and submit for TB culture (even if no lesions are evident). Submit sections in formalin for histopathology. Carefully palpate the lobes of both lungs from the apices to the caudal borders to detect any firm (nodular size) lesions. Due to the missing pleural elephants intrathoracal handling is required for the lung removal during necropsy. Please, wear a mouth and nose protector for this procedure. Take sections of any suspicious lesions. Open the trachea and look for nodules or plaques and process as above. Regional thoracic and tracheal lymph nodes should also be examined and processed accordingly. Split the trunk from the tip to its insertion and take samples of any plaques, nodules or suspicious areas for TB diagnosis as above. Look for and collect possible extra-thoracic TB lesions, particularly if there is evidence of advanced pulmonary TB.

Transmissible Diseases Handbook

XI. AVIAN INFLUENZA J.D.W. Philippa WAZA/EAZA/EAZWV

1. VACCINATION OF NON-DOMESTIC AVIAN SPECIES AGAINST HIGHLY PATHOGENIC AVIAN INFLUENZA (HPAI) VIRUSES 2. WAZA/EAZA/EAZWV RECOMMENDATIONS ON AVIAN INFLUENZA

1. Vaccination Of Non-Domestic Avian Species Against Highly Pathogenic Avian Influenza (HPAI) Viruses ADAPTED FROM: “VACCINATION OF NON-DOMESTIC ANIMALS AGAINST EMERGING INFECTIOUS DISEASES” - PHD THESIS ERASMUSMC/ROTTERDAM ZOO, 2007

J.D.W. Philippa

Introduction Avian influenza viruses (AIV) are type A influenza viruses and belong to the Orthomyxoviridae family. They can be classified according to the antigenicity of its surface proteins haemagglutinin (H) and neuraminidase (N). Currently 16H (H1-16) and 9N (N1-9) subtypes have been described in avian species (Fouchier et al., 2005). Furthermore the subtypes can be classified on the basis of their pathogenicity in chickens after intravenous inoculation. Highly pathogenic avian influenza (HPAI, formerly termed fowl plague), an acute generalised disease in which mortality in chickens may be as high as 100%, is restricted to subtypes H5 and H7, although most viruses of these subtypes have low pathogenicity, and do not cause HPAI. Low pathogenic avian influenza (LPAI) virus strains cause more variable morbidity and mortality (ranging from sub-clinical to fatal) but are generally associated in poultry with a mild, primarily respiratory disease with loss of egg production (Capua & Alexander, 2004), or mild enteric disease in non-domestic birds. In certain cases (in poultry flocks) the LPAI virus phenotype (of subtype H5 or H7) may mutate into the HPAI virus phenotype by the introduction of basic amino acid residues (arginine or lysine) at the cleavage site of the precursor haemagglutinin (HAO) (Banks et al., 2001), which facilitates systemic virus

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replication. H5 and H7 subtypes with an amino acid sequence at the HA0 cleavage site comparable to those that have been observed in virulent AI viruses are considered HPAI viruses, even when mortality in chickens is low (Office International d'Epizooties., 2004). However, the two forms of avian influenza (HPAI and LPAI) are distinctly different and should be regarded as such. Avian influenza viruses have a worldwide distribution and are infectious to all avian species (commercial, domestic and wild), with variable morbidity per virus isolate and species. Aquatic avian species, mainly those of the taxonomic orders Anseriformes and Charadriiformes are considered the main natural reservoir of all avian influenza viruses, including the LPAI ancestral viruses of HPAI strains (Munster et al., 2005; Munster et al., 2007). Waterfowl were generally considered resistant to infection with HPAI virus until 2002. However, in 2002 an outbreak of HPAI H5N1 virus occurred in wild migratory avian species and resident waterfowl (Sturm-Ramirez et al., 2004). Since then, this particular HPAI virus subtype has made an unprecedented spread from South East Asia throughout Asia and into Europe and Africa, with morbidity and mortality not only in domestic poultry, but in more than 130 non-domestic avian species from various taxonomic orders: Anseriformes, Charadriiformes, Ciconiiformes, Columbiformes, Falconiformes, Galliformes, Gruiformes, Passeriformes, Pelecaniformes, Phoenicopteriformes, Strigiformes, Struthioniformes, Psittaciformes, and Podicipediformes (USGS National Wildlife Health Center, 2008). Additionally, this virus strain has has caused mortality in a large number of mammalian species, and has caused 403 human cases with 254 deaths to date (27 January 2009) (World Health Organisation, 2009). Documented outbreaks of Asian lineage H5N1 HPAI virus in captive non-domestic birds have been limited to 6 cases: Penfold Park, Hong Kong, (People’s Republic of China, 2002), Kowloon Park, Hong Kong (People’s Republic of China, 2002), Phnom Tamao wildlife rescue centre (Cambodia, 2004), Ragunan Zoo, Jakarta (Indonesia, 2005), Dresden Zoo (Germany, 2006) and Islamabad Zoo (Pakistan, 2007). Large felids with H5N1 infection have been reported in Suphanburi Zoo (Thailand, 2003), and Sri Racha tiger zoo (Thailand, 2004). To curtail these outbreaks, a combination of increased bio-security measures (isolation and quarantine of infected animals, disinfection of the area), feeding of cooked poultry only, treatment of infected animals in quarantine areas, selective culling, extensive surveillance of migratory and captive birds and vaccination were used. Vaccination Vaccination is a useful means of reducing the horizontal spread of AIV in poultry (Capua et al., 2004; van der Goot et al., 2005) (Ellis et al., 2004). Vaccination protects against disease and mortality, but does not always prevent infection and virus spread. However, the dose required for infection is much higher, and vaccinated birds shed far less field virus after infection than unvaccinated birds (Brugh et al., 1979; Karunakaran et al., 1987). Protective antibodies produced in response to infection or vaccination, are directed against the H and N surface proteins. Vaccine-induced antibody responses are species-, dose-, and vaccine strain-dependent, e.g. the antibody responses upon AIV vaccination are generally higher in chickens than in other poultry species (Higgins, 1996). Published minimum serum antibody titres measured by HI test in vaccinated chickens that correlate with protection after challenge with HPAI virus are 1:10 (Swayne et al., 2006), or 1:16 (Ellis et al., 2004; Tian et al., 2005). However, domestic ducks with very low or undetectable antibody titres post vaccination have been shown to be protected from HPAI virus challenge (Middleton et al.,

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2006; Webster et al., 2006). Duration of protection from HPAI virus challenge may vary between species: chickens for up to 40 weeks after one dose of vaccine, domestic ducks for more than 52 weeks after 2 doses, while domestic geese which received 3 doses were protected for 34 weeks (Tian et al., 2005). The degree of homology of the H protein will largely affect the level of cross-protection and therefore efficacy of the vaccine (Swayne et al., 2000). A so-called Differentiation of Infected from Vaccinated Animal (DIVA) strategy, with a heterologous vaccine (using the same H subtype as the field virus, but a different N subtype), is recommended to differentiate between vaccinated and field-virus infected animals (Capua et al., 2003). However, in housing systems where birds are not housed permanently indoors (e.g., in zoos), contact with free-ranging birds can result in LPAI virus infections that go by unnoticed, but which may interfere with the DIVA principle. In the European Union routine vaccination of poultry against avian influenza viruses is currently not practised as this would interfere with stamping-out policies and international trade agreements. Instead, eradication measures during an outbreak in poultry include (longterm) confinement, large-scale culling and safe disposal of carcasses of all poultry on the infected farm, and, depending on the poultry density in the area and the epidemiological situation, pre-emptive culling of poultry on neighbouring farms and emergency vaccinations (Directive 92/94/EEC). Since 2003, more than 300 million birds have been culled to eradicate HPAI outbreaks. Vaccination in European Zoos The standard measures used to prevent and eradicate HPAI virus outbreaks in poultry (longterm confinement and large scale culling) would be detrimental to the welfare, conservation status and breeding programmes of zoo birds, which often are irreplaceable, valuable and endangered avian species (IUCN Red list, http://www.iucnredlist.org/). Directive 2005/94/EC foresees a derogation from culling of birds provided the birds can be brought inside and are subjected to virus detection tests (after the last death/positive finding, 2 tests at an interval of 21 days have to be performed according to the diagnostic manual Decision 2006/437/EC). However, most zoos do not have the capability to suitably confine their entire bird collections for extended time, and many species would not be able to adjust to confinement and increased stress with subsequent welfare problems and increased exposure to pathogens resulting in disease (e.g. aspergillosis and bumblefoot) (McMillian & Petrak, 1989; Redig, 2000; Harcourt-Brown, 1996). Instead of confinement, vaccination of zoo birds against HPAI virus was allowed as an additional preventive measure (while reducing confinement measures) in Belgian, Dutch and German zoos during an outbreak of HPAI H7N7 virus in poultry in 2003 (Decision 2003/291/EC). Similarly, in 2005, Decision 2005/744/EC allowed vaccination in European zoos against the encroaching H5N1 subtype. These campaigns were subject to rigorous surveillance and control requirements. Vaccination against HPAI H7N7 in zoos During the HPAI H7N7 outbreak in poultry in 2003, birds in Dutch zoos were vaccinated twice with six weeks interval using a whole inactivated oil-adjuvanted vaccine, based on influenza virus A/chicken/Italy/473/99 (H7N1), with high homology to the field strain HPAI H7N7 A/chicken/Netherlands/1/03 (97.4% nucleotide and 98.7 % amino acid sequence

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identity). This resulted in the induction of antibody titres ≥ 40 (used as a correlate of protection in this study) in 81.5% of the vaccinated birds, with an overall GMT of 190. Birds of the taxonomic orders Anseriformes, Galliformes and Phoenicopteriformes showed higher GMT, and larger percentages developed a serum HI antibody titre ≥ 40 than those of the other orders. Furthermore, a decrease in antibody response with an increase in body weight > 1.5 kg was shown. The high agreement between post vaccination antibody titres determined by serum HI test (using the vaccine strain), and VN titres (using the field strain), was used as a further measure of immunogenicity. The broad efficacy demonstrated in a large variety of taxonomic orders illustrated the benefits of vaccination as an additional preventive measure against HPAI virus infection (Philippa et al., 2005). Vaccination against HPAI H5N1 in zoos In 2005, the Dutch zoos were the first to implement Decision 2005/744/EC to provide protection against the encroaching HPAI H5N1 subtype. Birds were vaccinated with an inactivated adjuvanted H5N2 vaccine with vaccine doses adapted to mean body weight per species, using data collected during the H7N1 vaccination campaign. The vaccine strain (A/duck/Pottsdam/1402/86) had a homology of 90% to the HA gene of the H5N1 field strain (A/turkey/Turkey/1/05) on the basis of nucleotide sequence (1530 base pairs, including basic cleavage site), and 92.4% on the basis of amino acids. Vaccination was safe, and proved immunogenic throughout the range of species tested, with some variations between and within taxonomic orders. After booster vaccination the overall homologous GMT to the vaccine strain, measured in 334 birds, was 190 (95% CI:152–236), and 80.5% of vaccinated birds developed a titre of ≥ 40. Titres to the HPAI H5N1 virus followed a similar trend, but were lower (GMT: 61 (95% CI: 49–76); 61%≥ 40) (Philippa et al., 2007). The breadth of the immune response was further demonstrated by measuring antibody titres against prototype strains of four antigenic clades of currently circulating H5N1 viruses. Antigenic distances to the prototype strains were determined using antigenic cartography (Smith et al., 2004). Antigenic cartography uses the antigenic properties of influenza viruses combination with epidemiological and genetic data, and is used to select virus strains for use as human pre-pandemic (H5N1) vaccine candidates (World Health Organisation (WHO), 2006). Influenza vaccines whose haemagglutinins are antigenically similar to circulating strains provide the highest level of protection from infection in humans (Subbarao, 1999). The birds clustered in two groups based on the breadth of antibody responses. Group 1 (Anseriformes, Galliformes, Phoenicopteriformes, Psittaciformes and Struthioniformes) showed a very broad response to vaccination, with predicted protection against future strains up to 12 antigenic units from the current vaccine. Group 2 (Ciconiiformes, Gruiformes, Pelecaniformes and Sphenisciformes) had low HI antibody titres against the prototype strain of the most antigenically distant clade (A/Indonesia/5/05). In 2006, a working group of Animal health and Welfare experts was established by the European Food Safety Authority (EFSA)(European Food Safety Authority (EFSA), 2007), to provide a scientific assessment of the preventive vaccination against avian influenza of H5 and H7 subtypes carried out in zoos in Member States (MS). The total number of birds vaccinated, as reported by 12 MS, was 44721. Individual data from 4718 birds (374 species from 19 taxonomic orders) were submitted. Not all of these could be used for every evaluation: pre-vaccination titres could be evaluated for 3039 birds; titres after first vaccination were evaluated for 1429 birds, and post-second vaccination titres for 2296 birds.

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Differences in vaccination schedules, doses and routes, differences in methodology and antigens used in the HI tests between laboratories (due to the absence of international reference standards, and the absence of inter-laboratory standardisation of methodology), the use of different vaccines1 in different taxonomic orders and the sometimes incomplete reporting of results, limited the evaluation of some of the data provided by EU MS. Cut-off points varied with laboratory, and titres considered a measure of adequate immune response were 8, 16, 32, 40, and 64. Most countries used dilution series starting at 4 or 8, therefore results were evaluated for titres 16 and 32 [documented surrogate markers for protection in chickens (Ellis et al., 2004; Office International d'Epizooties., 2004; Swayne et al., 2006; Tian et al., 2005)], and undetectable titres were regarded as 4 for calculation of GMT. In the absence of (and unfeasibility of obtaining) vaccination/challenge data in often endangered zoo bird species, the evaluation had to be based on extrapolation of serological data from poultry and limited other bird species. The H5 vaccines registered for poultry in the EU showed differences in efficacy, measured as serum HI antibodies induced by two doses of vaccine (Table 1.). Three of the five vaccines evaluated induced relatively high GMT and high percentage seroconversion in the vast majority of vaccinated birds. The HI titres induced by vaccination showed marked differences between and within taxonomic orders. Both routes of vaccination (i.m. and s.c.) were effective in inducing HI serum antibody responses, and for most avian species the poultry dose was suitable. In some larger species higher doses adjusted to body weight, induced higher serum antibody titres. (e.g. for ostriches a 10-fold increase of the poultry dose (10 x 0.25ml). However, extremely high doses at a single site of injection (e.g. vaccination of ostriches with 10 ml of vaccine) appeared to have a negative effect on the induction of serum antibody titres, and induced local adverse reactions. There were indications that one vaccination was sufficient to induce high serum antibody titres in at least two taxonomic orders of birds (Anseriformes and Galliformes). However, a second vaccine administration ensured seroconversion in the majority of birds of most species. Limited data indicated that antibody titres persisted in several species for six months after vaccination. Adverse effects and mortality associated with vaccination were low and were mainly attributable to handling stress or trauma. Differences in adverse effects reported from different zoos highlight the importance of proper skilled handling. Longevity of antibody titres One year after vaccination with the H5N2 vaccine, birds in Dutch zoos were revaccinated with the same vaccine. Antibody titres one year after the initial two vaccinations and the effect of one booster vaccination at this time were evaluated. In Rotterdam Zoo, 72 previously vaccinated birds could be evaluated for the effect of one booster vaccination (Philippa et al, in press). For 44 birds, serum samples were available from 4 weeks after the initial two vaccinations the previous year, at the time of revaccination, and 4 weeks later. Birds which had been vaccinated with the H7 vaccine two years prior to the H5N2 revaccination were additionally tested for the presence of H7-specific antibodies.

1

Vaccine A: H5N9 (A/turkey/Wisconsin/68). Vaccine B: H5N2 (A/duck/Pottsdam/1402/86). Vaccine C: H5N2 (A/chicken/Mexico/232/94/CPA) Vaccine D: H5N9 (A/chickenk/Italy/22A/98). Vaccine E: H5N9 (A/chicken/ltaly/22A/H5N9/1998).

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Serum antibody titres of the birds tested in Rotterdam Zoo had clearly decreased in one year time: while 80% of birds had a positive titre (≥ 8) and 68% a high positive titre (≥ 32) after 2 vaccinations, these figures were 61% and 30% respectively one year later. Four weeks after re-vaccination these figures increased to 93% and 77% respectively. Although a larger percentage of these 44 birds had a serum HI antibody titre ≥ 32 after re-vaccination, the GMT was lower than GMT after 2 vaccine doses one year before (88 vs 66). Birds from 4 out of 8 taxonomic orders did not have a GMT > 5 one year after vaccination, and only one order (Phoenicopteriformes had a GMT > 40. Four weeks after one revaccination 6/8 taxonomic orders tested had a GMT > 40. GMTs had decreased even further two years after vaccination, as was shown by the H7 specific serum HI antibody titres. As these birds were not revaccinated with an H7 vaccine, the effect of revaccination two years after the initial vaccinations is not known.

Conclusions Bio-security measures remain the first line of protection of zoo birds against the introduction of AI viruses and should be implemented in zoos. These bio-security measures should include strict hygiene and quarantine measures, but should also exclude the possibility of introducing AI viruses through feed animals such as day old chicks, other poultry or their products. Continuous clinical monitoring of captive and wild birds in zoos should be practiced, for early detection of introduced viruses by wild birds, domestic birds, or their products. Strict bio-security measures will also reduce the risk of subsequent infection of wild birds from zoo birds. Wild birds have been documented to be susceptible to HPAI virus infection, and could potentially play a role in the spread of HPAI virus, although the majority of avian influenza viruses detected in free-ranging birds have been LPAI viruses. If biosecurity measures cannot sufficiently protect zoo birds from exposure to HPAI viruses coming from wild birds (based on an overall risk assessment which includes welfare aspects) vaccination with vaccines against HPAI of H5 and H7 subtypes authorised for use in poultry should be used to protect these zoo birds. In designing AI vaccination programmes and schedules for zoo birds, recent data on wild bird migration and prevalence of AI viruses should be taken into account. Vaccination against AI viruses of the H5 and H7 subtypes with current inactivated oil-adjuvanted poultry vaccines is safe and, in most taxonomic orders of zoo birds, effective in terms of inducing HI serum antibody titres. AI vaccines should be administrated in a way that elicits high HI antibody titres in the vast majority of the zoo birds vaccinated, i.e., by adjusting dose to average body weight. Although there are indications that one vaccination might suffice for some species, a second vaccine dose ensures high titres in the vast majority of species. Unless it is demonstrated that one vaccine administration is sufficient, two administrations are recommended. The H5 and H7 vaccines currently registered for poultry in the EU show differences in the performance in terms of HI response in zoo birds after two doses. There appears to be no difference in route of vaccination (s.c. or i.m.), so route can vary depending on the bird species to be vaccinated. In order to maintain high titres in the captive populations in zoological collections, annual revaccination seems to be required, as antibody titres decrease significantly in most taxonomic orders, and high titres are seen after a single annual booster dose. Mortality and adverse effects were low in all zoos evaluated in EU MS, and mainly attributed to handling stress and trauma. Zoos can, and should therefore try to minimise these losses in the execution of HPAI vaccination programmes. To minimise indirect losses due to

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decreased breeding results, AI vaccination during breeding seasons should be avoided whenever possible. Mortality due to catching and handling stress can be reduced by handling the birds less. Once the efficacy of a vaccination protocol has been validated for certain species using certain vaccines, measurement of post-vaccination HI serum antibody titres should no longer be mandatory by the EU. These birds will then only have to be handled for vaccination, and not 4 weeks later. Further research should be carried out to establish effective vaccination schedules, route, and dose regimen in different zoo bird species. This may, amongst others, lead to a reduction in the number of booster vaccinations needed in certain species. Novel generation vaccines which may be administered in the form of an aerosol (as is used in vaccination of poultry against Newcastle disease virus) may prove to be useful in non-domestic species, and would eliminate the need for handling the birds. The vaccination campaigns against HPAI virus have focused on protecting birds in zoological collections. However, a large number of mammalian species, including tigers and leopards, have also been documented with HPAI virus infection with recent H5N1 subtypes. There is currently no commercial vaccine available to protect mammals from HPAI H5N1 virus infection. A recombinant fowlpox-vectored vaccine expressing the H5 gene has been shown to produce high antibody titres against heterologous H5N1 virus antigen in cats after booster vaccination (Karaca et al., 2005), and may prove to be useful in prophylactic vaccination programs of mammals in the future. Until then, these animals have to be protected by excluding the introduction of AIV through raw poultry used as feed. The broad vaccine efficacy in the different avian taxonomic orders illustrates that vaccination against avian influenza is a useful tool for the protection of non-domestic avian species in zoos, which allows for an alleviation of confinement measures – and is therefore beneficial to the health and welfare of these birds. However, increased bio-security measures in combination with virological monitoring remain imperative in combating outbreaks of HPAI.

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References Banks,J., Speidel,E.S., Moore,E., Plowright,L., Piccirillo,A., Capua,I., Cordioli,P., Fioretti,A., Alexander,D.J., 2001. Changes in the haemagglutinin and the neuraminidase genes prior to the emergence of highly pathogenic H7N1 avian influenza viruses in Italy. Arch. Virol. 146:963-973. Brugh,M., Beard,C.W., Stone,H.D., 1979. Immunization of chickens and turkeys against avian influenza with monovalent and polyvalent oil emulsion vaccines. Am. J. Vet. Res. 40:165-169. Capua,I., Alexander,D.J., 2004. Avian influenza: recent developments. Avian Pathol. 33:393404. Capua,I., Terregino,C., Cattoli,G., Mutinelli,F., Rodriguez,J.F., 2003. Development of a DIVA (Differentiating Infected from Vaccinated Animals) strategy using a vaccine containing a heterologous neuraminidase for the control of avian influenza. Avian Pathol. 32:47-55. Capua,I., Terregino,C., Cattoli,G., Toffan,A., 2004. Increased resistance of vaccinated turkeys to experimental infection with an H7N3 low-pathogenicity avian influenza virus. Avian Pathol. 33:158-163. Ellis,T.M., Leung,C.Y., Chow,M.K., Bissett,L.A., Wong,W., Guan,Y., Malik Peiris,J.S., 2004. Vaccination of chickens against H5N1 avian influenza in the face of an outbreak interrupts virus transmission. Avian Pathol. 33:405-412. European Food Safety Authority (EFSA). Opinion of the Scientific Panel AHAW related with the vaccination against avian influenza of H5 and H7 subtypes as a preventive measure carried out in Member States in birds kept in zoos under Community approved programmes. 2007. Fouchier,R.A., Munster,V., Wallensten,A., Bestebroer,T.M., Herfst,S., Smith,D., Rimmelzwaan,G.F., Olsen,B., Osterhaus,A.D., 2005. Characterization of a novel influenza A virus hemagglutinin subtype (H16) obtained from black-headed gulls. J. Virol. 79:2814-2822. Harcourt-Brown,N.H. 1996. Bumblefoot. Pages 126-131 in Medicine. Harcourt Publishers Ltd, London.

Samour,J.H. editor. Avian

Higgins,D.A. 1996. Comparative immunology of avian species. Pages 149-205 in Davison,T.F., Payne,L.N., Morris,T.R. editors. Poultry Immunology. Carfax publishing co., Abingdon. Karaca,K., Swayne,D.E., Grosenbaugh,D., Bublot,M., Robles,A., Spackman,E., Nordgren,R., 2005. Immunogenicity of fowlpox virus expressing the avian influenza virus H5 gene (TROVAC AIV-H5) in cats. Clin. Diagn. Lab Immunol. 12:1340-1342. Karunakaran,D., Newman,J.A., Halvorson,D.A., Abraham,A., 1987. Evaluation of inactivated influenza vaccines in market turkeys. Avian Dis. 31:498-503. McMillian,M., Petrak,M., 1989. Retrospective study of aspergillosis in pet birds. J. Avian Med Surg211-215. Middleton,D., Bingham,J., Selleck,P., Lowther,S., Gleeson,L., Lehrbach,P., Robinson,S., Rodenberg,J., Kumar,M., Andrew,M., 2006. Efficacy of inactivated vaccines against H5N1 avian influenza infection in ducks. Virology.

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Munster,V.J., Baas,C., Lexmond,P., Waldenstrom,J., Wallensten,A., Fransson,T., Rimmelzwaan,G.F., Beyer,W.E., Schutten,M., Olsen,B., Osterhaus,A.D., Fouchier,R.A., 2007. Spatial, temporal, and species variation in prevalence of influenza A viruses in wild migratory birds. PLoS. Pathog. 3:e61. Munster,V.J., Wallensten,A., Baas,C., Rimmelzwaan,G.F., Schutten,M., Olsen,B., Osterhaus,A.D.M.E., Fouchier,R.A.M., 2005. Mallards and highly pathogenic avian influenza ancestral viruses, northern europe. Emerg. Infect. Dis. 11:1545-1551. Office International d'Epizooties. 2004. Avian Influenza.OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. Philippa,J., Baas,C., Beyer,W., Bestebroer,T., Fouchier,R., Smith,D., Schaftenaar,W., Osterhaus,A., 2007. Vaccination against highly pathogenic avian influenza H5N1 virus in zoos using an adjuvanted inactivated H5N2 vaccine. Vaccine 25:3800-3808. Philippa,J.D.W., Munster,V.J., Bolhuis,H., Bestebroer,T.M., Schaftenaar,W., Beyer,W.E., Fouchier,R.A.M., Kuiken,T., Osterhaus,A.D.M.E., 2005. Highly pathogenic avian influenza (H7N7): Vaccination of zoo birds and transmission to non-poultry species. Vaccine 23:57435750. Redig,P.T. 2000. Aspergillosis. Pages 275-287 in Samour,J. editor. Avian Medicine. Mosby, Philadelphia. Smith,D.J., Lapedes,A.S., de Jong,J.C., Bestebroer,T.M., Rimmelzwaan,G.F., Osterhaus,A.D., Fouchier,R.A., 2004. Mapping the antigenic and genetic evolution of influenza virus. Science 305:371-376. Sturm-Ramirez,K.M., Ellis,T., Bousfield,B., Bissett,L., Dyrting,K., Rehg,J.E., Poon,L., Guan,Y., Peiris,M., Webster,R.G., 2004. Reemerging H5N1 influenza viruses in Hong Kong in 2002 are highly pathogenic to ducks. J. Virol. 78:4892-4901. Subbarao,K., 1999. Influenza vaccines: present and future. Adv. Virus Res. 54:349-373. Swayne,D.E., Lee,C.W., Spackman,E., 2006. Inactivated North American and European H5N2 avian influenza virus vaccines protect chickens from Asian H5N1 high pathogenicity avian influenza virus. Avian Pathol. 35:141-146. Swayne,D.E., Perdue,M.L., Beck,J.R., Garcia,M., Suarez,D.L., 2000. Vaccines protect chickens against H5 highly pathogenic avian influenza in the face of genetic changes in field viruses over multiple years. Vet. Microbiol. 74:165-172. Tian,G., Zhang,S., Li,Y., Bu,Z., Liu,P., Zhou,J., Li,C., Shi,J., Yu,K., Chen,H., 2005. Protective efficacy in chickens, geese and ducks of an H5N1-inactivated vaccine developed by reverse genetics 6. Virology 341:153-162. USGS National Wildlife Health Center. Referenced reports of highly pathogenic avian influenza H5N1 in wildlife and domestic animals. USGS . 2008. 31-3-2009. van der Goot,J.A., Koch,G., de Jong,M.C., van,B.M., 2005. Quantification of the effect of vaccination on transmission of avian influenza (H7N7) in chickens. Proc. Natl. Acad. Sci. U. S. A. Webster,R.G., Webby,R.J., Hoffmann,E., Rodenberg,J., Kumar,M., Chu,H.J., Seiler,P., Krauss,S., Songserm,T., 2006. The immunogenicity and efficacy against H5N1 challenge of reverse genetics-derived H5N3 influenza vaccine in ducks and chickens. Virology 351:303311.

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World Health Organisation. Cumulative Number of Confirmed Human Cases of Avian Influenza A/(H5N1) Reported to WHO. World Health Organisation . 2009. World Health Organisation (WHO). Antigenic and genetic characteristics of H5N1 viruses and candidate H5N1 vaccine viruses developed for potential use as pre-pandemic vaccines. 2006.

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2. WAZA/EAZA/EAZWV Recommendations on Avian Influenza

Background Highly pathogenic avian influenza (HPAI) is an extremely contagious viral disease of domestic birds that can affect all species of birds. It is also known to have infected a range of wild and domestic species of mammals (including civets, ferret, beech marten, domestic cat, clouded leopard, leopard, tiger) and humans, and has experimentally been transmitted to mice, rats, rabbits and crab-eating macaques. The disease is caused by the Influenza A virus (Family Orthomyxoviridae), which can be divided into subtypes on the basis of distinct haemagglutinin antigens (H1–H16) and neuraminidase antigens (N1–N9). It is a notifiable disease under the Terrestrial Animal Health Code of the International Organisation for Animal Health (OIE), thus notifiable by law in the 167 Member Countries of the OIE. The Code defines notifiable avian influenza as an infection of poultry (i.e. not of wild birds!) caused by any influenza A virus of the H5 or H7 subtypes or by any AI virus with an intravenous pathogenicity index (IVPI) greater than 1.2 (or as an alternative at least 75% mortality). Notifiable avian influenza can be divided into highly pathogenic and low pathogenic notifiable avian influenza. Highly pathogenic avian influenza (HPAI) may result in a mortality rate of 100% within a poultry flock. These viruses have been restricted to subtypes H5 and H7, although not all viruses of these subtypes cause HPAI. All other viruses cause a much milder low pathogenicity avian influenza (LPAI) consisting primarily of mild respiratory disease and egg production problems in laying birds, or being completely inapparent. From 1959 to 1998, only 24 distinct outbreaks of HPAI from domestic poultry have been reported. The total number of birds involved in all outbreaks over this 40-year period was approximately 23 million. But the years 1999-2004 have witnessed six outbreaks that resulted in a considerable socio-economic impact (e.g. 1999/2000 in Italy, 2003 in the Netherlands and in Germany) and the unprecedented outbreak currently affecting several Asian countries. The USA, Canada, Chile and some African countries have also recently experienced HPAI disease outbreaks. From 1999-2004 over 200 million domestic birds have been affected by the disease. With the exception of some countries, where the disease in domestic poultry is not yet under control, a rigorous stamping out policy was applied by national authorities. This strategy included in general the rapid culling of infected poultry and those suspected of being infected together with the implementation of movement restrictions for live poultry and poultry products, increased monitoring and biosecurity measures. HPAI is considered to have been spread between countries by a number of different known vectors, including through the movement of avian livestock and bird by-products, legal and illegal trade in birds, equipment associated with these respective industries, movement of people, and migration of waterbirds. The genetic pool for AI viruses is primarily in the aquatic birds responsible for the perpetuation of these viruses in nature. It still is a matter of debate whether HPAI viruses are normally present in wild bird populations, or whether they arise only as a result of mutation after H5 or H7 LPAI viruses have been introduced to poultry from wild birds. The unusual situation of endemicity which is present in some Asian countries suggests a spill over of HPAI of the H5N1 subtype to the wild bird population. This situation has never occurred in

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the past, and therefore the consequences of this epidemiological situation are unpredictable. Until now, the limited number of known human infections with H5N1 virus has been through close contact with, or possibly by consumption of, infected poultry and none through contact with wild birds. If however the virus were to change its characteristics and to spread between humans, the result could be a pandemic with millions of victims. However, nobody can predict whether such changes will happen. Although, traditionally, legislation of most countries did not address avian influenza in wild birds, veterinary services, noting the pandemic potential of the H5N1 strain, have started monitoring the disease in wild birds and implementing protection measures also in the case of the presence of the virus in other species than farmed poultry. In Europe the legal basis for the new approach is Council Directive 2005/94/EC of 20 December 2005 on Community measures for the control of avian influenza, which repealed the previous Directive 92/40/EEC. Current approaches to control the disease may refer to both HPAI and LPAI infections and include: surveillance of bird species other than domestic poultry; early detection of possible spread of avian influenza to mammals; notification of surveillance measures and outbreaks; emergency measures applicable to holdings where outbreaks are suspected; killing of all poultry and other captive birds, disposal of carcases and eggs, and disinfection of the premises under official supervision, in holdings where outbreaks are confirmed; establishment of protection, surveillance and further restricted zones around affected holdings. Routine vaccination, although technically possible and effective, continues to be prohibited in many countries. Such countries may, however, have provisions for emergency vaccinations, grant certain derogations, or introduce preventive vaccinations on the basis of a risk assessment. Recognising that avian influenza has always occurred, and will continue to occur, in wild birds (although the incidence is usually very low), that WHO, FAO and OIE have concluded that attempts to eliminate HPAI in wild bird populations through lethal responses such as culling are not feasible and may exacerbate the problem by causing further dispersion of infected birds, that it is technically impossible to keep wild birds out of zoological institutions, and that zoological institutions need to keep birds of wild species in order to play their important role in conservation communication, education and public awareness (as stressed by resolutions of the Conferences of the Parties of the Ramsar Convention and Convention on Migratory Species), legislation tends to be somewhat more flexible with regard to zoos than legislation relating to other highly contagious diseases, such as foot-and-mouth disease, African swine fever and so on. It may thus be possible for a zoo to be granted certain derogations (e.g. to be allowed to vaccinate preventively, or not have to kill the entire collection in the case of an outbreak at or near the zoo) under specified conditions. These conditions may include, among other things, restrictions of the transfer of vaccinated birds to other holdings.

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References Convention on Migratory Species (2005). Res. 8.27 Migratory Species and Highly Pathogenic Avian Influenza, adopted at Eighth Meeting of the Conference of the Parties to CMS (COP 8), 20 - 25 November 2005, UNEP Headquarters Gigiri, Nairobi. European Association of Zoo and Wildlife Veterinarians (2004) Transmissible Diseases Handbook, 2nd edition. Houten NL. European Food Safety Authority – AHAW (2005). Scientific Report - Animal health and welfare aspects of Avian Influenza. Annex to the EFSA Journal (2005) 266, 1-21. European Union (2005) Commission Decision 2005/731/EC of 17 October 2005 laying down additional requirements for the surveillance of avian influenza in wild birds. OJ L 27/93 of 20.10.2005 European Union (2006) Council Directive 2005/94/EC of 20 December 2005 on Community measures for the control of avian influenza and repealing Directive 92/40/EEC. OJ L10/16 of 14.1.2006 OIE (2005) Terrestrial Animal Health Code. 14th edition. Paris.

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Avian Influenza

WAZA Council Resolution on Avian Influenza (11th April 2006) RECOGNIZING the potential risk of transmission of Highly Pathogenic Avian Influenza (HPAI) from wild birds, in particular waterfowl, to birds kept in zoological institutions; MINDFUL that the zoos’ role in environmental education may require to exhibit birds under conditions, contact with wild birds cannot effectively be excluded; CONCERNED about the impact an outbreak of HPAI in or near a zoological institution could have on the well-being of the birds in the collection, the functioning of conservation breeding programmes and the economy of the institution affected; AWARE that legislation to control HPAI may provide for derogations or special conditions applicable to zoological institutions; CONVINCED that, in spite of the limited knowledge about the efficacy and safety of current vaccines in non-domesticated species, appropriate preventive vaccination has more advantages than disadvantages, and that vaccination with an inactivated vaccine should be applied where allowed and appropriate to protect zoo birds likely to enter into contact with possibly HPAI infected wild birds; THE WORLD ASSOCIATION OF ZOOS AND AQUARIUMS RECOMMENDS: ASSOCIATION MEMBERS should  maintain a regular dialogue with the national veterinary services of their respective countries with a view of providing input into the legislation process and exploring the possibility of regulations being applied to zoological institutions that will take into account the special situation under which these institutions have to operate2;  inform their constituencies of current regulations and any existing derogations for zoological institutions;  advise their constituencies to head for common approaches and to coordinate such approaches as appropriate. INSTITUTION MEMBERS should  assume that all bird species are susceptible to HPAI and not only those that may be addressed by current legislation;  maintain a regular dialogue with the veterinary authority responsible for supervising the institution;  implement, and have approved by the competent veterinary authority, a health surveillance plan covering the entire collection;  apply strict quarantine measures when introducing birds into the collection, even in the event of these birds not being imported from a third country;  explore the possibility of the institution being subdivided into different compartments for the purpose of animal health measures;  apply biosecurity measures as appropriate within the institution;  take special precaution to avoid the spreading of any infection from one compartment to another, in case the institution has been subdivided;

2

EAZA should, in collaboration with EAZWV act accordingly with regard to the European Commission.

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Avian Influenza

develop and have agreed by the competent veterinary authority contingency plans to be implemented in the event of a HPAI outbreak in or in the neighbourhood of the institution; explore the possibility and assess the advantages and disadvantages of vaccinating those birds in their collection that are likely to enter into close contact with wild birds, in particular waterfowl.

Transmissible Diseases Handbook

XI. VACCINATION OF NON-DOMESTIC AVIAN SPECIES AGAINST HIGHLY PATHOGENIC AVIAN INFLUENZA (HPAI) VIRUSES ADAPTED FROM: “VACCINATION OF NON-DOMESTIC ANIMALS AGAINST EMERGING INFECTIOUS DISEASES” - PHD THESIS ERASMUSMC/ROTTERDAM ZOO, 2007

J.D.W. Philippa

Introduction Avian influenza viruses (AIV) are type A influenza viruses and belong to the Orthomyxoviridae family. They can be classified according to the antigenicity of its surface proteins haemagglutinin (H) and neuraminidase (N). Currently 16H (H1-16) and 9N (N1-9) subtypes have been described in avian species (Fouchier et al., 2005). Furthermore the subtypes can be classified on the basis of their pathogenicity in chickens after intravenous inoculation. Highly pathogenic avian influenza (HPAI, formerly termed fowl plague), an acute generalised disease in which mortality in chickens may be as high as 100%, is restricted to subtypes H5 and H7, although most viruses of these subtypes have low pathogenicity, and do not cause HPAI. Low pathogenic avian influenza (LPAI) virus strains cause more variable morbidity and mortality (ranging from sub-clinical to fatal) but are generally associated in poultry with a mild, primarily respiratory disease with loss of egg production (Capua & Alexander, 2004), or mild enteric disease in non-domestic birds. In certain cases (in poultry flocks) the LPAI virus phenotype (of subtype H5 or H7) may mutate into the HPAI virus phenotype by the introduction of basic amino acid residues (arginine or lysine) at the cleavage site of the precursor haemagglutinin (HAO) (Banks et al., 2001), which facilitates systemic virus replication. H5 and H7 subtypes with an amino acid sequence at the HA0 cleavage site comparable to those that have been observed in virulent AI viruses are considered HPAI viruses, even when mortality in chickens is low (Office International d'Epizooties., 2004). However, the two forms of avian influenza (HPAI and LPAI) are distinctly different and should be regarded as such. Avian influenza viruses have a worldwide distribution and are infectious to all avian species (commercial, domestic and wild), with variable morbidity per virus isolate and species. Aquatic avian species, mainly those of the taxonomic orders Anseriformes and Charadriiformes are considered the main natural reservoir of all avian influenza viruses, including the LPAI ancestral viruses of HPAI strains (Munster et al., 2005; Munster et al., 2007). Waterfowl were generally considered resistant to infection with HPAI virus until 2002. However, in 2002 an outbreak of HPAI H5N1 virus occurred in wild migratory avian species and resident waterfowl (Sturm-Ramirez et al., 2004). Since then, this particular HPAI virus subtype has made an unprecedented spread from South East Asia throughout Asia and into Europe and Africa, with morbidity and mortality not only in domestic poultry, but in more than

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Vaccination of Non-domestic Avian Species

130 non-domestic avian species from various taxonomic orders: Anseriformes, Charadriiformes, Ciconiiformes, Columbiformes, Falconiformes, Galliformes, Gruiformes, Passeriformes, Pelecaniformes, Phoenicopteriformes, Strigiformes, Struthioniformes, Psittaciformes, and Podicipediformes (USGS National Wildlife Health Center, 2008). Additionally, this virus strain has has caused mortality in a large number of mammalian species, and has caused 403 human cases with 254 deaths to date (27 January 2009) (World Health Organisation, 2009). Documented outbreaks of Asian lineage H5N1 HPAI virus in captive non-domestic birds have been limited to 6 cases: Penfold Park, Hong Kong, (People’s Republic of China, 2002), Kowloon Park, Hong Kong (People’s Republic of China, 2002), Phnom Tamao wildlife rescue centre (Cambodia, 2004), Ragunan Zoo, Jakarta (Indonesia, 2005), Dresden Zoo (Germany, 2006) and Islamabad Zoo (Pakistan, 2007). Large felids with H5N1 infection have been reported in Suphanburi Zoo (Thailand, 2003), and Sri Racha tiger zoo (Thailand, 2004). To curtail these outbreaks, a combination of increased bio-security measures (isolation and quarantine of infected animals, disinfection of the area), feeding of cooked poultry only, treatment of infected animals in quarantine areas, selective culling, extensive surveillance of migratory and captive birds and vaccination were used. Vaccination Vaccination is a useful means of reducing the horizontal spread of AIV in poultry (Capua et al., 2004; van der Goot et al., 2005) (Ellis et al., 2004). Vaccination protects against disease and mortality, but does not always prevent infection and virus spread. However, the dose required for infection is much higher, and vaccinated birds shed far less field virus after infection than unvaccinated birds (Brugh et al., 1979; Karunakaran et al., 1987). Protective antibodies produced in response to infection or vaccination, are directed against the H and N surface proteins. Vaccine-induced antibody responses are species-, dose-, and vaccine strain-dependent, e.g. the antibody responses upon AIV vaccination are generally higher in chickens than in other poultry species (Higgins, 1996). Published minimum serum antibody titres measured by HI test in vaccinated chickens that correlate with protection after challenge with HPAI virus are 1:10 (Swayne et al., 2006), or 1:16 (Ellis et al., 2004; Tian et al., 2005). However, domestic ducks with very low or undetectable antibody titres post vaccination have been shown to be protected from HPAI virus challenge (Middleton et al., 2006; Webster et al., 2006). Duration of protection from HPAI virus challenge may vary between species: chickens for up to 40 weeks after one dose of vaccine, domestic ducks for more than 52 weeks after 2 doses, while domestic geese which received 3 doses were protected for 34 weeks (Tian et al., 2005). The degree of homology of the H protein will largely affect the level of cross-protection and therefore efficacy of the vaccine (Swayne et al., 2000). A so-called Differentiation of Infected from Vaccinated Animal (DIVA) strategy, with a heterologous vaccine (using the same H subtype as the field virus, but a different N subtype), is recommended to differentiate between vaccinated and field-virus infected animals (Capua et al., 2003). However, in housing systems where birds are not housed permanently indoors (e.g., in zoos), contact with free-ranging birds can result in LPAI virus infections that go by unnoticed, but which may interfere with the DIVA principle. In the European Union routine vaccination of poultry against avian influenza viruses is currently not practised as this would interfere with stamping-out policies and international

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Vaccination of Non-domestic Avian Species

trade agreements. Instead, eradication measures during an outbreak in poultry include (longterm) confinement, large-scale culling and safe disposal of carcasses of all poultry on the infected farm, and, depending on the poultry density in the area and the epidemiological situation, pre-emptive culling of poultry on neighbouring farms and emergency vaccinations (Directive 92/94/EEC). Since 2003, more than 300 million birds have been culled to eradicate HPAI outbreaks. Vaccination in European Zoos The standard measures used to prevent and eradicate HPAI virus outbreaks in poultry (longterm confinement and large scale culling) would be detrimental to the welfare, conservation status and breeding programmes of zoo birds, which often are irreplaceable, valuable and endangered avian species (IUCN Red list, http://www.iucnredlist.org/). Directive 2005/94/EC foresees a derogation from culling of birds provided the birds can be brought inside and are subjected to virus detection tests (after the last death/positive finding, 2 tests at an interval of 21 days have to be performed according to the diagnostic manual Decision 2006/437/EC). However, most zoos do not have the capability to suitably confine their entire bird collections for extended time, and many species would not be able to adjust to confinement and increased stress with subsequent welfare problems and increased exposure to pathogens resulting in disease (e.g. aspergillosis and bumblefoot) (McMillian & Petrak, 1989; Redig, 2000; Harcourt-Brown, 1996). Instead of confinement, vaccination of zoo birds against HPAI virus was allowed as an additional preventive measure (while reducing confinement measures) in Belgian, Dutch and German zoos during an outbreak of HPAI H7N7 virus in poultry in 2003 (Decision 2003/291/EC). Similarly, in 2005, Decision 2005/744/EC allowed vaccination in European zoos against the encroaching H5N1 subtype. These campaigns were subject to rigorous surveillance and control requirements. Vaccination against HPAI H7N7 in zoos During the HPAI H7N7 outbreak in poultry in 2003, birds in Dutch zoos were vaccinated twice with six weeks interval using a whole inactivated oil-adjuvanted vaccine, based on influenza virus A/chicken/Italy/473/99 (H7N1), with high homology to the field strain HPAI H7N7 A/chicken/Netherlands/1/03 (97.4% nucleotide and 98.7 % amino acid sequence identity). This resulted in the induction of antibody titres ≥ 40 (used as a correlate of protection in this study) in 81.5% of the vaccinated birds, with an overall GMT of 190. Birds of the taxonomic orders Anseriformes, Galliformes and Phoenicopteriformes showed higher GMT, and larger percentages developed a serum HI antibody titre ≥ 40 than those of the other orders. Furthermore, a decrease in antibody response with an increase in body weight > 1.5 kg was shown. The high agreement between post vaccination antibody titres determined by serum HI test (using the vaccine strain), and VN titres (using the field strain), was used as a further measure of immunogenicity. The broad efficacy demonstrated in a large variety of taxonomic orders illustrated the benefits of vaccination as an additional preventive measure against HPAI virus infection (Philippa et al., 2005). Vaccination against HPAI H5N1 in zoos In 2005, the Dutch zoos were the first to implement Decision 2005/744/EC to provide protection against the encroaching HPAI H5N1 subtype. Birds were vaccinated with an inactivated adjuvanted H5N2 vaccine with vaccine doses adapted to mean body weight per

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Vaccination of Non-domestic Avian Species

species, using data collected during the H7N1 vaccination campaign. The vaccine strain (A/duck/Pottsdam/1402/86) had a homology of 90% to the HA gene of the H5N1 field strain (A/turkey/Turkey/1/05) on the basis of nucleotide sequence (1530 base pairs, including basic cleavage site), and 92.4% on the basis of amino acids. Vaccination was safe, and proved immunogenic throughout the range of species tested, with some variations between and within taxonomic orders. After booster vaccination the overall homologous GMT to the vaccine strain, measured in 334 birds, was 190 (95% CI:152–236), and 80.5% of vaccinated birds developed a titre of ≥ 40. Titres to the HPAI H5N1 virus followed a similar trend, but were lower (GMT: 61 (95% CI: 49–76); 61%≥ 40) (Philippa et al., 2007). The breadth of the immune response was further demonstrated by measuring antibody titres against prototype strains of four antigenic clades of currently circulating H5N1 viruses. Antigenic distances to the prototype strains were determined using antigenic cartography (Smith et al., 2004). Antigenic cartography uses the antigenic properties of influenza viruses combination with epidemiological and genetic data, and is used to select virus strains for use as human pre-pandemic (H5N1) vaccine candidates (World Health Organisation (WHO), 2006). Influenza vaccines whose haemagglutinins are antigenically similar to circulating strains provide the highest level of protection from infection in humans (Subbarao, 1999). The birds clustered in two groups based on the breadth of antibody responses. Group 1 (Anseriformes, Galliformes, Phoenicopteriformes, Psittaciformes and Struthioniformes) showed a very broad response to vaccination, with predicted protection against future strains up to 12 antigenic units from the current vaccine. Group 2 (Ciconiiformes, Gruiformes, Pelecaniformes and Sphenisciformes) had low HI antibody titres against the prototype strain of the most antigenically distant clade (A/Indonesia/5/05). In 2006, a working group of Animal health and Welfare experts was established by the European Food Safety Authority (EFSA)(European Food Safety Authority (EFSA), 2007), to provide a scientific assessment of the preventive vaccination against avian influenza of H5 and H7 subtypes carried out in zoos in Member States (MS). The total number of birds vaccinated, as reported by 12 MS, was 44721. Individual data from 4718 birds (374 species from 19 taxonomic orders) were submitted. Not all of these could be used for every evaluation: pre-vaccination titres could be evaluated for 3039 birds; titres after first vaccination were evaluated for 1429 birds, and post-second vaccination titres for 2296 birds. Differences in vaccination schedules, doses and routes, differences in methodology and antigens used in the HI tests between laboratories (due to the absence of international reference standards, and the absence of inter-laboratory standardisation of methodology), the use of different vaccines1 in different taxonomic orders and the sometimes incomplete reporting of results, limited the evaluation of some of the data provided by EU MS. Cut-off points varied with laboratory, and titres considered a measure of adequate immune response were 8, 16, 32, 40, and 64. Most countries used dilution series starting at 4 or 8, therefore results were evaluated for titres 16 and 32 [documented surrogate markers for protection in chickens (Ellis et al., 2004; Office International d'Epizooties., 2004; Swayne et al., 2006; Tian et al., 2005)], and undetectable titres were regarded as 4 for calculation of GMT. In the absence of (and unfeasibility of obtaining) vaccination/challenge data in often endangered 1

Vaccine A: H5N9 (A/turkey/Wisconsin/68). Vaccine B: H5N2 (A/duck/Pottsdam/1402/86). Vaccine C: H5N2 (A/chicken/Mexico/232/94/CPA) Vaccine D: H5N9 (A/chickenk/Italy/22A/98). Vaccine E: H5N9 (A/chicken/ltaly/22A/H5N9/1998).

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zoo bird species, the evaluation had to be based on extrapolation of serological data from poultry and limited other bird species. The H5 vaccines registered for poultry in the EU showed differences in efficacy, measured as serum HI antibodies induced by two doses of vaccine (Table 1.). Three of the five vaccines evaluated induced relatively high GMT and high percentage seroconversion in the vast majority of vaccinated birds. The HI titres induced by vaccination showed marked differences between and within taxonomic orders. Both routes of vaccination (i.m. and s.c.) were effective in inducing HI serum antibody responses, and for most avian species the poultry dose was suitable. In some larger species higher doses adjusted to body weight, induced higher serum antibody titres. (e.g. for ostriches a 10-fold increase of the poultry dose (10 x 0.25ml). However, extremely high doses at a single site of injection (e.g. vaccination of ostriches with 10 ml of vaccine) appeared to have a negative effect on the induction of serum antibody titres, and induced local adverse reactions. There were indications that one vaccination was sufficient to induce high serum antibody titres in at least two taxonomic orders of birds (Anseriformes and Galliformes). However, a second vaccine administration ensured seroconversion in the majority of birds of most species. Limited data indicated that antibody titres persisted in several species for six months after vaccination. Adverse effects and mortality associated with vaccination were low and were mainly attributable to handling stress or trauma. Differences in adverse effects reported from different zoos highlight the importance of proper skilled handling. Longevity of antibody titres One year after vaccination with the H5N2 vaccine, birds in Dutch zoos were revaccinated with the same vaccine. Antibody titres one year after the initial two vaccinations and the effect of one booster vaccination at this time were evaluated. In Rotterdam Zoo, 72 previously vaccinated birds could be evaluated for the effect of one booster vaccination (Philippa et al, in press). For 44 birds, serum samples were available from 4 weeks after the initial two vaccinations the previous year, at the time of revaccination, and 4 weeks later. Birds which had been vaccinated with the H7 vaccine two years prior to the H5N2 revaccination were additionally tested for the presence of H7-specific antibodies. Serum antibody titres of the birds tested in Rotterdam Zoo had clearly decreased in one year time: while 80% of birds had a positive titre (≥ 8) and 68% a high positive titre (≥ 32) after 2 vaccinations, these figures were 61% and 30% respectively one year later. Four weeks after re-vaccination these figures increased to 93% and 77% respectively. Although a larger percentage of these 44 birds had a serum HI antibody titre ≥ 32 after re-vaccination, the GMT was lower than GMT after 2 vaccine doses one year before (88 vs 66). Birds from 4 out of 8 taxonomic orders did not have a GMT > 5 one year after vaccination, and only one order (Phoenicopteriformes had a GMT > 40. Four weeks after one revaccination 6/8 taxonomic orders tested had a GMT > 40. GMTs had decreased even further two years after vaccination, as was shown by the H7 specific serum HI antibody titres. As these birds were not revaccinated with an H7 vaccine, the effect of revaccination two years after the initial vaccinations is not known.

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Vaccination of Non-domestic Avian Species

Conclusions Bio-security measures remain the first line of protection of zoo birds against the introduction of AI viruses and should be implemented in zoos. These bio-security measures should include strict hygiene and quarantine measures, but should also exclude the possibility of introducing AI viruses through feed animals such as day old chicks, other poultry or their products. Continuous clinical monitoring of captive and wild birds in zoos should be practiced, for early detection of introduced viruses by wild birds, domestic birds, or their products. Strict bio-security measures will also reduce the risk of subsequent infection of wild birds from zoo birds. Wild birds have been documented to be susceptible to HPAI virus infection, and could potentially play a role in the spread of HPAI virus, although the majority of avian influenza viruses detected in free-ranging birds have been LPAI viruses. If biosecurity measures cannot sufficiently protect zoo birds from exposure to HPAI viruses coming from wild birds (based on an overall risk assessment which includes welfare aspects) vaccination with vaccines against HPAI of H5 and H7 subtypes authorised for use in poultry should be used to protect these zoo birds. In designing AI vaccination programmes and schedules for zoo birds, recent data on wild bird migration and prevalence of AI viruses should be taken into account. Vaccination against AI viruses of the H5 and H7 subtypes with current inactivated oil-adjuvanted poultry vaccines is safe and, in most taxonomic orders of zoo birds, effective in terms of inducing HI serum antibody titres. AI vaccines should be administrated in a way that elicits high HI antibody titres in the vast majority of the zoo birds vaccinated, i.e., by adjusting dose to average body weight. Although there are indications that one vaccination might suffice for some species, a second vaccine dose ensures high titres in the vast majority of species. Unless it is demonstrated that one vaccine administration is sufficient, two administrations are recommended. The H5 and H7 vaccines currently registered for poultry in the EU show differences in the performance in terms of HI response in zoo birds after two doses. There appears to be no difference in route of vaccination (s.c. or i.m.), so route can vary depending on the bird species to be vaccinated. In order to maintain high titres in the captive populations in zoological collections, annual revaccination seems to be required, as antibody titres decrease significantly in most taxonomic orders, and high titres are seen after a single annual booster dose. Mortality and adverse effects were low in all zoos evaluated in EU MS, and mainly attributed to handling stress and trauma. Zoos can, and should therefore try to minimise these losses in the execution of HPAI vaccination programmes. To minimise indirect losses due to decreased breeding results, AI vaccination during breeding seasons should be avoided whenever possible. Mortality due to catching and handling stress can be reduced by handling the birds less. Once the efficacy of a vaccination protocol has been validated for certain species using certain vaccines, measurement of post-vaccination HI serum antibody titres should no longer be mandatory by the EU. These birds will then only have to be handled for vaccination, and not 4 weeks later. Further research should be carried out to establish effective vaccination schedules, route, and dose regimen in different zoo bird species. This may, amongst others, lead to a reduction in the number of booster vaccinations needed in certain species. Novel generation vaccines which may be administered in the form of an aerosol (as is used in vaccination of poultry against Newcastle disease virus) may prove to be useful in non-domestic species, and would eliminate the need for handling the birds. The vaccination campaigns against HPAI virus have focused on protecting birds in zoological collections. However, a large number of mammalian species, including tigers and leopards, have also been documented with HPAI virus infection with recent H5N1 subtypes. There is currently no commercial vaccine available to protect mammals from HPAI H5N1 virus

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Vaccination of Non-domestic Avian Species

infection. A recombinant fowlpox-vectored vaccine expressing the H5 gene has been shown to produce high antibody titres against heterologous H5N1 virus antigen in cats after booster vaccination (Karaca et al., 2005), and may prove to be useful in prophylactic vaccination programs of mammals in the future. Until then, these animals have to be protected by excluding the introduction of AIV through raw poultry used as feed. The broad vaccine efficacy in the different avian taxonomic orders illustrates that vaccination against avian influenza is a useful tool for the protection of non-domestic avian species in zoos, which allows for an alleviation of confinement measures – and is therefore beneficial to the health and welfare of these birds. However, increased bio-security measures in combination with virological monitoring remain imperative in combating outbreaks of HPAI.

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Vaccination of Non-domestic Avian Species

References Banks,J., Speidel,E.S., Moore,E., Plowright,L., Piccirillo,A., Capua,I., Cordioli,P., Fioretti,A., Alexander,D.J., 2001. Changes in the haemagglutinin and the neuraminidase genes prior to the emergence of highly pathogenic H7N1 avian influenza viruses in Italy. Arch. Virol. 146:963-973. Brugh,M., Beard,C.W., Stone,H.D., 1979. Immunization of chickens and turkeys against avian influenza with monovalent and polyvalent oil emulsion vaccines. Am. J. Vet. Res. 40:165-169. Capua,I., Alexander,D.J., 2004. Avian influenza: recent developments. Avian Pathol. 33:393404. Capua,I., Terregino,C., Cattoli,G., Mutinelli,F., Rodriguez,J.F., 2003. Development of a DIVA (Differentiating Infected from Vaccinated Animals) strategy using a vaccine containing a heterologous neuraminidase for the control of avian influenza. Avian Pathol. 32:47-55. Capua,I., Terregino,C., Cattoli,G., Toffan,A., 2004. Increased resistance of vaccinated turkeys to experimental infection with an H7N3 low-pathogenicity avian influenza virus. Avian Pathol. 33:158-163. Ellis,T.M., Leung,C.Y., Chow,M.K., Bissett,L.A., Wong,W., Guan,Y., Malik Peiris,J.S., 2004. Vaccination of chickens against H5N1 avian influenza in the face of an outbreak interrupts virus transmission. Avian Pathol. 33:405-412. European Food Safety Authority (EFSA). Opinion of the Scientific Panel AHAW related with the vaccination against avian influenza of H5 and H7 subtypes as a preventive measure carried out in Member States in birds kept in zoos under Community approved programmes. 2007. Fouchier,R.A., Munster,V., Wallensten,A., Bestebroer,T.M., Herfst,S., Smith,D., Rimmelzwaan,G.F., Olsen,B., Osterhaus,A.D., 2005. Characterization of a novel influenza A virus hemagglutinin subtype (H16) obtained from black-headed gulls. J. Virol. 79:2814-2822. Harcourt-Brown,N.H. 1996. Bumblefoot. Pages 126-131 in Samour,J.H. editor. Avian Medicine. Harcourt Publishers Ltd, London. Higgins,D.A. 1996. Comparative immunology of avian species. Pages 149-205 in Davison,T.F., Payne,L.N., Morris,T.R. editors. Poultry Immunology. Carfax publishing co., Abingdon. Karaca,K., Swayne,D.E., Grosenbaugh,D., Bublot,M., Robles,A., Spackman,E., Nordgren,R., 2005. Immunogenicity of fowlpox virus expressing the avian influenza virus H5 gene (TROVAC AIV-H5) in cats. Clin. Diagn. Lab Immunol. 12:1340-1342. Karunakaran,D., Newman,J.A., Halvorson,D.A., Abraham,A., 1987. Evaluation of inactivated influenza vaccines in market turkeys. Avian Dis. 31:498-503. McMillian,M., Petrak,M., 1989. Retrospective study of aspergillosis in pet birds. J. Avian Med Surg211-215. Middleton,D., Bingham,J., Selleck,P., Lowther,S., Gleeson,L., Lehrbach,P., Robinson,S., Rodenberg,J., Kumar,M., Andrew,M., 2006. Efficacy of inactivated vaccines against H5N1 avian influenza infection in ducks. Virology.

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Munster,V.J., Baas,C., Lexmond,P., Waldenstrom,J., Wallensten,A., Fransson,T., Rimmelzwaan,G.F., Beyer,W.E., Schutten,M., Olsen,B., Osterhaus,A.D., Fouchier,R.A., 2007. Spatial, temporal, and species variation in prevalence of influenza A viruses in wild migratory birds. PLoS. Pathog. 3:e61. Munster,V.J., Wallensten,A., Baas,C., Rimmelzwaan,G.F., Schutten,M., Olsen,B., Osterhaus,A.D.M.E., Fouchier,R.A.M., 2005. Mallards and highly pathogenic avian influenza ancestral viruses, northern europe. Emerg. Infect. Dis. 11:1545-1551. Office International d'Epizooties. 2004. Avian Influenza.OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. Philippa,J., Baas,C., Beyer,W., Bestebroer,T., Fouchier,R., Smith,D., Schaftenaar,W., Osterhaus,A., 2007. Vaccination against highly pathogenic avian influenza H5N1 virus in zoos using an adjuvanted inactivated H5N2 vaccine. Vaccine 25:3800-3808. Philippa,J.D.W., Munster,V.J., Bolhuis,H., Bestebroer,T.M., Schaftenaar,W., Beyer,W.E., Fouchier,R.A.M., Kuiken,T., Osterhaus,A.D.M.E., 2005. Highly pathogenic avian influenza (H7N7): Vaccination of zoo birds and transmission to non-poultry species. Vaccine 23:57435750. Redig,P.T. 2000. Aspergillosis. Pages 275-287 in Samour,J. editor. Avian Medicine. Mosby, Philadelphia. Smith,D.J., Lapedes,A.S., de Jong,J.C., Bestebroer,T.M., Rimmelzwaan,G.F., Osterhaus,A.D., Fouchier,R.A., 2004. Mapping the antigenic and genetic evolution of influenza virus. Science 305:371-376. Sturm-Ramirez,K.M., Ellis,T., Bousfield,B., Bissett,L., Dyrting,K., Rehg,J.E., Poon,L., Guan,Y., Peiris,M., Webster,R.G., 2004. Reemerging H5N1 influenza viruses in Hong Kong in 2002 are highly pathogenic to ducks. J. Virol. 78:4892-4901. Subbarao,K., 1999. Influenza vaccines: present and future. Adv. Virus Res. 54:349-373. Swayne,D.E., Lee,C.W., Spackman,E., 2006. Inactivated North American and European H5N2 avian influenza virus vaccines protect chickens from Asian H5N1 high pathogenicity avian influenza virus. Avian Pathol. 35:141-146. Swayne,D.E., Perdue,M.L., Beck,J.R., Garcia,M., Suarez,D.L., 2000. Vaccines protect chickens against H5 highly pathogenic avian influenza in the face of genetic changes in field viruses over multiple years. Vet. Microbiol. 74:165-172. Tian,G., Zhang,S., Li,Y., Bu,Z., Liu,P., Zhou,J., Li,C., Shi,J., Yu,K., Chen,H., 2005. Protective efficacy in chickens, geese and ducks of an H5N1-inactivated vaccine developed by reverse genetics 6. Virology 341:153-162. USGS National Wildlife Health Center. Referenced reports of highly pathogenic avian influenza H5N1 in wildlife and domestic animals. USGS . 2008. 31-3-2009. van der Goot,J.A., Koch,G., de Jong,M.C., van,B.M., 2005. Quantification of the effect of vaccination on transmission of avian influenza (H7N7) in chickens. Proc. Natl. Acad. Sci. U. S. A. Webster,R.G., Webby,R.J., Hoffmann,E., Rodenberg,J., Kumar,M., Chu,H.J., Seiler,P., Krauss,S., Songserm,T., 2006. The immunogenicity and efficacy against H5N1 challenge of reverse genetics-derived H5N3 influenza vaccine in ducks and chickens. Virology 351:303311.

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World Health Organisation. Cumulative Number of Confirmed Human Cases of Avian Influenza A/(H5N1) Reported to WHO. World Health Organisation . 2009. World Health Organisation (WHO). Antigenic and genetic characteristics of H5N1 viruses and candidate H5N1 vaccine viruses developed for potential use as pre-pandemic vaccines. 2006.

Transmissible Diseases Handbook

XII. VACCINATION OF NON-DOMESTIC CARNIVORES: A REVIEW Joost Philippa, DVM Institute of Virology, Erasmus MC, Rotterdam, The Netherlands

Since the introduction of vaccinia by Jenner 200 years ago, vaccines have been the first line of defense in controlling infectious diseases in man and domestic animals. Large scale domestic dog vaccination programmes against canine distemper virus (CDV) and canine adenovirus (CAV) became possible in the late 1940’s, when egg-adapted vaccines became available on a commercial basis, followed by tissue culture adapted vaccines in the late 1950’s (Piercy 1961). Before this time these devastating diseases had to be controlled through quarantine and vigilance in capturing feral domestic animals (Dolensek et al. 1977). Historically there have always been two major types of vaccines based on the living state of the antigens. Modified live (MLV) vaccines use attenuated pathogens which reproduce in the vaccinate, thereby elliciting an immunologic response without causing disease (a “controlled” infection). The other major type of vaccine uses antigens that are non-living or inert – killed vaccine (KV). These vaccinations are preferred when safety information is not available, as they do not replicate and are therefore incapable of causing an infection. The proces of inactivation however may be damaging enough to modify immunogenicity, usually resulting in an immune response that is shorter in duration, narrower in antigenic spectrum, weaker in cell-mediated and mucosal immune responses, and possibly less effective in totally preventing viral entry (Murphy et al. 1999). Recent advances in immunology, molecular biology and biochemistry have allowed the construction of subunit vaccines based on viral or bactererial recombinants, peptides, or plasmid vectors, which may lead to safer, more efficacious vaccines that can also be used in exotic species. Vaccines cannot be absolutely guaranteed to provide protection against disease. The principal objective of vaccination is to mimic the protective immune response induced by natural infection, ie to elicit a high titre of neutralising antibodies of the appropriate class, IgG and/or IgA, directed against the relevant epitopes on the virion (Murphy et al. 1999). Immunity induced by vaccination or infection in domestic animals has been evaluated mainly by measuring the levels of serum antibodies. The immunologic response that provides protection against infectious agents involves a cellular and a humoral response. For certain infections (e.g. CDV, CAV, canine parvovirus (CPV), feline panleukopenia virus (FPV) or Borrelia burghdorferi) the humoral responses, although not the only mechanism involved, tend to correlate with level of protection from clinical disease, and therefore may be a useful indicator of the immune status. Other agents (eg, Bordetella bronchiseptica, canine coronavirus (CCoV), feline enteric coronavirus (FCoV), canine parainfluenzavirus (CPIV) and Chlamydia psittaci) all replicate and cause damage on mucosal surfaces, and might require a mucosal immune response for protection. As a consequence, serum antibody titres do not necessarily correlate with protection (Pfizer 1998). A high concentration of antibodies in an

XII.

Vaccination of Non-domestic Carnivores

animal implies that the animal is probably protected but also could indicate that it may not be possible to stimulate an additional immune response in that animal. Cattle vaccinated with baculovirus-expressed haemaglutinin (H) and fusion (F) proteins of Rinderpest virus (RPV) were not protected against disease caused by challenge exposure despite having detectable antibody titres (Bassiri et al. 1993). For some diseases, caused by persistent intracellular pathogens (viral diseases or intracellular bacteria) the neutralising antibodies and complement may play a less important role than the cellular immunity. Jones et al. (1993, 1997) demonstrated a lack of detectable antibody titre to CDV in ferrets and to peste des petites ruminants (PPRV) in goats after vaccination with a RPV recombinant vaccine based on fowlpox expressing the F and H genes. When these animals were challenged they survived, suggesting that protection against clinical disease may be cell mediated rather than humoral. A comparable study by Fisher et al (2003) using a DNA vaccine showed clear protection after challenge with CDV, with only limited virus neutralising antibody titres. High antibody concentrations do serve to inhibit the spread of virus between cells and thus promote host resistance (Tizard & Ni 1998). Infection with feline rhinotracheitis virus, as all herpesviruses, requires local and cell-mediated immunity, and one study suggests that there is no correlation with protective immunity and antibody titre (Johnson & Povey 1985). Later studies (eg Scott & Geissinger 1999, Dawson et al. 2001) use antibody titres as indicators of protective effect. In general it can be stated that theoretically, cell-mediated immunity is the most effective arm of the immune system in controlling, if not eliminating latent/persistent infections such as those caused by herpesviruses and retroviruses (Murphy et al. 1999). The usefulness of antibody titres to measure immunity is thus limited to only a few disease agents, and in order to obtain a more complete view of the immunologic status of an animal one therefore needs to look at the humoral and cellular responses. One should also keep in mind that an animal that has mounted an immune response after vaccination will possess memory T and B cells, which will remain for years after the antibody titre has declined. These memory cells rapidly differentiate during a subsequent infection into effector cells that can eliminate an infection before clinical signs appear, although the exact mechanism responsible for this longevity is unknown (Ahmed & Gray 1996). One can only know if the measured level of immunity is protective by challenging the vaccinated animal with the pathogen. One of the principal causes of vaccination failures in domestic dogs is maternal antibody interference. MLV vaccines differ in their ability to evade antibodies, and may sometimes be prevented from inducing an immunologic response in the vaccinate when the antibody level is high. The duration of passive immunity is directly correlated with the metabolic size of the animal. Therefore immunoglobulins will persist longer in a larger animal (Armstrong et al. 1942). The “window of vulnerability” during which the pup is vulnerable to infection with virulent virus, but unresponsive to attenuated vaccine virus (Pollock and Carmichael 1990), has been shown to range from 2-5 weeks for parvovirus infection in domestic dogs, but varies between pathogens and vaccinate species. Females close to parturition may be hyperimmunised with an inactivated parvovirus vaccine, so that high levels of maternal antibody delay the window of vulnerability until the offspring are older and better able to withstand the effects of parvovirus infection. The recent debate in veterinary medicine concerning issues related to vaccine efficacy and safety as well as duration of immunity induced by the currently available vaccines (Smith et al. 1995, Schultz et al. 1998, Kruth & Ellis 1998, Tizard & Ni 1998, McCaw et al. 1998, Gumley et al. 1999, Hustead et al. 1999, Twark et al. 2000) has resulted in the need for more objective and scientific data and an increase in research in domestic animals. In non-domestic animals however, there have been only few controlled studies of vaccination (Heerden et al. 1980, Halbrooks et al. 1981, Behlert et al. 1981, Barker et al. 1983, Montali et al. 1983, Green et al. 1984, Bush et al. 1985, Hoover et al. 1985, Paul-Murphy et al. 1985,

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Vaccination of Non-domestic Carnivores

Briggs et al. 1986, Tham et al. 1987, Follmann et al. 1988, Spencer et al. 1991, 1992, Goodrich et al. 1994, Harrenstien et al. 1995, Schubert et al. 1995, Williams et al. 1996, Henke et al. 1997, Kadoi et al. 1998, Bingham et al. 1999, Harthorn et al. 1999, Pare et al. 1999, Wimsatt et al. 1999, 2003, Maack et al. 2000, Blasco et al. 2001, Federhoff et al. 2001, Lambot et al. 2001, Maia et al. 2001, van Heerden et al. 1998, 2002). These are usually restricted to measuring the (humoral) immune response and extrapolating the data from those known in domestic animals, as subjecting (endangered) zoo animals to challenge infections is generally not an option. Challenge studies in dogs have shown a range of reported protective titres – these may differ due to the variety of techniques and standards used. In humans there is a general standardisation of assay methods to measure antibody titres. Non-standardisation of serologic tests makes comparisons between laboratories of questionable use (Luff et al. 1987): Table 1: Reported protective titres in domestic dogs: Virus Protective Reference titre CAV Cole et al. 1998 30 CDV

>2 20 24 30 >50 96 100

Gillespie et al. 1965; Ackerman & Siebel 1974. Gillespie et al. 1958, 1972;Gorham et al. 1966; Prydie 1966; Krakowka et al. 1978; Cooper et al. 1991. Jones et al. 1997. Gillespie 1966. Olson et al. 1988. Mc Caw et al. 1998. Appel 1969; Krakowka et al. 1975, Montali et al. 1983; Carmichael et al. 1999.

CPV

80

Olson et al. 1988; Carmichael et al. 1983, 1994, 1997; Appel et al. 1979; Pollock and Carmichael 1982; Meunier et al. 1985.

RV

20

Bunn et al. 1984

Twark et al. (2000) report that CDV titres above 5 are indicative of an adequate antibody response in domestic dogs, although level of protection is unknown, and recommend CDV revaccination when titres are below 32. Carmichael et al. (1983) demonstrated that dogs with a CPV titre below 100 were not protected. Titres of 200-800 were protective in some dogs, titres above 1600 appeared to be protective in all dogs. There is still no general consensus on how often domestic animals need to be revaccinated for most vaccines there is little information on the duration of immunity. It is, however, recognised that protective immunity to CDV following MLV vaccination is of long duration, perhaps even lifelong. For other viruses or components of combination vaccines this duration may not be of such long duration. Vaccines used in domestic animals are approved for use in specific animals under specific conditions, and any other use is therefore extra-label. Most of the vaccines are not approved for non-domestic species, therefore there is always a potential liability to such use (Bittle 1993). MLV vaccines have been designed to be minimally virulent, while retaining maximal immunogenicity in their domestic counterparts. When used in other species or delivered by another route the residual virulence may cause disease (Tizard et al. 1990). It is not unusual to observe side-effects such as elevated temperature, swelling, and irritation at the site of injection, or systemic anaphylactic reactions like hyperaemia, hypersalivation, or vomiting (Greenacre 2003) that may in some cases be severe (Karesh et al. 1983).

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Vaccination of Non-domestic Carnivores

Several viruses induce a supression of the immune system, and it is known that some attenuated virus strains may still be able to cause immunosuppression, e.g. MLV CPV (Krakowka et al 1982). Sometimes the individual vaccine strains are not detectably immunosuppressive, but when combined in a combination vaccine they may induce a suppression of blood lymphocyte counts (Phillips et al. 1989a). Enhanced virulence of canine distemper virus produced in canine cell-cultures has been reported when used in combination with CAV-1 and live CCV vaccines (Carmichael et al. 1983, Martin et al. 1985, Wilson et al. 1986), and may lead to encephalitis. Table 2: There are many examples of vaccine-induced disease: Virus CAV CDV

Species Domestic dog (Canis familiaris) Maned wolf(Chrysocyon brachyurus) Domestic ferret (Mustela putorius furo) Kinkajous (Potus flavus) Red panda (Ailurus fulgens) Black-footed ferrets (Mustela nigripes) African wild dogs (Lycaon pictus): Maned wolf Bush dog (Speothos venaticus) European mink (Mustela lutreola)

RV

FeLV FHV FPV

Gray fox (Urocyon cinereoargenteus) Fennec fox (Fennecus zerda) Domestic dog Domestic cat Skunk (Mephitis mephitis) Raccoon (Procyon lotor) Cheetah Domestic cats Pallas cat Felidae: Cheetah (Acinonyx jubatus)

Reference Appel et al. 1978. Thomas-Baker et al. 1985. AVMA 1966; Wimsatt et al. 2001. Kazacos et al. 1981. Bush et al. 1976; Itakura et al. 1979. Carpenter et al. 1976; Pearson et al. 1977. Mc Cormick et al. 1983; Montali et al. 1983; Durchfeld et al. 1990. Thomas-Baker et al. 1985. Mc Innes et al. 1992. Sutherland-Smith et al. 1997. Ek-Kommonen et al. 2003 Halbrooks et al. 1981. Mehren et al. 1984. Pedersen et al. 1978; Whetstone et al. 1984. Erlewein et al. 1981. Debbie et al. 1979. 1978. Briggs et al. 1986. North et al. 1978 Wallach & Boever 1983 Behlert et al. 1981 Crawshaw et al. 1996

Preventive medicine is especially important in the management of non-domestic animals for several reasons: the ability to mask or hide illness or distress as a means of survival means that by the time signs of disease are exhibited, the underlying disease condition may have advanced to a critical stage. Another reason is that the use of anaesthesia is required to perform a proper physical examination. Active immunisation is only one of the factors (eg nutrition, parasite control, hygiene) associated with preventive medicine. Recommendations for use in exotic mammals are generally based on tradition, anecdotal/personal experiences or taken from limited precise, published data. This has led to a plethora of differing opinions and therefore the use of many different protocols in zoological collections. In general, inactivated viral or bacterial vaccines are preferred for use in exotic animals. The type, serial number, and source of product should be recorded in the medical records (Joslin et al. 1990). Use of polyvalent vaccines containing unnecessary antigen (one the species is not susceptible to) should be avoided where possible. Animals with active clinical illness should not be vaccinated. In the event of a viral disease outbreak in an animal collection, all susceptible species should be vaccinated immediately and boostered 10-14 days later, regardless of age and last time of immunization (Phillips 1989). Some drugs, such as tetracycline, chloramphenicol, dapsone, clindamycin, griseofulvin, nalidixic acid and sulphamethoxypyradizine have been associated with an inadequate response to vaccination (Kruth – Ellis 1998). Vaccination should also be avoided in animals undergoing glucocorticoid therapy, although challenge studies have been performed which show that “immunosuppressive” doses given at the time of vaccination do not significantly affect the level of

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Vaccination of Non-domestic Carnivores

post-vaccinial immunity to canine distemper or rabies (Dhein et al. 1986). When using remote delivery systems one must be sure that a full dose has been delivered. Syringe darts may rebound quickly on impact and fail to deliver the dose required to elicit a satisfactory immune response (Aiello 1998). Any vaccination programme should also take the current local prevalence of the pathogen into account, upon which the decision can be made if vaccination is warranted. Table 3: Disease susceptibility CAV CDV + + Canidae + Felidae + + Ursidae + Procyonidae + Mustelidae + Viverridae + + Hyanidae + + Pinnipedia

FPV ++ + + + + + -

FeLV + -

FHV + -

FCaV + -

Rabies + + + + + + + +

Lepto + + + + + + + +

Toxo + ++ + + + ? ? +

Canine distemper virus (CDV) All families of the order Carnivora are susceptible to CDV, and it is among the most significant infections of many species. Christensen (1963) recommended vaccination against CDV in susceptible zoo animals, with the addition that vaccination of young animals at an earlier stage than in dogs is preferred. CDV vaccination is recommended in all members of the Canidae, Procyonidae and Mustelidae by all authors. There is no mention of vaccination of felids against CDV in literature until several outbreaks occurred among large cats in zoos and the wild (Fix et al. 1989, Appel et al. 1994, Munson et al. 1995, Wood et al. 1995, Roelke-Parker et al. 1996, Meehan et al. 1998, Cameron et al. 1998, Miller & Anderson 2000). Following this, the vaccination of large cats is mentioned as being possible, but not recommended – unless in high risk situations - as risk of exposure is generally low, and vaccination carries some risk (Junge et al. 1995, Kennedy-Stosskopf 1996, Aiello 1998, Miller & Anderson 2000, Woodford 2001, www.5tigers.org). The susceptibility of members of the Ursidae and Hyaenidae to canine distemper virus is deemed questionable by some authors, and therefore not recommended by these authors (Fraser 1991, Aiello 1998). Although clinical disease as a result of CDV infection is rare in ursids (Poston & England 1992), serologic surveys have shown the presence of CDV specific antibodies (Munson et al. 1995, Marsilio 1997, Dunbar et al. 1998, Maack et al. 2000). Clinical disease and presence of CDV specific antibodies has been documented in spotted hyaenas (Crocuta crocuta) (Montali et al. 1987, Alexander et al. 1995, Haas et al. 1996, Harrison et al. 2002) and a palm civet (Paguma larvata) (Machida et al. 1992), therefore vaccination is recommended in these species by other authors (Fowler 1986, Phillips 1989, Miller 1989, Burroughs 1992, Junge 1995, Woodford 2001). Miller & Anderson (2000) recommend vaccination of Viverridae, do not recommend vaccination of Ursidae, and omit Hyanidae from their article. In general, when vaccination is recommended, the same regime is used in the different families. Due to the possibility of immunosuppressive effects of multivalent vaccines, it is recommended that the distemper vaccine be given separately at a reasonable interval from the other components (Montali et al. 1994) A problem faced in the prophylaxis of distemper in exotic carnivores is the variation between and within species in their reaction to MLV vaccines, with possible lethal consequences. There are two distinct types of MLV CDV vaccine, one produced in avian cells and the other in canine kidney cells, and neither is safe for use in all potential target species. Chicken embryo-adapted live virus distemper vaccines attenuated for ferrets have protected them from challenge with an infective dose of virulent virus within 48 hours after vaccination (Baker

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Vaccination of Non-domestic Carnivores

et al. 1952). Fromm D (Solvay Animal Health Inc.) appeared to be safe and efficacious for use in maned wolves, bush dogs and fennec foxes (Montali et al. 1983). But avian cell vaccines have caused disease in mink, ferrets and foxes (Sutherland-Smith et al. 1997, Carpenter et al. 1976, Henke et al. 1997). Fervac-D, also of avian origin caused fatal disease in 1 of 8 red pandas (Montali et al. 1994). Vaccine of canine origin has been responsible for vaccine induced distemper in a large number of species (see table ). Non-domestic canine pups can be vaccinated with a MLV measles vaccine (Klös & Lang 1982, Frankenhuis & Visee 1985) - measles and canine distemper virus are antigenically closely related, and the measles virus is not neutralised by the maternal antibodies in 6 week old puppies of domestic dogs (Appel et al. 1984). The second and all subsequent vaccinations should be with a MLV CDV vaccine. Until 1983 the use of MLV is mentioned without warning (Klös & Lang 1982, Wallach & Boever 1983, Miller 1989). After this KV are recommended for use in exotic species (Fowler 1986, Gabrisch & Zwart 1987, Franke et al. 1989, Fraser 1991, Aiello 1998) even though the efficacy of KV has been questioned (Appel et al. 1984, Sikarski et al. 1991). Currently there are no killed CDV vaccines commercially available, because there is no demand for its use in domestic dogs, and the market for zoo animals is too small (Appel & Montali 1994). Between the different commercially available MLV vaccines there is a clear difference in vaccine efficacy, as has been demonstrated in domestic dogs (Appel et al. 1987, Rikula et al. 1996, Kommonen et al. 1997). Considering the high incidence of vaccine induced disease, Miller & Anderson (2000) mention that in some cases strict isolation may be preferable to vaccination. The large range of (highly susceptible) host species in zoos for which vaccination is recommended underpins the need for the production of a safe and efficacious vaccine. An experimental subunit vaccine incorporating the CDV F and H proteins into immuno stimulating complexes (ISCOM) has been developed for use in dogs and seals (de Vries et al. 1988, Visser et al. 1992), and proved to be capable of producing humoral and cellular immunity. Although the immunity achieved was not sterile (infection of the upper respiratory tract occurred), the vaccinated seals were protected from a potentially lethal challenge with phocid distemper virus (Visser et al. 1992). The ISCOM vaccine has since been used experimentally in several European zoos (Schaftenaar pers. comm), and its potential use in non-domestic felids has been proposed in special cases (Kennedy-Stoskopf 1996). An experimental adjuvanted killed CD vaccine produced by M.J.G. Appel has been used in red pandas and giant pandas in several zoos. The vaccine appeared to be safe and effacacious, but produced low titres with inadequate durability, requiring booster vaccinations two to three times annually (Montali et al. 1994). This vaccine is no longer produced. In Germany a small amount of inactivated vaccine is produced for use in zoos (Geyer and Matern pers. comm. 2001). In 1997 a recombinant virus vectored CDV vaccine was introduced (Stephenson et al. 1997) and tested for its safety and efficacy along with MLV components (Pardo et al. 1997). Following vaccination experiments in mice using a vaccine expressing the H and F protein in mice, a vaccine containing the H, F and nucleocapsid (N) constructs produced highly encouraging results in domestic dogs after challenge with virulent virus, although the mechanism of protection was not clear (Cherpillod et al. 2000). Recently a monovalent canarypox-vectored vaccine expressing the H and F surface antigens of CDV has become commercially available in the US, and its efficacy and safety in domestic ferrets has been demonstrated (Wimsatt et al. 2001). In black-footed ferrets (Mustela nigripes) x Siberian polecat (Mustela eversmanni) hybrids the use of this vaccine has produced a good immune response (Williams & Montali 1998), and has since been used and evaluated in a large number of exotic species (Montali pers. comm.). This vaccine, Purevax (Merial) is registered for use in domestic ferrets, but its off-label use in all susceptible species in zoos is recommended by the American Association for Zoo Veterinarians (AAZV) and Woodford (2001). In the European Union its use is not permitted. The main advantage of avipoxvectored vaccines is their safety, the foreign genes in the vector are expressed, inducing

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Vaccination of Non-domestic Carnivores

protective cellular and humoral immunity in the absence of the complete virus, and therefore eliminating the possibility of infection. Members of the Avipox genus are distinguished by their host restriction to avian species, eliminating the potential for dissemination of the vector within the vaccinate and therefore the spread of the vector to nonvaccinated contacts or the environment (Paoletti 1996). Virus replication is blocked at a late stage of morphogenesis in mammalian cells, importantly leaving the synthesis of viral proteins unimpaired (Sutter & Moss 1992). For unknown reasons, canarypox virus appears to be superior to fowlpox virus in the induction of immune responses in mammals (Moss 1996). Recent research has shown that replication-competent CAV-2 recombinant vaccines expressing the H and F antigens of CDV triggered both a significant seroconversion and protective immunity in puppies born to CDV and CAV-2 immune dams, thereby overcoming the passive immunity (Fischer 2002). CDV Vaccination regime recommended in domestic dogs: 

Initial at 6 weeks, repeat every 2 weeks until 12 weeks. Booster annually with MLV vaccine (Appel et al. 1999, Carmichael 1999, Dodds et al. 1999).

Table 4: CDV vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster All MLV Initial at 3-6 weeks, at 9 weeks biannually susceptible combination vaccine. Single vaccination after 12 weeks Mustelidae MLV 10 weeks * All KV or Initial at 5-6 weeks repeat every 2 annually susceptible MLV weeks until 15 weeks Mustelidae KV for Initial at 8 weeks, repeat after 2-3 * blackfoot weeks & initial Procyonidae KV Initial at 6-8 weeks, repeat every 2-3 annually weeks until 14 weeks All Initial at 8 weeks, repeat after 2-3 annually in affected susceptible weeks areas, otherwise every 2-3 years All KV Initial distemper/measles at 6-8 annually susceptible (unavaila weeks, at 12-14 weeks combination ble) vaccine. Procyonidae Initial at 8 weeks, repeat at 12 and 16 * or 18 weeks of age All Single dose after weaning im, monthly annually susceptible booster up to 4 months All Initial at 6-8 weeks, repeat every 2-3 annually susceptible weeks with a total of 3 vaccinations, in special cases (ie early weaning, ill juveniles, high probability of exposure to disease) extended to 4 or 5. All ISCOM 8, 11, 14 weeks of age annually susceptible or KV (unavaila ble)

Reference Klös & Lang 1982

Klös & Lang 1982 Wallach & Boever 1985 Wallach & Boever 1985 Wallach & Boever 1985 Frankenhuis & Visee 1985 Burroughs 1992

Paré et al. 1999 Fraser 1991, Aiello 1998 Phillips 1989, Cubas et al. 1996

Blijdorp

Parvovirus infections Infections with Feline Panleukopeniavirus (FPV) have been reported in captive felids since the 1930’s (Hindle & Findlay 1932, Goss et al. 1942), and have since been reported in a large number of feline species. All members of the Canidae, Felidae, Viverridae, Mustelidae (except for the domestic ferret, Parrish et al. 1987), Procyonidae and Ursidae are susceptible and vaccination is recommended. Although there is some difference of opinion on the susceptibility of Hyaenidae, vaccination is recommended (Junge 1995). Vaccination regimes

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Vaccination of Non-domestic Carnivores

reported by these authors are as those for the exotic cats. In 1947 a new viral gastroenteritis was observed in farmed mink (Mustela vison) in Canada (Schofield 1949). That virus, closely related biochemically and serologically to FPV, was later named mink viral enteritis, and currently probably occurs wherever mink are farmed (Pollock and Larsen 1990). In 1978 a virus infecting canine species emerged with clinical similarities to FPV infection in cats (Appel 1979). This virus, referred to as canine parvovirus type 2 (CPV-2) is closely antigenically and pathogenically related to FPV. Members of the Canidae, Mustelidae, Viverridae and Procyonidae are considered to be susceptible, but the virus has not got the ability to replicate in Felidae. In 1979 and around 1984 new antigenic types of CPV (CPV-2a and CPV-2b) emerged with an increased host range including both domestic and large cats (Steinel et al. 2000), although the large cats appear to have a higher susceptibility for these virus types (Steinel et al. 2001). For all susceptible species vaccination is recommended (Fraser 1991, Aiello 1998, Steinel et al. 2001). Other authors recommend vaccination of canidae only (Wallach & Boever 1985, Cubas et al. 1996). Canidae may be vaccinated with a FPV vaccine, and Felidae may be vaccinated with CPV vaccine due to the close antigenic relationship (Pollock & Carmichael 1983). In general, when vaccinating non-domestic species, the vaccine selected should be based on the similarity of the hosts (e.g. CPV in coyotes) or the known or probable virus susceptibility of the host to be vaccinated (e.g. FPV vaccine in raccoons) (Barker and Parrish 2001). It is recommended to vaccinate members of the Canidae with CPV-2 vaccines, and considering the high incidence of infections with CPV2a and CPV-2b in large cats an inactivated vaccine containing these types rather than CPV-2 would be desirable, but is not yet commercially available. There is debate over vaccine dose in relation to size of species, and the use of KV and MLV vaccines. Povey and Davis (1977) suggest that it is probably unnecessary to increase the dose or antigen mass when using KV in large species, due to the antigenic efficiency of parvoviruses and the use of adjuvants. Increased or double doses have been recommended for the vaccination of large cats >50 kg, which will subsequently develop higher antibody titres (Fowler 1977, Wallach - Boever 1983, Fraser 1991, Aiello 1998). In other studies antibody titre does not appear to be dose dependent (Wack 1991), large felids do not require larger doses than the 1ml dose used in domestic cats to develop protective titres (Bush et al. 1981). MLV vaccines probably overcome the consideration of animal size in relation to dose of vaccine (Povey and Davis 1977), although the increased potency may result in decreased safety. Some MLV vaccines reported to be safe for one species may be insufficiently attenuated for use in another species, and have caused vaccine-induced disease (Klös & Lang 1982, Wallach & Boever 1983, Frankenhuis & Visee 1985, Appel et al., Fraser 1991, Aiello 1998, Woodford 2001). Christensen (1963) reported abortive cases of depression and diarrhoea with recovery after 1 or 2 days’ symptoms following the introduction of systematic FPV vaccination. Visee (pers. comm. 1974 & 1975) reported clinical panleukopenia in 3 vaccinated Siberian tigers and 1 leopard (panthera pardus), of which 2 fatal. Therefore the use of KV is generally recommended in exotic species, although the production of antibodies is not as effective and the duration of immunity is shorter. KV FPV vaccine used in bush dogs (Speothos venaticus) did not protect them from infection – protective titres were not reached before 23 weeks of age during which period they were susceptible to infection (Janssen et al. 1982). Fowler (1977) reports that MLV vaccines have been used in different wild felid species, and are apparently safe. Many authors after this recommend the use of the highly antigenic KV vaccine (Klös & Lang 1982, Wallach & Boever 1983, Appel et al., Fraser 1991, Aiello 1999), or at least KV for the initial vaccinations which can then be boostered with MLV (Frankenhuis & Visee 1985, Fowler 1986, Gabrisch & Zwart 1987). Phillips (1989) recommends the use of KV without components for the feline respiratory viruses in mustelidea, procyonidae and viverridae. MLV vaccine should never be used in in pregnant felidae, foetal infection results in cerebellar hypoplasia with clinical ataxia in the kitten (Fowler 1986).

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Many different regimes have been tried, in which the timing of the initial vaccination is one of the main variables. Interference with primary immunisation by maternal antibodies is the commonest cause of “vaccine failure” in both domestic (Greene 1990) and non-domestic carnivores (Janssen et. al. 1982; Wack et al. 1993). Vissee (pers. com. 1975) reorted that after a change in the vaccination regime (6, 12, 26 weeks to 12, 16, 20 weeks) no clinical signs were reported following MLV vaccination, suggesting that maternal antibodies interfere with vaccination during the first 3 months. No controlled studies on the half-life of maternal antibodies to FPV in non-domestic species have been performed, and it should not be directly extrapolated from the 9.5 days determined in domestic cats (Scott 1970). Therefore the vaccination recommendations for non-domestic species are the same as those used in domestic species during maternal immunity. Combination vaccines containing MLV CDV, CAV-2, CPI-3, and FPV/CPV have been used in wild canidae without adverse effects, but with variable results, especially among non-canine species. Burroughs (1992) recommends its use in canidae, viverridae and hyaenidae. A multivalent KV vaccine (Fel-O-Vax PCT, Fort Dodge Lab Inc.) provides good antibody titres to the three major infectious viral diseases of domestic cats (panleukopenia, rhinotracheitis and calicivirus). The European Endangered Species Program (EEP) recommends 1 ml of Fel-O-Vax be used for boosters in adults; juveniles should be vaccinated at 8, 12, and 16 weeks, repeated at 6 months, and then given annual boosters. This is the same schedule used in domestic cats. Another regime recommended is repeated vaccinations with 2 week intervals for 3 injections or until 16 weeks of age (Woodford 2001). When cheetah cubs are vaccinated every 4 weeks from 8-16 weeks they may not develop and maintain protective antibody titres during their first year of life. When these cubs are vaccinated every 2 weeks from 8-16 weeks will develop protective titres. A booster may need to be given at the age of 40 weeks to insure that titres are maintained during the first year of life. There is much individual variation – boosters every 3 months may be warranted in high risk situations (Wack et al. 1993) - but a single annual vaccination may be adequate to maintain protective antibody titres (Wack 1991). FPV vaccination recommendation in domestic cats: 

8, 12, 16 weeks, booster at 6 months, then annually (Fel-O-Vax, Fort Dodge)

Table 5: FPV vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster All MLV 6, 12, 26 weeks susceptible All MLV or KV 12, 16, 20 weeks susceptible All KV or MLV 4, 8, 12, 16 weeks annually when using susceptible KV All 4, 8, 16 weeks (bi-)annually susceptible All 6-8 weeks, repeat every 2-3 weeks annually susceptible until 14 weeks of age All 1-2 doses KV, then after 3 weeks annually in affected susceptible MLV. MLV at 8, 11 weeks. areas, otherwise every 3 years All KV 2, 4, 8, 12, 16 weeks (colostrum annually (KV or MLV) susceptible deprived). 8, 12, 16 weeks (naturally weaned) All 3 x KV with 2 weeks interval, then 2 x (bi-)annually susceptible MLV with 4 weeks interval (colostrum deprived), 3 x MLV with 3-4 weeks interval starting at 7-8 weeks. Skunks Initial at 8-10 weeks, repeat after 3 weeks

Reference Visee pers comm 1974 Visee pers comm 1975 Fowler 1977 Klös & Lang 1982 Wallach & Boever 1983 Frankenhuis & Visee 1985 Fowler 1986

Gabrisch Zwart 1987

&

Gabrisch Zwart 1987

&

XII. All susceptible Felidae

All susceptible All susceptible All susceptible All susceptible All susceptible

KV, (unavailabl e)

KV KV

Vaccination of Non-domestic Carnivores

pregnant females should be boosted during gestation Initial KV at 3 months, MLV combination vaccine at 4 months

annually

Sacramento zoo 1990

Initial at weaning and repeated at least twice with 2 week intervals 8 and 12 weeks

annually

Junge 1995

annually

2 x with 2 weeks interval

6-12 months

8, 10, 12, 14, 16 weeks, repeat at 6 months 8, 11, 14 weeks

annually

Meltzer et al. 1996 Fraser 1991, Aiello 1998 Miller & Anderson 2000 Blijdorp

annually

annually

Table 6: CPV vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster All Initial at 8-12 weeks, 2nd vaccination 3 annually, every 6 susceptible weeks later months in endemic areas All As in domestic dogs susceptible All MLV Initial at 3 months annually susceptible All Initial at 8 weeks, repeat 3-4 times with susceptible 2-3 week intervals All 8, 11, 14 weeks annually susceptible

Reference Wallach Boever 1985

&

Frankenhuis & Visee, 1985 Burroughs 1992 Cubas et 1996 Blijdorp

al.

Infectious canine hepatitis / Canine Adenovirus-1 (CAV-1) Canidae, Ursidae and Mustelidae are susceptible to canine adenoviruses (Mann et al. 1980, Whetstone et al. 1988). Annual vaccination of Ursidae is recommended by some authors (Sacramento zoo 1990, Fraser 1991, Aiello 1998), depending on risk of exposure (Fowler 1986, Appel 1987) or not at all (Wallach & Boever 1983, Frankenhuis & Visee 1985, Junge et al. 1995, Miller & Anderson 2000). Burroughs (1992) recommends vaccination of Canidae and Hyaenidae. KV has been recommended for use in genera other than Canidae, as MLV may prove virulent (Klös & Lang 1982, Appel 1987). Currently there is no KV commercially available. The diluent portion of Adenomune-7 however, contains a killed CAV-2 antigen that may be used in highly susceptible species like maned wolves (Woodford 2001). Commercial combination vaccines that include CDV and MLV CAV-1 or CAV-2 are generally used. CAV1 is the causative agent of of infectious canine hepatitis, CAV-2 is the causative agent of respiratory disease. The two viruses are closely antigenically related, and vaccination with CAV-2 provides cross-protection against CAV-1 without causing adverse postvaccinial reactions like corneal opacity, common in domestic dogs after vaccination with MLV CAV-1, and reported in maned wolves by Thomas-Baker et al. (1985). Although clinically dramatic, the oedema usually resolves after a few days without consequence. Another reason was the diminition of postvaccinial encephalitis which was noted after subtitution of CAV-2 for CAV-1 in combination vaccines (Carmichael 1999). Table 7: CAV vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster All Initial at 3-6 weeks bi-annually susceptible All MLV or KV Initial at 11-12 weeks, repeat annually susceptible after 2-3 weeks All As in domestic dogs susceptible

Reference Klös & Lang 1982 Wallach & Boever 1983 Frankenhuis & Visee 1985

XII. All susceptible All susceptible Alls usceptible

MLV (caution) KV CAV-2 MLV MLV combination

Vaccination of Non-domestic Carnivores

As in domestic dogs Initial at weaning, monthly booster up to 4 months of age Initial at 3 months

annually or prior possible exposure annually annually

to

Appel 1987 Fraser 1991, Aiello 1998 Burroughs 1992

Herpesvirus infections All smaller exotic Felidae are susceptible to feline herpesvirus (FHV), larger Felidae have no or mild symptoms. Vaccinations currently available are KV or MLV, but KV is recommended. Vaccines commercially available (eg Fel-O-Vax) are usually in combination with other agents (eg FPV, FCaV, Chlamydia). Humoral titres are usually short-lived and boosters every 3 months may be required in high-risk situations (Wack et al. 1993). Klös and Lang (1982) recommended vaccination of Mustelidae and Viverridae, all other authors regard Felidae as the only susceptible animals. Fatal phocine herpesvirus type 1 (PhHV-1) infections have been reported in young and immunocompromised harbour seals in rehabilitation centres (Osterhaus et al. 1985, Borst et al. 1986, Gulland et al. 1997, Harder et al. 1997). An experimental sub-unit vaccine using the gB protein of PhHV-1 has been produced that proved to be safe and provided protective immunogenicity after challenge infection in domestic cats. Humoral and cellular immunogenicity was produced in harbour seals (Martina et al. 2001; 2003). Table 8: FHV vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster All Initial at 9 weeks, repeat after 3 after 6 months susceptible weeks. Adult cat twice with 2-3 weeks interval MLV Initial at 8 weeks, repeat every 3-4 annually, in high Felidae weeks until 14 weeks of age incident areas every 6 months As in domestic cats Felidae

Wallach & Boever 1983 Frankenhuis & Visee 1985 Gabrisch & Zwart 1987

Felidae

MLV

Naturally weaned: Initial at 7-8 weeks, repeat 3 times with 3-4 weeks interval. Not naturally weaned: Initial at 7-8 weeks, repeat 2 times with 3-4 weeks interval.

Felidae

MLV

Pregnant female should be boosted

Felidae

KV or MLV combinatio n combinatio n KV KV

Single dose at weaning. Monthly intervals until 4 months of age

annually

Repeat at 2 week intervals for 3 injections or 16 weeks of age 8, 11, 14 weeks

annually

Woodford 2001

annually

Blijdorp

All susceptible All susceptible

annually, endangered cats and cheetahs every 6-9 months. With KV booster, larger cats need 2-5 x dose annually

Reference Klös & Lang 1982

Sacramento Zoo 1990 Fraser 1991, Aiello 1998

Feline Leukemia virus (FeLV) Has been reported in exotic felids, and efficacy of a subunit vaccine has been demonstrated (Briggs et al. 1986, Citino ety al. 1988, Pettan et al. 1992). It is recommended to serologically test all Felidae for exposure. In 1986 vaccination with the then recently developed Leucocell vaccine was recommended by Gabrisch & Zwart. Due to the low prevalence in exotic cats, and the interference with serologic antibody screening, vaccination is not generally done (Citino et al. 1988, Phillips 1989, Junge et al. 1995, Kennedy-Stosskopf 1996), but may be done when there is close contact with feral cats (Miller & Anderson 2000).

XII.

Vaccination of Non-domestic Carnivores

Leptospirosis Canidae, Procyonidae, Ursidae, Mustelidae, and Pinnipedia are susceptible (Shotts 1981). A recent study in Rio de Janeiro Zoo, Brazil revealed an antibody prevalence of 37.7% of all animals tested – carnivores and non-carnivores - belonging to 10 families, out of which seroprevalence was most common in Canidae and Procyonidae (Lilenbaum et al. 2002). Raccoons, opossums, and rodents can act as reservoirs and concurrently transmit infection to zoo animals. Vaccination is highly serovar specific: the carnivores should be vaccinated with bacterins that contain immunogens against Leptospira interrogans serovar canicola and icterohaemorrhagiae. Vaccinations may influence the immune response in young animals vaccination of pups younger than 9-10 weeks of age is not recommended (Appel et al. 1999). Vaccination does not necessarily prevent shedding of the organism (Aiello 1998). Table 9: Leptospirosis vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster Initial at 12 weeks, repeat after 2-3 weeks annually Canidae Initial at 12-14 weeks annually Procyonidae / Mustelidae All As in domestic dogs susceptible Initial at 8 weeks, repeat at 12 and 16 weeks of annually Ursidae age All annually susceptible All 1-2 ml dose im or sc at 6-8 weeks, repeat after bi-annual susceptible 2 weeks bi-annual Canidae/ Procyonidae

Reference Wallach & Boever 1983 Wallach & Boever 1985 Frankenhuis & Visee 1985 Sacramento zoo 1990 Junge et al. 1995 Fraser 1991, 1998 Woodford 2001

Aiello

Rabies All mammal species are susceptible. Previously rabies vaccination was recommended in all circumstances (Klös & Lang 1982, Wallach & Boever 1983, Fowler 1986, Appel 1987). More recently recommendations are to vaccinate depending on location, risk of exposure, or possible outbreak (Fraser 1991, Junge et al. 1995, Aiello 1998). In areas where the incidence of rabies in local wildlife (skunks, raccoons, foxes) is high, vaccination is recommended. Local veterinary authorities should be conctacted regarding the legal aspects of extra-label vaccination, as some areas may have restrictions. Klös & Lang (1982) recommended use of MLV, except for younger animals. All other authors agree that nondomestic animals should be vaccinated with KV only. MLV vaccines have not been available commercially after rare occasions where the vaccine caused rabies-like disease in dogs. The efficacy of KV vaccines were questioned by Fowler (1986), but a KV vaccine (Imrab, Pitman Moore) has proven to be efficacious and safe, and been approved for use in domestic ferrets (Rupprecht et al. 1990). A great reduction in wildlife rabies has been accomplished by oral immunization (see below). Table 10: RV vaccination Regimes recommended for use in non-domestic species Species Vaccine Regime Booster All KV Initial at 4-6 months annually susceptible Felidae

KV

Initial at 6 months

annually

All susceptible All susceptible

KV

Initial at 6 months

annually

KV

Initial at 3-4 months

annually

Reference Wallach & Boever 1983; Junge et al. 1995 Wallach & Boever 1983 Sacramento zoo 1990 Fraser 1991, 1998

Aiello

XII. All susceptible

Initial at 3-6 months

Vaccination of Non-domestic Carnivores tri-annually, during outbreak more often

Burroughs 1992

Toxoplasmosis Many species of zoo mammals (but also birds) are highly susceptible to Toxoplasma Gondii infections, especially New World monkeys and marsupials, although members of the Felidae family are the definitive host, and the only animals that pass the oocyst in the faeces. This protozoan parasite has been reported in several species of non-domestic felidae, of which the Pallas cat appears to be highly susceptible (Dubey et al. 1988, Dorny et al. 1989, Ocholi et al. 1989, Swanson et al. 1999, Silva et al. 2001). There are a number of different vaccines, of which New Zealand S48 vaccine containing live tachyzoites, proved to be capable of inducing acute, fatal toxoplasmosis and is therefore considered to be unsuitable for use in macropods (Lynch et al. 1993). An alternate study suggested that oral vaccination with Hammondia Hammondi oocysts – a closely related non-pathogenic protozoan - may offer partial protection to the clinical effects, but does not prevent infection by T. gondii (Reddacliff et al. 1993). Recently an experimental recombinant FHV vaccine expressing the ROP2 antigen of T.gondii has been developed and proven efficacious in domestic cats (Mishima et al. 2002), but this is not yet commercially available. Vaccination of free-ranging wildlife Should free-ranging wildlife be vaccinated? It is an interference of the natural selection, and therefore a topic under discussion. Re-introduced or translocated animals will not have been challenged under natural conditions with the local pathogens when young (and maternal immunity is still present), and therefore need to be vaccinated to obtain a level of immunity against these pathogens. Recently IUCN guidelines for the reintroduction of captive animals into the wild have been published (Woodford 2001). Pre-release vaccination of animals to be rehabilitated or translocated should be considered by the veterinarian after evaluating the immunological status, and the risk of infection upon release into the destined area. When vaccinating wildlife it is of utmost importance to consider the fact that MLV vaccines may not be sufficiently attenuated for exotic species, and that vaccine induced disease or shedding of virulent virus may occur, thereby infecting free-living populations. Another problem faced is the difficulty to booster under field conditions, so that the level of immunity may not be sufficient. It is therefore recommended to complete the vaccination regime before release when possible. When this vaccination is carried out during the “preparation stage”, sufficient time is allowed to develop the required immunity and detect possible adverse effects. Following the phocine distemper (PDV) epidemic of 1988 (Osterhaus et al. 1988, Osterhaus & Vedder 1988, Kennedy et al. 1988) and the development of an experimental ISCOM (Osterhaus et al. 1989, Visser et al. 1992), all rehabilitated seals from the rehabilitation and research centre Pieterburen have been vaccinated before release. The duration of protective immunogenicity following this vaccination is unknown, and is intended to last for the duration of stay in the rehabilitation centre. The discussion of whether to start vaccinating the wild population flared up during the recent PDV epidemic of 2002, but was not considered a viable option (Trilateral seal expert meeting 2002, DEFRA 2002). Vaccination of seals with a MLV vaccine is contraindicated (Kennedy et al. 2001). Vaccination of endangered species to infectious diseases may aid in the survival of these species. African wild dog (Visee et al. 2001) and black-footed ferret (Williams & Thorne 1999) conservation projects have been severely affected by CDV outbreaks, and much work

XII.

Vaccination of Non-domestic Carnivores

is done on the production of a safe and efficacious vaccine (Montali et al. 1998, Visee 2001, van de Bildt et al. 2002). Vaccination of a wild population of Mediterranean monk seals (Monachus monachus) was considered during a morbillivirus epizootic in the mediterranean in 1992 (Osterhaus et al. 1992). During the PDV outbreak among Northern European Harbour seals in the summer of 2002, vaccination was also considered in the UK, but not deemed feasible (DEFRA 2002). The zoonotic and economic aspects of rabies infection have resulted in prophylactic immunization of domestic dogs and the eradication of canine rabies in several countries (Bögel 1982). Following this achievement attention was focused on free-ranging vector species, which were much more difficult to vaccinate. The development of a MLV vaccine which vaccinated foxes by the oral route (Baer 1971) was a major step in the right direction, which proved its value when an advancing epidemic was stopped by the vaccination zone (Steck et al. 1982). This vaccine has since been replaced by a vaccinia recombinant vaccine (Pastoret 1997), which has proven to be efficacious, and safe for the target species, the fox, as well as for numerous non-target species (Black et al 1993). To be effective the vaccine must be brought into contact with oral and /or pharyngeal mucosa in a sufficient number of target animals via bait. Factors affecting uptake: size, texture, shape of bait and vaccine container, as well as physical and chemical characteristics. One should take into account bait density, distribution method (manual, aerial or both), sequence and frequency of bait distribution, season, selection of specific baiting areas, strategies for expansion of baiting areas and the overall duration of vaccination campaigns (Rupprecht et al. 2001). Research is being conducted on the development of ideal baits and baiting systems for different species (Steelman et al 1998, Knobel et al 2002, Linhart et al 2002). Oral vaccination may be used in the eradication of other diseases of wildlife in the future. Wimsatt (1999) reported potential oral efficacy of an experimental canarypox vectored recombinant CDV vaccine in a preliminary study, and this oral efficacy was demonstrated in Siberian polecats (Wimsatt 2003) but more research is needed to evaluate its efficacy and safety in other species.

XII.

Vaccination of Non-domestic Carnivores

References Ahmed, R., Gray, D. 1996. Immunologic memory and protective immunity: understanding their relation. Science 272(5258):54-60. Aiello, S.E. (ed) 1998. Vaccination of exotic mammals. In: Merck’s Veterinary Manual 8th edition: 1427-1431. Alexander, K.A., Kat, P.W., Frank, L.G., House, C., Appel, M.J.G. 1995. Evidence of canine distemper virus infection among free-ranging spotted hyaenas (Crocuta crocuta) in the Masai Mara, Kenya. Journal of Zoo and Wildlife Medicine 26(2):201-206. American Veterinary Medical Association, 1966. Conclusions and recommendations of the panel of the symposium on canine distemper immunisation. Journal of the American Veterinary Medical Association 149(5):714-8. Anand, R., Malik, P.K., Prasad, G., Mukherjee, S.K. 2000. Serosurvey for viral infections in endangered captive Asiatic lions and sympatric nondomestic carnivores from Western India. Proceedings of the American Association of Zoo Veterinarians: 443. Anonymous. Preventive medicine programme Sacramento Zoo 1990 Anonymous. 1978. Vaccine induced rabies in a raccoon. CDC Rabies Surveillance, Annual Summary 1978; 4 Appel, M.J.G. 1969. Pathogenesis of canine distemper. American Journal of Veterinary Research 30(7):1167-1182. Appel, M.J.G. 1978. CAV-2: a replacement for canine hepatitis vaccine. Norden News 53:46. Appel, M.J.G., Shek, W.R., Shesberadaran, H., Norrby, E. 1984. Measles virus and inactivated canine distemper virus induce incomplete immunity to canine distemper. Archives of Virology 82(1-2):73-82. Appel, M.J.G. 1987. Virus Infections of Carnivores, Amsterdam; Elsevier Science Publications BV. Appel, M.J.G., Scott, F.W., Carmichael, L.E. 1979. Isolation and immunization studies of a canine parvo-like virus from dogs with haemorrhagic enteritis. Veterinary Record 105(8):156-159. Appel, M.J.G., Yates, R.A., Foley, G.L., Bernstein, J.J., Santinelli, S., Spelman, L.H., Miller, L.D., Arp, L.H., Anderson, M., Pearce-Kelling, S., Summers, B.A. 1994. Canine distemper epizootic in lions, tigers, and leopards in North America. Journal of Veterinary Diagnostic Investigations 6(3):277-288. Appel, M.J.G., Montali, R.J. 1994. canine distemper and emerging morbillivirus diseases in exotic species. Proceedings of the American Association of Zoo Veterinarians Annual Conference:336-339. Appel, M.J.G., Schultz, R.D., 1999. Forty years of canine vaccination. Advances in Veterinary Medicine 41: 309-324. Armstrong, W.H., Anthony, C.H. 1942. An epizootic of canine distemper in a zoological park. Cornell Veterinarian 32:286. Baer, G.M., Abelseth, M.K. and Debbie, J.G. 1971. Oral vaccination of foxes against rabies. American Journal of Epidemiology 93:487-490. Baker GA et al. 1952. Immune response of ferrets to vaccination with egg-adapted distemper virus: I. Time of development of resistance to virulent distemper virus. Veterinary Medicine 47:463. Barker, I.K., Povey, R.C., Voigt, D.R. 1983. Response of mink, skunk, red fox and raccoon to innoculation with mink virus enteritis, feline panleukopenia and canine parvovirus and prevalence of antibody to parvorvirus in wild carnivores in Ontario. Canadian Journal of Comparative Medicine 47(2):188-197. Barker, I.K., and Parrish, C.R. 2001. Parvovirus infections. In: Williams, E.S. and Barker, I.K. (eds): Infectious diseases of wild mammals 3rd edition. Iowa State University Press:131146.

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Barrett, T. 1999. Morbillivirus infections, with special emphasis on morbilliviruses of carnivores. Veterinary Microbiology 69(1-2):3-13. Bassiri, M. Ahmad, S., Giavedoni, L., Jones, L., Saliki, J.T., Mebus, C., Yilma, T. 1993. Immunological responses of mice and cattle to Baculovirus-expressed F and H proteins of Rinderpest virus: lack of protection in the presence of neutralizing antibody. Journal of Virology 67(3):1255-1261. Behlert, O., Behlert, C. 1981. Antikörperbesimmungen in einem Felidbestand nach vakzination mit zwei verschiedenen Panleukopenie-Impfstoffen. Verh ber Erkr Zootiere, 23:47-51. Bildt, van de M.W.G., Kuiken, T., Visee, A.M., Lema, S., Fitzjohn, T.R, Osterhaus, A.D.M.E. 2002. Distemper and its effect on African wild dog conservation. Emerging infectious diseases 8(2):211-213. Bengis, R.G., Kock, R.A., Fischer, J. 2002. Infectious animal diseases: the wildlife/livestock interface. Revue Scientifique et Technique Office International des Epizooties 21(1):53-65. Bingham, J., Schumacher, C.L., Hill, F.W., Aubert, A. 1999. Efficacy of SAG-2 oral rabies vaccine in two species of jackal (Canis adustus and Canis mesomelas) Vaccine 17(6):551558. Bittle, J.M. 1993: use of vaccines in exotic animals. Journal of Zoo and Wildlife Medicine 24(3):352-356. Black, D.N., Pastoret, P.P., Gielkens, A.L.J., Skinner, M.A., Gaskell, R.M. 1993. Assesment of environmental impact from the use of live recombinant virus vaccines. EC-sponsored research on safety of genetically modified organisms: http://europa.eu.int/comm/research/quality-of-life/gmo/08-vaccines/08-07-project.html Blasco, E., Lambot, M., Barrat, J., Cliquet, F., Brochier, B., Renders, C., Krafft, N., Bailly, J., Munier, M., Pastoret, P.P., Aubert, M.F. 2001. Kinetics of humoral immune response after rabies VR-G oral vaccination of captive fox cubs (Vulpes vulpes) with or without maternally derived antibodies against the vaccine. Vaccine 19(32):4805-15. Bögel, K., Andral, L., Beran, G., Schneider, L.G., Wandeeler, A. 1982. Dog rabies elimination: a trend analysis and programme proposal prepared by a WHO working group. International Journal of Zoonoses 9(2):97-112. Borst, G.H.A., Osterhaus, A.D.M.E., Walvoort, H.C., Reijnders, P.J.H., Van der Kamp, J.S. 1986. An outbreak of herpesvirus infection in harbour seals (Phoca vitulina). Journal of Wildlife Diseases 22(1):1-6. Briggs, M.B., Ott, R.L. 1986. Feline leukemia infection in a captive cheetah and the clinical and antibody response of six captive cheetahs to vaccination with a subunit feline leukemia virus vaccine. Journal of the American Veterinary Medical Association 189(9):1197-9. Briggs, M.B., 1986. Possible feline leukemia virus infection in a captive cheetah and the response of captive cheetahs to feline leukemia vaccination. Proceedings of the American Association of Zoo Veterinarians. Budd, J. 1981. Distemper. In: Davis, Karstad and Trainer (eds), Infectious diseases of wild mammals 2nd edition. The Iowa State University press, Ames, IA:31-44. Bunn, T.O., Ridpath, H.D., Beard, P.D. 1984. The relationship between rabies antibody titres in dogs and cats and protection from challenge. Rabies Information Exchange 11: 9-13. Burroughs, R.E.J. 1992. Care of carnivores in captivity. In: McKenzie, A.A (ed): The capture and care manual. Wildlife decision support services, South Africa. Bush, M., Montali, R.J., Brownstein, O., James, A.E., Appel, M.J.G. 1976; Vaccine-induced canine distemper in a lesser panda. Journal of the American Veterinary Medical Association 169(9):959-960 Bush, M., Povey, R.C., Koonse, H. 1981. Antibody response to an inactivated vaccine for rhino- tracheitis, caliciviral disease, and panleukopenia in non-domestic felids. Journal of the American veterinary Medical Association 179(11):1203-05. Bush, M., Montali, R.J., Reid, F.L., sumner, J.W., Yager, P.A., Phillips, L.G. 1985. Antibody response in zoo animals to a killed rabies vaccine. Proceedings of the American Association of Zoo Veterinarians Annual Conference:30.

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Cameron, K.N., Campbell, M.K., Long, P.H., Munson, L., Chambers, A. Kroll, J.L., Summers, B.A. 1998. Canine distemper in a captive Indochinese tiger (Panthera tigris corbetti). Proceedings of the American Association of Zoo Veterinarians:201-202 Carmichael, L.E., Joubert, J.C., Pollock, R.V.H. 1983. A modified live canine parvovirus vaccine. II. Immune response. Cornell Veterinarian 73: 13-29. Carmichael, L.E., Schlafer, D.H., Hashimoto, A. 1994. Minute virus of canines (MVC, canine parvovirus type-1): pathogenicity for pups and seroprevalence estimate. Journal of Veterinary Diagnostic Investigation. 6(2):165-74. Carmichael, L.E. 1999. Canine vaccines at a turning point-a personal perspective. Advances in Veterinary Medicine 41:289-307. Carpenter, J.W., Appel, M.J.G., Erickson, R.C., Novilla, M.N. 1976; Fatal vaccine-induced distemper virus infection in black-footed ferrets. Journal of the American Veterinary Medical Association 169(9):961-964. Castro, A.E. 2001. Other Herpesviruses. In: Williams E.S., Barker, I.K. (eds): Infectious Diseases of Wildlife 3rd edition. Iowa State University Press:175-178. Chaffee, P.S. 1972. An outbreak of fox encephalitis in vaccinated wild canidae. Journal of Zoo Animal Medicine 3:5. Chalmers, W.S.K., Baxendale, W. 1994. A comparison of canine distemper vaccine and measles vaccine for the prevention of canine distemper in young puppies. Veterinary Record 135:349-353. Cherpillod, P. Tipold, A., griot-Wenk, M., Cardozo, C., Schmid, I, fatzer, R,. Schobesberger, M., Zurbriggen, R., Bruckner, L., Roch, F., VandeVelde, M., Wittek, R., Zurbriggen, A. 2000. DNA vaccine encoding nucleocapsid and surface proteins of wild type canine distemper virus protects its natural host against distemper.Vaccine 18(26):29927-2936. Christensen, N.O. 1964. Some observations on diseases in carnivores in the Copenhagen zoo. Proceedings 5th International Symposium on Diseases in Zoo-Animals 1963. Tijdschrift voor Diergeneeskunde 89 suppl. I. Citino, S.B. 1986. Transient FeLV viremia in a clouded leopard. Journal of Zoo Animal Medicine 17:5-7. Citino, S.B. 1988. Use of a subunit feline leukemia virus vaccine in exotic cats. Journal of the American Veterinary Medical Association 192(7):957. Cole, R. 1998. Rethinking canine vaccinations. Veterinary Forum jan:52-57. Collins, J.E., Leslie, P., Johnson, D., Nelson, D., Peden, W., Boswell, R., Draayer, H. 1984. Epizootic of adenovirus infection in American black bears. Journal of the American Veterinary Medical Association 185(11):1430-1432. Cooper PE et al 1991. Comparaison de léfficacite des differents vaccins du chien, utilises sous forme monovalente ou associee, par evaluation des responses serologiques et apres epreuves virulentes 12, 22 et 26 mois apres vaccination. Bulletin Mensuel de la Societe Veterinaire Pratique de France 75:131-152. Crawshaw, G.J., Mehren, K.G., Pare, J.A. 1996. Possible vaccine induced viral disease in cheetahs. Proceedings of the American Association of Zoo Veterinarians Annual conference:557-560. Cubas, Z.S. 1996. Special challenges of maintaining wild animals in captivity in South America 1996. Review of Science and Technology 15(1):267-87. Curlee, J.F. 1999. Cross-species vaccination in wild and exotic animals. Advances in veterinary Medicine 41:551- 556. Davis, J.W., Karstad, L.H., Trainer, D.O. 1981. Infectious Diseases of Wild Mammals 2nd edition. Iowa State University Press. Dawson, S., Willoughby, K., Gaskell, R.M., Wood, G., Chalmers, W.S. 2001. A field trial to assess the effect of vaccination against feline herpesvirus, feline calicivirus and feline panleukopenia virus in 6 week old kittens. Journal of Feline Medicine and Surgery 3(1):1722. Debbie, J.G. 1979. Vaccine induced rabies in a pet skunk. Journal of the American Veterinary Medical Association 175(4):376-7).

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Pettan, K.C.B., Jessup, D.A., Lowenstine, L.J., Pedersen, N.C. 1992. Feline leukemia virus infection in a free ranging cougar. Proceedings of the American Association of Zoo Veterinarians: 136-138. Pfizer 1998. Duration of Immunity in Companion Animals after natural infection and vaccination. www.pfizer.com Phillips, L.G. 1989. Preventive Medicine in non-domestic carnivores. In: Kirk RW (ed): Current Veterinary Therapy X– Small Animal Practice. Philidelphia; Saunders Co: 727-734. Phillips, T.R., Jensen, J.L., Rubino, M.J., Yang, W.C., Schultz, R.D. 1989a. Effects of vaccines on the canine immune system. Canadian Journal of Veterinary Research 53(2):154-160. Piercy, S.E. 1961. An appraisal of the value and method of use of living attenuated canine distemper vaccines. Veterinary Record 73:944-949. Pollock, R.V.H. & Carmichael, L.E. 1990. Enteric viruses. In: Green CE (ed): Infectious diseases of the dog and cat. Saunders, Philadelphia: 226-283. Pollock, R.V.H. & Carmichael, L.E. 1983. Use of modified live feline panleukopenia virus vaccine to immunize dogs against canine parvovirus. American Journal of Veterinary Research 44(2):169-175. Pollock, R.V.H. & Carmichael, L.E. 1982. Maternally derived antibody to canine parvovirus: transfer, decline and interference with immunization. Journal of the American Veterinary Medical Association 180(1):37-43. Porter, D.D., and Larsen, A.E. 1990. Mink parvovirus infections. In: Thijssen, P. (ed): CRC handbook of parvoviruses, vol 2, Boca Raton, Fl: 87-101. Poston, R.P., England, J.J. 1992. Veterinary Diagnostic Virology, Mosbey Year Book, St Lewis: 135-138. Povey, R.C., Davis, E.V. 1977. Panleukopenia and respiratory infections in wild felids. In: The World’s cats, Eton, R.L. (ed). The carnivore research institute, Burke museum of seattle, WA 3:120-128. Prydie J. 1966 Persistence of antibodies following vaccination against canine distemper and the effect of re-vaccination. Veterinary Record 78(14):486-8. Reddacliff, G.L., Parker, S.J., Dubey, J.P., Nicholls, P.J., Johnson, A.M., Cooper, D.W. 1993. An attempt to prevent acute toxoplasmosis in macropods by vaccination with Hammondia hammondi. Australian Veterinary Journal 70(1): 33-35. Rikula, U., Nuotio, L., Sihvonen, L. 2000. Canine distemper virus neutralising antibodies in vaccinated dogs. Veterinary Record 147(2):598-603 Rikula, U, Sihvonen, L., Voipio, H.M., Nevalainen, T. 1996. Serum antibody response to canine distemper virus vaccines in beagle dogs. Scandinavian Journal of Laboratory Animal science 23(1):31-33 Roelke-Parker, M.E., Munson, L., Packer, C., Kock, R., Cleaveland, S., Carpenter, M., O’Brien, S.J., Pospischil, A., Hofmann-Lehmann, R., Lutz, H., et al. 1996. A canine distemper virus epidemic in Serengeti lions (Panthera leo). Nature 379(6564):441-445. Rupprecht, C.E., Gilbert, J., Pitts, R., Marshall, K.R., Koprowski, H. 1990. Evaluation of an inactivated rabies vaccine in domestic ferrets. Journal of the American Veterinary Medical Association 196(10):1614-1616. Rupprecht, C.E. 1996. International meeting on research advances and rabies control in the Americas. Emerging Infectious diseases 2(3):243. Rupprecht, C.E., Stohr, K., Meredith, C. 2001. Rabies. In: Williams, E.S., Barker, I.K: Infectious diseases of wild mammals, 3rd edition, Iowa State University Press: 3-36. Schofield, F.W. 1949. Virus enteritis in mink. North American Veterinarian 30:651-654. Schroeder, H.D. 1974. Latest developments in immunoprophylaxis for zoo animals. Verhandlungsbericht XVI Internationalen Symposiums Ueber Erkrankungen Zootiere, Erfurt. Berlin: Akademie Verlag :155-165. Schubert, C.A. 1995. Effect of CDV on an urban raccoon population. Proceedings of the American Association of Zoo Veterinarians Annual Conference:493. Schultz, R.D. 1998. Current and future canine and feline vaccination programs. Veterinary Medicine 93(3):233-254.

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Scott, F.W. & Geissinger, C.M. 1999. Long-term immunity in cats vaccinated with an inactivated trivalent vaccine. American Journal of Veterinary Research 60(5):652-8. Scott, W.A., Csiza, C.K., Gillespie, J.H. 1970. Maternally derived immunity to feline panleukopenia. Journal of the American Veterinary Medical Association 156(4): 439-453. Scott, W.A. 1979. Use of vaccines in exotic species. Veterinary Record 104(9):199. Sedgewick, C.J. 1966. An introduction to zoo practice. Modern veterinary Practice 37:39. Sedgewick, C.J., Young, W.A. 1968. Distemper outbreak in a zoo. Modern Veterinary Practice 49:39. Shotts, E.B. 1981. Leptospirosis. In: Davis JW, Karstad LH, Trainer DO (eds): Infectious diseases of wild mammals, second edition. Ames, IA; Iowa State University press:97-101. Sikarski, J.G., Lowrie, C., Kennedy, F., Brady, G. 1991. Canine distemper in a vaccinated red panda (Ailuris fulgens fulgens). Proceedings of the Association of American Zoo Veterinarians Annual Conference:292-3. Silva, J.R., Ogassawara, S., Marvulo, M.F.V., Ferreira-Netto, J.S., Dubey, J.P. 2001. Toxoplasma gondii antibodies in exotic wild felids from Brazilian zoos. Journal of Zoo and Wildlife Medicine 32(3):349-351. Smith, C.A. 1995. Are we vaccinating too much? Journal of the American Veterinary Medical Association 207(4):421-425. Spencer, J.A., Burroughs, R. 1991. Antibody response of captive cheetahs to modified live feline virus vaccine. Journal of Wildlife Diseases 27(1):578-83. Spencer, J.A., Burroughs, R. 1992. Antibody response to canine distemper vaccination in African wild dogs. Journal of Wildlife Diseases 28(3):443-4. Spencer, J.A. 1991. Antibody response to modified-live canine adenovirus vaccine in African hunting dogs (Lycao pictus). Zentralblatt fur Veterinarmedizin 38(6):477-9. Spencer, J.A., Burroughs, R. 1992. Decline in maternal immunity and antibody response to vaccine in captive cheetah (Acinonyx jubatus) cubs. Journal of Wildlife Diseases 28(1):102104. Steck, F., Wandeler, A., Bichsel, P., Capt, S., Schneider, L. 1982. Oral immunization of foxes against rabies. A field study. Zentralblatt Veterinaermedizin 29(5):372-396. Steelman HG, Henke SE, Moore GM. 1998. Gray fox response to baits and attractants for oral rabies vaccination. Journal of Wildlife Diseases 34(4):764-70 Steinel, A., Munson, L., van Vuuren, M., Truyen, U. 2000. Genetic characterization of feline parvovirus sequences from various carnivores. Journal of General Virology 81(Pt 2):345-50. Steinel, A., Parrish, C.R., Bloom, M.E., Truyen, U. 2001. Parvovirus infections in wild carnivores. Journal of Wildlife Diseases 37(3):594-607. Stephensen, C.B., Welter, J., Thaker, S.R., Taylor, J., Tartaglia, J., Paoletti, E. 1997. Canine distemper virus (CDV) infection of ferrets as a model for testing morbillivirus vaccine strategies: NYVAC- and ALVAC-based CDV recombinants protect against symptomatic infection. Journal of Virology 71(2);1506-1513. Sutherland-Smith, M.R., Rideout, B.A., Mikolon, A.B., Appel, M.J.G., Morris, P.J., Shima, A.L., Janssen, D.J. 1997. Vaccine induced canine distemper in European mink (Mustela lutreola). Journal of Zoo and Wildlfe Medicine 28:312-319. Sutter, G. Moss, B. 1992. Nonreplicating vaccinia vector efficiently expresses recombinant genes. Proceedings of the National Academy of Sciences, USA 89:10847-10851. Swanson 1999. Toxoplasmosis and neonatal mortality in Pallas’ cats: a survey of North American zoological institutions. Proceedings of the American Association of Zoo Veterinarians. Taylor, D.R., Martin, R.B. 1987. Effects of veterinary fences on wildlife conservation in Zimbabwe. Environmental management 11:327-334. Tham, K.M., Studdert, M.J., 1987. Antibody and cell-mediated response to feline herpesvirus1 following inactivated vaccine and challenge. Journal of Veterinary Medicine. 34:585-597. Thomas-Baker, B. 1985. Vaccination induced distemper in maned wolves, vaccination induced corneal opacity in a maned wolf. Proceedings of the American Association of Zoo Veterinarians Annual Conference:53.

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Tiger information centre: www.5tigers.org/zoos/husbandry_manual/husman2a.htm Tizard, I. 1990. Risks associated with use of live vaccines. Journal of the American Veterinary Medical Association 196(11):1851-1858. Tizard, I., Ni, Y. 1998. The use of serologic testing to assess immune status of companion animals. Journal of the American Veterinary Medical Association 213(1):54-60. Tizard, I. 1999. Grease, Anthraxgate, and kennel cough : a revisionist history of early veterinary vaccines. Advances in Veterinary medicine 41:7-24. Trilateral seal expert meeting 06.06.2002. http://cwss.www.de/news/news/Seals/Reports/TrilateralSealMeeting.pdf Twark, L., Dodds, W.J. 2000. Clinical use of serum parvovirus and distemper virus antibody titres for determining revaccination strategies in healthy dogs. Journal of the American veterinary Medical Association 217(7):1021-4 Visee, A.M., Zwart, P., Haagsma, J. 1974. Two saces of infectious enteritis in maned wolves (Chrysocyon brachyurus). Verhandlungsbericht XVI Internationalen Symposiums Ueber Erkrankungen Zootiere, Erfurt. Berlin: Akademie Verlag :67-69. Visee, A. 2001. Distemper, Rabies and Parvovirus vaccination in a captive-breeding programme for the African wild dog (Lycaon pictus) in Northern Tanzania. Proceedings Erkrankungen der zootiere 40:243-250. Visser, I.K.G., Vedder, E.J., Bildt van de, M.W., Groen, J., Orvell, C., Raga, J.A., Osterhaus, A.D.M.E. 1992. Canine distemper virus ISCOMs induce protection in harbour seals against phocid distemper but still allow subsequent infection with phocid distemper virus-1. Vaccine 10(7):435-438. Vries de, P., UytdeHaag, F.G., Osterhaus, A.D.M.E. 1988. Canine distemper virus (CDV) immune stimulating complexes (ISCOMS), but not measles virus ISCOMS protect dogs against CDV infection. Journal of General Virology 69(8):2071-2083. Wack, R.F., Kramer, L.W., Cupps, W.L., Katz, S. 1990. Antibody titre response of cheetah (acinnyx jubatus) cubs to vaccination. Proceedings Of the American Association of Zoo Veterinarians Annual Conference:147-149. Wack, R.F. 1991. the vaccination of cheetahs (acinonyx jubatus). Proceedings of the American Association of Zoo Veterinarians Annual Conference:294-7. Wack RF et al 1993. The response of cheetahs to routine vaccination. Journal of Zoo and Wildlife Medicine 109-117. Whetstone, C.A., Draayer, H., Collins, J.E. 1988. Characterization of canine adenovirus type 1 isolated from American black bears. American Journal of Veterinary Research 49(6):778-80. Whetstone, C.A., Bunn, T.O., Emmons, R.W., Wiktor, T.J. 1984. Use of monoclonal antibodies to confirm vaccine-induced rabies in ten dogs two cats and one Fox. Journal of the American Veterinary Medical Association 185(3):285-8. Williams, E.S., Anderson, S.L., Cavender, J., list, K., Heam, C., Appel, M.J.G. 1996. Vaccination of black-footed ferret (mustela nigripes) x siberian polecat (mustela eversmann) hybrids and domestic ferrets (mustela putorius furo) against canine distemper. Journal of Wildlife Diseases 32(3):417-423 Williams ES, Montali RJ, 1998. Vaccination of black-footed ferret x siberian polecat hybrids against canine distemper with recombinant and modified live virus vaccines. Proceedings of the Annual Conference of the Wildlife Disease Association 1998:107. Williams, E.S., Thorne, E.T. 1999. Veterinary contribution to the black-footed ferret conservation programme. In: Fowler, M.E. & Miller, R.E.: Zoo and wild animal medicine, Current therapy 4. Saunders Co: 460-463. Wilson, R.B., Kord, C.E., Holladay, J.A. 1986. A neurologic syndrome associated with use of a canine coronavirus-parvovirus vaccine in dogs. Compendium on Continuing Education for the Practicing Veterinarian 8:117-124. Wimsatt, J., Biggins, D., Taylor, B., Innes, K., Garell, D. 1999 Preliminary experimental canarypox vectored recombinant canine distemper vaccine evaluation in the siberian polecat (Mustela eversmanni). Proceedings of the American Association of Zoo Veterinarians Annual Conference:351-352.

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Wimsatt, J. Jay, M.T., Innes, K.E., Jessen, M., Collins, J.K. 2001. Serologic evaluation, efficacy, and safety of a commercial modified-live canine distemper vaccine in domestic ferrets. American Journal of Veterinary Reasearch 62(5):736-740. Wimsatt, J., Biggins, D., Innes, K., Taylor, B., Garell, D. 2003. Evaluation of oral and subcutaneous delivery of an experimental canarypox recombinant canine distemper vaccine in the Siberian polecat (Mustela eversmanni). Journal of Zoo Wildlife Medicine 34(1):25-35 Wood, S.L. et al 1995. Canine distemper virus-like infection in a captive African lioness. Canadian Veterinary Journal 36:34-35. Woodford, M.H. (ed), 2001. Quarantine and health screening protocols for wildlife prior to translocation and release into the wild. Published jointly by the IUCN Species Survival Commission’s Veterinary Specialist Group, Gland, Switzerland, the Office International des Epizooties (OIE), Care for the Wild, UK, and the European Association of Zoo and Wildlife Veterinartians, Switzerland: 39-44. Woolf, A., Swart, J. 1974. An outbreak of feline panleukopenia. Journal of Zoo Animal Medicine 5:32.

Transmissible Diseases Handbook

XIII. REFERENCE LABORATORIES FOR ANIMAL DISEASES This chapter includes listings of OIE and European Union reference laboratories for animal diseases. Updated versions of these lists can be found at: http://www.oie.int/eng/oie/organisation/en_listeLR.htm and http://ec.europa.eu/food/animal/diseases/laboratories/index_en.htm#list The World Health Organization (WHO) also has Collaborating Centers for many diseases relevant to zoo and wild animals. An up to date database of these centers can be searched at: http://www.who.int/whocc/Default.aspx

Community Reference Laboratories African horse sickness Laboratorio de sanidad y producción animal Ministerio de Agricultura, Pesca y Alimentación 28110 Algete, Madrid Spain African swine fever Centro de Investigación en Sanidad Animal Ctra. De Algete a El Casar, Valdeolmos 28130, Madrid Spain Avian Influenza Veterinary Laboratories Agency New Haw, Weybridge Surrey KT 15 3NB United Kingdom Bivalve molluscs diseases The Ifremer Laboratory B.P. 133 17390 La tremblade France Bluetongue Institute for Animal Health Pirbright Laboratory Pirbright, Woking Surrey GU24 ONF United Kingdom Bovine tuberculosis VISAVET Laboratorio de vigilancia veterinaria, Facultad de Veterinaria, Universidad Complutense de Madrid

XIII. Reference Laboratories for Animal Diseases

Avda. Puerta de Hierro, s/n. Ciudad Universitaria 28040. Madrid Spain Brucellosis AFSSA, Nancy Laboratoire d’études sur la rage et la pathologie des animaux sauvages Domaine de Pixérécourt, BP 9 F-54220 Malzéville France Classical swine fever Institut für Virologie der Tierarztlichen Hochschule Hanover Bischofscholer Damm 15 D-3000 Hannover 1 Germany Crustacean diseases Centre for Environment, Fisheries & Aquaculture Science (Cefas) Weymouth Laboratory The Nothe, Barrack Road, Weymouth Dorset DT4 8UB United Kingdom Equine diseases other than African Horse Sickness Agence Française de Sécurité Sanitaire des aliments (AFSSA) Laboratoire d'études et de recherches en pathologie animale et zoonoses 23, avenue du Général de Gaulle F-94706 MAISONS-ALFORT Cedex France Fish diseases State Serum Laboratory Hangovej 2 8200-Aarhus Denmark Foot and mouth disease Institut für Virologieder Tierarztlichen Hochschule Hanover Bischofscholer Damm 15 D-3000 Hannover 1 Germany Newcastle disease Veterinary Laboratories Agency New Haw, Weybridge Surrey KT 15 3NB United Kingdom Rabies AFSSA, Nancy Laboratoire d’études sur la rage et la pathologie des animaux sauvages Domaine de Pixérécourt, BP 9 F-54220 Malzéville France

XIII. Reference Laboratories for Animal Diseases

Swine vesicular disease Institute for Animal Health Pirbright Laboratory Pirbright, Woking Surrey GU24 ONF United Kingdom Zootechnics (bovine breeding) INTERBULL Centre Department of Animal Breeding and Genetics Swedish University of Agricultural Sciences Box: 7023; S-750 07 Uppsala Sweden

XIII. Reference Laboratories for Animal Diseases

Organisation Mondiale de la Santé Animale World Organisation for Animal Health Organización Mundial de Sanidad Animal

OIE Reference Experts and Laboratories in Europe African horse sickness  Dr G.H. Gerdes Onderstepoort Veterinary Institute Private Bag X05, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.91.14 Fax: (27.12) 529.94.18 Email: [email protected]  Dr Concepción Gómez-Tejedor Ortiz (1) Laboratorio Central de Veterinaria de Algete Carretera de Algete, km 8, 28110 Algete, (Madrid) SPAIN Tel: (34.91) 347.92.82/92.77 Fax: (34.91) 347.82.96 Email: [email protected]  Prof. P.S. Mellor Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 Fax: (44.1483) 23.24.48 Email: [email protected]  Dr J.M. Sánchez-Vizcaíno (2) Facultad de Veterinaria, Laboratorio de Vigilancia Sanitaria (VISAVET), HCV Planta sótano, Universidad Complutense Avda. Puerta de Hierro s/n, 28040 Madrid SPAIN Tel: (34.91) 394.40.82 Fax: (34.91) 394.39.08 Email: [email protected]

African swine fever  Dr J.M. Sánchez-Vizcaíno (2) Facultad de Veterinaria, Laboratorio de Vigilancia Sanitaria (VISAVET), HCV Planta sótano, Universidad Complutense Avda. Puerta de Hierro s/n, 28040 Madrid SPAIN Tel: (34.91) 394.40.82 Fax: (34.91) 394.39.08 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

 Dr Chris Oura Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 Fax: (44.1483) 23.24.48 Email: [email protected]  Dr Baratang Alison Lubisi Diagnostic Virology, Molecular Epidemiology and Diagnostics Programme, Onderstepoort Veterinary Institute, Agricultural Research Council Private Bag X5, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.95.85 Fax: (27.12) 529.95.95 Email: [email protected] American foulbrood of honey bees  Dr Adriana M. Alippi Laboratorio de Loque Americana de la Unidad de Bacteriología del Centro de Investigaciones en Fitopatología (CIDEFI) calle 60 y 119 s/n c.c. 31, 1900 La Plata ARGENTINA Tel: (+54-221) 423.67.48 ext. 423 Fax: (+54-221) 425.23.46 Email: [email protected] Email: [email protected] Anthrax  Dr G. Harvey National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.65 Fax: (1.515) 663.75.69 Email: [email protected]  Dr B. Golsteyn-Thomas Canadian Food Inspection Agency, Lethbridge Laboratory P.O. Box 640, Township Road 9-1, Lethbridge, Alberta T1J 3Z4 CANADA Tel: (1.403) 382.55.51 Fax: (1.403) 381.12.02 Email: [email protected] Antimicrobial resistance  Dr Chris Teale VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44-1743) 46.76.21 Fax: (44-1743) 44.10.60 Email: [email protected] Aujeszky's disease  Dr S.L. Swenson National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA

XIII. Reference Laboratories for Animal Diseases

Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected]  Dr P. Vannier AFSSA Ploufragan, Laboratoire d'études et de recherches avicoles et porcines, UR Station de pathologie porcine Zoopôle Beaucemaine-Les Croix, BP 53, 22440 Ploufragan FRANCE Tel: (33 (0)2) 96.01.62.22 Fax: (33 (0)2) 96.01.62.23 Email: [email protected]  Dr A.T.J. Bianchi Central Veterinary Institute of Wageningen UR P.O. Box 2004, 8203 AA Lelystad THE NETHERLANDS Tel: (31.320) 23.88.00 Fax: (31.320) 23.86.68 Email: [email protected] Avian chlamydiosis  Dr Konrad Sachse Friederich-Loeffer Institute, Institute of Molecular Pathogenesis Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.43.34 Fax: (49.3641) 80.42.28 Email: [email protected] Avian mycoplasmosis (Mycoplasma gallisepticum, M. synoviae)  Dr N. Ferguson-Noel The University of Georgia, Poultry Diagnostic and Research Centrer 953 College Station Rd, Athens, Georgia 30602-4875 UNITED STATES OF AMERICA Tel: (1.706) 542.30.86 Fax: (1.706) 542.56.30 Email: [email protected] Avian tuberculosis  Dr I. Pavlik Veterinary Research Institute Hudcova 70, 62132 Brno CZECH (Rep.) Tel: (420.5) 33.33.16.01 Fax: (420.5) 33.33.12.29 Email: [email protected] Bacterial kidney disease (Renibacterium salmoninarum)  Dr James R. Winton Western Fisheries Research Center 6505 N.E. 65th Street, Seattle, Washington 98115 UNITED STATES OF AMERICA Tel: (1.206) 526.65.87 Fax: (1.206) 526.66.54 Email: [email protected] Bee diseases  Monsieur Jean-Paul Faucon

XIII. Reference Laboratories for Animal Diseases

AFSSA Sophia Antipolis, Unité Pathologie de l'abeille, Laboratoire de Pathologie des Petits Ruminants et des Abeilles 105 route des Chappes, BP 111, 06902 Sophia Antipolis FRANCE Tel: (33 (0)4) 92.94.37.00 Fax: (33 (0)4) 92.94.37.01 Email: [email protected]  Dr W. Ritter Chemisches und Veterinäruntersuchungsamt Freiburg P.O.B. 100462, 79123 Freiburg GERMANY Tel: (49.761) 150.21.75 Fax: (49.761) 150.22.99 Email: [email protected]

Bluetongue  Dr G.H. Gerdes Onderstepoort Veterinary Institute Private Bag X05, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.91.14 Fax: (27.12) 529.94.18 Email: [email protected]  Dr E.N. Ostlund Diagnostic Virology Laboratory, National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected]  Dr Giovanni Savini Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise 'G. Caporale' Via Campo Boario, 64100 Teramo ITALY Tel: (39.0861) 33.24.40 Fax: (39.0861) 33.22.51 Email: [email protected]  Dr Peter Daniels CSIRO, Australian Animal Health Laboratory (AAHL) 5 Portarlington Road, Private Bag 24, Geelong 3220, Victoria AUSTRALIA Tel: (61.3) 52.27.50.00 Fax: (61.3) 52.27.55.55 Email: [email protected]  Prof. P.S. Mellor Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 Fax: (44.1483) 23.24.48 Email: [email protected] Bovine babesiosis  Prof. Ikuo Igarashi

XIII. Reference Laboratories for Animal Diseases

National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine Inada-cho Nishi 2-13, Obihiro, Hokkaido 080-8555 JAPAN Tel: (81.155) 49.56.42 Fax: (81.155) 49.56.43 Email: [email protected] Bovine genital campylobacteriosis  Dr Jaap Wagenaar Animal Sciences Group (ASG), Division of Infectious Diseases P.O. Box 65, 8200 AB Lelystad THE NETHERLANDS Tel: (31.320) 23.81.57 Fax: (31.320) 23.89.61 Email: [email protected]  Dr Jaap Wagenaar Faculty of Veterinary Medicine (FVM), Department of Infectious Diseases and Immunology P.O. Box 80.165, 3508 TD Utrecht THE NETHERLANDS Tel: (31.30) 253.12.42 Fax: (31.30) 253.31.99 Email: [email protected]

Bovine spongiform encephalopathy  Dr Marion Simmons VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.75.64 Fax: (44.1932) 35.78.05 Email: [email protected] Web: http://www.defra.gov.uk/corporate/vla/science/science-tse-rl-web.htm  Prof. Andreas Zurbriggen Institute of Animal Neurology, University of Bern Bremgartenstrasse 109A, 3012 Bern SWITZERLAND Tel: (41.31) 631.25.09 Fax: (41.31) 631.25.38 Email: [email protected]  Dr Stefanie Czub Canadian Food Inspection Agency, Lethbridge Laboratory Township Road 9-1, Post Office Box 640, Lethbridge, Alberta CANADA Tel: (1.403) 382.55.49 Fax: (1.403) 382.55.83 Email: [email protected]  Dr Takashi Yokoyama Prion Diseases Research Unit, National Institute of Animal Health, National Agricultural Research Organization 3-1-5 Kannondai, Tsukuba, Ibaraki 305-0856 JAPAN Tel: (81.298) 38.77.40 Fax: (81.298) 38.83.32 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

 Dr Francisco Javier Blanco Viera Laboratorio Nacional de Referencia (LNR) para Encefalopatías Espongiformes Transmisible animales, Institutos de Patobiología y Virología, Centro de Investigaciones en Ciencias Veterinarias y Agronómicas (CICV), Instituto Nacional de Tecnología Agropecuaria (INTA) Castelar, Casilla de Correo 77, 1708 Morón, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 46.21.12.89 Fax: (54.11) 46.21.17.43 Email: [email protected] Email: [email protected] Bovine tuberculosis  Dr Bernardo Alonso Gerencia de Laboratorios (GELAB) del Servicio Nacional de Sanidad y Calidad, Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martinez - Pcia de Buenos Aires ARGENTINA Tel: (54.11) 48.36.00.36 Fax: (54.11) 48.36.00.36 Email: [email protected] Email: [email protected]  Mme María Laura Boschiroli-Cara AFSSA Alfort, Unité Zoonoses Bactériennes, Laboratoire d'études et de recherches en pathologie animale et zoonoses 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 Email: [email protected]  Dr Debby V. Cousins Australian Reference Laboratory for Bovine Tuberculosis, Agriculture Western Australia Locked Bag N° 4, Bentley Delivery Centre, Bentley WA 6983 AUSTRALIA Tel: (61.8) 93.68.34.51 Fax: (61.8) 94.74.18.81 Email: [email protected]  Prof. Glyn Hewinson VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 34.11.11 Fax: (44.1932) 34.70.46 Email: [email protected] Bovine viral diarrhoea  Dr D. Deregt (1) Canadian Food Inspection Agency, Animal Diseases Research Institute P.O. Box 640, Lethbridge, Alberta T1J 3Z4 CANADA Tel: (1.403) 382.55.00 Fax: (1.403) 381.12.02 Email: [email protected]  Dr T.W. Drew Head of Virology Department, VLA Weybridge Woodham Lane, New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM

XIII. Reference Laboratories for Animal Diseases

Tel: (44.1932) 35.76.37 Fax: (44.1932) 35.72.39 Email: [email protected]  Dr Peter Kirkland Elizabeth Macarthur Agriculture Institute (EMAI), Virology Laboratory PMB 8, Camden NSW 2570 AUSTRALIA Tel: (61-2) 46.40.63.31 Fax: (61-2) 46.40.64.29 Email: [email protected] Brucellosis (Brucella melitensis)  Dr Henrich Neubauer Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Bacterial Infections and Zoonoses Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.42.00 Fax: (49.3641) 80.42.28 Email: [email protected]  Dr Ana Maria Nicola Gerencia de Laboratorios (GELAB), Servicio Nacional de Sanidad y Calidad Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martínez, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 48.36.19.92 Fax: (54.11) 48.36.19.92 Email: [email protected]  Dr J.A. Stack VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.76.10. Fax: (44.1932) 35.72.16 Email: [email protected]  Dr B. Garin-Bastuji AFSSA Alfort, Unité Zoonoses Bactériennes, Lab. OIE/FAO de référence pour la brucellose animale, Laboratoire d'études et de recherches en pathologie animale et zoonoses 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 Email: [email protected]  Dr K. Nielsen Canadian Food Inspection Agency, Animal Diseases Research Institute P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 ext. 48.04 Fax: (1.613) 228.66.69 Email: [email protected]  Dr Massimo Scacchia CESME, Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise 'G. Caporale', National Centre for Exotic Diseases Via Campo Boario, 64100 Teramo ITALY Tel: (390.861) 33.24.05 Fax: (390.861) 33.22.51

XIII. Reference Laboratories for Animal Diseases

Email: [email protected]  Dr Menachem Banai Kimron Veterinary Institute Department of Bacteriology, P.O. Box 12, Beit Dagan 50250 ISRAEL Tel: (972.3) 968 16 98 Fax: (972.3) 968 17 53 Email: [email protected] Brucellosis (Brucella suis)  Dr Henrich Neubauer Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Bacterial Infections and Zoonoses Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.42.00 Fax: (49.3641) 80.42.28 Email: [email protected]  Dr Ana Maria Nicola Gerencia de Laboratorios (GELAB), Servicio Nacional de Sanidad y Calidad Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martínez, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 48.36.19.92 Fax: (54.11) 48.36.19.92 Email: [email protected]  Dr J.A. Stack VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.76.10. Fax: (44.1932) 35.72.16 Email: [email protected]  Dr B. Garin-Bastuji AFSSA Alfort, Unité Zoonoses Bactériennes, Lab. OIE/FAO de référence pour la brucellose animale, Laboratoire d'études et de recherches en pathologie animale et zoonoses 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 Email: [email protected]  Dr K. Nielsen Canadian Food Inspection Agency, Animal Diseases Research Institute P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 ext. 48.04 Fax: (1.613) 228.66.69 Email: [email protected]  Dr Massimo Scacchia CESME, Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise 'G. Caporale', National Centre for Exotic Diseases Via Campo Boario, 64100 Teramo ITALY Tel: (390.861) 33.24.05 Fax: (390.861) 33.22.51 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Brucellosis Brucella abortus  Dr Henrich Neubauer Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Bacterial Infections and Zoonoses Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.42.00 Fax: (49.3641) 80.42.28 Email: [email protected]  Dr Ana Maria Nicola Gerencia de Laboratorios (GELAB), Servicio Nacional de Sanidad y Calidad Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martínez, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 48.36.19.92 Fax: (54.11) 48.36.19.92 Email: [email protected]  Dr J.A. Stack VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.76.10. Fax: (44.1932) 35.72.16 Email: [email protected]  Dr B. Garin-Bastuji AFSSA Alfort, Unité Zoonoses Bactériennes, Lab. OIE/FAO de référence pour la brucellose animale, Laboratoire d'études et de recherches en pathologie animale et zoonoses 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 Email: [email protected]  Dr K. Nielsen Canadian Food Inspection Agency, Animal Diseases Research Institute P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 ext. 48.04 Fax: (1.613) 228.66.69 Email: [email protected]  Dr Massimo Scacchia CESME, Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise 'G. Caporale', National Centre for Exotic Diseases Via Campo Boario, 64100 Teramo ITALY Tel: (390.861) 33.24.05 Fax: (390.861) 33.22.51 Email: [email protected]  Dr Menachem Banai Kimron Veterinary Institute Department of Bacteriology, P.O. Box 12, Beit Dagan 50250 ISRAEL Tel: (972.3) 968 16 98 Fax: (972.3) 968 17 53 Email: [email protected]  Dr Suk-chan Jung

XIII. Reference Laboratories for Animal Diseases

Zoonosis Laboratory, Bacteriology and Parasitology Division, National Veterinary Research and Quarantine Service (NVRQS), Ministry of Food, Agriculture, Forestry, and Fisheries (MIFAFF) 480 Anyang 6-dong, Manan-gu, Anyang-si, Kyunggi-do CORÉE (RÉP. DE) Tel: (82.31) 467.17.65 Fax: (82.31) 467.17.78 Email: [email protected] Camelpox  Professor Ulrich Wernery Central Veterinary Research Laboratory P.O. Box 597, Dubai UNITED ARAB EMIRATES Tel: (971.4) 337.51.65 Fax: (971.4) 336.86.38 Email: [email protected] Campylobacteriosis  Dr Jaap Wagenaar Animal Sciences Group (ASG), Division of Infectious Diseases P.O. Box 65, 8200 AB Lelystad THE NETHERLANDS Tel: (31.320) 23.81.57 Fax: (31.320) 23.89.61 Email: [email protected]  Dr Jaap Wagenaar Faculty of Veterinary Medicine (FVM), Department of Infectious Diseases and Immunology P.O. Box 80.165, 3508 TD Utrecht THE NETHERLANDS Tel: (31.30) 253.12.42 Fax: (31.30) 253.31.99 Email: [email protected] Caprine arthritis/encephalitis  Dr Stephen Valas Laboratoire d'étude et de recherches caprines 60 rue du Pied de Fond, B.P. 3081, 79000 Niort FRANCE Tel: (33 (0)5 49.79.61.28 Fax: (33 (0)5 49.79.42.19 Email: [email protected]  Dr D.P. Knowles, Jr Animal Diseases Research Unit,USDA, ARS, Washington State University Pullman, Washington 99164-7030 UNITED STATES OF AMERICA Tel: (1.509) 335.60.22 Fax: (1.509) 335.83.28 Email: [email protected] Channel catfish virus disease  Dr Larry A. Hanson College of Veterinary Medicine, Fish Diagnostic Laboratory, Mississippi State University P.O. Box 6100, Spring Street, Mississippi 39762 UNITED STATES OF AMERICA Tel: (1.662) 325.12.02 Fax: (1.662) 325.10.31 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Chronic wasting disease  Dr Aru Balachandran Canadian Food Inspection Agency, Ottawa Laboratory 3851 Fallowfield Road, P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 ext. 4854 Fax: (1.613) 228.61.03 Email: [email protected] Classical swine fever  Dr John Pasick Canadian Food Inspection Agency, National Centre for Foreign Animal Disease 1015 Arlington Street, Winnipeg, Manitoba R3E 3M4 CANADA Tel: (1.204) 789.20.13 Fax: (1.204) 789.20.38 Email: [email protected]  Dr Shunji Yamada National Institute of Animal Health 6-20-1 Josui-honcho, Kodaira, Tokyo, 187-0022 JAPAN Tel: (81.42) 321.14.41 Fax: (81.42) 325.51.22 Email: [email protected]  Prof. V. Moennig Institute of Virology, Hannover Veterinary School Bünteweg 17, 30559 Hannover GERMANY Tel: (49.511) 953.88.40 Fax: (49.511) 953.88.98 Email: [email protected]  Prof. Dr Z. Pejsak National Veterinary Research Institute Partyzantow str. 57, 24-100 Pulawy POLAND Tel: (48.81) 889.30.30 Fax: (48.81) 886.25.95 Email: [email protected]  Dr T.W. Drew Head of Virology Department, VLA Weybridge Woodham Lane, New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.76.37 Fax: (44.1932) 35.72.39 Email: [email protected] Contagious agalactia  Dr Robin A.J. Nicholas Mycoplasma Group, Department of Statutory and Exotic Bacterial Diseases, VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 34 11 11 Fax: (44.1932) 34 70 76 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Contagious bovine pleuropneumonia  Dr F. Poumarat (1) AFSSA Lyon, Laboratoire de pathologie bovine 31 avenue Tony Garnier, BP 7033, 69342 Lyon Cedex 07 FRANCE Tel: (33 (0)4) 78.72.65.43 Fax: (33 (0)4) 78.61.91.45 Email: [email protected]  Dr Ana Rosa Pombo Botelho Laboratório Nacional de Investigaçâo Veterinária (LNIV) Estrada de Benfica 701, 1500 Lisboa PORTUGAL Tel: (351.21) 711.53.33/39/40 Fax: (351.21) 711.52.36 Email: [email protected]  Dr A. Pini CESME, Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise 'G. Caporale' Via Campo Boario, 64100 Teramo ITALY Tel: (39.0861) 33.24.81 Fax: (39.0861) 33.22.51 Email: [email protected]  Dr F. Thiaucourt (2) UMR15 CIRAD-INRA, Control of exotic and emerging animal diseases Campus international de Baillarguet TA A-15/G, 34398 Montpellier Cedex 5 FRANCE Tel: (33(0)4) 67.59.37.24 Fax: (33(0)4) 67.59.37.98 Email: franç[email protected] Contagious caprine pleuropneumonia  Dr F. Thiaucourt (2) UMR15 CIRAD-INRA, Control of exotic and emerging animal diseases Campus international de Baillarguet TA A-15/G, 34398 Montpellier Cedex 5 FRANCE Tel: (33(0)4) 67.59.37.24 Fax: (33(0)4) 67.59.37.98 Email: franç[email protected] Contagious equine metritis  Dr Brenda Morningstar-Shaw National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.65 Fax: (1.515) 663.75.69 Email: [email protected]  Mr Paul Todd VLA Bury St Edmunds Rougham Hill, Bury St Edmunds, Suffolk IP33 2RY UNITED KINGDOM Tel: (44-1284) 72.44.99 Fax: (44-1284) 72.45.00 Email: [email protected]  Dr Hendrik-Jan Roest

XIII. Reference Laboratories for Animal Diseases

Central Veterinary Institute of Wageningen UR, Bacteriology Department P.O. Box 65, 8203 AA Lelystad THE NETHERLANDS Tel: (31.320) 23.80.26 Fax: (31.320) 23.81.53 Email: [email protected] Control of Veterinary Medicinal Products in Sub-Saharan Africa  Dr Assiongbon Teko-Agbo Ecole Inter-Etats de Science et Médecine Vétérinaire (EISMV) BP 5077, Dakar SENEGAL Tel: (221) 33.865.10.08 Fax: (221) 33.825.42.83 Email: [email protected] Crayfish plague (Aphanomyces astaci)  Dr Birgit Oidtmann The Centre for Environment, Fisheries, and Aquaculture Science (CEFAS), Weymouth Laboratory Barrack Road, The Nothe, Weymouth, Dorset, DT4 8UB UNITED KINGDOM Tel: (44.1305) 20.66.61 Fax: (44.1305) 20.66.01 Email: [email protected]  Dr S. Viljamaa-Dirks Finnish Food Safety Authority Evira Kuopio, Neulaniementie 4, FIN-70210 Kuopio FINLANDE Tel: (358) 207.72.49.62 Fax: (358) 207.72.49.70 Email: [email protected] Crimean Congo haemorrhagic fever  Dr Michèle Bouloy Unité de génétique moléculaire des Bunyavirus, Département de Virologie, Institut Pasteur 25 rue du Dr Roux 75724 Paris cedex 15 FRANCE Tel: (33.1) 40.61.31.34 Fax: (33.1) 40.61.32.56 Email: E-mail: [email protected] Dourine  Prof. V.T. Zablotsky All-Russian Research Institute for Experimental Veterinary Medicine (VIEV), Veterinary Department, Laboratory of Equine Viral Diseases 24-1, Ryazanskiy prosp., 109428 Moscow RUSSIA Tel: (7.495) 785.84.27 Fax: (7.495) 970.03.69 Email: [email protected] Echinococcosis/hydatidosis  Prof. Masao Kamiya Laboratory of Environmental Zoology, Department of Biosphere and Environmental Sciences, Faculty of Environmental Systems, Rakuno-Gakuen University Midori-machi 582, Ebetsu 069-8501 JAPAN

XIII. Reference Laboratories for Animal Diseases

Tel: (81.11) 388.49.09 Fax: (81.11) 388.49.09 Email: [email protected] Web: http://www.k3.dion.ne.jp/~fea/  Dr P.S. Craig Cestode Zoonoses Research Group, Biosciences Research Institute, University of Salford Manchester M5 4WT UNITED KINGDOM Tel: (44.161) 295.54.88 Fax: (44.161) 295.52.15 Email: [email protected]  Prof. A. Dakkak Institut Agronomique et Vétérinaire Hassan II, Département de Parasitologie BP 6202, Rabat-Instituts MOROCCO Tel: (212.37) 77.64.32 Fax: (212.37) 77.64.32 Email: [email protected] Enteric septicaemia of catfish (Edwardsiella ictaluri)  Dr Larry A. Hanson College of Veterinary Medicine, Fish Diagnostic Laboratory, Mississippi State University P.O. Box 6100, Spring Street, Mississippi 39762 UNITED STATES OF AMERICA Tel: (1.662) 325.12.02 Fax: (1.662) 325.10.31 Email: [email protected] Enzootic abortion of ewes (ovine chlamydiosis)  Dr Konrad Sachse Friederich-Loeffer Institute, Institute of Molecular Pathogenesis Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.43.34 Fax: (49.3641) 80.42.28 Email: [email protected]  Dr Nicole Borel Institute for Veterinary Pathology (IVPZ), Vetsuisse Faculty, University of Zurich Winterhurerstrasse 268, CH-8057 Zurich SWITZERLAND Tel: (41.44) 635.85.71 Fax: (41.44) 635.89.34 Email: [email protected] Email: [email protected] Enzootic bovine leukosis  Mr C. Venables VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 34.11.11 Fax: (44.1932) 34.70.46 Email: [email protected]  Dr Thomas W. Vahlenkamp Friedrich-Loeffler-Institute, Institut für Molekularbiologie Südufer 10, 17493 Greifswald-Insel Riems

XIII. Reference Laboratories for Animal Diseases

GERMANY Tel: (49-38351) 7.172 Fax: (49-38351) 7.151 Email: [email protected]  Dr Jacek Kuzmak National Veterinary Institute SPartyzantow str. 57, 24-100 Pulawy POLAND Tel: (48-81) 889.31.14 Fax: (48-81) 886.25.95 Email: [email protected] Epizootic haematopoietic necrosis  Dr A. Hyatt (1) Australian Animal Health Laboratory, CSIRO Livestock Industries 5 Portarlington Road, Private Bag 24( Ryrie Street), Geelong, Victoria 3220 AUSTRALIA Tel: (61.3) 52.27.00.00 Fax: (61.3) 52.27.55.55 Email: [email protected]  Dr Richard Whittington (2) Chair Farm Animal Health, Faculty of Veterinary Science, University of Sydney 425 Werombi Road, Private Bag 3, Camden NSW 2570 AUSTRALIA Tel: (61.2) 93.51.16.19 Fax: (61.2) 93.51.16.18 Email: [email protected] Epizootic ulcerative syndrome  Dr S. Kanchanakhan Aquatic Animal Health Research Institute (AAHRI), Department of Fisheries, Kasetsart University Campus Paholyothin Road, Chatuchak, Bangkok 10900 THAILAND Tel: (66.2) 579.41.22 Fax: (66.2) 561.39.93 Email: [email protected] Equine encephalomyelitis (Eastern)  Dr E.N. Ostlund Diagnostic Virology Laboratory, National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected] Equine encephalomyelitis (Western)  Dr E.N. Ostlund Diagnostic Virology Laboratory, National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected] Equine infectious anaemia

XIII. Reference Laboratories for Animal Diseases

 Dr E.N. Ostlund Diagnostic Virology Laboratory, National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected]  Dr K. Murakami Viral disease Section, National Institute of Animal Health 3-1-5 Kannondai, Tsukuba, Ibaraki 305-0856 JAPAN Tel: (81.29) 838.78.41 Fax: (81.29) 838.79.07 Email: [email protected] Equine influenza  Dr W. Eichhorn Institute for Medical Microbiology, Infectious and Epidemic Diseases, Veterinary Faculty, LudwigMaximilians-University Veterinärstrasse 13, 80539 München GERMANY Tel: (49.89) 21.80.25.31 Fax: (49.89) 21.80.59.03 Email: [email protected]  Dr Jennifer A. Mumford Cambridge Infectious Diseases Consortium, Department of Veterinary Medicine Madingley Road, Cambridge CB3 0ES UNITED KINGDOM Tel: (44.1223) 76.49.64 Fax: (44.1223) 76.46.67 Email: [email protected]  Dr T.M. Chambers Maxwell H. Gluck Equine Research Center, Dept of Veterinary Science, University of Kentucky 108 Gluck Equine Research Center, Lexington, Kentucky 40546-0099 UNITED STATES OF AMERICA Tel: (1.859) 257.47.57 Fax: (1.859) 257.85.42 Email: [email protected]  Prof. Ann Cullinane Irish Equine Centre Johnstown, Naas, Co. Kildare IRLANDE Tel: (353.45) 86.62.66 Fax: (353.45) 86.62.73 Email: [email protected] Equine piroplasmosis  Prof. Ikuo Igarashi National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine Inada-cho Nishi 2-13, Obihiro, Hokkaido 080-8555 JAPAN Tel: (81.155) 49.56.42 Fax: (81.155) 49.56.43 Email: [email protected] Equine rhinopneumonitis

XIII. Reference Laboratories for Animal Diseases

 Dr Jennifer A. Mumford Cambridge Infectious Diseases Consortium, Department of Veterinary Medicine Madingley Road, Cambridge CB3 0ES UNITED KINGDOM Tel: (44.1223) 76.49.64 Fax: (44.1223) 76.46.67 Email: [email protected]  To be decided Maxwell H. Gluck Equine Research Center, Dept of Veterinary Science, University of Kentucky 108 Gluck Equine Research Center, Lexington, Kentucky 40546-0099 UNITED STATES OF AMERICA Tel: (1.859) 257.47.57 ext. 81119 Fax: (1.859) 257.85.42 Email:  Prof. Konstantin P. Yurov All-Russian Research Institute for Experimental Veterinary Medicine (VIEV), Laboratory of Equine Viral Diseases 24-1 Ryazanskiy prosp. 109428 Moscow RUSSIA Tel: (7.495) 995.88.63 Fax: (7.495) 970.03.69 Email: [email protected] Equine viral arteritis  Dr Peter J. Timoney Maxwell H. Gluck Equine Research Center, Chair and Director, Dept of Veterinary Science, University of Kentucky, 108 Gluck Equine Research Center, Lexington, Kentucky 40546-0099 UNITED STATES OF AMERICA Tel: (1.859) 257.47.57 ext. 81084 Fax: (1.859) 257.85.42 Email: [email protected]  Dr Takashi Kondo Epizootic Research Center, Equine Research Institute, The Japan Racing Association, K1400-4, Shiba, Shimotsuke-shi, Tochigi 329-0412 JAPAN Tel: (81.285) 44.00.90 Fax: (81.285) 40.10.64 Email: [email protected]  Dr T.W. Drew Head of Virology Department, VLA Weybridge Woodham Lane, New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.76.37 Fax: (44.1932) 35.72.39 Email: [email protected] Escherichia coli  Dr John Morris Fairbrother The Echerichia coli Laboratory (EcL) 3200 Sicotte Saint-Hyacinthe, Québec, J2S 7C6 CANADA Tel: (1.450) 773.85.21 Fax: (1.450) 778.81.08 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Foot and mouth disease  Dr Jef. M. Hammond Institute for Animal Health Ash Road, Pirbright, Woking, Surrey GU24 0NF UNITED KINGDOM Tel: (44.1483) 23.12.11 Fax: (44.1483) 23.26.21 Email: [email protected]  Dr O.G. Matlho Botswana Vaccine Institute, Department of Animal Health and Production Broadhurst Industrial Site, Lejara Road, Private Bag 0031, Gaborone BOTSWANA Tel: (267) 391.27.11 Fax: (267) 395.67.98 Email: [email protected] Web: www.bvi.co.bw  Dr Ingrid Bergmann Centro Panamericano de Fiebre Aftosa OPS/OMS Av. President Kennedy 7778, Sao Bento, Duque de Caxias, ZC 20054-40 Rio de Janeiro BRAZIL Tel: (55.21) 36.61.90.56 Fax: (55.21) 36.61.90.01 Email: [email protected]  Dr V.M. Zakharov Federal Governmental Institute, Centre for Animal Health, FGI-ARRIAH 600900 Vladimir, Yur'evets RUSSIA Tel: (4922) 26.19.14/26.06.14/26.38.77 Fax: (4922) 26.19.14/26.06.14/26.38.77 Email: [email protected] Web: http://www.arriah.ru Web: http://www.arriah.ru/portal/en  Dr Eduardo D. Maradei Laboratorio de Fiebre Aftosa de la Dirección de Laboratorios y Control Técnico Av. Sir. Alexander Fleming 1653, Martínez (1640), Buenos Aires ARGENTINA Tel: (+54-11) 48.36.19.95 Fax: (+54-11) 48.36.19.95 Email: [email protected] Email: [email protected]  Dr R.M. Dwarka Onderstepoort Veterinary Institute, Team leader: Exotic Animal Health, Exotic Diseases Division Private Bag X05, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.95.89 Fax: (27.12) 529.95.43 Email: [email protected]  Dr Wilai Linchongsubongkoch Department of Livestock Development Pakchong, Nakhon Ratchasima 30130 THAILAND Tel: (66.44) 27.91.12 Fax: (66.44) 31.48.89 Email: [email protected] Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Glanders  Dr Henrich Neubauer Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Bacterial Infections and Zoonoses Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.42.00 Fax: (49.3641) 80.42.28 Email: [email protected]  Professor Ulrich Wernery Central Veterinary Research Laboratory P.O. Box 597, Dubai UNITED ARAB EMIRATES Tel: (971.4) 337.51.65 Fax: (971.4) 336.86.38 Email: [email protected] Gyrodactylosis (Gyrodactylus salaris)  Dr T.A. Mo National Veterinary Institute, Section for Parasitology P.O. Box 750 Sentrum, 0106 Oslo NORWAY Tel: (47) 23.21.61.10 Email: [email protected] Heartwater  Dr Dominique Martinez CIRAD-EMVT Head of Research Unit, Control of exotic and emerging animal diseases, TA30/G Campus international de Baillarguet, 34398 Montpellier Cedex 5 FRANCE Tel: (33(0)4) 67.59.37.12 Fax: (33(0)4) 67.59.37.98 Email: [email protected] Hendra and Nipah virus diseases  Dr P. Daniels CSIRO, Australian Animal Health Laboratory (AAHL) 5 Portarlington Road, Private Bag 24, Geelong 3220, Victoria AUSTRALIA Tel: (61.3) 52.27.50.00 Fax: (61.3) 52.27.55.55 Email: [email protected] Highly pathogenic avian influenza and low pathogenic avian influenza (poultry)  Dr John Pasick Canadian Food Inspection Agency, National Centre for Foreign Animal Disease 1015 Arlington Street, Winnipeg, Manitoba R3E 3M4 CANADA Tel: (1.204) 789.20.13 Fax: (1.204) 789.20.38 Email: [email protected]  Dr Hualan Chen National Avian Influenza Reference Laboratory, Animal Influenza Laboratory of the Ministry of

XIII. Reference Laboratories for Animal Diseases

Agriculture, Harbin Veterinary Research Institute, CAAS 427 Maduan Street, Harbin 150001 CHINA (People's Republic of) Tel: (86-451) 85.93.50.79 Fax: (86-451) 82.73.31.32 Email: [email protected] Email: [email protected] Web: http://hvri.ac.cn  Dr Timm C. Harder Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Diagnostic Virology Boddenblick 5a, 17493 Greifswald - Insel Riems GERMANY Tel: (49.383) 51.71.52 Fax: (49.383) 51.72.75 Email: [email protected]  Dr Ian Brown VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.73.39 Fax: (44.1932) 35.72.39 Email: [email protected]  Dr Paul W. Selleck CSIRO, Australian Animal Health Laboratory (AAHL) 5 Portarlington Road, Private Bag 24, Geelong 3220, Victoria AUSTRALIA Tel: (61.3) 52.27.50.00 Fax: (61.3) 52.27.55.55 Email: [email protected]  Dr B. Panigrahy National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected]  Dr Ilaria Capua Istituto Zooprofilattico Sperimentale delle Venezie, Laboratorio Virologia Via Romea 14/A, 35020 Legnaro, Padova ITALY Tel: (39.049) 808.43.79 Fax: (39.049) 808.43.60 Email: [email protected]  Dr H. Kida Graduate School of Veterinary Medicine, Hokkaido University, Department of Disease Control Kita-18, Nishi-9, Kita-ku, Sapporo 060-0818 JAPAN Tel: (81.11) 706.52.07 Fax: (81.11) 706.52.73 Email: [email protected]  Dr S.C. Dubey High Security Animal Disease Laboratory, Indian Veterinary Research Institute, Indian Council of Agricultural Research Anand Nagar, Bhopal 462021, Madhya Pradesh INDIA Tel: (91.7552) 26.94.87 Fax: (91.7552) 26.94.87

XIII. Reference Laboratories for Animal Diseases

Email: [email protected] Infection with Batrachochytrium dendrobatidis  Dr A. Hyatt (1) Australian Animal Health Laboratory, CSIRO Livestock Industries 5 Portarlington Road, Private Bag 24( Ryrie Street), Geelong, Victoria 3220 AUSTRALIA Tel: (61.3) 52.27.00.00 Fax: (61.3) 52.27.55.55 Email: [email protected]  Dr Richard Whittington (2) Chair Farm Animal Health, Faculty of Veterinary Science, University of Sydney 425 Werombi Road, Private Bag 3, Camden NSW 2570 AUSTRALIA Tel: (61.2) 93.51.16.19 Fax: (61.2) 93.51.16.18 Email: [email protected] Infection with Bonamia exitiosa  Dr Isabelle Arzul IFREMER, Laboratoire de Génétique et Pathologie BP 133, 17390 La Tremblade FRANCE Tel: (33-5) 46.76.26.10 Fax: (33-5) 46.76.26.11 Email: [email protected] Infection with Bonamia ostreae  Dr Isabelle Arzul IFREMER, Laboratoire de Génétique et Pathologie BP 133, 17390 La Tremblade FRANCE Tel: (33-5) 46.76.26.10 Fax: (33-5) 46.76.26.11 Email: [email protected] Infection with Haplosporidium nelsoni  Dr E.M. Burreson Director for Research and Advisory Services, Virginia Institute of Marine Science, College of William and Mary P.O. Box 1346, Gloucester Point, VA 23062 UNITED STATES OF AMERICA Tel: (1.804) 684.70.15 Fax: (1.804) 684.77.96 Email: [email protected] Infection with Marteilia refringens  Dr Isabelle Arzul IFREMER, Laboratoire de Génétique et Pathologie BP 133, 17390 La Tremblade FRANCE Tel: (33-5) 46.76.26.10 Fax: (33-5) 46.76.26.11 Email: [email protected] Infection with Marteilia sydneyi

XIII. Reference Laboratories for Animal Diseases

 Dr Isabelle Arzul IFREMER, Laboratoire de Génétique et Pathologie BP 133, 17390 La Tremblade FRANCE Tel: (33-5) 46.76.26.10 Fax: (33-5) 46.76.26.11 Email: [email protected] Infection with Mikrocytos mackini  Dr S. Bower Department of Fisheries and Oceans Pacific Biological Station 3190 Hammond Bay Road, Nanaimo, British Columbia V9T 6N7 CANADA Tel: (1.250) 756.70.77 Fax: (1.250) 756.70.53 Email: [email protected] Infection with Perkinsus marinus  Dr E.M. Burreson Director for Research and Advisory Services, Virginia Institute of Marine Science, College of William and Mary P.O. Box 1346, Gloucester Point, VA 23062 UNITED STATES OF AMERICA Tel: (1.804) 684.70.15 Fax: (1.804) 684.77.96 Email: [email protected] Infection with Perkinsus olseni  Dr E.M. Burreson Director for Research and Advisory Services, Virginia Institute of Marine Science, College of William and Mary P.O. Box 1346, Gloucester Point, VA 23062 UNITED STATES OF AMERICA Tel: (1.804) 684.70.15 Fax: (1.804) 684.77.96 Email: [email protected] Infection with ranavirus  Dr A. Hyatt (1) Australian Animal Health Laboratory, CSIRO Livestock Industries 5 Portarlington Road, Private Bag 24( Ryrie Street), Geelong, Victoria 3220 AUSTRALIA Tel: (61.3) 52.27.00.00 Fax: (61.3) 52.27.55.55 Email: [email protected]  Dr Richard Whittington (2) Chair Farm Animal Health, Faculty of Veterinary Science, University of Sydney 425 Werombi Road, Private Bag 3, Camden NSW 2570 AUSTRALIA Tel: (61.2) 93.51.16.19 Fax: (61.2) 93.51.16.18 Email: [email protected] Infection with Xenohaliotis californiensis  Prof. Carolyn Friedman Friedman Shellfish Health Laboratory, School of Aquatic and Fishery Sciences, University of Washington

XIII. Reference Laboratories for Animal Diseases

Box 355020, Seattle, Washington 98195 UNITED STATES OF AMERICA Tel: (1.206) 543.95.19 Fax: (1.206) 616.86.89 Email: [email protected] Infectious bovine rhinotracheitis/infectious pustular vulvovaginitis  Dr Martin Beer National Referance Laboratory for Bovine herpesvirus type 1, Institute of Diagnostic Virology, Federal Research Centre for Virus Diseases of Animals (BFAV) Insel Riems, Boddenblick 5a, 17493 Greifswald - Insel Riems GERMANY Tel: (49) 383.517.223 Fax: (49) 383.517.275 Email: [email protected]  Dr M. Banks VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 34.11.11 Fax: (44.1932) 34.70.46 Email: [email protected] Infectious bursal disease (Gumboro disease)  Dr Y.M. Saif Food Animal Health Research Program, Ohio Agricultural Research and Development Center, The Ohio State University 1680 Madison Avenue, Wooster, OH 44691-4096 UNITED STATES OF AMERICA Tel: (1.330) 263.37.43 Fax: (1.330) 263.36.77 Email: [email protected]  Dr N. Eterradossi AFSSA Ploufragan, Unité de virologie, immunologie et parasitologie aviaires et cunicoles BP 53, 22440 Ploufragan FRANCE Tel: (33 (0)2) 96.01.62.22 Fax: (33 (0)2) 96.01.62.63 Email: [email protected] Infectious haematopoietic necrosis  Dr James R. Winton Western Fisheries Research Center 6505 N.E. 65th Street, Seattle, Washington 98115 UNITED STATES OF AMERICA Tel: (1.206) 526.65.87 Fax: (1.206) 526.66.54 Email: [email protected] Infectious hypodermal and haematopoietic necrosis  Prof. Donald V. Lightner Aquaculture Pathology Laboratory, Department of Veterinary Science and Microbiology, University of Arizona Building 90, Room 202 Pharmacy/Microbiology, Tucson, AZ 85721 UNITED STATES OF AMERICA Tel: (1.520) 621.84.14 Fax: (1.520) 621.48.99 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Infectious myonecrosis  Prof. Donald V. Lightner Aquaculture Pathology Laboratory, Department of Veterinary Science and Microbiology, University of Arizona Building 90, Room 202 Pharmacy/Microbiology, Tucson, AZ 85721 UNITED STATES OF AMERICA Tel: (1.520) 621.84.14 Fax: (1.520) 621.48.99 Email: [email protected] Infectious salmon anaemia  Dr F. Kibenge Atlantic Veterinary College, Department of Pathology and Microbiology, Faculty of Veterinary Medicine, University of Prince Edward Island 550 University Avenue, Charlottetown, Prince Edward Island, C1A 4P3 CANADA Tel: (1.902) 566.09.67 Fax: (1.902) 566.08.51 Email: [email protected]  Dr B. Dannevig National Veterinary Institute P.O. Box 750, Sentrum., 0106 Oslo NORWAY Tel: (47.23) 21.64.04 Fax: (47.23) 21.63.01 Email: [email protected] Koi herpesvirus disease  Dr Motohiko Sano National Research Institute of Aquaculture, Fisheries Research Agency 422-1 Nakatsuhamaura, Minami-ise, Mie 516-0193 JAPAN Tel: (81.599) 66.18.30 Fax: (81.599) 66.19.62 Email: [email protected]  Dr Keith Way The Centre for Environment, Fisheries and Aquaculture Science (CEFAS), Weymouth Laboratory Barrack Road, The Nothe, Weymouth, Dorset, DT4 8UB UNITED KINGDOM Tel: (44.1305) 20.66.39 Fax: (44.1305) 20.66.01 Email: [email protected] Leptospirosis  Dr R.A. Hartskeerl Royal Tropical Institute, N.H. Swellengrebel Laboratory of Tropical Hygiene, Division of Health, Department of Biomedical Research Meibergdreef 39, 1105 AZ Amsterdam THE NETHERLANDS Tel: (31.20) 566.54.38 Fax: (31.20) 697.18.41 Email: [email protected]  Dr W.A. Ellis Department of Agriculture, Veterinary Sciences Division Stoney Road, Stormont, Belfast BT4 3SD, Northern Ireland UNITED KINGDOM

XIII. Reference Laboratories for Animal Diseases

Tel: (44.2890) 51.94.41 Fax: (44.2890) 52.57.55 Email: [email protected]  Dr L. Samartino (1) Instituto de Bacteriología, Centro de Investigaciones en Ciencias Veterinarias (CICV), Instituto Nacional de Tecnología Agropecuaria (INTA) Castelar, Casilla de Correo 77, 1708 Morón, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 46.21.12.89 Fax: (54.11) 46.21.17.12 Email: [email protected]  Dr Gleyre T.D. de Mazzonelli (2) Gerencia de Laboratorios (GELAB), Servicio Nacional de Sanidad y Calidad Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martínez, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 48.36.19.92 Fax: (54.11) 48.36.19.92 Email: [email protected]/[email protected]  Dr Mark Wilson National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.73.42 Fax: (1.515) 663.76.73 Email: [email protected]  Dr L.D. Smythe WHO/FAO/OIE Collaborating Centre for Reference and Research on Leptospirosis, Western Pacific Region, Queensland Health Scientific Services 39 Kessels Road, Coopers Plains, P.O. Box 594, Archerfield, Queensland 4108 AUSTRALIA Tel: (61.7) 32.74.90.64 Fax: (61.7) 32.74.91.75 Email: [email protected] Lumpy skin disease  Dr G.H. Gerdes Onderstepoort Veterinary Institute Private Bag X05, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.91.14 Fax: (27.12) 529.94.18 Email: [email protected]  Dr Eeva Tuppurainen Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 Fax: (44.1483) 23.24.48 Email: [email protected] Maedi-visna  Dr Stephen Valas Laboratoire d'étude et de recherches caprines 60 rue du Pied de Fond, B.P. 3081, 79000 Niort

XIII. Reference Laboratories for Animal Diseases

FRANCE Tel: (33 (0)5 49.79.61.28 Fax: (33 (0)5 49.79.42.19 Email: [email protected]  Dr D.P. Knowles, Jr Animal Diseases Research Unit,USDA, ARS, Washington State University Pullman, Washington 99164-7030 UNITED STATES OF AMERICA Tel: (1.509) 335.60.22 Fax: (1.509) 335.83.28 Email: [email protected] Marek's disease  Dr K. Venugopal Institute for Animal Health, Compton Laboratory, Compton Newbury, Berkshire RG20 7NN UNITED KINGDOM Tel: (44.1635) 57.84.11 Fax: (44.1635) 57.72.63 Email: [email protected]  Dr Aly M. Fadly USDA, ARS, Avian Disease and Oncology Laboratory 33606 East Mount Hope Roas, East Lansing, Michigan 48823 UNITED STATES OF AMERICA Tel: (1.517) 337.68.29 Fax: (1.517) 337.67.76 Email: [email protected] Newcastle disease  Dr Christian Grund Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Diagnostic Virology Boddenblick 5a, 17493 Greifswald - Insel Riems GERMANY Tel: (49.383) 51.71.96 Fax: (49.383) 51.72.75 Email: [email protected]  Dr Ian Brown VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.73.39 Fax: (44.1932) 35.72.39 Email: [email protected]  Dr Paul W. Selleck CSIRO, Australian Animal Health Laboratory (AAHL) 5 Portarlington Road, Private Bag 24, Geelong 3220, Victoria AUSTRALIA Tel: (61.3) 52.27.50.00 Fax: (61.3) 52.27.55.55 Email: [email protected]  Dr B. Panigrahy National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48

XIII. Reference Laboratories for Animal Diseases

Email: [email protected]  Dr Ilaria Capua Istituto Zooprofilattico Sperimentale delle Venezie, Laboratorio Virologia Via Romea 14/A, 35020 Legnaro, Padova ITALY Tel: (39.049) 808.43.79 Fax: (39.049) 808.43.60 Email: [email protected] New world screwworm (Cochliomyia hominivorax)  Dr Agustin Sagel COPEG Panama/US Commission for the Eradication and Prevention of NWS) Apartado Postal 0816-07636 PANAMA Tel: (507) 296.00.06 Fax: (507) 296.09.69 Email: [email protected] Email: [email protected] Email: [email protected] Oncorhynchus masou virus disease  Dr M. Yoshimizu Laboratory of Biotechnology and Microbiology, Graduate School of Fisheries Sciences, Hokkaido University, Faculty of Fisheries, Microbiology Department 3-1-1 Minato-cho, Hakodate, Hokkaido 041-8611 JAPAN Tel: (81.138) 40.88.10 Fax: (81.138) 40.88.10 Email: [email protected] Ovine epididymitis (Brucella ovis)  Dr Henrich Neubauer Federal Research Centre for Virus Diseases of Animals (BFAV), Institute of Bacterial Infections and Zoonoses Naumburger Str. 96a, 07743 Jena GERMANY Tel: (49.3641) 80.42.00 Fax: (49.3641) 80.42.28 Email: [email protected]  Dr Ana Maria Nicola Gerencia de Laboratorios (GELAB), Servicio Nacional de Sanidad y Calidad Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martínez, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 48.36.19.92 Fax: (54.11) 48.36.19.92 Email: [email protected]  Dr J.A. Stack VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.76.10. Fax: (44.1932) 35.72.16 Email: [email protected]  Dr B. Garin-Bastuji

XIII. Reference Laboratories for Animal Diseases

AFSSA Alfort, Unité Zoonoses Bactériennes, Lab. OIE/FAO de référence pour la brucellose animale, Laboratoire d'études et de recherches en pathologie animale et zoonoses 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 Email: [email protected]  Dr K. Nielsen Canadian Food Inspection Agency, Animal Diseases Research Institute P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 ext. 48.04 Fax: (1.613) 228.66.69 Email: [email protected]  Dr Massimo Scacchia CESME, Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise 'G. Caporale', National Centre for Exotic Diseases Via Campo Boario, 64100 Teramo ITALY Tel: (390.861) 33.24.05 Fax: (390.861) 33.22.51 Email: [email protected]  Dr Menachem Banai Kimron Veterinary Institute Department of Bacteriology, P.O. Box 12, Beit Dagan 50250 ISRAEL Tel: (972.3) 968 16 98 Fax: (972.3) 968 17 53 Email: [email protected] Paratuberculosis  Dr Jacek Gwozdz Johne's Disease Laboratory, Research and Development Division, Department of Primary Industries 475 Mickleham Road, Attwood, Victoria 3049 AUSTRALIA Tel: (61.3) 92.17.42.00 Fax: (61.3) 92.17.42.99 Email: [email protected]  Dr Bernardo Alonso Gerencia de Laboratorios (GELAB) del Servicio Nacional de Sanidad y Calidad, Agroalimentaria (SENASA) Av. Alexander Fleming 1653, 1640 Martinez - Pcia de Buenos Aires ARGENTINA Tel: (54.11) 48.36.00.36 Fax: (54.11) 48.36.00.36 Email: [email protected] Email: [email protected]  Dr I. Pavlik Veterinary Research Institute Hudcova 70, 62132 Brno CZECH (Rep.) Tel: (420.5) 33.33.16.01 Fax: (420.5) 33.33.12.29 Email: [email protected]  Mme María Laura Boschiroli-Cara

XIII. Reference Laboratories for Animal Diseases

AFSSA Alfort, Unité Zoonoses Bactériennes, Laboratoire d'études et de recherches en pathologie animale et zoonoses 23 avenue du Général de Gaulle, 94706 Maisons-Alfort Cedex FRANCE Tel: (33 (0)1) 49.77.13.00 Fax: (33 (0)1) 49.77.13.44 Email: [email protected] Peste des petits ruminants  Prof. Tom Barrett Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 direct 213 10 09 Fax: (44.1483) 23.24.48 Email: [email protected]  Dr Geneviève Libeau CIRAD-BIOS, Control of Exotic and Emerging Animal Diseases Programme Santé animale, TA A-15/G Campus international de Baillarguet, 34398 Montpellier Cedex 5 FRANCE Tel: (33 (0)4) 67.59.37.98 Fax: (33 (0)4) 67.59.38.50 Email: [email protected] Porcine reproductive and respiratory syndrome  Dr Tomasz Stadejek National Veterinary Research Institute, Department of Swine Diseases Partyzantow Str. 57, 24-100 Pulawy POLAND Tel: (48-81) 889.30.99 Fax: (48-81) 886.25.95 Email: [email protected] Rabbit haemorrhagic disease  Dr L. Capucci Istituto Zooprofilattico Sperimentale della Lombardia e dell'Emilia Romagna, 'B. Ubertini' Via A. Bianchi n° 7/9, 25124 Brescia ITALY Tel: (39.030) 229.06.17 Fax: (39.030) 229.05.59 Email: [email protected] Rabies  Dr A. Wandeler Centre of Expertise for Rabies, Animal Diseases Research Institute 3851 Fallowfield Road, P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 Fax: (1.613) 228.66.69 Email: [email protected]  Dr J. Barrat AFSSA-LERPAS, Laboratoire d'études sur la rage et la pathologie des animaux sauvages Domaine de Pixérécourt, BP 9, 54220 Malzéville FRANCE Tel: (33 (0)3) 83.29.89.50 Fax: (33 (0)3) 83.29.89.58

XIII. Reference Laboratories for Animal Diseases

Email: [email protected]  Mme F. Cliquet AFSSA-LERPAS, Laboratoire d'études sur la rage et la pathologie des animaux sauvages Domaine de Pixérécourt, BP 9, 54220 Malzéville FRANCE Tel: (33 (0)3) 83.29.89.50 Fax: (33 (0)3) 83.29.89.58 Email: [email protected]  Dr T. Müller Institute of Epidemiology, Friedrich-Loeffler Institut, Federal Research Institute for Animal Health Seest. 55, 16868 Wustherhausen/Dosse GERMANY Tel: (49.33) 97.98.01.86 Fax: (49.33) 97.98.02.00 Email: [email protected]  Dr Claude Taurai Sabeta Onderstepoort Veterinary Institute, Rabies Unit Private Bag X05, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.94.39 Fax: (27.12) 529.93.90 Email: [email protected]  Dr Anthony Fooks Rabies and Wildlife Zoonoses Group, Virology Department, VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44.1932) 35.78.40 Fax: (44.1932) 35.72.39 Email: [email protected] Red sea bream iridoviral disease  Dr K. Nakajima National Research Institute of Fisheries Science, Fisheries Research Agency 2-12-4 Fukuura, Yokohama-shi, Kanagawa 236-8048 JAPAN Tel: (81.45) 788.76.15 Fax: (81.45) 788.50.01 Email: [email protected] Rift Valley fever  Dr G.H. Gerdes Onderstepoort Veterinary Institute Private Bag X05, Onderstepoort 0110 SOUTH AFRICA Tel: (27.12) 529.91.14 Fax: (27.12) 529.94.18 Email: [email protected]  Dr Michèle Bouloy Unité de génétique moléculaire des Bunyavirus, Département de Virologie, Institut Pasteur 25 rue du Dr Roux 75724 Paris cedex 15 FRANCE Tel: (33.1) 40.61.31.34 Fax: (33.1) 40.61.32.56 Email: E-mail: [email protected]

XIII. Reference Laboratories for Animal Diseases

Rinderpest  Prof. Tom Barrett Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 direct 213 10 09 Fax: (44.1483) 23.24.48 Email: [email protected]  Dr Geneviève Libeau CIRAD-BIOS, Control of Exotic and Emerging Animal Diseases Programme Santé animale, TA A-15/G Campus international de Baillarguet, 34398 Montpellier Cedex 5 FRANCE Tel: (33 (0)4) 67.59.37.98 Fax: (33 (0)4) 67.59.38.50 Email: [email protected] Salmonellosis  Dr R.H. Davies VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB UNITED KINGDOM Tel: (44-1932) 35.73.61 Fax: (44-1932) 35.75.95 Email: [email protected]  Dr M. Hartung Bundesinstitut für Risikobewertung (Federal Institute for Risk Assessment) P.O. Box 330013, 14191 Berlin GERMANY Tel: (49.30) 84.12.22.12 Fax: (49.30) 84.12.29.52 Email: [email protected] Web: http://www.bfr.bund.de  Dr Cornelius Poppe Laboratory for Foodborne Zoonoses, Guelph Laboratory, Health Canada, Public Health Agency of Canada 110 Stone Road West, Guelph, Ontario, N1G 3W4 CANADA Tel: (1.519) 822.33.00 Fax: (1.519) 822.22.80 Email: [email protected]  Dr Antonia Ricci Istituto Zooprofilattico Sperimentale delle Venezie, National Reference LaboratoryL for Salmonella Viale Dell'Università 10, 35020 Legnaro (PD) ITALY Tel: (39.049) 808.42.96 Fax: (39.049) 883.02.68 Email: [email protected] Scrapie  Dr Marion Simmons VLA Weybridge New Haw, Addlestone, Surrey KT15 3NB

XIII. Reference Laboratories for Animal Diseases

UNITED KINGDOM Tel: (44.1932) 35.75.64 Fax: (44.1932) 35.78.05 Email: [email protected] Web: http://www.defra.gov.uk/corporate/vla/science/science-tse-rl-web.htm  Prof. Andreas Zurbriggen Institute of Animal Neurology, University of Bern Bremgartenstrasse 109A, 3012 Bern SWITZERLAND Tel: (41.31) 631.25.09 Fax: (41.31) 631.25.38 Email: [email protected]  Dr Aru Balachandran Canadian Food Inspection Agency, Ottawa Laboratory 3851 Fallowfield Road, P.O. Box 11300, Station H, Nepean, Ontario K2H 8P9 CANADA Tel: (1.613) 228.66.98 ext. 4854 Fax: (1.613) 228.61.03 Email: [email protected]  Dr Francisco Javier Blanco Viera Laboratorio Nacional de Referencia (LNR) para Encefalopatías Espongiformes Transmisible animales, Institutos de Patobiología y Virología, Centro de Investigaciones en Ciencias Veterinarias y Agronómicas (CICV), Instituto Nacional de Tecnología Agropecuaria (INTA) Castelar, Casilla de Correo 77, 1708 Morón, Pcia. de Buenos Aires ARGENTINA Tel: (54.11) 46.21.12.89 Fax: (54.11) 46.21.17.43 Email: [email protected] Email: [email protected] Sheep pox and goat pox  Dr H.R. Varshovi RAZI Vaccine and Serum Research Institute P.O. Box 31975/148, Hessarak, Karadj, Teheran IRAN Tel: (98.21) 311.79.08 Fax: (98.261) 455.31.94 Email: [email protected] Email: [email protected]  Dr Eeva Tuppurainen Institute for Animal Health, Pirbright Laboratory Ash Road, Pirbright, Woking, Surrey GU24 ONF UNITED KINGDOM Tel: (44.1483) 23.24.41 Fax: (44.1483) 23.24.48 Email: [email protected] Spherical baculovirosis (Penaeus monodon-type baculovirus)  Prof. Donald V. Lightner Aquaculture Pathology Laboratory, Department of Veterinary Science and Microbiology, University of Arizona Building 90, Room 202 Pharmacy/Microbiology, Tucson, AZ 85721 UNITED STATES OF AMERICA Tel: (1.520) 621.84.14 Fax: (1.520) 621.48.99 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

 Dr Grace Lo Department and Institute of Zoology, National Taiwan University 1, Sec. Roosevelt Road, Taipei CHINESE TAIPEI Tel: (886.2) 23.63.35.62 Fax: (886.2) 23.63.81.79 Email: [email protected] Spring viraemia of carp  Dr Peter Dixon The Centre for Environment, Fisheries and Aquaculture Science (CEFAS), Weymouth Laboratory Barrack Road, The Nothe, Weymouth, Dorset, DT4 8UB UNITED KINGDOM Tel: (44.1305) 20.66.42 Fax: (44.1305) 20.66.01 Email: [email protected] Surra (Trypanosoma evansi)  Dr Filip Claes Institute of Tropical Medicine Antwerp, Department of Parasitology Nationalestraat 155, B-2000 Antwerpen BELGIUM Tel: (32.3) 247.65.34 Fax: (32.3) 247.63.73 Email: [email protected]  Prof. Noboru Inoue National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine Inada-cho Nishi 2-13, Obihiro, Hokkaido 080-8555 JAPAN Tel: (81.155) 49.56.47 Fax: (81.155) 49.56.43 Email: [email protected] Swine vesicular disease  Dr Jef. M. Hammond Institute for Animal Health Ash Road, Pirbright, Woking, Surrey GU24 0NF UNITED KINGDOM Tel: (44.1483) 23.12.11 Fax: (44.1483) 23.26.21 Email: [email protected]  Dr E. Brocchi Istituto Zooprofilattico Sperimentale della Lombardia e dell'Emilia Romagna 'B. Ubertini' Via A. Bianchi n° 9, 25124 Brescia ITALY Tel: (39.030) 229.03.10 Fax: (39.030) 229.03.69 Email: [email protected] Taura syndrome  Prof. Donald V. Lightner Aquaculture Pathology Laboratory, Department of Veterinary Science and Microbiology, University of Arizona Building 90, Room 202 Pharmacy/Microbiology, Tucson, AZ 85721 UNITED STATES OF AMERICA

XIII. Reference Laboratories for Animal Diseases

Tel: (1.520) 621.84.14 Fax: (1.520) 621.48.99 Email: [email protected] Tetrahedral baculovirosis (Baculovirus penaei)  Prof. Donald V. Lightner Aquaculture Pathology Laboratory, Department of Veterinary Science and Microbiology, University of Arizona Building 90, Room 202 Pharmacy/Microbiology, Tucson, AZ 85721 UNITED STATES OF AMERICA Tel: (1.520) 621.84.14 Fax: (1.520) 621.48.99 Email: [email protected] Theileriosis  Dr Dirk Geysen Department of Animal Health, Institute of Tropical Medicine Nationalestraat 155, 2000 Antwerpen BELGIQUE Tel: (32.3) 247.62.66 Fax: (32.3) 247.62.68 Email: [email protected] Transmissible gastroenteritis  Dr Linda J. Saif Food Animal Health Research Program, Ohio Agricultural Research and Development Center, The Ohio State University 1680 Madison Avenue, Wooster, OH 44691-4096 UNITED STATES OF AMERICA Tel: (1.330) 263.37.44 Fax: (1.330) 263.36.77 Email: [email protected] Trichinellosis  Dr E. Pozio Istituto Superiore di Sanita, Laboratorio di Parasitoligia Viale Regina Elena 299, 00161 Roma ITALY Tel: (39.06) 49.90.23.04 Fax: (39.06) 49.90.35.61 Email: [email protected]  Dr A. Gajadhar Canadian Food Inspection Agency, Centre for Animal Parasitology 116 Veterinary Road, Saskatoon, Saskatchewan S7N 2R3 CANADA Tel: (1.306) 975.53.44 Fax: (1.306) 975.57.11 Email: [email protected] Trypanosomosis (tsetse-transmitted)  Dr M. Desquesnes CIRAD-EMVT Programme Santé animale, TA30/G Campus international de Baillarguet, 34398 Montpellier Cedex 5 FRANCE Tel: (33(0)4) 67.59.37.24 Fax: (33(0)4) 67.59.37.98 Email: [email protected]

XIII. Reference Laboratories for Animal Diseases

Tularemia  Dr T. Mörner National Veterinary Institute, Department of Wildlife 751 89 Uppsala SWEDEN Tel: (46.18) 67.40.00 Fax: (46.18) 67.46.90 Email: [email protected] Turkey rhinotracheitis  Dr N. Eterradossi AFSSA Ploufragan, Unité de virologie, immunologie et parasitologie aviaires et cunicoles BP 53, 22440 Ploufragan FRANCE Tel: (33 (0)2) 96.01.62.22 Fax: (33 (0)2) 96.01.62.63 Email: [email protected] Venezuelan equine encephalomyelitis  Dr E.N. Ostlund Diagnostic Virology Laboratory, National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected] Vesicular stomatitis  Dr Ingrid Bergmann Centro Panamericano de Fiebre Aftosa OPS/OMS Av. President Kennedy 7778, Sao Bento, Duque de Caxias, ZC 20054-40 Rio de Janeiro BRAZIL Tel: (55.21) 36.61.90.56 Fax: (55.21) 36.61.90.01 Email: [email protected]  Dr S.L. Swenson National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected] Viral encephalopathy and retinopathy  Dr G. Bovo Istituto Zooprofilattico Sperimentale delle Venezie, Dipartimento di Ittioviroligia, Via Romea 14/A, 35020 Legnaro PD ITALY Tel: (39.049) 808.42.48 Fax: (39.049) 808.43.92 Email: [email protected]  Dr T. Nakai Hiroshima University, Graduate School of Biosphere Science, Laboratory of Fish Pathology Higashi Hiroshima 739-8528 JAPAN

XIII. Reference Laboratories for Animal Diseases

Tel: (81.82) 424.79.47 Fax: (81.82) 424.43.80 Email: [email protected] Viral haemorrhagic septicaemia  Dr Niels Jørgen Olesen National Veterinary Institute, Technical University of Denmark Hangøvej 2, 8200 Aarhus N DENMARK Tel: (45) 72.34.68.31 Fax: (45) 72.34.69.01 Email: [email protected] West Nile Fever  Dr E.N. Ostlund Diagnostic Virology Laboratory, National Veterinary Services Laboratories P.O. Box 844, Ames, IA 50010 UNITED STATES OF AMERICA Tel: (1.515) 663.75.51 Fax: (1.515) 663.73.48 Email: [email protected] White spot disease  Prof. Donald V. Lightner Aquaculture Pathology Laboratory, Department of Veterinary Science and Microbiology, University of Arizona Building 90, Room 202 Pharmacy/Microbiology, Tucson, AZ 85721 UNITED STATES OF AMERICA Tel: (1.520) 621.84.14 Fax: (1.520) 621.48.99 Email: [email protected]  Dr Grace Lo Department and Institute of Zoology, National Taiwan University 1, Sec. Roosevelt Road, Taipei CHINESE TAIPEI Tel: (886.2) 23.63.35.62 Fax: (886.2) 23.63.81.79 Email: [email protected] White tail disease  Dr A. Sait Sahul Hameed C. Abdul Hakeem College, Aquaculture Biotechnology Division, Department of Zoology Melvisharam-632 509, Vellore Dt. Tamil Nadu INDIA Tel: (91.4172) 26.94.87 Fax: (91.4172) 26.94.87 Email: [email protected] Yellow head disease  Dr P. Walker CSIRO, Aquaculture and Aquatic Animal Health (AAHL) 5 Portarlington Road, Private Bag 24, Geelong 3220, Victoria AUSTRALIA Tel: (61.3) 52.27.54.65 Fax: (61.3) 52.27.55.55 Email: [email protected]

EAZWV Transmissible Disease Fact Sheet

Sheet No.

DISEASE NAME ANIMAL GROUP AFFECTED

TRANSMISSION

CLINICAL SIGNS

FATAL DISEASE ?

TREATMENT

PREVENTION & CONTROL In houses

in zoos

Fact sheet compiled by

Last update

Fact sheet reviewed by Susceptible animal groups Causative organism Zoonotic potential Distribution Transmission Incubation period Clinical symptoms Post mortem findings Diagnosis Material required for laboratory analysis EU Reference Laboratory OIE Reference Laboratories Relevant diagnostic laboratories Treatment Prevention and control in zoos Suggested disinfectant for housing facilities Notification Guarantees required under EU Legislation Guarantees required by EAZA Zoos

1

EAZWV Transmissible Disease Fact Sheet

Sheet No.

Measures required under the Animal Disease Surveillance Plan Measures required for introducing animals from non-approved sources Measures to be taken in case of disease outbreak or positive laboratory findings Conditions for restoring disease-free status after an outbreak Contacts for further information References

2

Transmissible Diseases Handbook

INSTRUCTIONS FOR AUTHORS 1) If you plan to write a fact sheet or a general chapter on a certain topic, please INFORM THE SECRETARIAT first. In this way we can avoid that two people work on the same disease/subject at the same time. 2) ONE (OR MORE) AUTHOR(S) SHOULD BE LISTED on each fact sheet. This author will be the reference/contact person responsible for redaction and corrections. 3) Each fact sheet has to be REVIEWED BY TWO PEOPLE. Reviewers are not necessarily members of IDWG but should be specialists or at least have a minimal experience with the disease concerned. Both reviewers should be listed on the concerned fact sheet. The author has to find reviewers himself and should SUBMIT HIS FACT SHEETS TO THE SECRETARIAT AFTER REVIEW. 4) Be short in your descriptions! Each fact sheet should NOT EXCEED 4 PAGES (including references and laboratories!). 5) USE THE EMPTY FACT SHEET MODEL to write your text (write directly in the model). To become the model in an electronic format, please contact the secretariat. 6) As a text format, use ARIAL standard, size 10. 7) Don't forget to send the fact sheets to the IDWG-secretary in an ELECTRONIC FORMAT (email attachement or floppy disc). 8) NAME YOUR FILE according to the following model: "disease name" and "date of the text version". Example: Shigellosis 24.08.01 9) REFERENCE LABORATORIES: The EU and the OIE established a list of reference laboratories. National laboratories are the official labs recognized by the EU. For diseases which don’t need to be notified, no lab is mentioned in the official lists. However, if there is an important lab to recommend for the analysis, it can be mentioned under “Relevant Laboratories”. 10) The 3 points "Measures required under the Animal Disease Surveillance Plan", "Measures required for introducing animals from non-approved sources" and "Measures to be taken in case of a disease outbreak or positive laboratory findings" are conditional to the adoption of the revised Annex C of the Balai directive. Ask the secretariat for the actual situation before submitting your fact sheet. 11) AVOID LISTING TOO MANY REFERENCES. It might be more helpful to suggest a few good books for further readings. 12)

List the LITERATURE according to the existing fact sheets. Basically, the presentation follows the GUIDELINES OF THE JOURNAL OF ZOO AND WILD ANIMAL MEDICINE.

__________ May 2004

Transmissible Diseases Handbook

4th Edition, February 2010

Coordination of the work & Editor of the Handbook

Jacques Kaandorp

Co-editors

Norin Chai Ayla Bayens

Pictures (cover page)

Nico Schoemaker

Authors

In alphabetical order: Gian Lorenzo d’Alterio Luca Bacciarini Hester van Bolhuis Debra Bourne Manfred Brack J. Brandt V. Briones Norin Chaï R. De Deken Peter Dollinger Gerry Dorrestein Marein van der Hage Jesus Fernandéz Morán Scott Fitzgerald Edmund Flach Klaus Gunther Friedrich Kai Frölich Chris Furley Fabricio Gamberale S. Geerts Manuel Garcia Hartmann Ursula Höfle Jacques Kaandorp Marja Kik Antoine Leclerc Alexis Lécu Matti Kiupel Michael Lierz Rachel Marschang Marian Mensink (continued on next page)

Authors

(continuation) Joost Philippa Willem Schaftenaar Stefanie Sanderson Elvira Schettler Gabrielle Stalder Martin Straube S. Téllez Werner Tschirch Francis Vercammen Chris Walzer Marno Wolters

Reviewers

In alphabetical order: R. Adone, J. Alunda, S. Bergmann, G. Bertoni, S. Bolin, S. Blahak, M. Brack, J. Brandt, C. Buonavoglia, J. Caird, M. Coosemans, T. Cornish, A. Cunningham, O. Czupalla, R. De Deken, G. Dorrestein, P. Duff, C. Feliu, G. Ferrari, M. Fox, C. Furley, D. Geysen, G. Goodman, A. Gröne, O. Haenen, H. Hafez, J. Hatt, W. Jakob, T. Jauniaux, J. Kaandorp, E. Kaleta, M. Kiupel, T. Kuiken, A. Lécu, J. Lewis, H. Li, N. Majó i Masferrer, R. Marschang, B. Martina, R. McLean, T. Mettenleiter, D. Miller, R. Montali, J. Mortelmans, B. Neurohr, G. Paiba, T.Papp, F. Pasmans, P. Pasquali, M. Popoff, A. Pospischil, C.Reid, L. Richman, W. Rietschel, W. Schaftenaar, J. Sikarskie, J. Slingenbergh, G. Stevenson, G. Strauß, P. Sutmöller, H. Thiel, U. Truyen, F. Vercammen, W. Vosloo, T. Wahli, J. Wellehen, O. Werner, E. Williams, L. Woods, P. Zwart