These technologies offer rapid results with potentially high sensitivity and specificity, at relatively low cost. Recogn...
DNA-basedMolecular DiagnosticTechniques ResearchNeedsforStandardizationand ValidationoftheDetectionofAquatic AnimalPathogensandDiseases
FAO FISHERIES TECHNICAL PAPER
395 AUSTRALIAN CENTRE FOR INTERNATIONAL AGRICULTURAL RESEARCH
DEPARTMENT FOR INTERNATIONAL DEVELOPMENT
NETWORK OF AQUACULTURE CENTRES IN ASIA-PACIFIC
COMMONWEALTH SCIENTIFIC AND INDUSTRIAL RESEARCH ORGANIZATION
DNA-based Molecular Diagnostic Techniques:
FAO FISHERIES TECHNICAL PAPER
395
Research Needs for Standardization and Validation of the Detection of Aquatic Animal Pathogens and Diseases
Edited by Peter Walker CSIRO, Australia and Rohana Subasinghe FAO, Rome
Report and Proceedings of the Expert Workshop on DNA-based Molecular Diagnostic Techniques: Research Needs for Standardization and Validation of the Detection of Aquatic Animal Pathogens and Diseases. Bangkok, Thailand, 7-9 February 1999
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DFID
Department for International Development
The designations employed and the presentation of material in this publication do not imply the expression of any opinion whatsoever on the part of the Food and Agriculture Organization of the United Nations concerning the legal status of any country, territory, city or area or of its authorities, or concerning the delimitation of its frontiers or boundaries.
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FAO 2000
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PREPARATION OF THIS DOCUMENT This document contains the report, including recommendations, and thirteen papers presented at the Expert Workshop on DNA-based Molecular Diagnostic Techniques: Research Needs for Standardization and Validation of the Detection of Aquatic Animal Pathogens and Diseases, held in Bangkok, Thailand, from 7-9 February 1999. The Expert Workshop was jointly organized by FAO Inland Water Resources and Aquaculture Service, Network of Aquaculture Centres in Asia-Pacific (NACA), Centre for Scientific and Industrial Research for Australia (CSIRO), Australian Centre for International Agriculture Research (ACIAR), and the Department for International Development of the United Kingdom (DFID) and was held at the NACA Headquarters in Bangkok. The editing, publishing, and distribution of the document were undertaken by FAO, Rome.
Distribution: Aquatic animal health personnel Ministries and Directorates of Fisheries Participants of the Expert Workshop FAO Fishery Regional and Sub-Regional Officers FAO Fisheries Department NACA, CSIRO, ACIAR, DFID
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Walker, P. and Subasinghe, R. (eds.) DNA-based molecular diagnostic techniques: research needs for standardization and validation of the detection of aquatic animal pathogens and diseases. Report and proceedings of the Joint FAO/NACA/CSIRO/ACIAR/DFID Expert Workshop. Bangkok, Thailand, 7-9 February 1999. FAO Fisheries Technical Paper. No.395. Rome, FAO. 2000. 93p. ABSTRACT In efforts to limit trans-boundary movement of pathogens and reduce the economic and socioeconomic impact of disease in aquaculture, there is considerable scope for more effective use of DNA-based methods of pathogen detection. These technologies offer rapid results with potentially high sensitivity and specificity, at relatively low cost. Recognition of these advantages has led to rapid adoption of available DNA-based tests, particularly in shrimp culture for which histological procedures lack specificity and culture-based methods have not been possible. However, few if any of the available tests have been assessed appropriately against other diagnostic methods or standardized and validated for specified applications. In fish and shrimp, type or strain specificity of most tests for pathogens in the Asian region is poorly understood and, in molluscs, there is little information on the significant pathogens and few tests of any kind have been developed. Furthermore, tests presently available are frequently conducted by technicians who may not be sufficiently aware of the need for stringent test protocols or the meaning and limitations of the data generated. Implementation of standardized practices that produce reliable, useful and comparable data will require a significant investment in research, training and infrastructure development. Effective implementation will also be assisted by enhanced communication between aquatic animal health practitioners in the region and scientists with expertise in molecular diagnostic technologies. This review recommends development by FAO/NACA of 2 programs of managed cooperative research to assist more effective use of DNA-based detection tests. Program A should focus on improving the knowledge base by identification of new and emerging pathogens, relating pathogens in the region to those described elsewhere, and defining the extent of genetic variation between related pathogens in the region. Program B should draw on information currently available or obtained from Program A to develop suitably specific DNA-based diagnostic methods and to evaluate and validate the methods for disease diagnosis and pathogen screening programs. To increase the availability of scientists and technicians with skills in pathology and molecular diagnostic technologies, the review also recommends development of FAO/NACA-sponsored training programs for staff from key laboratories in the region. Training priorities should be in: i) the use of standard histopathological methods for health screening of fish and molluscs; and ii) the use of standard DNAbased methods for pathogen detection including sample collection, application of test protocols and the analysis and interpretation of test results. Because of the urgency of disease problems and the availability of suitable tests, training in DNA-based methods should focus initially on detection of shrimp pathogens. The review also recommends the development of a laboratory accreditation program in order to achieve standardization of sampling methods and test procedures. The establishment of reference laboratories will assist accreditation for each of the major pathogens. Laboratory accreditation and training programs should complement the activities of OIE in obtaining internationally agreed test standards for molecular diagnostic technologies.
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CONTENTS 1
BACKGROUND........................................................................................................................................... 2 1.1 1.2 1.3 1.4
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EXPERT WORKSHOP ............................................................................................................................... 4 2.1 2.2 2.3
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GLOBAL AQUACULTURE DEVELOPMENT .................................................................................................... 2 IMPACT OF DISEASE ................................................................................................................................... 2 ROLE OF DNA-BASED TECHNOLOGIES IN DIAGNOSIS AND PATHOGEN DETECTION .................................... 2 IMPEDIMENTS TO THE USE OF DNA-BASED DIAGNOSTIC TECHNIQUES ...................................................... 3
TERMS OF REFERENCE ............................................................................................................................... 4 PARTICIPANTS ........................................................................................................................................... 4 WORKSHOP PROCESS ................................................................................................................................. 5
PATHOGEN FOCUS GROUP REPORTS................................................................................................ 5 3.1 SHRIMP PATHOGENS .................................................................................................................................. 5 3.1.1 Status of research and identification of research needs ............................................... 5 3.1.2 Standardization of DNA-based diagnostic tests ........................................................... 6 3.1.3 Networking and communication ................................................................................. 7 3.1.4 Training and extension .............................................................................................. 7 3.2 MOLLUSC PATHOGENS............................................................................................................................... 8 3.3 FINFISH PATHOGENS .................................................................................................................................. 9 3.3.1 Diseases requiring application of DNA-based technologies .......................................... 9 3.3.2 Evaluation of needs for rapid detection techniques ...................................................... 9 3.3.3 Status of research..................................................................................................... 9 3.3.4 Key constraints to establishment of international standards........................................ 10 3.3.5 Recommendations for research programs ................................................................ 10
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ISSUES RELATING TO THE ADOPTION OF INTERNATIONAL STANDARDS FOR DISEASE DIAGNOSIS AND HEALTH CERTIFICATION................................................................................... 11
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KEY OUTCOMES AND RECOMMENDATIONS ................................................................................ 11 5.1 5.2 5.3 5.4 5.5
GENERAL COMMENTS .............................................................................................................................. 11 RESEARCH NEEDS .................................................................................................................................... 12 TRAINING NEEDS ..................................................................................................................................... 13 COMMUNICATION NEEDS ......................................................................................................................... 13 INTERNATIONAL STANDARDISATION ....................................................................................................... 14
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REFERENCES ........................................................................................................................................... 14
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CONTRIBUTED PAPERS AND REVIEWS........................................................................................... 16
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ANNEX I - LIST OF PARTICIPANTS.................................................................................................... 92
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Background
1.1 Global aquaculture development In 1995, global production from aquaculture reached 27.8 million tones and was valued at US$ 42,300 million. Developing countries contributed over 87% of total production, of which 90.1% was from Asia. China contributed 63.4% of total world production. Over the past decade, global aquaculture production has grown at an average annual rate of 9.6% compared to 3.1% for livestock meat and 1.6% for capture fisheries. Between 1984 and 1995, growth in aquaculture production in low-income food deficit countries (LIFDCs) was over five times faster than in developed countries (Rana, 1997).
1.2 Impact of disease Disease outbreaks are recognized as a significant constraint to aquaculture production and trade, affecting both the economic development and socioeconomic revenue of the sector in many countries in the world. According to Chamberlain (in press), disease is a primary limiting factor for shrimp farming today and the risk of disease losses is likely to increase as the shrimp sector continues to grow. Economic loss attributed to outbreaks of disease in developing countries in the Asian region was estimated to be at least US$ 1,400 million in 1990 (ADB/NACA, 1991). The cost of lost production in China alone was approximately US$ 1,000 million in 1993. In Thailand, the loss in 1996 due to yellow head virus (YHV) and white spot syndrome virus (WSSV) was estimated to be 40% of total production (70,000 tones) valued at over US$ 500 million (Alday-Sanz and Flegel, 1997). Recent estimates, based on farm level surveys in 16 Asian countries, suggest that disease and environment-related problems have caused annual losses of more than US$ 3,000 million to aquaculture production (ADB/NACA, in press). Serious financial losses have also been recorded in other regions of the world. In 1993, Ecuador lost 28,000 tones of shrimp production in due to an epizootic of Taura syndrome virus. Salmon farming in many countries also faced serious disease problems that resulted in significant production losses. Various factors have been related to the apparent increased incidence of disease. Environmental factors and poor water quality, sometimes resulting from increased self-pollution due to effluent discharge and pathogen transfer via movements of aquatic organisms appear to be an important underlying cause of such epizootics.
1.3 Role of DNA-based technologies in diagnosis and pathogen detection The effective control and treatment of diseases of aquatic animals requires access to diagnostic tests that are rapid, reliable and highly sensitive. In many cases, post-mortem necropsy and histopathology have been the primary methods for the diagnosis of fish and shellfish diseases. However, these methods often lack specificity and many pathogens are difficult to detect when present in low numbers or when there are no clinical signs of disease (Ambrosia and De Wall, 1990). Direct culture of pathogens is also widely used. However, these methods are time-consuming and costly, and, for shrimp and other crustaceans, cell lines suitable for virus culture have not been available. Efforts to overcome these problems have led to the development of immunoassay and DNA-based diagnostic methods including fluorescent antibody tests (FAT), enzyme-linked immunosorbent assays (ELISA), radioimmunoassay (RIA), in situ hybridization (ISH), dot blot hybridization DBH) and polymerase chain reaction (PCR) amplification techniques. The use of DNA-based methods derives from the premise that each species of pathogen carries unique DNA or RNA sequences that differentiate it from other organisms. The techniques offer high sensitivity and specificity, and
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diagnostics kits allowing rapid screening for the presence of pathogen DNA are moving rapidly from development in specialized laboratories to routine application. DNA probes are expected to find increasing use in routine disease monitoring and treatment programs in aquaculture, in field epidemiology and in efforts to prevent the international spread of pathogens (national quarantine and certification programs). DNA-based methods have been used in diagnosis and for detection of many economically important viral pathogens of cultured finfish and penaeid shrimp. For finfish, tests have been developed for pathogens such as channel catfish virus (CCV), infectious hematopoietic necrosis virus (IHNV), infectious pancreatic necrosis virus (IPNV), viral hemorrhagic septicemia virus (VHSV), viral nervous necrosis virus (VNNV) and Renibacterium salmoninarum (see Muroga, 1997; Plumb, 1997). PCR has been used in Japan to screen striped jack (Pseudocaranx dentex) broodstock for VNNV, permitting selection of PCR-negative spawners as an effective means of preventing vertical transmission of this pathogenic virus to the larval offspring (Muroga, 1997). DNA-based detection methods for detection of penaeid shrimp viruses are now used routinely in a number of laboratories around the world. These include probes for such diseases as white spot syndrome virus (WSSV), yellow head virus (YHV), infectious hematopoietic and infectious hypodermal and haematopoeitic necrosis virus (IHHNV) and Taura syndrome virus (TSV) which pose the greatest threat to world shrimp culture production (Lotz, 1997). DNA probes have also been developed for an intracellular parasites and bacteria infecting shrimp. DNA-based techniques will have an important role to play in efforts to develop sustainable shrimp culture in Asia and elsewhere. Production facilities in Thailand are currently using PCR techniques to screen shrimp post-larvae for WSSV. Culturing such larvae in closed (biosecure) or semi-closed culture systems can prevent or minimize viral infections, leading to a viable shrimp industry. The development of specific pathogenfree shrimp stocks will also depend on the use of such techniques. The further development and use of DNA-based diagnostic techniques will also assist international efforts to control the introduction of exotic diseases into new geographic areas. Reliable and rapid techniques are needed by national and regional diagnostic laboratories to screen imported fish and shellfish for important pathogens. The Office International des Epizootics (OIE) or World Animal Health Organization, is a veterinary organization with 147 member countries. The OIE (through its Fish Diseases Commission) is responsible for tracking diseases of fish and shellfish that have a serious economic impact on aquaculture and capture fisheries. There is considerable potential to apply DNA-based methods for OIE testing if they can meet the stringent criteria of a standardized, validated, accurate, reliable and accessible diagnostic technique.
1.4 Impediments to the use of DNA-based diagnostic techniques Although offering considerable potential, the routine use of DNA-based diagnostic techniques is hampered by a number of potential problems (Chanratchakool et al., 1998). x The extreme sensitivity of these methods allows the detection of target DNA present at very low levels. However, positive results provide little quantitative assessment of the infection level, and do not indicate whether the pathogen is replicating or causing disease in the species tested. Thus, carrier status and viability of the pathogen are not determined using DNA-probes. x The extremely high specificity of these tests, coupled with the ability of many viruses to rapidly change in genetic structure, can result in failure to detect a virus that has altered its genetic profile.
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x Large differences in sensitivity are related to the PCR method used (e.g., 1-step PCR or 2-step PCR with nested primers). x PCR methodologies are highly susceptible to contamination. Contamination during processing may result in false positives, particularly in 2-step PCR methods. PCR tests must be conducted in very well managed, clean laboratories. x "False negatives" are easily caused by the selection of inappropriate host tissue sources for detection of the pathogen in question, incorrect choice of DNA extraction method, or low pathogen prevalence in the population sampled. DNA-based detection and diagnostic methods have the potential for widespread application of in aquaculture. As the technology is already being adopted rapidly in developing countries in Asia, there is an urgent need to address these issues and to develop an action plan for research and training activities that will facilitate more effective utilization.
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Expert Workshop
2.1 Terms of reference The terms of reference were as follows: x Identify and prioritize research areas where the introduction and use of nuclear and related technologies is likely to have the most significant impact on improving disease diagnosis, with emphasis on diseases affecting aquaculture, in developing countries. x Evaluate needs for rapid diagnostic techniques for the principal diseases of cultured fish and shellfish. x Review the status of research towards meeting these needs. x Identify problems and key constraints related to establishing international standards for protocols and procedures for such tests and make recommendations towards their solution x Make recommendations for programs of research to be developed jointly by IAEA, FAO, and other interested and concerned agencies and institutions, to assist developing countries to develop, standardize and validate nuclear related, DNA-based rapid diagnostic tools for major aquatic animal pathogens.
2.2 Participants Dr Alexandra Adams, University of Stirling, Scotland. Dr Franck Berthe, IFREMER, France. Dr Eugene Burreson, Virginia Institute of Marine Science, Virginia, USA. Dr Pornlerd Chanratchakool, Aquatic Animal Health Research Institute, Bangkok, Thailand. Dr Supranee Chinabut, Aquatic Animal Health Research Institute, Thailand. Mr Dan Fegan, Natl. Centre for Genetic Engineering and Biotechnology, Bangkok, Thailand. Prof. Timothy Flegel, Maihidol University, Bangkok, Thailand. Dr Barry Hill, Fish Diseases Commission, OIE, Weymouth, England. Dr Mike Hine, National Institute of Water and Atmospheric Research, New Zealand. Dr Maura Hiney, National University of Ireland, Galway, Ireland.
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Dr Indrani Karunsagar, University of Agricultural Sciences, Mangalore, India. Prof. Donald Lightner, Arizona State University, Tucson, USA. Dr James Lilley, Institute of Aquaculture, University of Stirling, Scotland. Dr Sharon McGladdery, Gulf Fisheries Centre, New Brunswick, Canada. Dr Gary Nash, Shrimp Culture Research and Development Company, Thailand. Dr Michael Phillips, NACA, Bangkok, Thailand. Dr Krishen Rana, FAO, Rome, Italy. Dr Melba Reantaso, NACA, Bangkok, Thailand. Prof. Mohamed Shariff, University Putra Malaysia, Serdang, Malaysia. Dr Rohana Subasinghe, FAO, Rome, Italy. Dr Kamonporn Tonguthai, Aquatic Animal Health Research Institute, Bangkok, Thailand. Dr Peter Walker, CSIRO Tropical Agriculture, Brisbane, Australia. See Annex I for details.
2.3 Workshop process The participants assembled a team of experts currently working on the development of DNA-based rapid diagnostic techniques for the detection of aquatic animal pathogens, and representatives from other concerned agencies. With assistance from several cooperating agencies (FAO, NACA, ACIAR, CSIRO, and DFID), all experts participated in a workshop at NACA Headquarters (Bangkok, Thailand) on 7-9 February, 1999. The workshop comprised a series of papers on issues related to the use and limitations of DNA-based diagnostic technologies and related research needs, and a series of selected focus groups considering finfish, mollusc and shrimp pathogens. In March and April 1999, Dr Franck Berthe (IFREMER, France) and Dr Peter Walker (CSIRO, Australia) conducted consultancies at FAO, Rome to consider the outputs of the workshop and to assemble this report.
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Pathogen focus group reports
3.1 Shrimp pathogens Ponlerd Chanratchakool, Dan Fegan, Tim Flegel, Indrani Karunsagar, Don Lightner, Gary Nash, Mohamed Shariff, Peter Walker.
3.1.1 Status of research and identification of research needs Shrimp pathogens in the Asia-Pacific region presently listed by NACA include WSSV and YHV (notifiable), INNHV, GAV, MBV, BMNV and SMV (significant pathogens). HPV also may have significant effects on production. The range of histopathological and molecular techniques available for detection of these agents has recently been reviewed (Lightner and Redman, 1998). Research needs pertaining to the application of DNA-based technologies vary, reflecting the stage of development of the technology and the relative importance of the pathogens. In view of their ongoing impact on shrimp aquaculture in Asia and OIE-notifiable status, this report has focussed primarily on the research status and needs for WSSV, YHV and viruses in the YHV complex. The urgency of implementing measures to control these viruses is presently the dominant concern for shrimp health in Asia.
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WSSV. A number of PCR, nested-PCR and hybridization tests has been developed for WSSV detection. The tests use a range of different PCR primers and hybridization probes targeted to different and poorly defined sites in the WSSV genome. Methods of DNA sample preparation and PCR test protocols vary and there has been no objective comparison of the sensitivity and specificity of the tests. A recent analysis of PCR products amplified from WSSV samples obtained from a wide geographic range has indicated remarkable uniformity. However, the data suggested that at least one WSSV-related virus in the USA may be distinct (Lo et al., 1999). Other reports also suggest some variability in the detection specificity of PCR tests (Park et al., 1998; R.A.J. Hodgson and P.J. Walker, unpublished data). There is evidence that a range of crustaceans and other arthropods test positive by PCR (Lo et al., 1996; Alday-Sanz and Flegel, 1997; Maeda et al., 1998), but the significance of these results for the epidemiology WSSV infection in shrimp is unclear. A more extensive analysis of WSSV variation should be conducted to determine the implications for detection and disease diagnosis. The proliferation of PCR tests and protocols for WSSV detection also presents problems for comparative validation. The one-step and nested PCR primers and procedures described by Lo et al. (1996) are well documented in the published literature and the utility of the tests has been demonstrated for a range of applications. The test appears to be reliable and not subject to commercial restrictions, and would be suitable as a primary reference for standardization purposes. Estimation of the level of WSSV infection may have important applications in disease management strategies and this should be more clearly defined. It may be useful to adopt a quantitative PCR test as a secondary reference standard. YHV. The YHV complex constitutes a group of related agents which includes yellow head virus (YHV), gill-associated virus (GAV) and lymphoid organ virus (LOV). YHV and GAV are closely related but distinct pathogens; LOV is a variant of GAV which occurs in healthy shrimp. Yellow head disease has been reported from many countries in the Asian region but, in most cases, the agents have not been clearly defined. To date, YHV has been shown to occur only in Thailand and GAV/LOV only in Australia. An RT-PCR test is available for YHV but the test does not detect GAV. RT-PCR and nested RT-PCR tests are available for GAV. The GAV RT-PCR also detects at least some isolates of YHV but the test will not distinguish GAV and LOV. In situ hybridization probes have also been developed for YHV and GAV. The YHV probe detects both YHV and GAV. All tests developed to date have targeted sequences in the polymerase gene. More research is required to determine the range and distribution of YHV complex viruses in the region, to identify other possible members of the complex and to develop both pan-specific and typespecific detection tests. If possible, tests should also be developed to differentiate pathogenic from non-pathogenic variants. There may be value in the adoption of multiplex PCR tests for YHV that allow discrimination of viruses in the complex.
3.1.2 Standardization of DNA-based diagnostic tests A program of laboratory accreditation is proposed in order to achieve reliability and comparability of DNA-based test results in the Asia-Pacific region. The program should seek primarily to provide an improved regional capacity for effective disease control. However, if adequate and common levels of test performance can be obtained, the program may also provide a mechanism for eventual adoption of molecular-based methods for health certification by OIE.
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It is proposed that FAO/NACA accreditation should be issued by reference laboratories designated for each pathogen. The reference laboratory should publish a detailed standard protocol including procedures for sample collection, DNA/RNA preparation, PCR reagent preparation and storage, PCR amplification and sample analysis, and preparation and use of control reagents. Adoption of the standard protocol will be facilitated by publication in local languages. The reference laboratories should also be responsible for: x maintaining an agreed test as the primary reference standard against which other testing protocols should be assessed; x maintaining standard PCR reagents including primers and suitable positive and negative controls; x monitoring standards and providing technical advise to accredited laboratories; x providing definitive diagnosis in difficult or unusual cases; x retaining and archiving virus isolates for reference; x ongoing assessment and Research and Development of DNA-based testing protocols. Accreditation should initially focus on WSSV diagnosis. Because of the proliferation of WSSV PCR testing protocols, a primary reference standard should be identified. It is proposed that the one-step and nested WSSV PCR tests described by Lo et al. (1996a; 1996b) should be adopted as the primary standard. Implementation of a regional accreditation program should proceed following the issue of standard protocols. A relative evaluation of the diagnostic capabilities of participating laboratories should be conducted using standard coded samples of extracted DNA and shrimp tissue. An epidemiologist should be involved in the design of the evaluation and data analysis. The evaluation should be used as a basis for the development of an accreditation protocol. Standardization and accreditation for YHV diagnosis is considered to be premature, as there is presently insufficient information available on the relationship of viruses that constitute the YHV complex. The establishment of a WSSV accreditation program will facilitate future accreditation of laboratories for diagnosis of YHV and other shrimp pathogens.
3.1.3 Networking and communication NACA should be the conduit for maintaining communication between regional diagnostic laboratories, preferably through an email network. The establishment of an accreditation program should involve an initial meeting of participating laboratories to set up comparison protocols.
3.1.4 Training and extension There is a need for training both at farm-level and of diagnostic practitioners. Practitioner training should be through intensive short courses which include both theory and practice of DNA-based technologies and high quality tertiary courses for veterinarians and fish pathologists. Farm-level training is best achieved through in-country training of extension officers to meet local needs and should include: i) instruction on sample collection and preparation methods; ii) accurate interpretation of PCR results; iii) limitations of PCR technology; and iv) basic epidemiology.
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Standard procedures for the application of DNA-based technologies for disease prevention should also be documented. These should include the use of PCR in hatcheries and on-farm, and for the selection of SPF broodstock.
3.2 Mollusc pathogens Franck Berthe, Eugene Burreson, Mike Hine, Sharon McGladdery, Mike Phillips, Melba Reantaso. With the possible exception of Australia and New Zealand, there is a lack of published or readily available information on molluscan parasites, pests and diseases in the Asia-Pacific region. Most countries lack dedicated expertise, facilities and infrastructures for molluscan health examination. However, in many countries, mollusc production (subsistence and aquaculture) is established and growing. Species under cultivation for food or secondary products include pearl oysters, edible oysters, mussels, scallops, abalone and capiz shell. Although there is no immediate application for use of DNA-based diagnosis methods on a routine basis for molluscs, tools already developed could be used for cross-checking the specific/generic identities of emerging parasites which appear related to known pathogens e.g. Haplosporidium spp., Marteilia spp. and Perkinsus spp. Unpublished data from Australia indicates the presence of Haplosporidium sp. in Pinctada maxima, and Perkinsus sp. in Saccostrea commercialis. In view of the presence of potential pathogens, the reported growth of mollusc aquaculture industries and the increasing pressure for live introductions and transfers, there is an urgent need for a survey of normal and diseased animals to obtain more extensive data on mollusc health. There is also a need to establish national and regional expertise through training. As a prerequisite to implementation, NACA should contact National Coordinators to: i) confirm support for a mollusc health program; ii) identify commercially significant species; and iii) designate at least two technicians who would be dedicated to the project. NACA should also establish a panel of experts (from within and outside the region) who will support the project and provide reagents and reference material. Initially, training will aim to establish an initial baseline of expertise in husbandry, sampling, gross observations, fixation, anatomy, and histology. This first step would provide the basis for national programs for mollusc health monitoring. Where resources are limited, this could include market samples. Initial training would be followed (3-4 months later) by an advanced session for the same participants covering histopathology and technical problem solving. During this session, each participant would bring examples of their own material for joint consultation and evaluation. In addition, contacts with mollusc pathology specialists and reference material from other collections (slides and guides) should be available for these participants. The goal of this training program would be to establish a capability for independent monitoring of mollusc health in each country. As a result of this training program, a clearer picture of the health status of molluscs and the diagnostic needs in the region should emerge. The connection between this initiative and the supporting network (comprising specialist mollusc pathology laboratories) will enable access to the advanced diagnostic tools discussed in detail in the full workshop forum (for finfish, shrimp and molluscan pathogens). The mollusc pathogen Focus Group determined that, in the Asia-Pacific region, these methods should be reserved for cross-checking material containing organisms resembling known pathogens for which advanced diagnostic methods are available in supporting laboratories (e.g., Marteilia spp. DNAprobes). They could also be used to determine the geographic and host distributions of these pathogens as they emerge. DNA sequence analysis should also be conducted (e.g. to confirm positive probe results) to validate application of available probes to pathogens in the region. The need to develop additional molecular diagnostic tools and other pathogen/pathogen-group research needs will be determined as pathogens emerge from this surveillance program.
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3.3 Finfish pathogens Alexandra Adams, Supranee Chinabut, Barry Hill, Maura Hiney, James Lilley, Kamonporn Tonguthai.
3.3.1 Diseases requiring application of DNA-based technologies There are 3 areas for which the introduction and use of DNA-based techniques is likely to have a significant impact on improving disease diagnosis in developing countries. Mycobacteriosis. As there is a zoonosis risk associated with M. marinum, there is a special need to identify mycobacterium infections to the species level. Currently available PCR methods should be standardized and validated to the extent that stocks can be certified (see paper by Adams, Appendix 1). Viral nervous necrosis (VNN). VNN is a potential threat to an important grouper industry in the region. DNA probes are available to particular strains in Europe and possibly Australia. A comparative study of available probes is required and a validated method for screening wild grouper broodstock should be developed. Epizootic ulcerative syndrome (EUS). EUS infections cannot presently be distinguished from occurrences of ulcerative mycosis in USA. There is a need for a standardized, validated in situ hybridization test that is specific for A. invadans. The development of PCR primers and/or hybridization probes for the EUS-associated rhabdovirus would also assist in understanding the disease syndrome (see Lilley and Chinabut, Appendix I)). In addition to these 3 areas, there is a need for basic research on potentially emerging diseases, including red spot disease and streptococcal infections.
3.3.2 Evaluation of needs for rapid detection techniques The majority of pathogens causing the principal diseases of fish in the Asia-Pacific region can be detected and identified using existing methodologies. These include visual observation and light microscopy for parasitic infections, and routine bacteriology and histology for bacterial pathogens. Specific staining methods and antibody-based techniques are also utilized. Cell culture is used for grown and identification of viral pathogens. The primary fungal infection (EUS) is detected by histology. However, DNA-based technologies would assist in controlling the spread of fish diseases disease if used for confirmation of diagnosis and for screening fish for specific pathogens such as M. marinum, VNNV and A. invadans.
3.3.3 Status of research Research is already underway on all three areas: Mycobacterium spp. Monoclonal antibodies (MAbs), PCR primers and DNA probes are available. The MAbs can discriminate M. marinum at species level but do not detect all isolates. New MAbs are currently being developed. Two PCR-based methods have been developed, one employing enzyme restriction analysis of the PCR product, and the other employing reverse cross-blot analysis. The later appears to be more specific and can detect all 3 species infecting fish (M. marinum, M. fortuitum and M. chelonei). This test is presently used to detect aquatic mycobacteria in fish tissue samples, water samples and in human clinical biopsies but it requires further standardization and field validation.
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VNNV. VNN is present in Australia and has recently been detected in grouper in Thailand. Diagnostic antibodies, PCR primers and DNA probes have been developed for VNNV isolates from the Mediterranean region where the disease causes widespread mortalities. These reagents should be used to determine their specificity for VNNV isolates from the Asia-Pacific region. EUS. PCR and in situ hybridization tests are currently being developed to detect A. invadans from Asia. These should be utilized to screen samples from different geographical locations for the presence of A. invadans. Primers should also be developed against the rhabdovirus which is often isolated in association with EUS.
3.3.4 Key constraints to establishment of international standards The need for a more formal set of protocols and procedures for PCR-based diagnostic assays has been recognized for a number of years. However, as in many other areas of diagnostics, there has been no serious attempt to set agreed international standards. For a technique with the potential power PCR, the lack of clear guidelines for the correct performance is a serious constraint to its routine use. A second and more serious constraint is the almost complete absence of validation studies undertaken on field samples for any of the currently available assays (see Hiney, Appendix I). Although validation programs are expensive and time consuming, and carefully designed laboratory studies can provide much of the data required to ensure that PCR-based assays perform with acceptable precision, information on the meaning of the results generated by non-culture-based diagnostic techniques can only be obtained through field validation programs. Therefore, collaborative projects to assess currently available PCR-based diagnostic techniques through comparative and predictive field validation studies are urgently required and should be actively supported by funding agencies.
3.3.5 Recommendations for research programs Adoption of PCR-based diagnostic assays, to replace more established methods is an attractive option for many laboratories. These assays are seen as more sensitive, rapid and logistically simple, and have the allure of high technology. However, in the absence of adequate internal (laboratory) and external (field) validation, interpretation of the meaning of the results generated by the PCR-based assays will remain problematic. While an assay may be acceptable in a research context it will not be acceptable in either a diagnostic or regulatory context and could generate misleading data. Therefore, the priority of any research program that aims to replace current diagnostic methods with PCR-based assays must be a carefully considered validation program that addresses the issue of interpretation of meaning. As a first approach, comparative validation programs should be conducted in which PCR-based assays are performed in parallel with established methods over a reasonable period of time and using statistically significant number of samples. This type of validation program would also allow standardization of the performance of the assay in the laboratory. Standard reference material (both positive and negative controls) supplied from a central reference laboratory would be important to ensure the precision of the results generated by such a study. More importantly, it is essential that results generated by PCR-based diagnostic assays can be related to field situations. Positive or negative results generated by the assay should be reliably related to actual disease episodes. It is only through such ‘predictive validation’ that interpretation of the results in relation to disease diagnosis is meaningful. Some components of a predictive validation study could be conducted retrospectively if field data of adequate quality were available. The 3 fish diseases (mycobacteriosis, VNN and EUS) highlighted in this report are recommended for future research.
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Issues relating to the adoption of international standards for disease diagnosis and health certification
In 1995, the World Trade Organisation (WTO) implemented the Agreement on the Application of Sanitary and Phyto-Sanitary Measures (SPS Agreement) to define the conditions under which countries can impose sanitary conditions on imports of animal and animal and plant products. This Agreement is intended to promote international trade by requiring all member states to use only international standards for reducing disease risks associated with imported products. For animals and animal products, the Agreement identifies the guidelines and recommendations established by the Office International des Epizooties (OIE) as the appropriate and preferred international standard. However, the Agreement also allows governments to use standards developed by other relevant international organisations whose membership is open to all WTO members, or to use higher standards if an appropriate risk assessment provides adequate scientific justification. In Europe, EU Member States have agreed common standards for the health conditions applying to intra-Community trade in aquaculture animals and their products, based on those recommended by OIE. Countries in the Asia-Pacific Region may also wish to agree on common standards of diagnostic and health assurance tests. However, if international standards established in a particular region differ significantly from those recommended by the OIE, trade with countries outside the region may be adversely affected. For example, if the standard adopted in a region is lower than the OIE standard, countries which apply the OIE standards could rightly refuse imports on the basis that they have insufficient health guarantees. Conversely, if the standard agreed in a region is higher than those of OIE, the same standard must be applied to imports from other regions, to ensure equivalence and protection of the regional health situation. Furthermore, if an individual country adopts higher health protection standards, the SPS Agreement requires scientific justification based on import risk assessment. For diseases not listed by the OIE, trading countries may agree on any mutually acceptable standards. If a regional training and laboratory accreditation program for molecular diagnosis is to be established in the Asia-Pacific region, the adoption of tests recommended in the OIE Diagnostic Manual for Aquatic Animal Diseases would assist in achieving universally agreed standards. However, as the Manual is revised only every 3 years or so, better methods may emerge from research before the next edition is published. In considering a case for adoption of new molecular diagnostic methods, the suitability must be rigorously assessed in consultation with independent experts. Acceptance is more likely if the method has been published, has received wide scientific acceptance and has been standardised, and preferably validated, in comparison with other standardised methods.
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Key outcomes and recommendations
5.1 General comments There is considerable scope for more effective use of DNA-based methods of pathogen detection and disease diagnosis in Asia-Pacific aquaculture. However, implementation of standardised practices that produce reliable, useful and comparable data will require a significant investment in research, training and infrastructure development. Effective implementation will also be assisted by enhanced communication between aquatic animal health practitioners in the region and scientists with expertise in disease diagnosis and pathogen detection.
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Although there are some common themes, it is also evident that there are significant differences in the current relevance of DNA-based methods of pathogen detection for the different aquaculture sectors. DNA-based methods are particularly suitable for detection and diagnosis of shrimp and mollusc pathogens because of the absence of an antibody response in invertebrates and lack of suitable cell lines for virus cultivation. In shrimp, the primary pathogens are well known and many DNA-based methods have already been developed. However, in molluscs there is very limited knowledge of pathogens and few diagnostic procedures of any kind are being employed in the Asia-Pacific region. In fish, antibody and culture-based diagnostic methods are available and considered to be robust and effective for routine diagnostic applications. As such, DNA-based methods in fish appear to be most suitable for confirmatory diagnosis and rapid screening of low level or unapparent infections. To achieve maximum impact, it is essential that research and training programs recognise these differences and are tailored to reflect current levels of knowledge and sector-specific needs. Where DNA-based tests are available and/or suitable, the most significant impediment to effective implementation is the lack of standardised methodologies that are validated for specific applications. There is a need for international agreement on methodologies that have been rigorously evaluated and accredited for specific applications in disease diagnosis and pathogen screening. There is also a need to ensure that tests are performed by trained staff with access to standardised reagents and suitably equipped laboratories. Because of existing limitations on the reliability and accessibility of the methods, international standards recommended by OIE do not presently include DNA-based methodologies. However, the potentially high sensitivity and specificity and relatively low cost of these tests has resulted in a surprisingly rapid adoption rate in Asia, particularly for shrimp pathogens. Therefore, it is essential that DNA-based tests are assessed on their merits against existing technologies and that programs to achieve improved performance and international standardisation should be developed. It is also essential that these programs should assist and complement the activities of OIE in obtaining internationally agreed test standards.
5.2 Research needs There are a number of pathogens for which DNA-based test methodologies are published or available commercially. However, in general, further research is required before standardised and validated DNA-based test protocols can be implemented for disease diagnosis and pathogen detection in the major aquaculture sectors in the Asia-Pacific region. Research needs vary for each pathogen depending on the existing knowledge base and state of the technology. Recommendation Programs of international research cooperation should be developed and coordinated by FAO/NACA. The research should be conducted by managed collaborative networks and provide the information and technology necessary for delivery of suitably specific and validated tests for pathogens of fish, shrimp and molluscs in the Asia-Pacific region. Two research programs are proposed: Program A: Identification and characterisation of potential pathogens of molluscs, shrimp and fish in the Asia-Pacific region. This program should focus on improving the knowledge base by identification of new and emerging pathogens (through health screening, epidemiological investigation and subsequent molecular characterisation), relating pathogens in the region to those described elsewhere, and defining the extent of genetic variation between related pathogens in the region. The program should include the following priority projects:
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x x x x x
Health screening and pathogen identification in molluscs; Characterization of WSSV and YHV strain and pathotype variation in prawns; Characterization of Haplosporidium, Marteilia and Perkinsus spp. infecting molluscs in Asia; Characterization of VNNV strain variation in grouper and other fish of economic importance; Characterization of emerging fish diseases including red spot and streptococcal infections.
Program B: Development and validation of DNA-based diagnostic and detection methods for diseases of aquaculture in the Asian region. This program should draw on information currently available or obtained from Program A to develop suitably specific DNA-based diagnostic methods and to evaluate and validate the methods for disease diagnosis and pathogen screening programs. The research program should include the following priority projects: x Standardisation and validation of group and strain-specific DNA-based detection tests for WSSV and YHV-complex viruses; x Development and validation of species and strain-specific DNA-based detection tests for mycobacteriosis, viral nervous necrosis and epizootic ulcerative syndrome in Asia-Pacific; x Development and validation of DNA-based detection tests for Haplosporidium, Marteilia and Perkinsus spp. in Asia-Pacific.
5.3 Training needs The implementation of effective DNA-based diagnosis is severely constrained by the availability of scientists and technicians with skills in pathology and molecular diagnostic technologies. Recommendation. FAO/NACA should develop training programs for staff from key laboratories in the region. Training is required in the following priority areas: x x
The use of standard histopathological methods for health screening of fish and molluscs. The use of standard DNA-based methods for pathogen detection including sample collection, application of test protocols and the analysis and interpretation of test results. Initially, training should focus on detection of shrimp pathogens.
5.4 Communication needs There is a need to improve communication links between practitioners and scientists with recognised expertise in disease diagnosis and pathogen detection. Recommendation. FAO/NACA should establish and maintain sector-based (fish, molluscs, shrimp) communication networks of diagnostic practitioners and internationally recognised experts in aquatic animal health. Activities of the networks should include: x Exchange on information pathogen distribution in the Asia-Pacific region and the availability of diagnostic tests and reagents; x Development of cooperative research projects and training programs; x Development of cooperative programs for test validation and laboratory accreditation.
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5.5 International standardisation Lack of standardisation of tests and test protocols is a major impediment to effective implementation of DNA-based methods in the Asia-Pacific region. Standardisation requires international agreement and cooperation in test selection, practitioner training and laboratory accreditation. Improvements in the reproducibility, validity and comparability of data resulting from accreditation will also assist OIE in assessing the suitability of DNA-based methods for detection of listed pathogens. Recommendation. FAO/NACA should develop a program of accreditation of standard DNA-based tests and laboratories with the required standards of operation and expertise to conduct the tests effectively. The program should be administered by NACA through pathogen-specific reference laboratories with the following functions: x x x x
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Maintain accredited tests and reagents including reference standards; Monitor standards and provide technical advise to accredited laboratories; Provide definitive diagnosis in difficult or unusual cases; Archive pathogens for future reference.
References
ADB/NACA. (1991). Fish Health Management in Asia-Pacific. Report on a regional study and workshop on fish disease and fish health management. ADB Agriculture Department Report Series No. 1. Network of Aquaculture Centres in the Asia Pacific, Bangkok, Thailand. ADB/NACA. Final Report on the Regional Study and Workshop on Aquaculture Sustainability and the Environment (RETA 5534). Asian Development Bank and Network of Aquaculture Centres in Asia-Pacific. NACA, Bangkok, Thailand. (in press). Alday-Sanz, V. and Flegel, T.W. (1997). The risk of introducing yellow-head and white-spot viral infections from Asia to the Americas. CD ROM Paper No. 1, IV Congreso Ecuatoriano de Acicultura, 22-27 October, 1997, Guayaquil, Ecuador, 9 p. Ambrosia, R.E. and De Wall, D.T. (1990). Diagnosis of parasitic disease. Reviews of Sci Techn., Office Intern. Epizool. 9, 759-778. Chamberlain, G.W. Sustainability of world shrimp farming. In: E.K. Pikitch, D.D. Huppert, and M.P. Sissenwine (eds.) In: Global Trends: Fisheries Management. American Fisheries Society Symposium 20, Bethesda, Maryland . (in press). Chanratchakool, P., Turnbull, J.F., Funge-Smith, S.J., MacRae, I.H. and Limsuwan, C. (1998). Health Management in Shrimp Ponds. Aquatic Animal Health Research Institute, Bangkok, Thailand, 152 p. Knowles, D.P. and Gorham, J. R. (1990). Diagnosis of viral and bacterial diseases. Reviews of Sci. Techn., Off. Intern. Epizool. 9, 733-757. Lightner, D.V. and Redman, R.M. (1998). Shrimp diseases and current diagnostic methods. Aquaculture 164, 201-220. Lo C.-F., Ho, C.-H., Peng, S.-E., Chen, C.-H., Hsu, H.-C., Chiu, Y.-L., Chang, C.-F., Liu, K.-F., Su, M.-F., Wang, C.-H. and Kou, G.-H. (1996a). White spot syndrome (WSBV) detected in cultured and captured shrimp, crabs and other arthropods. Diseases of Aquatic Organisms 27, 215-225. Lo C.-F., Leu, J.-H., Ho, C.-H., Chen, C.-H., Peng, S.-E., Chen, Y.-T. Chou, C.-M., Yeh, P.-Y., Huang, C.-J., Chou, H.-Y., Wang, C.-H. and Kou, G.-H. (1996b). Detection of baculovirus associated with white spot syndrome (WSBV) in penaeid shrimps using polymerase chain reaction. Diseases of Aquatic Organisms 25, 133-141. Lo C.-F., Hsu, H.-C., Tsai, M.-F., Ho, C.-H., Peng, S.-E., Kou, G.-H. and Lightner, D.V. (1999). Specific genomic DNA fragment analysis of different geographical clinical samples of shrimp white spot syndrome virus. Diseases of Aquatic Organisms 35, 175-185.
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Lotz, J.M. (1997). Special topic review: Viruses, biosecurity and specific pathogen-free stocks in shrimp aquaculture. World Journal of Microbiology and Biotechnology 13, 405-413. Muroga, K. (1997). Recent advances in infectious diseases of marine fish with particular reference to the case in Japan. p. 21-31. In: T.W. Flegel and I.H. MacRae (eds.) In: Diseases in Asian Aquaculture III. Fish Health Section, Asian Fish. Soc., Manila. Plumb, J.A. (1997). Trends in freshwater fish disease research. p. 35-47. In: T.W. Flegel and I.H. MacRae (eds.) In: Diseases in Asian Aquaculture III. Fish Health Section, Asian Fish. Soc., Manila. Rana, K.J. (1997). Recent trends in global aquaculture production: 1984-1995. 14-19p. FAO Aquaculture Newsletter. No. 16. August 1997, FAO, Rome. 28pp.
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CONTRIBUTED PAPERS AND REVIEWS
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Technological constraints to disease prevention and control in aquatic animals, with special rference to pathogen detection Sharon McGladdery Fisheries and Oceans Canada, Gulf Fisheries Centre, PO Box 5030 (343 Ave. Université Ave.), Moncton, NB E1C 9B6 (E1C 5K4), Canada
Introduction Effective disease management and risk analyses rely on accurate data and information. For aquatic organisms, much of this information has been derived from a relatively narrow array of diagnostic tools, most of which are either non-pathogen-specific or have undergone “the test of time” rather than methodical validation or standardization procedures. Recently, however, this range of diagnostic methods has expanded to encompass the molecular expertise pioneered by human health and agricultural food production needs. The complexity of many of these techniques, and their rapid adaptation to “field kits” for use by non-specialist personnel, has prompted a serious re-evaluation of what we use in aquatic animal health management and why (Hiney, 1997). This paper is aimed at determining which areas of aquatic animal health management are limited by diagnostic and pathogen detection technology, and which are adequately met by traditional methods. Specific disease examples are described elsewhere in these proceedings by the specialists working with them, thus, the points below are deliberately general and provided as “food for thought”.
The Issues Aquatic animal health management needs arise from two separate situations: 1. The proliferation of opportunistic pathogens in physiologically stressed or immunologically compromised host populations, requiring sensitive, early, detection of potential pathogens. 2. The spread of a primary infectious organisms between infected and uninfected individuals, stocks or populations, requiring accurate identification of the pathogens responsible for disease outbreaks, sensitive detection of pathogens in sub-clinical carriers or abnormal hosts and accurate differentiation between benign and significant infectious organisms.
Disease diagnosis - identification of the cause of a disease outbreak. Some diseases can be diagnosed in the field with minimal technology or the need to isolate the causative agent, e.g., bacterial gill disease in association with stressful rearing conditions (Thoesen, 1994). Others present clinical signs which defy rapid or conclusive diagnosis, e.g., Malpeque disease of American oysters, Crassostrea virginica (McGladdery, 1993). Yet other diseases are caused by a range of different infectious agents e.g., chitinolytic fungal and bacterial shell diseases of crustaceans (Brock and Lightner, 1990). These situations can lead to diagnostic confusion (misdiagnosis) and ineffective management. First time disease outbreaks may (and should) require sample referrals to laboratories or diagnosticians which have experience with the putative pathogen - experience often being as important as technology for rapid and accurate disease diagnosis.
What do we have? The tools available for disease diagnosis differ between the types of aquatic organisms being examined. For finfish, there is a relatively broad range of diagnostic techniques, many of which can be used as cross checks for diagnosis of a single disease. For example, epizootic haematopoietic necrosis
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of redfin perch (Perca fluviatilis) and rainbow trout (Oncorhynchus mykiss) can be confirmed by: i) conventional isolation on BF-2 (bluefin gill 2) or FHM (fathead minnow) cell lines with serological identification of the iridovirus agent; ii) an indirect immunofluorescence antibody test (IFAT); iii) an enzyme-linked immunosorbent assay (ELISA); or iv) polymerase chain reaction (PCR) amplification and subsequent sequencing of the iridovirus DNA, using two published primers (OIE, 1997). Viral encephalopathy and retinopathy (viral nervous necrosis) virus and related nodaviruses are detectable using a range of specific and less specific techniques including: i) ultrastructural confirmation of virus-induced histopathology; ii) immunohistochemistry; iii) DFAT; iv) ELISA; and v) PCR amplification and sequence analysis (OIE, 1997). Some diseases with a more limited range of diagnostic options can be diagnosed accurately using the techniques available e.g. whirling disease of salmonids, caused by Myxosoma cerebralis, is presumptively diagnosed by behaviour, with confirmatory observation of the myxosporean spores in cartilage digests or histology preparations. Furthermore, few finfish diseases with a single aetiology, have defied conclusive diagnosis for long periods. The role of multiple infectious agents in a disease can usually be resolved through experimental research and verification using Koch’s postulates. One example for finfish (which has an as yet unidentified aetiologic agent) is erythrocytic inclusion body syndrome (EIBS). Although the causative agent is believed to be viral in nature, secondary infections by bacteria and fungi can confound diagnosis (Thoesen, 1994; Jarp et al., 1996). Once the primary infectious agent for such diseases is identified, subsequent diagnosis is simplified and, generally, ignores the presence of the secondary pathogens. The classic example of one such multi-factorial disease is epizootic ulcerative syndrome (EUS) which is described in detail elsewhere in these proceedings. The range of diagnostic techniques available for molluscan and crustacean diseases is narrower than that for finfish. Most significantly, aquatic invertebrates lack self-replicating cell lines for isolation and identification of intracellular pathogens. Finfish cell lines can be used, but the nature of the isolated viruses is often subject to question, since they could be vertebrate contaminants rather than primary invertebrate pathogens (Hill et al., 1986). In addition, Koch’s postulates have rarely been fulfilled or replicated for molluscan or crustacean isolates from fish cellm lines. Thus, most intracellular infections which cause overt disease in crustaceans and molluscs require histopathology for presumptive diagnosis, with ultrastructural confirmation of viral or bacterial aetiology. Histology, although laborious, has the advantage that it provides a permanent record of the pathogen in situ and can be used to assess focal or systemic histopathology. Conversely, it is limited in sensitivity to infectious agents which can be detected, and identified, at the light microscope level, eliminating most viruses, many bacteria, protists and even some metazoan parasites (which require whole mount or adult-stage identification). As with finfish, multiple diagnostic techniques are available for a number of shrimp diseases (Lightner, 1996) but most clinical cases can be presumptively diagnosed using non-specific techniques (gross observation, histology and tissue smears). Confirmatory diagnosis is then achieved using culture e.g. crayfish plague (Alderman and Polglase, 1986) or electron microscopic examination of ultrastructural features (Lightner, 1996). Pathogen culture is rarely used for diagnosis of molluscan diseases with the exception of two groups of significant disease agents: i) Perkinsus spp. (Ray, 1966; Gauthier and Vasta, 1993; LaPeyre et al., 1993); and ii) Labyrinthuloides-like protists (Bower, 1987; Kleinschuster et al., 1998). In clinical cases, however, these are also readily diagnosed using standard histology. Most other significant disease agents of molluscs are difficult to culture, but can be isolated under certain conditions (Hervio et al., 1993; 1995) i.e. acute infections. However, pathogen isolation is normally reserved for development of more specific detection and identification techniques rather than clinical diagnoses.
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What are the limitations? Speed of diagnosis is always a concern, especially with acute losses relying on histopathology, ultrastructural confirmation or long periods of tissue/media culture. The time span required for confirmatory diagnosis is frequently overcome by remedial action being based on presumptive diagnoses such as tissue smears, gross pathology or behavioural changes. This is most effective in areas with a well defined history of the disease e.g. Denman Island disease which is caused by Mikrocytos mackini in Pacific oysters (Crassostrea gigas) on the west coast of Canada (Bower, 1988). First time disease outbreaks in new species to culture, or appearing at a location for the first time, can undergo protracted periods of non-diagnosis or, worse, misdiagnosis. Examples include a serious disease of hard shell clams, Mercenaria mercenaria, caused by an unidentified Labyrinthuloides-like organism, “QPX”. This may have been causing mortalities in pre-culture history (Drinnan and Henderson, 1963) but its significance was not fully realised until hatchery broodstock and cultured stock on grow-out beds started to die (Whyte et al., 1994; Ragone Calvo et al., 1997; Smolowitz et al., 1996). Similarly, infectious salmon anaemia of Atlantic salmon (Salmo salar) was described as haemorrhagic kidney syndrome (HKS) when first detected in Atlantic Canada (Getchell, 1997). It took over a year before the causative agent was recognised as a virus and identified as ISAV, an agent previously known only from Norway (Hastein, 1997).
Where can molecular techniques enhance disease diagnosis? Significant pathogens that require long, complex culture or histology-based confirmatory diagnosis are prime candidates for rapid, pathogen-specific diagnostic kits. This applies predominantly to microbial pathogens, but may be equally appropriate for protists which are difficult to distinguish morphologically at the light microscope level or which have a diverse host-range. Rapid, pathogenspecific diagnostics would be particularly appropriate for disease management and control when diseases emerge in new geographic locations or host species, as described under limitations. An additional application for molecular techniques is for research into the pathogenesis of a disease via non-lethal sampling e.g. of haemolymph, fin- or gill-clips. This would provide useful information on pathogen proliferation, haemolymph profiles etc. but negates examination of the physical hostpathogen interface.
Pathogen screening – detecting infectious agents in sub-clinical or healthy organisms. Screening for infectious agents in healthy hosts is probably the most controversial area of aquatic animal health management. This is due to: i) inconclusive negative results; ii) the potential disease risks; and iii) the difficulty of controlling a disease outbreak in naïve and/or open-water populations. Since pathogen screening is frequently a pivotal part of disease risk assessments prior to transboundary transfers, the techniques used can also be “politically sensitive”. More recently, pathogen and/or disease screening is being used to define aquatic zones (intra-national and, more rarely, international) based on health profiles. This is usually limited to specific pathogens of commercially important host species (OIE, 1977). These zones can then be used for management purposes, to allow movement of pathogen carriers between non-confluent waters where the pathogen is known to occur (“like-to-like” transfers). Pathogen detection in healthy (carrier) hosts never assures 100% confidence, statistically, therefore negative samples all have a level of error which can be directly related to the sensitivity of the screening technique(s) applied. Since disease risk analyses have been, and continue to be, well-debated (DeVoe, 1992), more effort has been focussed on epidemiological principals in an effort to quantify and standardise the broad range of qualitative-based risk evaluations (Hiney, 1997; Thorburn, 1999). This has revealed a broad gap between the probability of detection of a single pathogen in a given sample and the statistical confidence in that detection. This reflects non-survey-based assumptions for pathogen prevalence in wild or open-water populations, as well as detection sensitivity, since prevalence is one of the critical factors determining confidence of detection of a single pathogen in any given sample size (Ossiander and Wedemeyer, 1973).
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What do we have now? Pathogen screening of aquatic animals involves the same techniques described above for disease diagnosis. In addition to culture-based techniques, immunoassays (fluorescent antibody tests, agglutination tests, ELISA) and nucleic acid probes have been available for finfish pathogens for years, and some form the basis of kits now used for pathogen screening (e.g. Aeromonas salmonicida which causes furunculosis, and Renibacterium salmoninarum which causes bacterial kidney disease). Pathogen-sensitive techniques for molluscs pathogens have been developed more recently e.g. immunoassays for Perkinsus marinus (Dungan and Roberson, 1993), Bonamia ostreae (Mialhe et al., 1988), Vibrio tapetis (causative agent of brown ring disease of Manila and Portuguese carpet clams, Ruditapes philippinarum and R. descussatus, respectively) (Castro et al., 1995) and giant rickettisia of the sea scallop Pecten maximus (Le Gall et al., 1992). However, none of these techniques has yet been transformed from research to routine diagnostic application and histology remains the most broadly used detection/diagnostic method applied to molluscs. Detection of sub-clinical infections in shrimp is limited to infectious hypodermal and hematopoietic necrosis virus (IHHNV), using bioassays as well as in situ and dot blot hybridisation or PCR of viral product in haemolymph or tissue homogenates, and baculoviral midgut gland necrosis virus (BMNV) using bioassays in susceptible Penaeus japonicus and a fluorescent antibody test (Lightner, 1996). Stress-induced enhancement of infections is another procedure used to enhance some sub-clinical viral infections in shrimp (and other aquatic species) which cannot be detected by histology (Lightner, 1996). What are the limitations? For cryptic infectious agents (mainly microbial) routine diagnostic procedures on healthy animals, especially non-culture-based techniques, are particularly weak in detection sensitivity. Thoesen (1994) lists several diseases caused by primary pathogens for which there are no procedures documented for detecting sub-clinical infections (e.g. Vibrio salmonicida, cold-water vibriosis or “Hitra disease”; channel catfish virus, CCV; Haplosporidium nelsoni, MSX; and H. costale, SSO, of American oysters, Crassostrea virginica). Lightner (1996) lists very few pathogens of shrimp for which there no methods to detect sub-clinical infections (hepatopancreas parvovirus, HPV). However, some pathogens can only be detected following stress-testing (e.g. monodon baculoviruses, MBV). For most disease agents in sub-clinical or abnormal “carrier” hosts, this means that sample size or sampling frequency has to be increased to enhance the level of confidence in detection. For techniques such as histology and ultrastructure, this frequently involves compromise between sample size (confidence level) and resource capability (time and manpower). For other more sensitive techniques (tissue culture, immunoassays and nucleic acid probes), the compromise may involve time, expense and specialist resource factors. Where can molecular techniques enhance zonation establishment and surveillance or transfer disease risk analysis? As described above, most agents of significant infectious diseases are difficult to detect using routine diagnostic techniques in healthy, sub-clinical hosts. This means that establishing an area which is designated free of a specific pathogen, inherently, includes a degree of error. Molecular screening techniques for specific pathogens could reduce this error margin by increasing confidence of detection. This would be especially important for areas that export live aquatic animals on a regular basis (“uninfected” zone to “uninfected” zone transfers). However, the pathogen specificity of these screening techniques negates detection of any other pathogenic or potentially significant organisms in the same specimens. Additional non-specific, but less sensitive, screening techniques may, therefore be required to give a true health “profile”. In addition, full-scale molecular-based testing of populations for a given pathogen, especially where there has been no history of the disease, could meet with varying degrees of resistance on both a practical and political level. Interpretation of low positive results from such an area would be especially problematic and difficult to resolve. In conclusion, molecular techniques might best serve as confirmatory screening to reinforce/refute results from
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general screening methods both for establishing zones and for certifying stocks free of specific pathogens. This would reduce the sample size and frequency required for high-technology screening, making their application more practical and easy to justify. Ideally the confirmatory screening should be on the same specimens (or sub-samples) from the same collections to ensure cross-reference validity. Certification of stocks as free of a specific pathogen could also benefit from the application of molecular-based detection techniques, especially where transmission is direct and negative result accuracy is imperative to prevent the spread of an endemic disease. Again, however, use of molecular probes or other pathogen-specific assays would mean any other infectious organisms would be undetected. Thus, as with zonation and surveillance, this pathogen-specific technology may best be applied as a confirmatory detection method, especially for certification of transfers from disease endemic areas.
Epizootiological research – determining the factors that trigger pathogen transmission and proliferation. Since pathogen eradication is rarely achieved in open-water or flow-through production systems, this is a crucial area of scientific research. It provides the information essential for effective reduction of disease losses to a negligible or economically acceptable level. Epizootiology is a complex science, involving detailed research into host immunity, physiology, genetics and environmental influences. Therefore, it requires a complex battery of techniques that range in application from controlled laboratory experiments to field observations. What do we have now? The methods available for epidemiological research are the same as those described above for pathogen screening and disease diagnosis What are the current limitations to epizootiological research? The difficulty of direct observation, handling stress and duplication of environmental variables in laboratory investigations often complicates the process of quantifying qualitative clinical and subclinical disease observations. In addition, despite extensive and well studied physiological and immunological parameters for finfish host-environment-pathogen research (Thomas and Woo, 1995; van Muiswinkel, 1995), a lack of standardisation and validation of routine diagnostic procedures has negated their direct application to epidemiological investigations of finfish diseases (Klontz, 1993; Thorburn, 1999). Research into host-pathogen interactions is further complicated for molluscs and crustaceans, where molecular immunology has only come under close scrutiny relatively recently (Bachere et al., 1995). One notable exception is research on Gaffkemia (caused by Aerococcus viridans var. homari) of lobsters (Homarus americanus) (Stewart and Zwicker, 1972; 1974). Sadly, however, this case bears little extrapolation to other crustacean host-pathogen interactions since lobster and bacterium have a rather unique association, as summarised by Stewart (1984). Another limitation to epidemiological research into aquatic animal pathogens is the inability to easily detect abnormal hosts (carrier, reservoir, accidental) of significant pathogens, especially those with low or unknown host-specificity. Abnormal hosts may demonstrate non-characteristic lesions or harbour the agents in tissues that are not infected in the “normal” host. This makes both detection and identification difficult, or even impossible, using routine diagnostic techniques. Where can molecular techniques enhance epizootiological research? As for screening, detection of sub-clinical (pre- and post-clinical) infections is imperative for understanding the dynamics of the pathogen, the factors that trigger pathogenicity and determining optimum management strategies. This includes detection of the pathogen in the environment or “abnormal” host species. In order to improve confidence in screening such samples, pathogen-specific detection or isolation techniques are required. To date, few probes which show consistent sensitivity
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have been developed for such broad screening (Hiney, 1997) and this appears to be an area which merits further study and development (Stokes et al., 1997).
Conclusion In general the range of tools available and under development show different advantages and disadvantages for a range of different aquatic animal health applications. No one technique shows a replacement advantage over another, and none appear sufficient to merit “stand-alone” application, with the possible exception of pathogen-specific research.
References Alderman, D.J. and Polglase, J.L. (1986). Aphanomyces astaci: isolation and culture. Journal of Fish Diseases 9, 367-379. Bachere, E., Mialhe, E., Noel, D., Boulo, V., Morvan, A. and Rodriguez, J. (1995). Knowledge and research prospects in marine mollusc and crustacean immunity. Aquaculture 132, 17-32. Bower, S.M. (1987). Artificial culture of Labyrinthuloides haliotidis (Protozoa: Labyrinthomorpha), a parasite of juvenile abalone. Canadian Journal of Zoology 65, 2013-2020. Bower, S.M. (1988). Circumvention of mortalities caused by Denman Island oyster disease during mariculture of Pacific oysters. American Fisheries Society Special Publication 18, 246-248. Brock, J.A. and Lightner, D.V. (1990). Diseases of Crustacea. Diseases caused by microorganisms. In: Kinne, O. (ed.) Diseases of Marine Animals. Vol III: Introduction, Cephalopoda, Annelida, Crustacea, Chaetognatha, Echinodermata, Urochordata. Biologische Anstalt Helogoland, Hamburg, Germany, pp. 245-349. Castro, D., Luque, A., Santamaria, J.A., Maes, P., Martinez-Manzanares, E. and Borrego, J.J. (1995). Development of immunological techniques for the detection of the potential causative agent of the brown ring disease. Aquaculture 132, 97-104. DeVoe, R.M. (ed.). (1992). Introductions and Transfers of Marine Species: Achieving a Balance Between Economic Development and Resource Protection. South Carolina Sea Grant Consortium, Charleston, South Carolina. 198 pp. Drinnan, R.E. and Henderson, E.B. (1963). 1962 mortalities and a possible disease organism in Neguac quahaugs. In: Ellerslie Biological Station Annual Report 1962-63, B. Department of Fisheries and Oceans, Canada, 18-20. Dungan, C.F. and Roberson, B.S. (1993). Binding specificities of mono- and polyclonal antibodies to the protozoan oyster pathogen Perkinsus marinus. Diseases of Aquatic Organisms 15, 9-22. Gauthier, J.D. and Vasta, G.R. (1993). Continuous in vitro culture of the eastern oyster parasite Perkinsus marinus. Journal of Invertebrate Pathology 62, 321-323. Getchell, R. (1997). ISA investigation continues; new disease challenges fish diagnosticians. Fish Farming News 5, 12. Haastein, T. (1997). Infectious salmon anaemia (ISA): a historical and epidemiological review of the development and spread of the disease in Norwegian fish farms. Infectious Salmon Anaemia Workshop, St. Andrews, NB (Canada), Nov 26 1997. Hervio, D., Bower, S.M. and Meyer, G.R. (1993). Detection, isolation, and host specificity of Mikrocytos mackini, the cause of Denman Island disease in Pacific oysters Crassostrea gigas. Journal of Shellfish Research 12, 136. Hervio, D., Bachere, E., Boulo, V., Cochennec, N., Vuillemin, V., Le Coguic, Y., Cailletaux, G., Mazurie, J. and Mialhe, E. (1995). Establishment of an experimental infection protocol for the flat oyster, Ostrea edulis, with the intrahaemocytic protozoan parasite, Bonamia ostreae: Application in the selection of parasite-resistant oysters. Aquaculture 132, 183-194. Hill, B.J., Way, K. and Alderman, D.J. (1986). IPN-like birnaviruses in oysters: infection or contamination? In: Vivares, C.P., Bonami, J.-R. and Jaspers, E. (eds.) Pathology in Marine Aquaculture (Pathologie en Aquaculture Marine). European Aquaculture Society, Special Publication no. 9, Bredene, Belgium. p. 297 (abstract only). Hiney, M. (1997). How to test a test: Methods of field validation for non-culture-based detection techniques. Bulletin of the European Association of Fish Pathologists 17, 245-250.
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Jarp J., Taksdal T. and Torud B. (1996). Infectious pancreatic necrosis in Atlantic salmon Salmo salar in relation to specific antibodies, smoltification, and infection with erythrocytic inclusion body syndrome (EIBS). Diseases of Aquatic Organisms 27, 81-88. Kleinschuster, S. J., Smolowitz, R., and Parent, J. (1998). In vitro life cycle and propagation of quahog parasite unknown. Journal of Shellfish Research 17, 75-78. Klontz, G.W. (1993). Epidemiology. In: Stoskopf, M.K. (ed.) Fish Medicine. W.B. Saunders, Philadelphia, US. pp. 210-213. LaPeyre, J.F., Faisal, M. and Burreson, E.M. (1993). In vitro propagation of the protozoan Perkinsus marinus, a pathogen of the eastern oyster, Crassostrea virginica. Journal of Eukaryotic Microbiology 40, 304-310. Le Gall, G., Mourton, C., Boulo, V., Paolucci, F., Pau, B. and Mialhe, E. (1992). Monoclonal antibodies against a gill Rickettsiales-like organism of Pecten maximus (Bivalvia): application to indirect immunofluorescence diagnosis. Diseases of Aquatic Organisms 14, 213-217. Lightner, D.V. (1996). A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, Louisiana, US. (looseleaf non-paginated). McGladdery, S.E. (1999). Shellfish diseases (Viral, bacterial and fungal). In: Woo, P.T.K. and Bruno, D.W. (eds.) Fish Diseases and Disorders Vol. 3. Viral, bacterial and fungal infections. CAB International, Oxon, UK. pp. 723-842. McGladdery, S.E., Drinnan, R.E. and Stephenson, M.F. (1993). A manual of parasites, pests and diseases of Canadian Atlantic bivalves. Canadian Technical Report of Fisheries and Aquatic Science 1931, 123 pp. Mialhe, E., Boulo, V., Elston, R.A., Hill, B.J., Montes, J., van Banning, P. and Grizel, H. (1988). Serological analysis of Bonamia in Ostrea edulis and Tiostrea lutaria using polyclonal and monoclonal antibodies. Aquatic Living Resources 1, 67-69. OIE (1997). OIE Diagnostic Manual for Aquatic Animal Diseases. 2nd Edition. OIE, Paris, France. 251 pp. Ossiander, F.J. and Wedemeyer G. (1973). Computer program for sample size required to determine disease incidence in fish populations. Journal of the Fisheries Research Board of Canada 30, 1383-1384. Ragone Calvo, L.M., Walker, J.G. and Burreson, E.M. (1997). Occurrence of QPX, quahog parasite unknown in Virginia hard clams, Mercenaria mercenaria. Journal of Shellfish Research 16, 334. Ray, S.M. (1966). A review of the culture method for detecting Dermocystidium marinum, with suggested modifications and precautions. Proceedings of the National Shellfisheries Association 54, 55-69. Smolowitz, R., Leavitt, D. and Perkins, F. (1996). An important new disease of hard clams, Mercenaria mercenaria, in the Northeast United States. Journal of Shellfish Research 15, 460-461. Stewart, J.E. (1984). Lobster diseases. Helgolander Meeresuntersuchungen 37, 243-254. Stewart, J.E. and Zwicker, B.M. (1972). Natural and induced bacterial activities in the hemolymph of the lobster, Homarus americanus: products of hemocyte-plasma interaction. Canadian Journal of Microbiology 18, 1499-1509. Stewart, J.E. and Zwicker, B.M. (1974). Comparison of various vaccines for inducing resistance in the lobster Homarus americanus to the bacterial infection, Gaffkemia. Journal of the Fisheries Research Board of Canada 31, 1887-1892. Stokes, N.A., Flores, B.S., Burreson, E.M., Alcox, K.A., Guo, Ximing and Ford, S.E. (1997). Life cycle studies of Haplosporidium nelsoni (MSX) using PCR technology. Journal of Shellfish Research 16, 336. Thoesen, J.C. (ed.). (1994). Suggested procedures for the detection and identification of certain finfish and shellfish pathogens. 4th Edition, Version 1, Fish Health Section, American Fisheries Society, Bethesda, Maryland, USA (loose-leaf, non-paginated).
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Thomas, P.T. and Woo, P.T.K. (1995). Immunological approaches and techniques. In: Woo, P.T.K. (ed.) Fish Diseases and Disorders Vol. 1. Protozoan and Metazoan Infections. CAB International, Oxon, UK., pp. 751-771. Thorburn, M.A. (1999). Applying epidemiology to infectious diseases of fish. In: Woo, P.T.K. and Bruno, D.W. (eds.) Fish Diseases and Disorders Vol. 3. Viral, bacterial and fungal infections. CAB International, Oxon, UK. pp. 689-722. van Muiswinkel, W.B. (1995). The piscine immune system: Innate and acquired immunity. In: Woo, P.T.K. (ed.) Fish Diseases and Disorders Vol. 1. Protozoan and Metazoan Infections. CAB International, Oxon, UK., pp. 729-750. Whyte, S.K., Cawthorn, R.J. and McGladdery, S.E (1994). QPX (Quahaug Parasite X), pathogen of northern quahaug Mercenaria mercenaria from the Gulf of St. Lawrence, Canada. Diseases of Aquatic Organisms 19, 129-136.
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The absolute requirement for predictive validation of non-culture based detection techniques Maura Hiney Fish Disease Group, Department of Microbiology, National University of Ireland, Galway, Galway City, Ireland.
Quality assurance and validation Quality assurance. For diagnostic assays of any type, quality assurance strives to ensure that the results are repeatable and reproducible i.e. that the assay has an acceptable level of precision (Welac Working Group, 1993). The precision of results is assured by stipulation of the source of reagents and disposable materials, acceptability limits for instrument calibration, the generation of standard operating procedures (SOPs) for the performance of the assay and the incorporation of appropriate positive and negative controls into the performance of the assay. These controls should, if appropriately chosen, function as both safeguard and check against deviation in the performance of the assay that might be generated as a consequence of inhibition and contamination, sample to sample variation and variability arising from the use of the technique by different operators or laboratories. Standardisation also requires description of other parameters of the assay including specification of the sample type to be analysed and how that sample should be collected, stored and processed. It is assumed, for the purpose of this paper, that all the facets of quality assurance of molecular-based detection techniques would already be in place prior to the application of the technique in the field. Validation. Quality assurance is clearly vital in the performance of diagnostic assays upon which important management or regulatory decisions may be made. However, quality assurance cannot provide information on how to interpret assay results. This information can only be obtained by the process of validation, defined here as “investigation of the extent to which a technique can legitimately be used for a particular purpose”. Validation does not demonstrate that an assay will have standardised performance, but rather that it is appropriate for a given application. Therefore, validation is essential to determine the extent to which the potential of PCR-based techniques for detection aquatic animal pathogens will be realised in practice. Hiney and Smith (1999) have provided a framework for the validation of PCR-based detection techniques which outlines three major criteria (quantitivity, qualitivity and reliability) which should, where possible, be evaluated sequentially at four levels of increasing experimental complexity (in vitro, seeded samples, incurred samples and field samples). In vitro studies and studies in seeded and incurred samples can be performed in the laboratory. Although these laboratory-based validation studies form a vital part of the validation process, they can only provide information on the performance of the technique in the sample type to which it will be applied. In order to ascribe meaning to the results generated in the field, the last level of validation (i.e. field application) is the most important. Ascribing meaning to results. When assessing the data generated by a non-culture-based detection technique, either DNA- or immunological-based, the critical question to be addressed is: ‘what can the results validly be taken to mean”. The meaning that can be attributed to the results generated by any non-culture-based detection technique will be dependent on the application for which that technique is validated, and also on the context in which the results will be interpreted.
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a). Application With regard to the intended application of a detection technique, the sample type being examined may be either CLINICAL or ENVIRONMENTAL. Laboratory validation should, therefore, have demonstrated that the detection technique performs reliably in the sample type in which that technique will be used. In addition, the type of study being conducted will normally be designed to answer questions of either ECOLOGICAL or EPIZOOTIOLOGICAL relevance. Hiney (1997) outlined the difference in emphasis of the questions being asked in these two types of studies and the importance of designing a validation programme suitable for the study type. b). Context The meaning that will be attributed to results will also depend on the context of the interpretation. Contexts include research, disease diagnostics and regulation of fisheries operations which will require different levels of validity (Fig. 1). In the research context, the required level of validity need not be high provided the technique generates information of use for the formulation or confirmation of models of disease. On the other hand, the use of the technique for diseases diagnosis necessitates a much higher level of validity because of the therapeutic and/or management decisions that may rest on the outcome of the application of that technique. At the highest level of validity, the interpretation of results in a regulatory context may have far-reaching and serious implications for a fish farmer, region or country.
LOW Research Disease Diagnosis
Regulation HIGH Figure 1. Required levels of validity of detection techniques in the context of their application. The selectivity of non-culture-based detection techniques. Regardless of the application intended for a non-culture-base detection technique or the context of that application, the most important question that must be asked of any detection technique is whether the technique provides THE TRUTH, THE WHOLE TRUTH, AND NOTHING BUT THE TRUTH. In other words, does the technique detect the species (viral, bacterial or protozoan) of interest, does the technique detect all members of that species and is there cross-reaction with members of any non-target species? The property of ‘truth’ is often referred to as ‘specificity’ in scientific reports on detection techniques. However, in veterinary terms ‘specificity’ has another meaning, that is, the number of false-negatives in a population, which is clearly not the property we wish to describe. More correctly, the property of truth should be called ‘selectivity’. The Welac Working Group (1993) defined selectivity as “the extent to which a technique can detect a particular analyte in a complex mixture without interference from other components in the mixture”. In the case of microbial detection techniques, the mixture will be a sample, either clinical or environmental, and potential interfering components would include nontarget species and chemical components in that mixture which might generate false positive or negative responses.
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Non-culture-based techniques are proxy measurements. In determining the selectivity of nonculture-based detection techniques it must be born in mind that these techniques provide a proxy measurement of the presence of the target species i.e. they detect only a fragment of that cell. The use of proxy measurements is based on two important assumptions about the meaning of signals generated by non-culture-based detection techniques: i) that the SIGNAL = TARGET; and ii) that the SIGNAL = DISEASE RISK. The purpose of validation is to establish that these assumptions are true.
Assumption 1: signal = target The selectivity of non-culture-based detection techniques is established in the laboratory by the use of CONTROL PANELS i.e. collections of strains of the same species (the truth) that should be representative of all members of the target species (the whole truth), and collections of closely related and non-related strains (nothing but the truth). However, a number of problems can be identified for control panels. The first is that many of these control panels are badly designed, containing too few target organisms (especially in the case of heterogeneous species), inappropriate representatives of the target species or too few or badly chosen closely related non-target organisms. A second problem frequently observed in control panels is that they do not contain appropriate application-dependent non-target organisms. Application dependent panels should take account of the sample type in which the technique will be applied and should, therefore, contain the non-target organisms most likely to be present in that sample type, be they other pathogenic species that infect the same host or species indigenous to that environment. Even where these problems have been addressed, there still remains a fundamental problem with the design of control panels, especially for environmental applications (although clinical applications will also be effected), namely that ONLY ORGANISMS THAT CAN BE CULTURED CAN BE INCLUDED IN CONTROL PANELS. It has been variously estimated that only 0.1 - 1% of all organisms present in the environment have been cultured in the laboratory. This leaves a vast and unknowable reservoir of organisms whose potential for cross-reaction cannot be assessed.
Comparative validation In reality, no amount of internal or laboratory validation or standardisation can tell us how we should interpret results generated in the field. To do this we need to use external validation techniques, of which there are two main types, comparative validation and predictive validation. Comparative validation involves the comparison of results generated by two or more methods targeted against the same organism. There are a number of approaches that can be taken to comparative validation. a). Compare against method previously validated for the same application As there are currently no non-culture-based detection techniques adequately validated for diagnosis of fish diseases this option will for the moment remain theoretical. b). Compare against another unvalidated non-culture-based method The comparison of two methods based on the same detection principle, such as two PCR-based assays, is not ideal. This type of comparison cannot allow for inherent flaws (e.g. inhibition, matrix interaction) in the technique. Ideally the methods being compared should be based on different detection principles (genetic, immunological or culture-based) and should not, in theory, be inhibited by the same components of the test matrix or generate the same false positives from that matrix. If the degree of concordance in the results generated by these techniques is high when applied to the same samples, then the methods can be said to mutually co-validate each other.
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What results may comparative validation generate? There are a number of possible outcomes to a comparative validation programme: a). Both techniques valid A comparison should generate full concordance if the techniques have the same lower detection limit and are both valid for the application for which they have been developed. b). Both techniques valid but lower detection limits different In the case of the comparison of two or more valid techniques whose lower detection limits are different we would expect to see asymmetric concordance. A percentage of the samples will be positive by both techniques, but additional samples will also be positive by the assay with the better lower detection limit. c). One technique invalid When one of the methods being compared is invalid for the intended application, that is, generates either false positive or negative signals, we would expect low concordance in any comparative study. Unfortunately, it may not be possible to distinguish which method is invalid unless the results generated by it are at odds with more than one other method. d). Both techniques invalid If all of the methods being compared are invalid for a particular application then the results generated will have low concordance. Regardless of the results generated by a comparative validation, this approach can only provide us with information on the presence of the target per se (Assumption 1). It is still possible that what we are detecting are cell fragments or dead cells. Therefore, comparative validation cannot give us any information about what the presence of that target means in terms of disease (Assumption 2). So how could we interpret these results in any meaningful way?
Predictive validation One possible means of interpreting the results generated in the field by a non-culture-based detection technique is through predictive validation (i.e. ‘ESTABLISHING THE ABILITY OF A TECHNIQUE TO PREDICT A DISEASE EVENT’). Clearly, with regard to diagnostic techniques the most important event that can be predicted would be the occurrence of a disease episode in the host following the detection of a positive response through application of a non-culture-based detection technique to either host tissue or the environment of that host. However, ‘disease’ is a rather loose concept, defined by the World Health Organisation as “any divergence from a healthy state”. Therefore, the event to be predicted must be capable of being established empirically (e.g. that the detection of positive responses by a PCR assay would predict the future isolation of the pathogen of interest bacteriologically from host tissue). Equally, the prediction could be that the absence of a positive response would predict a reduction in the requirement for antibiotic therapy in the host population. Regardless of the predicted event, the ultimate objective of a predictive validation program is to establish that the detection of a positive signal by a non-culture-based technique has meaning in terms of disease, that is the SIGNAL = DISEASE (Assumption 2).
Meaning in context Getting back to the relevance of context in the interpretation of results a number of observations can be made. In terms of research, a great deal of interesting and useful data can generated by the use of techniques of poor validity or whose validity has not been adequately established through either comparative or predictive validation studies. However, as diagnostic tools, assays with poor validity or
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whose validity has not been adequately established may generate completely misleading data and should not be considered for application in this context until sufficient data on their performance in the field is available. Most seriously, from the regulatory viewpoint, the interpretation of the data obtained by assays with poor validity or whose validity has not been adequately established, to indicate a disease risk and warrant regulatory sanctions would be completely invalid. Therefore, the first priority of any programme that hopes to introduce non-culture-based detection techniques for detection of aquatic animal pathogens must be to establish an adequate validation programme which includes both comparative and predictive validation which take cognisance of the intended application (i.e. sample type, conditions, context). Only by such an approach can we have confidence that we are ascribing the correct meaning to the results we generate.
References Hiney, M. (1997). How to test a test: Methods of field validation for non-culture-based detection techniques. Bulletin of the European Association of Fish Pathologists 17, 245-250 Hiney, M.P. and Smith, P.R. (1999). Validation of polymerase chain reaction-based techniques for proxy detection of bacterial fish pathogens: Framework, problems and possible solutions for environmental applications. Aquaculture 162, 41-68. Welac Working Group (1993). Welac Eurochem Guidance Doc. No.1. Accreditation for chemical laboratories: Guidance on the interpretation of the EN45000 series of standards and ISO/IEC guide 25. Addition 1. April.
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Evaluation of diagnostic tests: the epidemiological approach1 Daniel F. Fegan BIOTEC, Thailand
The use of diagnostic tests is widespread in studies of disease in aquatic animals. These are often used in isolation and the results interpreted or applied to populations without sufficient regard to wider implications of the results. Much effort is devoted to the understanding of disease processes at the individual animal, organ, cellular and genetic levels, and the complex interplay between individuals in populations and the environment can be forgotten. At the population level the use of diagnostic tests is made more complicated by population effects such as prevalence of the pathogen, expression and impact of the disease on the population and potential for pathogen spread among others. The inadequacy of the Henle-Koch postulates in animal disease has long been recognised as they do not work well with multi-factorial causes of disease and the impact of predisposing factors. As a result, the familiar “epidemiological triad” concept (Host-Pathogen-Environment), illustrated in the famous diagram of Snieszko (1974) was introduced (Figure 1).
Figure 1: The epidemiological triad (Snieszko, 1974) This neatly illustrates the complex interplay of factors which result in disease at the individual and population levels. The existence of multiple contributing factors to disease outbreak is summarised in the epidemiological definition of the cause of a disease as “an event, condition or characteristic that plays an essential role in producing an occurrence of the disease” (Baldock, 1996). This implies that the presence of a pathogen may not, in itself, be sufficient to cause disease in the absence of other factors, a concept expressed in the statement that a pathogen is a necessary but not sufficient cause of a particular disease. This is classically seen in epizootic ulcerative syndrome (EUS) of fish where epidermal damage by a stress such as lowered pH is required to before infection with Aphanomyces sp. and the resultant characteristic lesions can occur. Application of these concepts requires a different approach to the interpretation of diagnostic test results, particularly where they will be applied to a decision-making process. This paper is intended to 1
This paper draws heavily on the information on diagnostic testing in Chapter 17 of Thrusfield (1995). This should be referred to for a fuller explanation than is possible here.
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briefly introduce the basic concepts of epidemiology as they relate specifically to diagnostic tests. [For a fuller treatment, the reader is referred to the textbooks of Thrusfield (1995) or Pfeiffer (1998).] In veterinary medicine, a diagnosis is a statement of an animal’s state of “normality” and represents an interpretation of one or several observations that form the basis for a decision on further action. The decision is based on a number of factors including factual knowledge, experience and intuition as well as clinical diagnostic tests and it is the correct use of all of these which increases the probability of correct diagnosis (Figure 2). This definition clearly identifies the uncertainty associated with diagnosis and the outcome of a given course of action taken as a result. Factual Knowledge Uncertainty
Diagnostic Tests
Diagnosis
Disease Status
Experience
Intuition Prevalence
Figure 2: Factors influencing veterinary diagnoses (from Pfeiffer, 1998) This differs somewhat from the classical concept of diagnosis in the Henle-Koch postulates as the consistent isolation and identification of a particular aetiological agent associated with a disease.
Definitions Unfortunately, some of the terms used in veterinary epidemiology are the same as those used in clinical pathology but with different definitions. The terms “sensitivity” and “specificity” in particular, have been the cause of considerable confusion. Some definitions of terms used in veterinary diagnosis are given above. Accuracy Bias Precision Sensitivity
Specificity
The accuracy of a test refers to the level of agreement between the test result and the “true” clinical state. Bias measures the systematic deviation from the “true” clinical state Represents the degree of fluctuation of a series of measurements around the central measurement. Proportion of animals with the disease which test positive (i.e. proportion of true positives). This equates to the laboratory definition where it means the ability of an analytical method to detect very small amounts of the analyte (such as an antibody or antigen). Thus a test which is highly “sensitive” from a laboratory perspective is also likely to be “sensitive” from an epidemiological perspective. Proportion of animals without the disease which test negative (i.e. proportion of true negatives). This equates to the laboratory definition where it means the ability of the test to react only when the particular analyte is present and not react to the presence of other compounds. Thus a test which is highly “specific” from a laboratory perspective is also likely to be “specific” from an epidemiological perspective.
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PPV
NPV
True prevalence
Apparent Prevalence
(Positive Predictive Value ) The probability (or likelihood) that an animal which returns a positive test result actually has the disease in question. (Negative Predictive Value) The probability (or likelihood) that an animal which returns a negative test result actually does not have the disease in question. Proportion of animals in the population which really do have the disease in question regardless of their test result. From a test result point of view, it includes the “true” positives and the “false” negatives. Proportion of animals in the population giving a positive test result regardless of their true status for the disease in question. From a test result point of view, it is all the test positive animals, some of whom will be “true” positives and some which are “false” positives.
Diagnostic testing Diagnostic tests are more or less objective methods which reduce the uncertainly factor in diagnosis. Diagnostic tests are often interpreted using a dichotomous outcome (normal/abnormal, diseased/healthy, treat/don’t treat) which poses less difficulty when the test itself is dichotomous (presence or absence of a pathogen) but can cause considerable difficulty in interpretation when it is continuous (e.g. serum antibody levels or cell counts). In such cases, the selection of an appropriate cut-off point to separate ‘positive’ and ‘negative’ results introduces a level of uncertainty. In most diagnostic tests false positives and false negatives occur. Some of the reasons for positive and negative results in serology, for example, are given in Table 1. Consequently any diagnostic test which does not directly identify the presence of the infection can only produce an estimate of the apparent prevalence of a disease (i.e. the proportion of animals giving a positive test result) and does not equate to the presence of infection. Estimates of true prevalence, however, can be made taking into account test sensitivity and specificity where these are known. Positive Results Actual infection Group cross-reaction Non-specific inhibitors Non-specific agglutinins Negative Results Absence of infection Natural/induced tolerance Improper timing Improper selection of test Non-specific inhibitors Toxic substances Antibiotic induced immunoglobulin suppression Incomplete or blocking antibody Insensitive tests
True positive False positives
True negative
False negative
Table 1: Reasons for positive and negative results from serological tests (from Stipes et al. 1982). Estimates of true prevalence, however, can be made taking into account test sensitivity and specificity. True prevalence
=
apparent prevalence + (specificity – 1) specificity + (sensitivity –1)
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Sensitivity and specificity are indicators of the validity of diagnostic tests (Thrusfield, 1995). When a cut-off point is used, sensitivity and specificity show an inverse relationship – as sensitivity increases, specificity decreases and vice versa. Estimation of the sensitivity and specificity requires testing of animals for which the disease status is known. This requires the use of an appropriate unequivocal diagnostic method as a “gold standard”. For example, in the case of the protistan oyster pathogen Haplosporidium nelsoni (MSX disease)2, data on which was presented at the workshop, the histological demonstration of the disease may be used as an estimation of true status (the “gold standard”) and to evaluate the PCR data obtained by constructing the following simple table.
True Status Test 1 +ve Test 1 –ve
+ve a c a+c
-ve b d b+d
a+b c+d a +b + c + d
In the table, “a” represents the true positives, “d” the true negatives and “b” and “c” the false positives and false negatives respectively. The various epidemiological values can also be simply calculated as follows: x Sensitivity = a/(a+c) x Specificity = d/(b+d) x PPV = a/(a+b) x NPV = d/(c+d) x Apparent prevalence = a+b/(a+b+c+d) x True prevalence = a/(a+b+c+d) Substituting the data for incidence of MSX:
Histology PCR +ve PCR –ve
+ve 74 2 76
-ve 55 127 182
129 129 258
Using the above formulae, the calculations are: x Sensitivity = 74/76 = 97% x Specificity = 127/182 = 70% x PPV = 74/129 = 57% for the particular prevalence of 29% x NPV = 127/129 = 98% for the particular prevalence of 29% x Apparent prevalence = 129/258 = 50% x True prevalence = 74/258 = 29% From the table, it appears that the PCR test has a high sensitivity but only a moderate specificity. In other words, 97% of animals with the disease test positive using PCR (a false negative rate of 3%) but only 70% of animals without the disease test negative with PCR (i.e. a false positive rate of 30%). Therefore, although the test would be useful for screening to reduce the possibility of introducing infected individuals into a population (for which false positives are not a major concern), it would not
2
Data used with kind permission of Dr. Eugene Burreson, Virginia Institute of Marine Science.
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be sufficient on its own to make a definitive diagnosis of the disease due to the high false positive rate, and would certainly not be appropriate as the basis for a decision on action to be taken. The selection of the appropriate level of sensitivity and specificity often depends upon the particular need. When screening for a disease or pathogen (for example, testing animals to eliminate infected individuals) we require a reliable positive result with few false negatives and a reasonable number of false positives (within an economically justifiable level of rejection). This would require a test with a high sensitivity and reasonable specificity. This type of test would be used in a quarantine situation, for example, to reduce the risk of disease introduction or when demonstrating absence of a disease to establish “disease-free” zones. On the other hand, if we need as few false positives as possible (e.g. to confirm a tentative diagnosis) a test with a high specificity and reasonable sensitivity is used. It is, however, important to note that the consequence of any diagnostic test with imperfect specificity (less than 100%) is that if a large number of tests are made on a single uninfected animal, there is a significant chance of finding a positive result.
Predictive values For a diagnostic decision, it is also useful to make some estimate of the predictive value of a diagnostic test. The predictive value quantifies the probability that a positive test result for a particular animal or sample correctly identifies the presence of infection and a negative test result correctly identifies the absence of infection. This requires knowledge of not only the sensitivity and specificity of the test but the prevalence of the condition. The effect of prevalence on predictive values is considerable. As prevalence increases, Positive Predictive Value (PPV) increases and Negative Predictive Value (NPV) decreases. Formulae for calculating predictive values are based on Bayes' theorem of conditional probability (Fleiss, 1981) and are as follows: Positive predictive value = a/(a+b) =
Prev x Se Prev x Se + (1-Prev) x (1-Sp)
Negative predictive value = d/(c+d) =
(1-Prev) x Sp (1-Prev) x Sp + Prev x (1-Se)
Se: sensitivity; Sp: specificity; Prev: pre-test probability of disease (or true prevalence). Predictive values are functions of prevalence and the test characteristics of sensitivity and specificity. As prevalence declines so does positive predictive value. The converse is true for negative predictive value (see Table 2). If the sensitivity and specificity of a diagnostic test are known for a particular target population, then predictive value graphs can be drawn for the range of all possible pre-test probabilities of disease from 0 to 1 (100%). Table 2: Effect of prevalence on positive predictive value (PPV) with a hypothetical serological test (Se and Sp = 0.95) Prevalence (%) PPV (%)
0.1
1
2
5
10
50
90
100
1.9
16.1
27.9
50.0
67.8
95.0
99.4
100
The important point that this table indicates is that despite using a good test (Se and Sp = 0.95) most reactors are non-infected (false positives) when the disease is present in the population at a low
34
prevalence. Of the 2 test properties, it can be shown that specificity exerts a greater influence on PPV than does sensitivity. On the other hand, sensitivity exerts a greater influence on negative predictive value (NPV). Again using the data for oysters, the PCR test has a good negative predictive value but poor positive predictive value (Proportion of PCR -ve animals which do not have disease = 98%; Proportion of PCR +ve animals which have the disease = 57%) where the true prevalence is 29%. In other words, the test is a poor predictor of disease occurrence and would be of limited use in confirming the existence of suspect disease. As a rule of thumb, highly specific tests should be used to confirm tentative diagnoses while highly sensitive tests should be used to rule out possible disease. Finally, the impact of the test on estimated prevalence is clearly seen. Because of the low specificity and PPV, the prevalence of infection is overestimated considerably (apparent prevalence is 50% compared with the true prevalence of 29%). This would diminish the usefulness of PCR as a diagnostic tool in this particular case. The PPV of a particular test can be improved by appropriate selection strategies (Baldock, 1996): 1. Testing of “high risk” groups (animals with clinical signs rather than normal animals) 2. For the same test using a higher cut-off with higher specificity or use a second test with a higher specificity) 3. Use of multiple tests for interpretation of results.
Population level test interpretation When dealing with testing a group of animals (such as a tank of shrimp postlarvae or a pond of fish) for disease rather than an individual, some additional factors have to be taken into account. In addition to the sensitivity and specificity of the test, the number of animals from the group which are tested (the sample size), the true prevalence and the number of positives required to classify the population as infected are important. At the group level we require high sensitivity and high specificity in a test, the same as for individual level tests. It is important to note, however, that individual and group level test characteristics are not equivalent. At a group level, sensitivity and specificity are influenced by sample size (as the sample size increases, so does sensitivity) and the number of positives required (as the number of positives required increases, there is a corresponding increase in specificity). Again, as with individual level tests, sensitivity and specificity are inversely related. It should be noted that even relatively good tests with high sensitivity and specificity will have a low predictive value at low levels of prevalence. For example, if a test with sensitivity of 99% and specificity of 99.9% was used at a high prevalence, say 10%, a single test conducted on 10 million animals would give 9,000 false positives and 990,000 true positives. On the other hand, if the prevalence were 0.01% the test would give 9,900 true positives and 9,990 false positives. This has important implications for eradication campaigns, quarantine screening and other situations where prevalence may change with time.
Evaluation of diagnostic techniques As previously explained, evaluation of diagnostic techniques requires some independent, valid measure of the true condition of the animal (the ‘gold standard’). The ‘gold standard’ may be a single unequivocal test (histological or post-mortem demonstration of the disease, for example) or a combination of alternative tests which, when simultaneously positive, identify animals which are true positives. The assessment or comparison of diagnostic tests requires their application, with the ‘gold standard’, to a sample of animals with a typical disease spectrum. The characteristics of the test are compared with the gold standard in terms of their sensitivity and specificity (see definitions).
35
Frequently, however, no ‘gold standard’ exists for a particular condition and it is necessary to evaluate the diagnosis by the level of agreement between different tests. This assumes that agreement between tests is evidence of validity, whereas disagreement suggests that the tests are not reliable. The kappa test can be used to measure the level of agreement beyond that which may be obtained by chance. The kappa statistic lies within a range between –1 and +1. The kappa test uses the same table as for calculation of epidemiological values with the observed agreement given by the formula: OA = (a + d)/(a + b + c + d). This is compared to the expected agreement which would be obtained by chance which is given by the formula: EA = [{(a + b)/n} x {(a + c)/n}] + [{(c + d)/n} x {(b + d)/n}] Kappa is the agreement greater than that expected by chance divided by the potential excess. (OA – EA) / (1-EA) The kappa values are evaluated according to arbitrary “benchmarks” as shown in Table 3.
Kappa value > 0.81 0.61 – 0.80 0.41 – 0.60 0.21 – 0.40 0.01 – 0.20 0.00
Evaluation Almost perfect agreement Substantial agreement Moderate agreement Fair agreement Slight agreement Poor agreement
Table 3: Evaluation of kappa statistic (Everitt, 1989)3 For example, again using the data from oysters used previously. OA = (74 + 127)/ 258 = 0.779 EA = [{129/258} x {76/258}] + [{129/258} x {182/258}] = (0.500 x 0.295) + (0.500 x 0.705) = 0.1475 + 0.3525 = 0.500 The maximum possible agreement beyond chance = 1 – 0.500 = 0.500 k = (0.779 – 0.5)/0.5 = 0.279/0.5 = 0.558 indicating moderate agreement between the two tests. It should be noted that the kappa value gives no indication which of the tests is better and that a good agreement may indicate that both tests are equally good or equally bad. Another important characteristic of a test is its repeatability or the consistency of the test results in two or more replicates on the same animal. For a test whose outcome is either positive or negative, the level of agreement will give an indication of the reliability of the test result. The statistical tests used are outside the scope of this paper and can be found in Thrusfield (1995) or standard statistical 3
Note that these interpretations are relatively arbitrary and that other authors may use different values for the level of agreement.
36
texts. However, if the test is repeated twice, then McNemar’s chi square test for related samples can be used, and for three or more, Cochran’s Q-test is used. If the proportion of positive and negative results are significantly different between the replicates, the repeatability of the test may be low.
Selection of diagnostic tests The selection of an appropriate diagnostic test depends upon the intended use of the results. If the intention is to rule out a disease, reliable negative results are required for which a test with high sensitivity (i.e. few false negatives) is used. If it is desired to confirm a diagnosis or find evidence of disease (i.e. to “rule in” the disease) we require a test with reliable positive results (i.e. high specificity). As a general rule of thumb, a test with at least 95% sensitivity and 75% specificity should be used to rule out a disease and one with at least 95% specificity and 75% sensitivity used to rule in a disease (Pfeiffer, 1998).
Conclusions The interpretation of diagnostic tests depends upon the definition of clinical disease and its distinction from the presence of the pathogen. It is the case in most disease outbreaks that the presence of the pathogen is a necessary but not sufficient cause of disease. This is because there are often other factors involved in the expression of the disease condition, an important consideration when making a diagnosis for a population in which a decision has to be made. In studying disease outbreaks, especially in populations, we need to look at them from both a pathological and epidemiological standpoint. Ideally, a diagnostic test can be evaluated based on a clear relationship with an unequivocal "gold standard" diagnosis. The analytical sensitivity of a method and its relationship with the epidemiological sensitivity, at a population level can change as prevalence increases, as sample size increases and depending upon the number of positive reactions we accept as sufficient on which to base a diagnosis. Highly sensitive (in the analytical sense) methods such as PCR may pick up early stages of a disease condition and this will often manifest itself by a change in the number of false positives over time. Thus, it can be the case that a simplified interpretation of data taken at one point in time may represent a snapshot view. However, as data accumulates, it should be possible to establish a more accurate picture. Pathologists and researchers involved in lab-based diagnostic work should consider the epidemiological approach required if such results are to be extrapolated to populations. The use of epidemiological methods in the planning and analysis of diagnosis, or better still, a greater cooperation between pathologists and epidemiologists, will assist greatly in the development and interpretation of better diagnostic tests.
References Thrusfield. M. (1995). Veterinary Epidemiology 2nd Edition. Publ. Blackwell Science Ltd., Oxford, UK. Baldock, C. (1996). Course notes from the Australian Centre for International Agricultural Research Workshop on “Epidemiology in Tropical Aquaculure” Bangkok, 1-12 July, 1996. Snieszko, S.F. (1974). The effects of environmental stress on outbreaks of infectious diseases of fishes. Journal of Fisheries Biology 6, 197-208. Pfeiffer, D. (1998). Veterinary Epidemiology. An Introduction. Institute of Veterinary, Animal and Biomedical Sciences. Massey University, Palmerston, New Zealand. Stites, D.P., Stobo, J.D., Fundenberg, H.H. and Wells, J.V. (1982). Basic and Clinical Immunology, 4th Edition. Lange Medical Publications, Los Altos, USA.
37
DNA-based diagnostic and detection methods for penaeid shrimp viral diseases Donald V. Lightner Department of Veterinary Science and Microbiology University of Arizona, Tucson, AZ 85721 USA
The most important diseases, in terms of economic impact, of cultured penaeid shrimp in Asia, the Indo-Pacific, and the Americas have infectious etiologies. Among the infectious diseases of cultured shrimp, certain virus-caused diseases stand out as the most significant. Since the first report of a penaeid shrimp virus disease by Couch in 1974, at least 20 more viruses have been described from the penaeids (Table 1). The earliest diagnostic methods developed for these pathogens included the traditional methods of morphological pathology (direct light microscopy, histopathology, and electron microscopy), as well as enhancement and bioassay methods. While tissue culture is considered to be a standard tool in medical and veterinary diagnostic labs, it has never been developed as a useable, routine diagnostic tool for shrimp pathogens. Likewise, there are few antibody-based diagnostic tests available for the penaeid virus diseases (Lightner and Redman, 1998). The need for rapid and sensitive diagnostic methods has led to the application of modern biotechnology to penaeid shrimp diseases. DNA-based detection methods for the most important viral pathogens (IHHNV, HPV, SMV, TSV, YHV, GAV/LOV, WSSV, MBV, and BP) have been reported in the literature and some DNAbased diagnostic methods are commercially available. PCR or RT-PCR methods are available for several of these viruses and some are in routine use by certain sectors of the industry. For others, specific DNA probes tagged with non-radioactive labels provide highly specific detection methods for application in dot blot formats with hemolymph or tissue extracts, and with routine histological sections using in situ hybridization (Lightner, 1996; OIE, 1997; Lightner and Redman, 1998). The OIE Fish Disease Commission at its September 1998 meeting voted to recommend to the OIE that three of the penaeid shrimp viruses diseases from the “listed” category be upgraded to the “notifiable” category. If the recommendations of the Fish Disease Commission are approved by the OIE’s General Assembly in May 1999, the notifiable and listed viral pathogens of crustacea will be those shown in Table 2. All of these viral diseases affect cultured penaeid shrimp. Before a disease may be included on the OIE lists of notifiable and listed diseases, several criteria must be met: 1) the etiological agent must be known, 2) reliable diagnostic(s) methods must be available, and 3) the disease must be a significant disease of local, regional, or international importance. The accompanying Tables in this report list the known penaeid shrimp viruses and summarize the available traditional and DNA-based diagnostic and detection methods available for the recently proposed OIE notifiable and listed pathogens of penaeid shrimp (OIE, 1997; OIE, unpublished report). The diagnostic and detection methods for these viruses are listed in Table 3. There is a growing need to standardize and validate the DNA-based diagnostic methods and the laboratories that use them. Standardization of DNA-based diagnostic methods is almost inherent in the nature of the tests. That is, a specific DNA probe, or a specific set of primers, that is used to demonstrate the presence of absence of a unique DNA or RNA sequence does not vary from batch to batch. Hence, with proper controls, these DNA-based methods are readily standardized (Reddington and Lightner, 1994). However, despite the growing dependence of the shrimp culture industry on DNA-based diagnostic methods, none of the tests that are available from commercial sources nor from the literature have been validated using controlled field tests. Likewise, there are no formal accreditation or certification programs yet in place to assure that test results from technicians and laboratories running the tests are indeed accurate and properly controlled (Lightner and Redman, 1998). The
38
implementation of a formal program by appropriate international agencies or professional societies is needed to validate new diagnostic methods and to periodically review the accreditation and certification of diagnosticians and diagnostic laboratories. The establishment of regional reference laboratories for DNA-based diagnostic methods of penaeid shrimp/prawn pathogens would fit well into such a program with the goal of making these methods uniform, reliable, and readily applicable to disease control and management strategies for viral diseases of cultured penaeids.
References Adams, J.R., and Bonami, J.R. Eds. (1991). Atlas of Invertebrate Viruses. CRC Press, Boca Raton, FL. 684 pp. Arimoto, M., Yamazaki, T., Mizuta, Y. and I. Furusawa. (1995). Characterization and partial cloning of the genomic DNA of a baculovirus from Penaeus japonicus (PjNOB = BMNV). Aquaculture 132, 213-220. Bonami, J.R., Brehelin, M., Mari, J., Trumper, B. and Lightner, D.V. (1990). Purification and characterization of IHHN virus of penaeid shrimps. Journal of General Virology 71, 2657-2664. Bonami, J.R., Lightner, D.V., Redman, R.M. and Poulos, B.T. (1992). Partial characterization of a togavirus (LOVV) associated with histopathological changes of the lymphoid organ of penaeid shrimps. Diseases of Aquatic Organisms 14, 145-152. Bonami, J.R., Bruce, L.D., Poulos, B.T., Mari, J. and Lightner, D.V. (1995). Partial characterization and cloning of the genome of PvSNPV (= BP-type virus) pathogenic for Penaeus vannamei. Diseases of Aquatic Organisms 23, 59-66. Bonami J.R., Hasson, K.W., Mari, J., Poulos, B.T. and Lightner, D.V. (1997). Taura syndrome of marine penaeid shrimp: characterization of the viral agent. Journal of General Virology 78, 313-319. Boonyaratpalin, S., Supamattaya, K., Kasornchandra, J., Direkbusaracom, S., Aekpanithanpong, U. and Chantanachooklin, C. (1993). Non-occluded baculo-like virus, the causative agent of Yellow Head Disease in the black tiger shrimp (Penaeus monodon). Fish Pathology 28, 103-109. Brock, J.A., Gose, R., Lightner, D.V. and Hasson, K.W. (1995). An overview on Taura syndrome, an important disease of farmed Penaeus vannamei. Pp. 84-94 In C.L. Browdy and J.S. Hopkins, editors. Swimming through troubled water, Proceedings of the special session on shrimp farming, Aquaculture '95. World Aquaculture Society, Baton Rouge, LA, USA. Brock, J.A., Gose, R. B., Lightner, D. V. and Hasson, K. (1997). Recent developments and an overview of Taura Syndrome of farmed shrimp in the Americas. Pp. 275-284 In: T.W. Flegel and I.H. MacRae (eds.) Diseases in Asian Aquaculture III. Fish Health Section, Asian Fisheries Society, Manila. Chang P.S., Lo, C.F., Kou, G.H. and Chen, S.N. (1993). Purification and amplification of DNA from Penaeus monodon-type baculovirus (MBV). Journal of Invertebrate Pathology 62, 116-120. Chantanachookin, C., Boonyaratpalin, S., Kasornchandra, J., Direkbusarakom, S., Ekpanithanpong, U., Supamataya, K., Sriurairatana, S. and Flegel, T.W. (1993). History and ultrastructure reveal a new granulosis-like virus in Penaeus monodon affected by `yellow head' disease. Diseases of Aquatic Organisms 17, 145-157. Chou, H.Y., Huang, C.Y., Wang, C.H., Chiang, H.C. and Lo, C.F. (1995). Pathogenicity of a baculovirus infection causing white spot syndrome in cultured penaeid shrimp in Taiwan. Diseases of Aquatic Organisms 23, 165-173. Couch, J.A. (1974a). Free and occluded virus similar to Baculovirus in hepatopancreas of pink shrimp. Nature 247, 229-231. Couch, J.A. (1974b). An enzootic nuclear polyhedrosis virus of pink shrimp: ultrastructure, prevalence, and enhancement. Journal of Invertebrate Pathology 24, 311-331. Durand, S., Lightner, D.V., Nunan, L.M., Redman, R.M., Mari, J. and Bonami, J.R. (1996). Application of gene probes as diagnostic tool for the white spot baculovirus (WSSV) of penaeid shrimps. Diseases of Aquatic Organisms 27, 59-66.
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Durand, S., Lightner, D.V., Redman, R.M. and Bonami, J.R. (1997). Ultrastructure and morphogenesis of white spot syndrome baculovirus (WSSV). Diseases of Aquatic Organisms 29, 205-211. Flegel, T.W., Sriurairtana, S., Wongteerasupaya, C., Boonsaeng, V., Panyim, S. and Withyachumnarnkul, B. (1995). Progress in characterization and control of yellow-head virus of Penaeus monodon. Pp. 76-83 In: C.L. Browdy and J.S. Hopkins, editors. Swimming Through Troubled Water, Proceedings of the Special Session on Shrimp Farming, Aquaculture '95. World Aquaculture Society, Baton Rouge, LA, USA. Fraser, C.A. and Owens, L. (1996). Spawner-isolated mortality virus from Australian Penaeus monodon. Diseases of Aquatic Organisms 27, 141-148. Hasson, K.W., Lightner, D.V., Poulos, B.T., Redman, R.M., White, B.L., Brock, J.A. and Bonami, J.R. (1995). Taura Syndrome in Penaeus vannamei: demonstration of a viral etiology. Diseases of Aquatic Organisms 23, 115-126. Huang, J., Song, X.L., Yu, J. and Yang, C.H. (1995). Baculoviral hypodermal and hematopoietic necrosis - study on the pathogen and pathology of the shrimp explosive epidemic disease of shrimp. Marine Fisheries Research 16, 1-10. Kasornchandra, J., Boonyaratpalin, S. and Itami, T. (1998). Detection of white-spot syndrome in cultured penaeid shrimp in Asia: microscopic observation and polymerase chain reaction. Aquaculture 164, 243-251. Kimura, T., Yamano, K., Nakano, H., Momoyama, K., Hiraoka, M. and Inouye, K. (1996). Detection of penaeid rod-shaped DNA virus (PRDV) by PCR. Fish Pathology 31, 93-98. Lightner, D.V. (1996). A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA. 304 p. Lightner, D.V., Redman, R.M., Bell, T.A. and Brock, J.A. (1983a). Detection of IHHN virus in Penaeus stylirostris and P. vannamei imported into Hawaii. Journal of the World Mariculture Society 14, 212-225. Lightner, D.V., Redman, R.M. and Bell, T.A. (1983b). Observations on the geographic distribution, pathogenesis and morphology of the baculovirus from Penaeus monodon Fabricius. Aquaculture 32, 209-233. Lightner, D.V. and Redman, R.M. (1993). A putative iridovirus from the penaeid shrimp Protrachypene precipua Burkenroad (Crustacea: Decapoda). Journal of Invertebrate Pathology 62, 107-109. Lightner D.V., Redman, R.M., Hasson, K.W. and Pantoja, C.R. (1995). Taura syndrome in Penaeus vannamei: histopathology and ultrastructure. Diseases of Aquatic Organisms 21, 53-59 Lightner, D.V. and Redman, R.M. (1998). Shrimp diseases and current diagnostic methods. Aquaculture 164, 201-220. Lo, C.F., Leu, J.H., Chen, C.H., Peng, S.E., Chen, Y.T., Chou, C.M., Yeh, P.Y., Huang, C.J., Chou, H.Y., Wang, C.H. and Kou, G.H. (1996a). Detection of baculovirus associated with white spot syndrome (WSBV) in penaeid shrimps using polymerase chain reaction. Diseases of Aquatic Organisms 25, 133-141. Lo, C.F.,. Ho, C.H, Peng, S.E., Chen, C.H., Hsu, H.C., Chiu, Y.L., Chang, C.F., Liu, K.F., Su, M.S., Wang, C.H. and Kou, G.H. (1997). White spot syndrome baculovirus (WSBV) detected in cultured and captured Shrimp, crabs and other arthropods. Diseases of Aquatic Organisms 27, 215-225. Lo, C.F., Kou, G.H., Hsu, H.C., Ho, C.H., Tsai, M.F. and Lightner, D.V. (1999). Specific genomic DNA fragment analysis of different geographical isolates of Shrimp white spot syndrome associated virus. Disease of Aquatic Organisms (in press). Lu, Y., and Loh, P.C. (1994). Viral structural proteins and genome analyses of the rhabdovirus of penaeid shrimp (RPS). Diseases of Aquatic Organisms 19, 187-192. Mari, J., Bonami, J.R. and Lightner, D. (1993). Partial cloning of the genome of infectious hypodermal and hematopoietic necrosis virus, an unusual parvovirus pathogenic for penaeid shrimps; diagnosis of the disease using a specific probe. Journal of General Virology 74, 2637-2643.
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Mari, J., Bonami, J.R. and Lightner, D.V. (1998). Taura syndrome of Penaeid shrimp: cloning of viral genome fragments and development of specific gene probes. Diseases of Aquatic Organisms 33, 11-17. Momoyama, K., and Sano, T. (1996). Infectivity of baculovirus midgut gland necrosis virus (BMNV) to larvae of 5 crustacean species. Fish Pathology 31, 81-85. Nadala, E.C.B., Jr., Lu, Y., Loh, P.C. and Brock, J.A. (1992). Infection of P. stylirostris (Boone) with a rhabdovirus isolated from Penaeus spp. Fish Pathology (Gyobyo Kenkyu) 27, 143-147. Nunan, L.M., Poulos, B.T. and Lightner, D.V. (1998). Reverse transcription polymerase chain reaction (RT-PCR) used for the detection of Taura Syndrome Virus (TSV) in experimentally infected shrimp. Diseases of Aquatic Organisms 34, 87-91. OIE. (1997). Diagnostic Manual for Aquatic Diseases. Office International des Epizooties, Paris, 251 p. Owens, L. (1993). Description of the first haemocytic rod-shaped virus from a penaeid prawn. Diseases of Aquatic Organisms 16, 217-221. Owens, L., DeBeer, S. and Smith, J. (1991). Lymphoidal parvovirus-like particles in Australian penaeid prawns. Diseases of Aquatic Organisms 11, 129-134. Reddington, J. and Lightner, D. (1994). Diagnostics and their application to aquaculture. World Aquaculture 25, 41-48. Spann, K.M., Vickers, J.E. and Lester, R.J.G. (1995). Lymphoid organ virus of Penaeus monodon. Diseases of Aquatic Organisms 26, 127-134. Spann, K.M., Cowley, J.A., Walker, P.J. and Lester, R.J.G. (1997). A yellow-head-like virus from Penaeus monodon cultured in Australia. Diseases of Aquatic Organisms 31, 169-179. Takahashi, Y., Itami, T., Kondo, M., Maeda, M., Fujii, R., Tomonaga, S., Supamattaya, K. and Boonyaratpalin, S. (1994). Electron microscopic evidence of bacilliform virus infection in Kuruma shrimp (Penaeus japonicus). Fish Pathology 29, 121-125. Takahashi, Y., Itami, T., Maeda, M., Suzuki, N., Kasornchandra, J., Supamattaya, K., Khongpradit, R., Boonyaratpalin, S., Kondo, M., Kawai, K., Hirono, I. and Aoki, T. (1996). Polymerase chain reaction (PCR) amplification of bacilliform virus (PV-PJ) DNA in Penaeus japonicus Bate and systemic ectodermal and mesodermal baculovirus (SEMBV) DNA in Penaeus monodon Fabricius. Journal of Fish Diseases 19, 399-403. Tang, K.F.J. and Lightner, D.V. (1999). A yellow-head virus gene probe: application to in situ hybridization and determination of its nucleotide sequence. Diseases of Aquatic Organisms (in press). Tsing, A. and Bonami, J.R. (1987). A new virus disease of the tiger shrimp Penaeus japonicus Bate. Journal of Fish Diseases 10, 139-141. Wang, C.H., Lo, C.F., Leu, J.H., Chou, C.M., Yeh, P.Y., Chou, H.Y., Tung, M.C., Chang, C.F., Su, M.S. and Kou. G.H. (1995). Purification and genomic analysis of baculovirus associated with white spot syndrome (WSBV) of Penaeus monodon. Diseases of Aquatic Organisms 23, 239-242. Wang, S.Y., Hong, C. and Lotz, J.M. (1996). Development of a PCR procedure for the detection of Baculovirus penaei in shrimp. Diseases of Aquatic Organisms 25, 123-131. Wang, Y.C., Lo, C.F., Chang, P.S. and Kou, G.H. (1998). White spot syndrome associated virus (WSSV) infection in cultured and wild decapods in Taiwan. Aquaculture 164, 221-231. Wongteerasupaya, C., Vickers, J.E., Sriurairatana, S., Nash, G.L., Akarajamorn, A., Boonsaeng, V., Panyim, S., Tassanakajon, A., Withyachumnarnkul, B. and Flegel, T.W. (1995). A nonoccluded, systemic baculovirus that occurs in cells of ectodermal and mesodermal origin and causes high mortality in the black tiger prawn, Penaeus monodon. Diseases of Aquatic Organisms 21, 69-77. Wongteerasupaya, C., Sriurairatana, S., Vickers, J.E., Anutara, A., Boonsaeng, V., Panyim, S., Tassanakajon, A., Withyachumnarnkul, B. and Flegel, T.W. (1995). Yellow-head virus of P. monodon is an RNA virus. Diseases of Aquatic Organisms 22, 45-50.
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Wongteerasupaya, C., Boonsaeng, V., Panyim, S., Tassanakajon, A., Withyachumnarnkul, B. and Flegel, T.W. (1997). Detection of yellow-head virus (YHV) of Penaeus monodon by RTPCR amplification. Diseases of Aquatic Organisms 31, 181-186. Table 1a. DNA Viruses of Penaeid Shrimp (as of February 1999; modified from Lightner, 1996; Lightner and Redman, 1998)
ACRONYM / FULL NAME
Key References
DNA VIRUSES PARVOVIRUSES: IHHNV HPV SMV LPV
= = = =
infectious hypodermal and hematopoietic necrosis virus hepatopancreatic parvovirus spawner-isolated mortality virus lymphoidal parvo-like virus
Lightner et al., 1983a,b Bonami et al., 1990 Adams and Bonami, 1991 Fraser and Owens, 1996 Owens et al., 1991
BACULOVIRUSES and BACULO-LIKE VIRUSES: BP-type = Baculovirus penaei type viruses (PvSNPV type sp.): BP strains from the Gulf of Mexico, Hawaii and Eastern Pacific MBV-type = Penaeus monodon-type baculoviruses (PmSNPV type sp.): MBV strains – East and SE Asia, Australia, the Indo-Pacific, and India BMN-type = baculoviral midgut gland necrosis type viruses: BMN = from Ma. Japonicus in Japan TCBV = type C baculovirus of P. monodon PHRV = hemocyte-infecting non-occluded baculo-like virus
Couch 1974a; 1974b Bonami et al., 1995 Adams and Bonami, 1990 Wang et al., 1996 Momoyama and Sano, 1996 Arimoto et al., 1995 Mari et al., 1993 Chang et al., 1993 Owens, 1993
WHITE SPOT SYNDROME VIRUSES (PmNOBII-type): SEMBV = systemic ectodermal and mesodermal baculo-like virus RV-PJ = rod shaped virus of Ma. japonicus PAV = penaeid acute viremia virus HNBV = hypodermal and hematopoietic necrosis baculo-like virus; agent of "SEEDS" (shrimp explosive epidermic disease) WSBV = white spot baculo-like virus PRDV = penaeid rod-shaped DNA virus
Wongteerasupaya et al., 1995 Takahashi et al., 1994; 1996 Huang et al., 1995 Wang et al., 1995, 1998 Lo et al., 1996; 1997; 1999 Durand et al., 1996, 1997 Chou et al., 1995 Kimura et al., 1996 Kasornchandra et al., 1998
IRIDOVIRUS: IRIDO
=
shrimp iridovirus
Lightner and Redman, 1993
42
Table 1b. RNA Viruses of Penaeid Shrimp (as of February 1999; modified from Lightner, 1996; Lightner and Redman, 1998). RNA VIRUSES ACRONYM / FULL NAME
Key References
PICORNAVIRUS:
Lightner et al., 1995 Brock et al., 1995; 1997 Hasson et al., 1995 Bonami et al., 1997 Mari et al., 1998 Nunan et al., 1998
TSV
=
Taura syndrome virus
Tsing and Bonami, 1987 Adams and Bonami, 1991
REOVIRUSES: REO-III and IV = reo-like virus type II and IV
Bonami et al., 1992 Lightner, 1996
TOGA-LIKE VIRUS: LOVV = lymphoid organ vacuolization virus
Nadala et al., 1992 Lu and Loh, 1994
RHABDOVIRUS: RPS
=
rhabdovirus of penaeid shrimp
YELLOW HEAD VIRUS GROUP: YHV/"YBV" = yellow head virus of P. monodon GAV = gill associated virus of P. monodon LOV = lymphoid organ virus of P. monodon
43
Chantanachookin et al., 1993 Boonyaratpalin et al., 1993 Wongteerasupaya et al., 95; 97 Tang et al., 1999 Flegel et al., 1995 Spann et al., 1995 Spann et al., 1997
Table 2. Proposed list of OIE notifiable and listed penaeid shrimp diseases and their current, presently known distribution in wild and cultured stocks (modified from Lightner, 1996; Lightner and Redman, 1998
Virus or Virus Group OIE Notifiable Viruses of Penaeid Shrimp: WSSV YHV TSV OIE Listed Viruses of Penaeid Shrimp: IHHNV BP MBV SMV
Eastern Hemisphere
Western Hemisphere
Wild and cultured Wild and cultured Not reported
Wild and cultured Not reported Wild and cultured
Wild and cultured Not reported Wild and cultured
Wild and cultured Wild and cultured Reported; not enzootic Not reported
Cultured
Table 3. Diagnostic and pathogen detection methods for the OIE notifiable and listed viral diseases of penaeid shrimp (modified from Lightner, 1996; Lightner and Redman, 1998
Method* Direct BF / LM / PH / DF Histopathology Bioassay TEM / SEM ELISA with PAb / Mab DNA Probes DBH / ISH PCR / RT-PCR
WSSV
IHHNV
BP
MBV
BMN
SMV
YHVgroup
TSV
++ ++ ++ + -
++ + + -
+++ ++ + + +
+++ ++ + -
++ ++ + + +
++ ++ -
++ +++ + + -
+ +++ ++ + ++
+++
+++
++
++
++
+++
+++
+++
+++
+++
+++
+
-
+++
+++
+++
* +
Definitions for each virus: = no known or published application of technique. = application of technique known or published, but not commonly practiced or readily available. ++ = application of technique considered by authors of present paper to provide sufficient diagnostic accuracy or pathogen detection sensitivity for most applications. +++ = technique provides a high degree of sensitivity in pathogen detection. Methods: BF = bright field LM of tissue impression smears, wet-mounts, stained whole mounts; LM = light microscopy; PH = phase microscopy, DF = dark-field microscopy, EM = electron microscopy of sections or of purified or semi-purified virus; ELISA = enzyme-linked immunosorbent assay; PAbs = polyclonal antibodies;
MAbs = monoclonal antibodies; DBH = dot blot hybridization, ISH = in situ hybridization
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Practical problems with PCR detection in Asia: The importance of standardization M. Shariff , S. Soon, K. L. Lee and L.T. Tan Aquatic Animal Health Unit, Faculty of Veterinary Medicine, University Putra Malaysia, 43400 Serdang, Selangor, Malaysia
Introduction The use of any DNA-based diagnostic technology faces some fundamental problems, as implementation requires knowledge in the field of molecular diagnostics. Such knowledge is obtained by experience gained from research on molecular detection methods. It is important that true expertise is established in molecular biology and that technologies are not introduced with a “jump in the bandwagon” mentality. It should also be recognised that interpretation of results can be problematic when strict controls, guidelines and problem solving are not adhered to. Furthermore, the implementation of DNA-based diagnostic programs must add real value. They should not be implemented for the sake of novelty, particularly when employed for diagnostic certification. The need for implementation needs to be determined in terms of practicality, ease of use, speed and technical capabilities of trained manpower. The programs must also be reliable and reproducible with very low frequency of false interpretation and variation between laboratories. Finally, the cost involved in running such programs should not be prohibitive as the outcome of results reflects the professionalism, responsibility and standards with which the tests were conducted.
Advantages of PCR diagnosis PCR is at the forefront of molecular diagnostic technology today. It is highly sensitive, with a capacity to amplify from even a single molecule of DNA. It is also very specific due to the nature and orientation of the oligonucleotide primers that are required to allow amplification to proceed. PCR is also very rapid. In only a few hours, millions of copies of a single DNA segment can be produced by standard procedures. In my laboratory, we have achieved similar results in less than 5 minutes by using rapid cycle amplification in capillary tubes. The potential for semi-automation of the procedure would make it even more attractive, allowing genetic information to be acquired more quickly. There is also potential for DNA sequence analysis of the PCR product to confirm the identity of a disease or pathogen and allow examination for genetic variation. Such advantages have been of great benefit to molecular diagnostics. In general, 2 areas that has benefited the most from this technology are disease gene mutation screening and of course the detection and characterisation of specific pathogens. Can there be any doubt that PCR should be used as a diagnostic tool when it can pick up a needle and amplify it in a haystack?
Potential obstacles to PCR standardization Why are there potential problems with the implementation of PCR for the diagnosis of WSSV? The reason is very simple. Diagnosing WSSV with PCR can go awfully wrong if the technology is applied casually. There are some contributing factors that could lead us to arrive at such a conclusion. Firstly, PCR requires some technical knowledge for effective implementation. Standardization between laboratories that may have no molecular biological research background will require the provision of support to develop technical capabilities. Secondly, due to the flexibility of PCR with its many amplification strategies, the selection of appropriate gold standards that will be critical for
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acceptance of diagnostic certification for WSSV requires careful consideration. The use of different strategies such as single or nested PCR leads to differences in the sensitivity of detection. There has been strong inclination for laboratories in Asia to adopt the nested approach, as the method is at least 100 to 1000 times more sensitive than single PCR. This extreme sensitivity comes with its own set of problems as will be discussed later. Another important factor that needs to be addressed is that of reproducibility between laboratories. The assay procedure not only consists of performing the PCR but also reproducing the same sensitivity, eliminating of false interpretation and implementing contamination control procedures. Questions have also been posed as to whether PCR results should be just a simple positive or negative. This issue stems from an interpretation that a PCR result represents an absolute presence or absence of WSSV in the test sample. Presently, routine PCR detection of WSSV can result in the same observation for both early and late stages of infection. The biological significance of such a PCR result is somewhat obscure. How do we assess the degree of infection? This problem has been demonstrated by the reports of farms having white spot infection but with varying degrees of mortality. The use of PCR in WSSV diagnosis is presently devoid of the ability to monitor the progress of the disease. There are ways to slightly overcome this problem, but the work and effort required for routine diagnosis may be prohibitive and the methods do not provide good accuracy in predicting the viral load of WSSV. Another area of serious concern is the high probability of false interpretation of results. This is largely due to ignorance in establishing, employing and following strict PCR controls for verification of results. For example, failure to use negative controls can lead to the interpretation of negative WSSV detection in a test which was actually a failed PCR! This is frequently caused by PCR inhibition – a factor which deserves greater attention in WSSV diagnosis as shrimp tissues frequently contain PCR inhibitory substances. False positive interpretation is almost exclusively due to contamination from post-PCR amplification products. While sample to sample contamination do exist, it is contamination from PCR products that causes greatest concern and headache. This problem is even more serious when procedures such as nested PCR are used for routine WSSV detection. In view of these problems, it will be important to ensure that, in addition to the use of standard PCR procedures, appropriate positive and negative controls are also employed. The use of proper controls is seldom emphasised, and if controls are emphasised obtaining them is not easy. Perhaps the greatest concern in the use of PCR for WSSV diagnosis is the lack of strict quality control on the results released. Without proper guidelines, consistent, standard and valid PCR diagnosis of WSSV will be virtually impossible to achieve.
Areas that require standardization With so many factors that could go wrong in PCR, one could wonder where to begin with standardization. Perhaps a general view of the process involved to set up a PCR based diagnostic assay would be useful. This is certainly not an exhaustive list but may help the difficult process of finding focal points in establishing standard and valid PCR assays for WSSV diagnosis in Asia. The areas we have identified that may lead to initiatives in standardization and validation programs are sample processing, PCR set-up, amplification strategies, controls, specificity and sensitivity. Sample processing. The initial stages before performing PCR analysis involve sample processing, and correct sampling methods. It is important to ensure the correct sample size for reliable detection of pathogens as possible low prevalence of the agents can render a false negative interpretation. In determining sample size for PCR analysis, the statistical confidence levels provided by Amos (1985) are a useful guide. However, a practical issue to consider is the sampling method by the farmer. For example, we have had cases in which hatchery operators have requested PCR analysis on post-larval
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samples that have been pooled from various tanks. This sampling practice jeopardises the accuracy and statistical confidence of the PCR results. The proper selection of DNA sources is also critical in obtaining accurate PCR results. From our own experience, use of gill, muscle and integument tissues yields good results, provided that the amount of DNA is appropriate. An excess of DNA may inhibit the PCR process. Haemolymph can also be used but it is usually not appropriate for routine diagnosis. PCR on post-larvae is sometimes more problematic as PCR inhibitory substances are present in the eyes which should be removed before DNA extraction. The inhibitory substances have not yet been properly identified but there is evidence suggesting the involvement of poryphrin which is a common component of pigment in shrimp. Little research has been conducted to resolve this problem. The DNA extraction procedure may be a good area in which to commence standardisation. There are a number of methods to obtain DNA for PCR analysis and these will be discussed later. The key factors to consider in the development of a standardized procedure are time/manpower, safety, reliability, contamination risks, inhibition and DNA quality. The significance time required to conduct the test is important, especially when a large sample number has to be screened. The availability of manpower and technical technical skills should also be considered. Worker safety when employing the procedures should not be compromised. The extraction procedure should be evaluated for its reliability and risk of contamination when processing a large number of samples. The extraction methods should minimise the use of PCR inhibitory substances as will be described later. There is also a need for some level of DNA quality for PCR analysis to generate accurate results. Methods which can be employed for extraction of WSSV DNA include alkaline lysis, proteinase K digestion, treatment with guanidinium salts (passive) and boiling. Alkaline lysis is rapid and provides good quality DNA for PCR. However, a boiling step to properly liberate DNA from tissues and a neutralisation step are required. Depending on how the samples are handled, contamination risks can increase when boiling is employed. Proteinase K digestion is relatively slow as it involves several subsequent steps and the protocol also usually involves the use of hazardous materials such as phenol and chloroform. Detergents are also commonly used for the liberation of DNA. However, ionic detergents such as SDS can inhibit Taq polymerase at certain concentrations. Residual phenol can also cause inhibition of Taq polymerase. Guanidinium salts have not been widely used but could provide some very useful advantages. Guanidinium salts are non-toxic and the method is rapid, requiring only a few steps involving ethanol washing and precipitation. The technique also allows passive lysis for release of DNA from tissues without the need to boil or homogenise. As such, contamination risks can be significantly reduced. Guanidinium salts also inhibit common DNA degrading enzymes so tissue can be stored in solution while being transferred or while awaiting extraction. The use of simple boiling methods in lysis buffers should be evaluated carefully in terms of contamination risks and DNA quality. It is our opinion that the disadvantages far outweigh its advantages of DNA extraction using this method. In summary, a good DNA extraction protocol after going through all the above discussion should ideally be rapid, non-toxic, economical, and have a low risk of contamination. PCR set-up. The second area for which standardisation is the set-up of the PCR laboratory. A properly equipped laboratory to perform PCR diagnostics is absolutely critical if routine WSSV diagnosis is to be conducted. The use of dedicated instruments such micro-pipettors and aerosol resistant tips for PCR must be strictly observed. A PCR facility must be clean and well managed, and the architecture of the laboratory must include separate rooms for pre- and post-amplification procedures. Samples must not be prepared in the same room as the PCR machine is located and PCR
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post-amplification analysis is performed. The laboratory in which PCR reagents are prepared should also clean benches and ideally should include a laminar flow cabinet with built-in UV lamps for decontamination procedures after every reaction set-up. The need for trained manpower is critical, not just to perform the PCR assay but to ensure the reliable management of a PCR diagnostic laboratory. PCR reagents must be properly prepared and troubleshooting skills must be available so that the quality of result test results is consistently and strictly controlled. Decontamination protocols should also be established and strictly followed. Amplification strategies. Despite all care, it is our experience that contamination can and will occur. For this reason, the use of low contamination risk amplification strategies is essential. The following PCR strategies are now in use. PCR may be conducted using single step or nested primer methodologies. In our experience, as shared by many other laboratories in Asia, nested PCR can reach 1000 times the sensitivity as compared to single step PCR. The extreme sensitivity of nested PCR works well in many cases but the test is highly susceptible to contamination as it involves more time for sample manipulation. The most common source of contamination is the product from a previous PCR. In fact, one molecule of a contaminating PCR template in the first step reaction may be sufficient to obtain a false positive result by nested PCR. The nested PCR is also more costly as it involves the use of 2 separate reactions to arrive at one result. The cost will increase dramatically if assays are repeated when contamination occurs. A new methodology which may give more biological significance to the test result is quantitative PCR. Qualitative PCR provides no useful quantitative assessment of the infection level or disease progression and does not indicate pathogen replication. Because of this deficiency, the application of primary cell lines have been suggested to be more appropriate pathogen detection than PCR, despite the difficulty in establishing shrimp cell lines. However, quantitative PCR technology has been available for some time and has been widely applied in diagnosis of human pathogens such as HIV and herpes simplex virus. The ability to quantify using PCR allows investigation of the molecular pathogenesis of an infection. Such information for WSSV has been very limited apart from studies conducted using DNA probes in Taiwan. In situ hybridisation (ISH) using DNA probes also available for disease investigation but the method is technically demanding and requires time for data interpretation. Although it does not provide a direct quantification of the virus, ISH may give information on the severity of infection. Quantification using PCR is more rapid and accurate, and can provide an absolute determination of the number or copies of the targeted DNA. The most reliable method for PCR quantification is by competitive PCR. This technique utilises a known amount of engineered internal standard which has the same primer binding sites as the target DNA or RNA. The internal standard is differentiated from the target by size on the basis of size by inclusion of a small deletion, or by including a single base mutation that allows separation after restriction enzyme digestion. Since the target DNA and internal standard are virtually identical, the efficiency of amplification should be equivalent, leading to a fair competition when co-amplified and an accurate measurement of the relative concentrations. This eliminate tube-to-tube and template variability. PCR quantification can be conducted in real time with specialised equipment which uses fluorescent dye quenching technology. The cost to purchase such equipment may be prohibitive for some laboratories. The procedure makes absolute quantification possible but is technically demanding in the initial stages of design and development of engineered standards for co-amplification. It is also more costly as it requires the amplification of multiple standards simultaneously and is less sensitive as compared to nested PCR. In order to develop an easy, sensitive and robust method with lower risk of contamination for the routine diagnosis of WSSV infection, we have been developing a single sample load, single-tube nested PCR. By this method, we can reliably detect 1-10 copies of double-stranded viral DNA target.
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The assay is much easier, more user friendly and faster with very little hands-on manipulation of the PCR reaction preparation and is less susceptible to sporadic contamination. The assay is also half the cost of our normal 2 tube nested PCR as only require one reaction tube and set of PCR reaction components are required. Semi-quantitative properties can also be incorporated into the system for an assessment of the WSSV infection level. The two tube nested PCR provides very little quantitative information. A positive PCR signal has very much the same overall intensity over a wide range of DNA target copies. The reamplification of the first step PCR product quickly masks differences in the concentrations of amplified products due to the tendency of the PCR process to reach a plateau after 30 or 40 cycles, regardless of the amount of the input target template. This is also due in part to the presence of renewed sources for PCR amplification in the second tube. The characteristics of the single-tube nested PCR approach are different. There is a direct relationship between the amount of the first step PCR product and the amount of the nested PCR product. At low target concentration, the nested PCR step does not proceed to reach a plateau as in the 2 tube nested PCR. This phenomenon occurs only when high template target is present. As such, different levels of PCR signal intensity can be observed, reflecting the severity of the infection. This permits a semi-quantitative approach to be applied to the assessment of a positive result. Although this requires extensive and intensive optimisation to achieve reliability, such approaches may be more beneficial in the long run. In our case, the use of more sophisticated PCR methods has been worth the investment in energy, money and manpower. Diagnostic controls. Standardisation is also required in the use of appropriate diagnostic controls. Without proper controls for result interpretation, no PCR result should ever be accepted as valid. Appropriate controls, which are essential for elimination of false negative results, involve the simultaneous amplification of a fragment of host DNA using primers targeting conserved sequences in the host genome. This control indicates that the PCR has been successful and that template quality was adequate for PCR amplification. It also allows the recognition of PCR inhibition factors in the sample. The process will require the use of multiplex PCR as it will be more reliable when the control is amplified in the same diagnostic reaction. Appropriate control reactions to indicate false positive results should also be performed. A negative control reaction without template must be used in every assay. The use of vapour barriers such a mineral oil overlay of the PCR reaction is advised even though most thermal cyclers now employ a heated bonnet to prevent vapour transfers. Strict adherence to contamination prevention and decontamination procedures must also be followed. The use of positive controls that can be differentiated from the target is also advantageous when it is necessary to distinguish a true positive result from contamination with a positive control. By knowing whether contamination is from a diagnostic target or from a positive control permits a more focused approach in eliminating the source of contamination. Weak positive controls (low amount of target) can also be employed if necessary to minimise the risk of contamination in the laboratory. A novel approach is the use of engineered controls that differ in properties from diagnostic targets. Differentiation can be accomplished by another round of PCR or by hybridisation with an internal probe that detects the difference in the PCR fragments. However, the fastest and easiest way is to use restriction enzymes that cut either the target or control fragments. By this method, the result can be determined by analysing the PCR products elctrophoretically. Diagnostic specificity. In establishing standard PCR protocols for WSSV diagnosis, the specificity of PCR primers for conserved regions of the genome also should be considered. WSSV variants may occur that cannot be detected by certain primer sequences. For example, Korean researchers (Park et al., 1998) could amplify PCR products from WSSV-diseased shrimp using primers developed for RVPJ but not when using primers developed by Lo and co-workers. It is also interesting that the Korean
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virus looks similar to WSSV found in Taiwan and not to RV-PJ from Japan. The sequence of the PCR product was identical to sequence of RV-PJ fragment. It seems likely that mutations in WSSV are occurring and this should be investigated. The possibility of having less virulent or virulent strains of WSSV should also be considered. This has been reported for yellow head viruses in research conducted in Australia (Dr. Peter Walker, CSIRO, Chiang Mai Meeting). However, as YHV is an RNA virus, it would be expected to mutate more frequently than WSSV which is a DNA virus. Confirmation of PCR specificity is quite straightforward. The nested PCR approach is already an accepted method for confirmation of PCR specificity based on the distinct size of the nested PCR product. Other techniques such as RFLP analysis and the use of an internal ISH probe are more laborious. Diagnostic sensitivity. Another area that sometimes evokes confusion that requires proper standardisation is the definition of PCR diagnostic sensitivity or the limit of PCR detection. Descriptions of detection limits currently vary from the number of DNA target copies to the number of virus particles. It has not been established for WSSV whether the identification and detection of single gene sequence reflects the detection of a single viable virus. It should be determined if sensitivity can be standardised with quantitative qualities such as infection status, disease progression, viral replication or antiviral therapies and strategies. The ideal WSSV diagnostic PCR assay would determine the infection status and allow more accurate monitoring of the progression of the disease. The monitoring of the disease has an important role in the management of WSSV infection on farms. There is clearly potential to reduce the economic impact of WSSV by developing disease management plans based on viral load information. In this approach, it will be important to determine whether the virus is replicating and causing the disease. The most important potential application of PCR technology is the possibility of developing new antiviral therapies or strategies against WSSV. It is impossible to assess these accurately using the techniques presently available. Conclusion How do we begin to validate PCR protocols then for WSSV diagnosis? Is it first through standardisation or it is more practical to identify areas for validation that would allow a better standardisation exercise? After evaluating all the problems and technical issues associated with the PCR process, validation steps can be perhaps grouped into a few general areas. Firstly, it is obvious that for a procedure to be valid, the protocol must be reliable, simple and sensitive enough to do the work required. It must be technically easy to perform to allow reliable replication between laboratories. This should encompass the entire process from DNA preparation to PCR amplification. Secondly, the use of proper controls must be employed consistently to totally eliminate false negative and false positive results. In other words, very strict quality control must be imposed on laboratories performing diagnosis. Thirdly, it may be important to include a quantitative element to add the biological significance dimension to diagnosis. This criterion has often been applied to diagnostic assays for infectious pathogens. In any case, it is easier to verify a result when degree of infection is incorporated into the assay. Again strict quality control must be implemented to allow the validation of a diagnostic result. This may also minimise legal concerns and certainly will reduce unnecessary economic losses on farms. The use of a PCR test kit may help in the validation process. In this way, individual laboratories would need only to meet certain requirements and avoid the need to develop diagnostic methods that
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meet the imposed standards. However, a suitable PCR diagnostic kit must first be developed. This will help encourage private companies to provide commercial clinical diagnostic services without a huge investment in RandD. The kit in our opinion must satisfy basic criteria such as being simple to perform and interpret, have a low contamination risk, include an appropriate false negative and false positive control/indicators, have appropriate sensitivity controls and the test should be semiquantitative. In short, the test should be designed to meet the expectations of the novice clinician for whom the use of PCR as a diagnostic tool is still foreign. The test should also be sufficiently complete in its features to satisfy the experienced investigator. The interpretation of results should be similarly and readily understood by both groups. The complete package should be sufficiently attractive to justify its use despite a potentially higher cost. The test must be the gold standard if is to be implemented as a service to our local farmers who depend on us in seeing through their crop and their livelihood.
References Botwell, D.D.L. (1987). Rapid isolation of eukaryotic DNA. Analytical Biochemistry 162, 463-465. Chang, P.S., Lo, C.F., Wang, Y.C. and Kou, G.H. (1996). Identification of white spot syndrome associated baculovirus (WSBV) target organs in shrimp Penaeus momnodon by in situ hybridization. Diseases of Aquatic Organisms 27, 131-139. Chomczynski, P., Mackey, K., Drews, R. and Wilfinger, W. (1997). DNAzol£: A reagent for the rapid isolation of genomic DNA. BioTechniques 22, 550-553. Clementi, M., Menzo, S., Bagnarelli, P., Manzin, A., Valenza, A. and Varaldo, P.E. (1993). Quantitative PCR and RT-PCR in virology. PCR Methods and Application 2, 191-196. Clementi, M., Menzo, S., Manzin, A. and Bagnarelli, P. (1995). Quantitative molecular methods in virology. Archives of Virology 140, 1523-1539. Flegel, T.W. (1997). Major viral diseases of the black tiger prawn (Penaeus monodon) in Thailand. World Journal of Microbiology and Biotechnology 13, 433-442. Lo, C.F., Leu, J.H., Ho, C.H., Chen, C.H., Peng, S,E., Chen, Y.T., Chou, C.M., Yeh, P.Y., Huang, C.J., Chaou, H.Y., Wang, C.H. and Kou, G.H. (1996). Detection of baculovirus associated with white spot syndrome (WSBV) in penaeid shrimps using polymerase chain reaction. Diseases of Aquatic Organisms 27, 131-139. Park, J-H., Lee, Y.S., Lee, S. and Lee, Y. (1998). An infectious viral disease of penaeid shrimp newly found in Korea. Diseases of Aquatic Organisms 34, 71-75. Rolfs, A., Schuller, I., Finckh, U. and Weber-Rolfs, I. (1992). PCR: Clinical diagnostics and research. Springer-Verlag Berlin Heidelberg. 268 pp. Saiki, R.K., Gelfand, D.H., Stoffel, S., Scharf, S.J., Higuchi, R., Horn, G.T., Mullis, K.B. and Erlich, H.A. (1988). Primer directed enzymatic amplification of DNA with a thermostable DNApolymerase. Science 239, 487-491. Wongteerasupaya, C., Wongwisansri, S., Boonsaeng, V., Panyim, S., Pratanpipat, P., Nash, G.L., Withyachumnarnkul, B. and Flegel, T.W. (1996). DNA fragment of Penaeus monodon baulovirus PmNOBII gives positive in situ hybridization with white spot viral infections in six penaeid shrimp species. Aquaculture 143, 23-32. Xu, H., Jevnikar, A.M. and Rubin-Kelly, V.E. (1990). A simple method for the prepartion of chromosomal DNA from cell. Culture. Nucleic Acids Research 18, 4943.
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Application of polymerase chain reaction for detection of shrimp pathogens in India Indrani Karunasagar Department of Fishery Microbiology, University of Agricultural Sciences, College of Fisheries, Mangalore - 575 002, India
Rapid detection of pathogens would be very essential for effective health management in aquaculture. While conventional microbiological isolation methods are used in case of bacterial pathogens, histopathology is widely used to detect viral infections. However, these methods are time consuming and lack sensitivities to detect latent pathogens. Polymerase chain reaction (PCR) on the other hand is highly sensitive and rapid and can be used to detect latent pathogens. In our laboratory, we have been studying the application of PCR for rapid detection of shrimp pathogens such as whitespot syndrome virus (WSSV) and Vibrio spp. WSSV is a serious pathogen that has caused extensive mortalities in shrimp culture systems in India (Karunasagar et al., 1997, 1998; Karunasagar and Karunasagar, 1999). We have studied the application of PCR primers reported by Lo et al. (1996) for detection of WSSV. Our results indicate that one step PCR is able to detect infection in case of clinically symptomatic animals. Two step PCR was necessary to detect WSSV in clinically asymptomatic shrimps and in other carrier animals (Otta et al., 1999). Using two step PCR, a large number of apparently healthy P.monodon post larval stages were screened for the presence of WSSV. Only 5% apparently healthy PL gave positive reaction in one step PCR whereas 48% showed positive reaction in two step PCR (Otta et al., 1999). These results suggest that two step PCR is very essential for detection of WSSV in asymptomatic animals and carriers. Using PCR, WSSV could be detected in carrier animals such as crabs, Acetes spp and even in water samples. An evaluation of the relation between PCR positivity and infectivity was studied in the case of Penaeus monodon showing clinical signs of white spot syndrome. Viral extracts from freshly harvested shrimp (22-25g) were highly infective causing clinical signs and mortality in healthy shrimp within 48 h. Viral extracts prepared from clinically symptomatic animals, which were stored under frozen conditions (-20qC) for two months, showed positive reaction in PCR. However, the viral extracts from the frozen specimens failed to induce clinical signs or mortality in healthy animals. This suggests that WSSV might lose infectivity during frozen storage in shrimp tissue. Therefore, PCR positivity in frozen shrimp should be interpreted with caution with respect to its potential to spread the virus. In our laboratory, PCR is also being used to detect Vibrio parahaemolyticus in shrimp. Comparision of culture and PCR methods show that PCR would be very useful in detecting this organism and the technique has potential to detect even atypical strains showing variation in biochemical reactions (Karunasagar et al., 1997). Thus PCR would be a very useful tool for rapid and sensitive detection of pathogens in shrimp. The technique has applications in diagnostic laboratories, for monitoring health, to identify environmental reservoirs of infections, to detect the presence of pathogens in animals in quarantine etc.
References Lo, C.F., Leu, J.H., Ho, C.H., Chen, C.H., Peng, S.E., Chen, Y.T., Chou, C.M., Yeh, P.Y., Huang, C.J., Chou, H.Y., Wang, C.H. and Kou, G.H. (1996). Detection of white spot syndrome baculovirus (WSSBV) in penaeid shrimp using polymerase chain reaction. Diseases of Aquatic Organisms 25, 133-141.
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Karunasagar, I., Otta, S.K. and Karunasagar, I. (1997). Histopathological and bacteriological study of white spot syndrome of Penaeus monodon along west coast of India. Aquaculture 153, 913. Karunasagar, I., Nayak, B.B. and Karunasagar, I. (1997). Rapid detection of Vibrio parahaemolyticus from fish by polymerase chain reaction. In Diseases in Asian Aquaculture III. T.W. Flegel,. et al (Ed). Pp. 119-122. Asian Fisheries Society, Manila.. Karunasagar, I., Otta, S.K. and Karunasagar, I. (1998). Disease problems affecting cultured penaeid shrimp in India. Fish Pathology 33, 413-419. Karunasagar, I. and Karunasagar, I. (1999). Diagnosis, treatment and prevention of microbial diseases of fish and shellfish. Current Science 76, 387-399. Otta, S.K., Shubha, G., Biju, J., Karunasagar, I. and Karunasagar, I. (1999). Polymerase chain reaction based detection of whitespot syndrome virus in cultured and wild crustaceans in India. Diseases of Aquatic Organisms (in press).
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Viral genetic variation: implications for disease diagnosis and detection of shrimp pathogens Peter J. Walker and Jeff A. Cowley Co-operative Research Centre for Aquaculture, CSIRO Tropical Agriculture, PMB3 Indooroopilly, Q 4068, Australia.
Introduction During the past 10 years, the shrimp farming industry in the Asia-Pacific region and in the Americas has experienced the devastating impact of successive panzootics of viral disease (Chamberlain, 1999). In the ongoing effort to control and prevent these diseases, molecular methods are finding increasing application for differential diagnosis, epidemiological investigations and screening of covert infections in hatcheries and on farms. Methods such as the polymerase chain reaction (PCR), dot-blot hybridisation (DBH) and in situ hybridisation (ISH) have now been developed for a wide range of shrimp viruses and a number of significant bacterial pathogens (Lightner and Redman, 1998). Although technically complex and requiring specialised analytical equipment, these molecular methods have been adopted at a suprisingly rapid rate by diagnosticians, researchers and industry. This is primarily due to their exquisite sensitivity and specificity compared to standard histological procedures. The rapid uptake has also been driven by the lack of other methods that are used commonly in diagnosis of animal viral infections. Because of the absence of adaptive immunity in invertebrates (Fearon and Locksley, 1996), serology cannot be used to detect existing or prior infections with shrimp viruses. Virus propagation in vitro has had very limited application because of the absence of suitable cultured cell lines. Antibody-based methods for viral detection (eg. ELISA, indirect immunofluorescence or immunoperoxidase tests) appear to offer considerable potential but have yet to be explored adequately for most shrimp viruses. Although molecular genetic methods can provide rapid and accurate information on the infection status of shrimp, there is a considerable risk of misdiagnosis if the various parameters that determine reaction specificity are not carefully monitored and controlled. In each of these methods, diagnostic specificity is determined by a hybridisation (annealing) reaction in which a DNA (or RNA) probe must bind to the target sequence in the infecting virus. The efficiency of this annealing reaction will be determined by various parameters such as temperature, ion concentration, the accessibility and integrity of the target, the size of the probe and the relative concentrations of target and probe. The test result will also be influenced by the accuracy of the match between the probe and target sequences. This is particularly a problem in PCR tests for which a single base mismatch can sometimes prevent primer efficient extension of the primer-template hybrid (Kwok et al., 1990; Sommer and Tautz, 1989). This paper will consider the inherent genetic variability of viruses and the implications of sequence variation for pathogen detection and diagnosis. The potential to exploit variations in the genetic sequence of viral isolates to determine disease epidemiology and to track the movements of aquatic animal pathogens will also be considered.
Biomolecular basis of viral variation Genetic variation is an essential feature of all living organisms. It provides the resource for natural selection and for the progressive adaptation of the population to a changing environment. Viruses face continuous environmental change as they pass from host to host. The most obvious and significant of these is the defensive or immunological response. Evasion of the host defences is a central feature of the survival strategy of all viruses. However, allelic variations in host genes or differences in their
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pattern of expression also present a changing environmental landscape that can determine susceptibility to infection or efficiency of replication (Gibbs et al. 1995; Morse, 1994). Viral variation can be generated by a number of mechanisms. Major rearrangements in genome structure and organisation can occur by genetic recombination. Gene duplications, gene exchanges and gene adoptions also occur. However, the most common form of variation is mutation by nucleotide substitution. This occurs as a consequence of polymerase error in reading the template during replication. As viruses replicate rapidly and prodigiously, viral variation has significant implications for diagnosis and epidemiology (Morse, 1994).
Genetic variation in RNA and DNA viruses From a genetic perspective, viruses can be classified according to whether the genome comprises RNA or DNA. RNA viruses are inherently hypervariable as RNA polymerases, which replicate the viral genome, lack proof reading and error editing functions that occur in cellular DNA polymerases (Steinhauer and Holland 1987; Steinhauer et al. 1992). The resulting rate of nucleotide misincorporation in RNA viruses (10-3-10-4) is at least 1000 times that of bacteria or eukaryotes, causing one or more base substitutions each time the viral genome replicates. Some mutations are lethal as they truncate or distort the resulting protein, rendering it non-functional. However, many mutations result in viable genomes that continue to replicate and contribute to the virus population. In this way, RNA viruses continually refine their genetic structure to accommodate the changing environment. Some RNA viruses may also undergo genetic rearrangements that allow exchange of corresponding genes or gene segments during mixed infections (Steinhauer and Holland, 1987). These recombination and reassortment events allow the most efficient and environmentally adapted combinations of genes to emerge from the available genetic pool, increasing the potential for viral survival. RNA viruses known to infect farmed shrimp include Taura syndrome virus (TSV), yellow head virus (YHV), gill-associated virus (GAV), lymphoid organ virus (LOV) and rhabdovirus of penaeid shrimp (RPS). Each of these viruses is likely to replicate with a high mutation frequency. Some may also have a capacity for genetic recombination. In DNA viruses, the mutation rate is usually far lower than in RNA viruses (Steinhauer and Holland, 1987). DNA polymerases, both cellular and viral, do employ proof reading and repair functions to reduce the intrinsic error rate. However, some small DNA viruses (eg. parvoviruses) appear to produce factors that suppress the repair function, generating an error rate similar to that of RNA viruses (Parrish et al., 1991). Even at the lower error frequency, the prodigious replication rate of DNA viruses generates mutations in the genome that can be observed over time. For example, during natural infections of palm beetles following experimental release, the Oryctes rhinoceros non-occluded baculovirus has been reported to mutate at the rate of approximately 0.05 % (or 100 nucleotides) per year. Over a monitoring period of 4 years, changes in genome structure such as insertions, point mutations and recombinations could easily be detected in molecular tests (Crawford and Zelany, 1990). DNA viruses have also been reported to produce sequence duplications and insert host DNA sequences into the viral genome. DNA viruses known to infect farmed shrimp include white spot syndrome virus (WSSV), monodon baculovirus (MBV), baculoviral midgut necrosis virus (BMNV), infectious hypodermal and haematopoeitic necrosis virus (IHHNV), spawner mortality virus (SMV) and hepatopancreatic parvovirus (HPV). A recent study of WSSV DNA from sources in different geographic locations has suggested little sequence variation between isolates, except in some samples obtained from the USA (Lo et al., 1999). It is not yet known if crustacean parvo-like viruses (HPV, SMV and IHHNV) have a capacity to suppress the error repair function of DNA polymerases.
55
Genetic variation in viral detection and disease diagnosis Observations of genetic variability in viruses proclaim the need for care in the use of molecular methods for disease diagnosis. Mutations in the nucleotide sequence can prevent binding of PCR primers to target sequences, cause primers to bind non-specifically to non-target sequences, or prevent PCR extension of the sequence from the primer site (Kwok et al., 1990; Sommer and Tautz, 1989). Sequence insertions or duplications can generate size variations in the PCR product. In each case, the result may appear falsely negative. At the protein level, mutations and other variations in sequence can affect the binding of diagnostic reagents such as monoclonal antibodies. Variations can also cause closely related strains to have significantly different biological properties such as pathogenicity, tissue tropism or host range. An understanding of these factors is important for accurate interpretation of data obtained for disease diagnosis, epidemiolgical investigation or screening for disease-free certification.
The YHV complex – a case study in viral variation At least three RNA viruses with very similar morphology infect farmed P. monodon in the Asia-Pacific region. As the first of these to be reported was yellow head virus, the term ‘YHV complex’ has been adopted here to encompass this group of related agents. An understanding of the relationship between the viruses in the YHV complex is now emerging from molecular genetic studies that illustrate the importance of viral variation in disease diagnosis and epidemiology. Yellow head virus (YHV) was first reported to be associated with mass mortalities of farmed P. monodon in Thailand in 1990 (Limsuwan, 1991). It now appears that YHV or related viruses may have been responsible for serious production losses in Taiwan, Indonesia, China, Malaysia and the Philippines since 1986 (Lightner, 1996). Yellow head disease affects juvenile to sub-adult prawns in which it usually causes a yellowish colouration of the cephalothorax and gills, and stimulates erratic swimming near the surface at the pond edge. YHV replicates in the cytoplasm of infected tissues that include lymphoid organ, haemocytes and gills. The virus infects a range of penaeid species but appears not to infect other crustaceans. Gill-associated virus (GAV) has been the primary cause of a yellow head-like disease and associated mortalities that have affected the industry in Australia since 1994. The virus is indistinguishable from YHV by TEM, infects a similar range of tissues, and causes similar histopathology (Spann et al., 1997). In moribund prawns, the lymphoid organ displays extensive structural degeneration and cellular necrosis. In GAV infections, mortality is usually preceded by varying degrees of red colouration of the body and pink to yellow colouration of the gills. There has been no evidence of pale body colouration or yellowing of the cephalothorax as described for YHV. Prior to the identification of GAV, a virus with similar morphology was observed to be common in healthy P. monodon in Australia (Spann et al., 1995). Lymphoid organ virus (LOV) causes the formation of distinct foci of hypertrophic cells (spheroids) in the lymphoid organ which otherwise remains structurally intact. ISH and TEM of lymphoid organ tissue indicate that LOV is contained only within spheroids. The virus has not been observed by TEM in other tissues but can be detected in haemolymph and gills by PCR tests. LOV infections appear to be non-pathogenic in uncompromised P. monodon. The complete nucleotide sequence of the GAV genome has now been obtained (JA Cowley and PJ Walker, unpublished data). Analysis of the sequence has indicated that GAV is most closely related to RNA viruses in the family Coronaviridae. In order to assess the genetic relationship between GAV and YHV, sequence comparisons were conducted in 3 regions of the RNA replicase (ORF1b) gene that had been used for the development of PCR and ISH tests (Wongteerasupaya et al., 1997; Tang and Lightner, 1998; Cowley et al., submitted). The comparisons over a total of 1780 nucleotides (approximately 6.0 % of the total genome) indicate that GAV and YHV vary by 17.6 % in nucleotide sequence and 10.7 % in amino acid sequence. This degree of variation is typical of closely related
56
RNA viruses that constitute distinct geographic topotypes (Cowley et al., 1999). As the sequence of viral polymerase genes usually is relatively conserved, more genetic variation between YHV and GAV might be expected in some other regions of the genome. Comparison of sequences amplified from the ORF1b gene of a large number of LOV isolates from healthy P. monodon has indicated that they vary from the prototype GAV nucleotide sequence by an average of d1.5 % (Cowley et al., 1999). This is within the range of variation expected within a single population of replicating RNA sequences and indicates that the GAV and LOV are pathogenic and non-pathogenic variants of the same virus. The available nucleotide sequence data has been used to develop primary and nested RT-PCR tests to detect GAV in infected prawn tissue (Cowley et al., submitted). Each of these GAV RT-PCR tests will detect both GAV and LOV, which cannot presently be distinguished genetically. The GAV RT-PCR test will also amplify the expected product from a Thai isolate of YHV. This occurs because the nucleotide sequences of GAV and YHV in the regions targeted by the PCR primers are sufficiently related to allow primer hybridization under the conditions of the test. An RT-PCR test has also been described for detection of YHV (Wongteerasupaya et al., 1997). However, due to significant differences in sequence from YHV at one of the primer binding sites, this test will not detect either GAV or LOV (Cowley et al., 1999). A comparison of the sequences of YHV and GAV isolates in the primer binding sites for each of these PCR tests is shown in Fig.1, illustrating the poor correspondence in sequence at the site of primer 144R. A first assessment of these tests might suggest that the GAV PCR is group-specific, detecting all 3 viruses, and that the YHV PCR discriminates between GAV and YHV. However, as the extent of variation among viruses associated with yellow head disease in Thailand and the Asian region is presently unknown, such a presumption is premature. It is also possible that other non-pathogenic LOV-like viruses are common in the region. We have observed that the prevalence of LOV in healthy P. monodon captured in northern Queensland is extremely high (Cowley et al., submitted) and it is likely that pathogenic YHV-complex viruses emerge from such a background of non-pathogenic infection. This clearly illustrates the need for care in the design and interpretation of PCR tests and the need for accreditation and standardisation of procedures. Ultimately, when adequate information on the nature and distribution of YHV complex viruses is available, sequence data could be used to devise a range of encompassing and discriminatory molecular tests.
Primer 10F YHV : ::: ::::: ::::: : GAV Primer 144R YHV :: :: : :: :: : GAV
5’-CCGCTAATTTCAAAAACTAAG-3’ ATGATAACTTCAAGAACTATG
5’-CTTCCTCGACATAACACCTT-3’ TCATCTTGATCTCACGCCCT
Figure 1. Comparison of the YHV and GAV sequences at the primer binding sites for YHV PCR primers 10F and 144R Wongteerasupaya et al. (1997). Dots indicate the location of homologous nucleotides. GAV sequences are described in Cowley et al. (1999).
Molecular epidemiology and the movement of aquatic animal pathogens Although presenting challenges for test design, variability in nucleotide sequence can be a very potent tool in understanding the epidemiology of disease. By applying nucleotide sequence analysis and other discriminating molecular techniques to the analysis of virus isolates, there is potential to trace the origin and movement of viruses on a local and regional basis. It may also be possible to discriminate between pathogenic and non-pathogenic strains that otherwise may be indistinguishable. Such
57
molecular approaches to epidemiology are now commonly used in the study of viruses infecting terrestrial animals and humans. In the case of YHV-complex viruses, sequence analysis of the 618 nucleotide product generated by the GAV PCR test has already demonstrated that LOV and GAV are variants in the same virus population of which individual isolates are genetically distinct (J.A. Cowley and P.J. Walker, unpublished data). It has also been possible to define the Australian viruses as a population that has evolved with a different lineage to that of YHV from Thailand. The accumulation of more comprehensive data from multiple domains of the genome and from YHV complex isolates obtained throughout the Asia-Pacific region will provide better understanding of these viruses, the origins of disease and the risk factors associated with farming practices. The principles illustrated here for the YHV complex are equally applicable to other RNA viruses and may well apply to DNA viruses infecting aquatic animals. Through the use of modern PCR and sequencing technology and the development of bioinformatics systems, the capability for rapid accumulation and analysis of nucleotide data is now a reality. If standard analytical procedures and appropriate security protocols can be agreed, there is obvious potential for such an approach to be a powerful tool in managing disease and in defining a more rational basis for controlling the movement of aquatic animal pathogens.
Acknowledgements Work reported in this paper was conducted as a co-operative project involving CSIRO Australia, and Mahidol University, NACA and the Aquatic Animal Health Research Institute in Thailand. This research has been supported by the Australian Centre for International Agricultural Research (ACIAR) and the National Centre for Genetic Engineering and Biotechnology (BIOTEC) in Thailand.
References Chamberlain, G.W. (1999). Sustainability of world shrimp farming. In: Global Trends: Fisheries Management. EK Pikitch, DD Huppert and MP Sissenwine, Eds American Fisheries Society Symposium 20, Bethesda, MD. Cowley, J.A., Dimmock, C.M., Spann, K.M. and Walker, P.J. Detection of Australian gill-associated virus (GAV) and lymphoid organ virus (LOV) of Penaeus monodon by RT-nested PCR. Diseases of Aquatic Organisms (submitted). Cowley, J.A., Dimmock, C.M., Wongteerasupaya, C., Boonsaeng, V., Panyim, S. and Walker, P.J. (1999). Yellow head virus from Thailand and gill-associated virus from Australia are closely related but distinct prawn viruses. Diseases of Aquatic Organisms 36, 153-157. Crawford, A.M. and Zelany, B. (1990). Evolution in Oryctes baculovirus: rate and types of genomic change. Virology 174, 294-298. Fearon, D.T. and Locksley, R.M. (1996). The instructive role of innate immunity in the aquired immune response. Science 272, 50-54. Gibbs, A.J., Calisher, C.H. and Garcia-Arenal, F. (Eds.) (1995). Molecular Basis of Viral Evolution. Cambridge University Press. Kwok, S., Kellogg, D.E., McKinney, N., Spasic, D., Goda, L., Levenson, C. and Sninsky, J.J. (1990). Effects of primer-template mismatches on the polymerase chain reaction: Human immunodeficiency virus type 1 model studies. Nucleic Acids Research 18, 999-1005. Lightner, D.V. (1996). A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA. 304 p. Lightner, D.V. and Redman, R.M. (1998). Shrimp diseases and current diagnostic methods. Aquaculture 16, 201-220. Limsuwan, C. (1991). Handbook for cultivation of black tiger prawns. Tansetakit Co. Ltd, Bangkok.
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Lo, C.-F., Hsu, H.-C., Tsai, M.-F., Ho, C.-H., Kou, G.-H. and Lightner, D.V. (1999). Specific genomic DNA fragment analysis of different geographical clinical samples of shrimp white spot syndrome virus. Diseases of Aquatic Organisms 35, 175-185. Morse, S.S. (1994). Towards an evolutionary biology of viruses. In The Evolutionary Biology of Viruses. (S.S. Morse Ed.). Pp. 1-28. Raven Press, New York. Parrish, C.R., Aquadro, C.F., Strassheim, M.L., Evermann, J.F., Sgro, J.-Y. and Mohammed, H.O. (1991). Rapid antigenic-type reeplacement and DNA sequence evolution of canine parvovirus. Journal of Virology 65, 6544-6552. Sommer, R. and Tautz, D. (1989). Minimal homology for PCR primers. Nucleic Acids Research 17, 6749. Spann, K.M., Vickers, J.E. and Lester, R.J.G. (1995). Lymphoid organ virus of Penaeus monodon from Australia. Diseases of Aquatic Organisms 23, 127-134. Spann, K.M., Cowley, J.A., Walker, P.J. and Lester, R.J.G. (1997). A yellow-head-like virus from Penaeus monodon cultured in Australia. Diseases of Aquatic Organisms 31, 169-179. Steinhauer, D.A., Domingo, E. and Holland, J.J. (1992). Lack of evidence fro proofreading mechanisms associated with RNA virus polymerase. Gene 122, 281-288. Steinhauer, D.A. and Holland, J.J. (1987). Rapid evolution of RNA viruses. Annual Review of Microbiology 41, 409-433. Tang, K.F-J. and Lightner, D.V. (1999). A yellow head virus gene probe: application to in situ hybridization and determination of its nucleotide sequence. Diseases of Aquatic Organisms 35, 165-173. Wongteerasupaya, C., Tongcheua, W., Boonsaeng, V., Panyim, S., Tassanakajon, A., Withyachumnarnkul, B. and Flegel, T.W. (1997). Detection of yellow-head virus of Penaeus monodon by RT-PCR amplification. Diseases of Aquatic Organisms 31, 181-186.
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The need for molecular tools in the study of mollusc pathogens P.M. Hine National Institute of Water and Atmospheric Research, PO Box 14-901, Wellington, New Zealand.
Introduction The OIE list of notifiable diseases of molluscs and the pathogens causing them comprises marteiliosis (Marteilia refringens, Marteilia sydneyi), bonamiosis (Bonamia ostreae, Bonamia sp.), mikrocytosis (Mikrocytos mackini, Mikrocytos roughleyi), haplosporidiosis (Haplosporidium nelsoni, Haplosporidium costale) and perkinsosis (Perkinsus marinus, Perkinsus olseni). This paper gives a brief overview of the areas in which molecular tools are needed to overcome problems associated with these diseases, and considers the needs of the Asian region.
Infection levels Moderate to heavy infections with Marteilia refringens, Marteilia sydneyi, both Bonamia spp., and both Haplosporidium spp. are relatively easy to detect by routine histology. Both Perkinsus spp. can be cultured using Ray’s Fluid Thioglycollate Medium (RTFM) (Bushek et al., 1994), allowing light infections to be amplified, and consequently detected. Mikrocytos spp. are much harder to detect. Mikrocytos roughleyi infects the haemocytes of Sydney rock oysters (Saccostrea commercialis) in eastern (Georges River) and western (Carnarvon, Albany) Australia, and ultrastructurally it has a single mitochondrion. Mikrocytos mackini infects connective tissue cells of Pacific oysters (Crassostrea gigas) off the coast of British Columbia, and ultrastructurally it lacks a mitochondrion. Currently it is thought that these two pathogens are not closely related, and M. roughleyi may be more closely related to Bonamia spp. Macroscopically both Mikrocytos spp. produce macroscopic pustular or abcess-like lesions in cases of heavy infection, and if occurring in the known range of these two pathogens, may allow presumptive diagnosis. However, microscopically both Mikrocytos spp. are only ~2 m in diameter, do not stain well, consequently they are difficult to detect in moderate to light infections (Hervio et al., 1996). Therefore probes are needed to detect light infections with all OIE listed pathogens except (Perkinsus spp.), and light to moderate infections with Mikrocytos spp.
The identification of species The inter-relationships of all the OIE listed notifiable pathogens are currently uncertain. Marteilia sydneyi was initially distinguished from the previously described Marteilia refringens, on the grounds that the latter possessed refringent granules, whereas the former did not. However, M. sydneyi does possess refringent granules (Roubal et al., 1989). Also it is unclear how the Marteilia sp. that caused a massive epizootic in calico scallops off the coast of Florida (Moyer et al., 1993) relates to described species. Although distinction on the basis of cleavage patterns during development seems to overcome these uncertainties, the distinction of Marteilia spp. is still being questioned (Bower et al., 1994). The two species of Bonamia have also not yet been clearly distinguished, although B. ostreae has dense forms that are seldom seen in Bonamia sp, and the latter has a vacuolated stage (Hine, 1991) that has not been reported from B. ostreae. The two species of Haplosporidium can be distinguished by spore size, and H. nelsoni can be distinguished from all other Haplosporidium spp. as it sporulates in epithelial cells of the digestive diverticulae, and the other species sporulate in connective tissue. Currently, H. costale cannot be distinguished reliably from other Haplosporidium spp. except H. nelsoni. Mikrocytos spp. do not resemble each other closely and M. roughleyi may be more closely related to Bonamia spp. (see above). Perkinsus olseni shows similarities to Perkinsus atlanticus
60
(Hamaguchi et al., 1998), but Perkinsus marinus also shows similarities to P. atlanticus (Robledo et al., 1997). Specific probes are currently available, or are being developed for, Marteilia refringens, Marteilia sydneyi, Bonamia spp., Haplosporidium nelsoni, and H. costale.
Life cycles Bonamia spp., Mikrocytos mackini, and Perkinsus spp., transmit directly from one host to another. Haplosporidium spp. and Marteilia spp. cannot be transmitted directly from one to another, and probably require an intermediate host (Roubal et al., 1989; Berthe et al., 1998). Probes currently developed or being developed for Haplosporidium spp. and Marteilia spp. will be used to identify the DNA of these pathogens in likely alternative hosts, such as filter feeding or detrivorous invertebrates.
The Asian region The OIE listed diseases of molluscs mainly infect bivalves in temperate regions. This is true of Bonamia spp. in temperate oysters (Ostrea, Tiostrea, Crassostrea), Mikrocytos spp. in temperate oysters (Crassostrea, Saccostrea), and Haplosporidium spp. of temperate oysters (Crassostrea). Marteilia refringens is also a parasite of temperate bivalves (Ostrea, Tiostrea, Crassostrea, Mytilus, Cerastoderma), and although Marteilia sydneyi occurs in the subtropics of southern Queensland, it is primarily a parasite of temperate oysters (Saccostrea). Marteilia refringens, Bonamia ostreae, Mikrocytos mackini, Haplosporidium costale and Perkinsus marinus have not been reported from the Asian region. Marteilia sydneyi, Bonamia sp., Mikrocytos roughleyi and Perkinsus olseni have been reported from Australia, and a Perkinsus sp. from clams (Tapes philippinarum) in Japan (Hamaguchi et al., 1998). The Japanese isolate had sequences intermediate between P. olseni in Australia, and P. atlanticus in Manila clams (Tapes philippinarum) from Spain. Perkinsus atlanticus from Ruditapes philippinarum and Ruditapes decussatus around Spain and Portugal, are closely related to P. olseni (Robledo et al., 1997). It may be therefore that P. olseni was moved from Asia in Manila clams to Europe, where it was described as P. atlanticus. If so, P. olseni/atlanticus may be widely distributed throughout Southeast Asia. Although Haplosporidium nelsoni has not been formally reported from Asia, a Haplosporidium sp., similar in size and pathology to H. nelsoni, occurs in Pacific oysters (Crassostrea gigas) in California and in Matsushima Bay, Japan, from which the Californian stocks derived (Friedman et al., 1991; Friedman, 1996). A sensitive and specific probe for H. nelsoni (Stokes and Burreson, 1995) labels the Californian and Japanese Haplosporidium, suggesting that it is also H. nelsoni, and that H. nelsoni was originally introduced into California in Japanese Pacific oysters. Therefore, some of the temperate OIE listed diseases occur in Australia and Japan, and it is likely that Haplosporidium nelsoni and Perkinsus olseni/atlanticus occur more widely in Asia than is currently realized. As bivalve health expertise becomes more widespread in Asia, other serious diseases of tropical, as well as temperate bivalves, are likely to emerge. One such pathogen may be a Haplosporidium sp. pathogenic in silverlip pearl oysters (Pinctada maxima) in northwestern Australia (Hine and Thorne, 1998). Also, an apparently infectious disease that has caused massive mortalities among akoya pearl oysters (Pinctada fucata) in Japan since 1994 (Miyazaki et al., 1998) may well prove to be a serious disease in Asia. Currently a parasite related to Marteilia, called Marteilioides chungmuensis, which parasitizes the ova of Pacific oysters (Crassostrea gigas) is having a serious impact on oyster production, and may also prove to be a problem in Asia. Once such problems have been identified, molecular tools will need to be developed to detect low infection levels, distinguish species and study life cycles, as for the currently listed diseases. The OIE protocols are designed to control spread of aquatic animal diseases, using a certification and reporting system that requires a national infrastructure, based in law, and a network of skilled and experienced aquatic animal health specialists, including technicians, inspectors and pathologists.
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Development of such systems occurs as industries to be serviced develop, and therefore are found where aquaculture industries are well established. Such established industries use hatchery production or natural spat settlement as their source of stock. Certification allows stock to be traded between farms, and the identification and establishment of disease free zones minimizes risk of disease spread. In developing bivalve farms, it is often necessary to initially acquire brood stock from the wild, or to move bivalves into an area to enhance natural spat settlement. Such movements are already taking place throughout Asia. This process must be undertaken with extreme care to minimize the risk of introducing disease onto the farm site. Studies on the parasites and diseases of wild bivalves in northwestern Australia have shown that the prevalence of potentially serious diseases in wild stocks may be extremely low (Table 1). Prevalences of ~0.1% are common. To detect such infection with 95% confidence, it is necessary to sample 2,994 animals, and even then a light infection may well be missed. Molecular tools are needed to detect such low levels of infection before stocks are moved.
References Berthe, F.C.J., Pernas, M., Zerabib, M., Haffner, P., Thébault, A. and Figueras, A.J. (1998). Experimental transmission of Marteilia refringens with special consideration of the life cycle. Diseases of Aquatic Organisms 34, 135-144. Bower, S.M., McGladdery, S.E. and Price, I.M. (1994). Synopsis of infectious diseases and parasites of commercially exploited shellfish. Annual Review of Fish Diseases 4, 1-199. Bushek, D., Ford, S.E. and Allen, S.K. (1994). Evaluation of methods using Ray’s fluid thioglycollate medium for diagnosis of Perkinsus marinus infections in the eastern oyster, Crassostrea virginica. Annual Review of Fish Diseases 4, 201-217. Friedman, C.S. (1996). Haplosporidian infections of the Pacific oyster, Crassostrea gigas (Thunberg), in California and Japan. Journal of Shellfish Research 15, 597-600. Friedman, C.S., Cloney, D.F., Manzer, D. and Hedrick, R.P. (1991). Haplosporidiosis of the Pacific oyster, Crassostrea gigas. Journal of Invertebrate Pathology 58, 367-372. Hamaguchi, M., Suzuki, N., Usuki, H. and Ishioka, H. (1998). Perkinsus protozoan infection in shortnecked clam Tapes (=Ruditapes) philippinarum in Japan. Fish Pathology 33, 473-480. Hervio, D., Bower, S.M. and Meyer, G.R. (1996). Detection, isolation, and experimental transmission of Mikrocytos mackini, a microcell parasite of Pacific oysters Crassostrea gigas (Thunberg). Journal of Invertebrate Pathology 67, 72-79. Hine, P.M. (1991). Ultrastructural observations on the annual infection pattern of Bonamia sp. in flat oysters Tiostrea chilensis. Diseases of Aquatic Organisms 11, 163-171. Hine, P.M. and Thorne, T. (1998). Haplosporidium sp. (Haplosporidia) in hatchery-reared pearl oysters, Pinctada maxima (Jameson, 1901), in north Western Australia. Journal of Invertebrate Pathology 71, 48-52. Miyazaki, T., Goto, K., Kobayashi, T. and Miyata, M. (1998). An emergent virus disease associated with mass mortalities in Japanese pearl oysters Pinctada fukata martensii. In: Proceedings of the VIIth International Colloquium on Invertebrate Pathology and Microbial Control. Sapporo, Japan, August 23-28th 1998, pp. 154-159. Moyer, M.A., Blake, N.J. and Arnold, W.S. (1993). An ascetosporan disease causing mass mortality in the Atlantic calico scallop Argopecten gibbus (Linnaeus, 1758). Journal of Shellfish Research 12, 305-310. Robledo, J.A.F., Wright, A.C., Coss, C.A., Vasta, G.R. and Goggin, C.L. (1997). Further studies of conserved genes from Perkinsus isolates. Journal of Shellfish Research 16, 342. Roubal, F.R., Masel, J. and Lester, R.J.G. (1989). Studies on Marteilia sydneyi, agent of QX disease in the Sydney rock oyster, Saccostrea commercialis, with implications for its life cycle. Australian J.ournal of Marine and Freshwater Research 40, 155-167. Stokes, N.A. and Burreson, E.M. (1995). A sensitive and specific DNA probe for the oyster pathogen Haplosporidium nelsoni. Journal of Eukaryote Microbiology 42, 350-357.
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Oyster Creek
n = 758
3 0 0 13 12 3 0
n = 22
0 0 0 0 0 0 0 0 0
Bivalve
Saccostrea commercialis
RLOs Marteilia sydneyi Perkinsus sp. Ancistrocomid ciliates Spenophyra-like ciliates Tylocephalum sp. Sporocysts
Saccostrea cuccullata
IVI RLOs Haplosporidium sp. Marteilia sp. Perkinsus sp. Ancistrocomid ciliates Nematopsis sp. Tylocephalum sp. Nematode larvae
1 1 121 1 0 3 1 0 4
769
0 0 0 0 0 13 1
50
ExmouthIslands
0 0 4 0 1 4 0 9 0
430
0 1 0 0 1 0 0
117
63
Dampier Archipelago
0 0 0 0 0 0 0 0 0
33
0 0 1 0 0 1 0
8
King Sound
-
0
-
0
DarwinBynoe
0.1% 0.1% 10.0% 0.1% 0.1% 0.6% 0.1% 0.7% 0.3%
1,254
0.3% 0.1% 0.1% 1.4% 1.4% 1.8% 0.1%
933
All areas
Table 1. Prevalence of parasites in Saccostrea spp. IVI = intranuclear virus-like inclusions. RLOs = Rickettsiales-like organisms.
Development and validation of DNA-based diagnostic techniques with particular reference to bivalve mollusc pathogens Franck C.J. Berthe IFREMER, Laboratoire de Génétique et Pathologie, Station de La Tremblade, BP 133, 17 390, Ronce les bains, France.
Introduction The International Aquatic Animal Health Code of the OIE (the World Organization for Animal Health) includes serious pathogens that have been causing important losses in the mollusc aquaculture industry throughout the world. This list also meets the ones established by the European Union regulation (Annex B of Directive 91/67/EC; Annex D of Directive 95/70/EC). In fact, one of the very few ways to reduce the impact of such pathogens on commercially exploited molluscs, is to establish effective programmes to prevent the transfer of infected stocks. Consequently, an area where molluscs are infected with any of these pathogens should not be allowed to export into another area free of this pathogen. Obviously, both country imports and abnormal mortality outbreaks in mollusc stocks should be examined for the presence of listed pathogens. This includes the detection of exotic diseases as well as emerging diseases. As a matter of fact, the effective control of diseases of bivalve molluscs requires an access to diagnostic tests that are rapid, reliable, accurate and sensitive. Techniques applicable to molluscan pathogens are limited and most investigations are based on histological and ultrastructural examinations. Arising from this, the development of molecular diagnostic tools will probably be one of the most important areas for research in the near future.
Potential detrimental consequences of transfers of molluscs and the need of accurate diagnostic tools There are very few ways to limit the detrimental effect of mollusc pathogens. Molluscs are usually reared in the open sea and this strongly limits the use of chemotherapy, because of the quantity required and thre consequent impact on the environment. On the other hand, vaccination is also limited, due to the fact that molluscs do not produce antibodies. Consequently, one of the few methods of controlling mollusc diseases is likely to be the establishment of effective programmes to prevent the transfer of infected stocks. This is of utmost importance if we consider that the introduction of molluscs from other geographic areas for aquaculture has frequently resulted in the introduction of devastating pathogens in native stocks. The risk associated with transfers of molluscs particularly serious when they occur over long distances or overseas. Unforseen dramatic consequences, due to pathogens described as being of no concern, may result from exposure of a naive population. For example, in the early 1970s, the Portuguese oyster, Crassostrea angulata, was dramatically affected by an iridovirus (Marteil, 1976). It has been speculated that uncontrolled transfer of Crassostrea gigas introduced this iridovirus to C. angulata, which was highly susceptible. Crassostrea angulata and C. gigas are two taxa of the same species (Boudry et al., 1998). Another example of particular interest is Bonamia ostreae which, in 1979, dramatically affected the flat oyster (O. edulis) industry in France (Pichot et al., 1979). This pathogen rapidly spread to almost all oyster farming areas in Europe, including Spain, Netherlands, Ireland and United Kingdom (Van Banning, 1982; Banister and Key 1982; Polanco et al., 1984; McArdle et al., 1991). A microcell disease similar to bonamiosis was also described in California in the 1960s and was known to occur in several populations of flat oysters from the western coast of North America (Elston et al., 1986). Bonamia ostreae was later identified as the causative agent of this disease on the basis of host susceptibility and ultrastructural characteristics. Moreover, the use of monoclonal antibodies
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demonstrated no antigenic differences between B. ostreae isolates originating from Europe and the USA (Mialhe et al., 1988). More recently, bonamiosis was recorded from the east coast of the USA, New Meadows River, Quahog Bay and Damariscotta River (Barber and Davis, 1994; Friedman and Perkins, 1994). These results and historical commercial records lead to the hypothesis that the Californian microcell disease was actually bonamiosis and that bonamiosis spread from California to Europe because of transfers of B. ostreae infected flat oysters (Elston et al., 1986; Cigarria and Elston, 1997). Decision makers responsible for supervising translocations of molluscs deal with a high risk situation. Risk analysis prior to transfers should help to minimise this risk. However, a serious limiting factor is a lack of scientific information on even basic biology of mollusc pathogens (Hine, 1996). Surveillance for mollusc pathogens is routinely performed by histology. This technique is time consuming and dependant of visula observation. In 1998, the Community Reference Laboratory proposed a ring test for the detection of two parasites (Bonamia ostreae and Marteilia refringens) by means of histology which is currently the standard method. The goal of this profeciency test was to establish that examination of a given sample lead to the same conclusions in any of the eight participating laboratories. The ring test was based on an itinerant collection of stained histological sections of Ostrea edulis. Statistical analysis of the results was based on the test of symetry and kappa coefficient. Significant discordance among the results obtained by participating laboratories was evident from the study. This clearly illustrated the need for training in histological diagnosis, particularly for exotic diseases, and the need for epidemiological surveillance programmes in order to prevent the transfer of diseases. However, using histological methods many pathogens are difficult to detect when present in low numbers. Recent efforts to overcome these problems have led to the development of immunoassay techniques and nucleic acid-based diagnostic methods. Serological methods for diagnostic purposes obviously cannot be applied to molluscs as they do not produce antibodies. Molecular probes, such as monoclonal antibodies or nucleic acid probes, may be used for direct detection of pathogenic agents. These techniques are expected to find increasing use in routine disease monitoring programs in aquaculture, in field epidemiology and in efforts to prevent the international spread of pathogens. Therefore, it is extremely important to develop, validate and standardize this type of diagnostic technique for major mollusc diseases and pathogens.
Few prerequisites to the development of molecular diagnostic methods When mortalities occur, various presumptive diagnostic methods can be used in addition to histology. This led us to consider three different levels of investigation which are: i) diagnostic procedures (standard methods for the assessment of a disease free status in a zone); ii) detection procedures (presumptive methods for the quick detection of a suspected pathogen); and iii) confirmatory procedures (methods for the specific identification of an encountered pathogen). Obviously, the required quality criteria of the selected method depend on the level of investigation for which it is to be used. For example, detection procedures require techniques that are easy to perform (e.g. smears or tissue imprints) or sensitive techniques that usually are based on an amplification step (e.g. culture of the pathogen or polymerase chain reaction- PCR). Choice of technique may then be based on the time required to obtain a result and this can range from few hours to few days. In the case of diagnostic procedures, specificity of the selected methods is obviously the most important criterion. Confirmatory procedures currently in use are ultrastructural observations by transmission electron microscopy. One should say that in the near future, with the increasing use of molecular technics, diagnostic procedures will increasingly be confirmatory.
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Increased sensitivity is often used as an argument for the use of new diagnostic methods. However, we should stress here that sampling strategy is of a central importance to ensure detection of a pathogen whatever detection method is used. The timing and frequency of sampling should be determined by the cycle of infection and pre-patent period. Also, the intensity of infection may increase following spawning due to loss of host condition and therefore molluscs should be sampled post-spawning. The recommanded sample size for each sampling site is 150 or a sufficient number to ensure detection at a 95% confidence level of pathogens at a prevalence of 2%. However, if molluscs are to be moved from natural beds onto a farm site or between natural beds in different zones, large numbers of molluscs must be sampled because of low parasite prevalence. In Western Australia, Perkinsus sp. occurs in isolated beds sometimes at very low prevalences (Hine, 1996). However, the probability of detecting an infection may be increased by holding the molluscs in quarantine for a long period, subjecting them to stress and examination of cohabitant species of molluscs that are highly susceptible to the infection. The use of PCR could help in such situations. However, widely recognized limitations of PCR methods include the false positive results (due to inhibiting substances in marine organisms, lack of target organs, pre-patent periods, etc) and false positive results (crossreaction with closely related organisms, laboratory contamination of samples, etc). This leads us to consider specificity as one of the main imput of molecular diagnostic methods currently developped. The problem of specificity in pathogen diagnosis is clearly illustrated by difficulties in dfferentiating Marteilia species. In Europe, Marteilia refringens has been observed in Ostrea edulis (Grizel et al., 1974) and also in Mytilus edulis and M. galloprovincialis (Tigé and Rabouin, 1976; Claver-Derqui, 1990; Villalba et al., 1993). However, Marteilia maurini has also been described in both Mytilus edulis and M. galloprovincialis from France (Comps et al., 1982; Auffret and Poder, 1985). In spite of numerous papers published on the genus Marteilia, the question of taxonomic relationships of these species remains unresolved. Differential diagnosis of M. refringens and M. maurini was based on ultrastructural characteristics and host specificity (Grizel et al., 1974; Comps et al., 1982) but host specificity was discarded when M. refringens was described in Mytilus galloprovincialis. Indeed, the species parasitizing mussels may not be truely different from M. refringens. Recently, the small subunit of the rRNA gene was sequenced and sequences confirmed that both Ostrea edulis and Mytilus edulis are hosts of M. refringens (Berthe et al., 1999). Current work is directed towards establishing the existing species among the genus Marteilia. Clarification of taxonomy of the targeted pathogens is of a central importance but is often underestimated as a problem in diagnosis. Perkinsus atlanticus is another well documented example of data gap in the field of taxonomy prior to the development of molecular tools. This organism is known to occur in both Europe (Azevedo, 1989) and Asia (Hamaguchi et al., 1998). In fact, more than 50 species of molluscs may harbour Perkinsus species from temperate to tropical waters of the Atlantic and Pacific oceans and Mediterranean Sea, apparently without harmful effect. Nucleotide sequence analysis of the internal transcribed spacers (ITS) of the ribosomal gene cluster (rDNA) has indicated that the Australian organism P. olseni is probably conspecific to Perkinsus atlanticus (Goggin, 1994). Taking this into account, the geographical distribution of the mollusc pathogen P. osleni could be wider than currently accepted. This should be urgently investigated because of the obvious consequences of such considerations. In summary, we would like to pinpoint the need of adequacy of the methods to be developed and validated, as well as the absolute need of a clear taxonomy of pathogens under consideration.
Development of DNA-based methods for the detection of mollusc pathogens: the example of Marteilia refringens In a preliminary study, the 18S gene of M. refringens was sequenced (Berthe et al., 1999). Apart of clarifying the controversial taxonomy of Marteilia refringens and its relatives, this gene is interesting from a detection point of view because it is present in a high copy number in the genome, and so provides increased sensitivity of detection when targeted. Furthermore it contains conserved and nonconserved regions interspaced in the sequence which allows the design of universal and specific PCR primers.
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After alignement of the Marteilia refringens rDNA SSU sequence with various eukaryotic organisms, PCR primers were designed (Le Roux et al., 1999). Specific primers were used to amplify DNA extracted from purified Marteilia refringens and infected hosts. The detection was also possible from paraffin embedded tissues which is a frequent source of biological material in the field of mollusc pathology. The specificity of amplified fragments was confirmed by Southern blotting with an oligoprobe. Furthermore, the sensitivity of the detection was increased by this method. In brief, the designed primers allow rapid and specific screening of numerous samples from different sources for the presence of Marteilia refringens with a good sensitivity. Universal primers provide an internal control for amplification experiments. Working with marine organisms, such an internal control of the PCR reaction is of a central importance as the reaction efficiency depends on various parameters including the presence of inhibitory factors and the quality and quantity of targeted DNA. In the present study, universal primers were designed and successfully used to amplify DNA from both Marteilia refringens and its hosts. These primers should be included in further use of PCR for M. refringens detection. For in situ hybridization, four probes were tested by Northern blotting for the specific detection of 18S RNA isolated from Marteilia refringens and other eukaryotic cells. The most specific probe was used successfully to detect Marteilia refringens by in situ hybridization. The selected probe produced consistent strong reactions when used for Marteilia refringens-infected Ostrea edulis and Mytilus edulis, as well as for Marteilia maurini-infected Mytilus galloprovincialis. A similar result was obtained with Marteilia sydneyi in Saccostrea commercialis. However, no cross-reaction was noted when the probe was tested against Marteilioides chungmuensis in Crassostrea gigas. It was concluded that the sequence of Smart 2 is shared partially, if not completely, by Marteilia spp. Similarly, specific primers designed from the 18S sequence of Marteilia refringens led to the amplification of specific fragment from European Marteilia-infected bivalves. No amplification was obtained when M. sydneyi DNA was targeted. Although the taxonomic relationships among the European species are not clearly established, PCR could be used to specificity discriminate Marteilia refringens from M. sydneyi. Repeatability and reproduceability were successfully tested. A study was commenced to validatethe in situ hybridization as a confirmatory method. Oysters originating from three different European zones (highly infected originating from Marennes-Oléron, France (n = 200); medium infection from Brittany, France (n = 200); and free of marteiliosis (n = 200) originationg from Lake Grevelingen, Netherlands) were processed by this method and compared with classical histology which is considered to be the standard method. Statistical analysis indicated a strong validation of the in situ hybridization method for the detection of Marteilia refringens. However, most of these resulsts were obtained on laboratory material stored in good condition, small size samples and were conducted by trained staff. Further work is underway in our laboratory to further validate of these tools.
Potential use of these new diagnostic tools The results presented here clearly demonstrate the growing interest in molecular methods. However, in the case of molluscs, histology provides a large amount of information and should be used initially, before and beside any other type of examination. It is particularly important because macroscopic examination usually gives no pathognomonic signs. Also, mortality may be due to several pathogens, or loss of condition following spawning, and this can only be determined by histology. In the case of Marteilia refringens, it is possible to recommand selected methods for the three different levels of investigation described above. The detection of Marteilia refringens by in situ hybridization could be used in addition to classical histological examination as a confirmatory method at a genus level. Histology and in situ hybridization can thus be used as a two step diagnostic procedure, and
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could become a standard method for the assessment of a disease free status in a zone. Detection procedures that require presumptive methods for rapid detection of a suspected pathogen could be cunducted by using digestive gland imprints. It is very important to keep in mind the multiple advantages of such a method (which can be applied in the course of sample preparation for histology) as it is cheap and provides an immediate answer. PCR tests, because of their specificity, could be proposed for the specific identification of encountered pathogens as a confirmatory procedure. However, standardization of protocols, including negative and positive controls, is required. Compared to transmission electron microscopy (the currently used confirmatory procedure), PCR provides a quick and specific answer. In the near future, the number and diversity of available methods should increase. In the case of Marteilia spp., oligoprobes targetting the ITS region have been developed and should be used for diagnostic purposes at a species level. Similarly, the sequencing of this region of the ribosomal gene cluster is currently demonstrating the possible existance of different strains within the species refringens. Further development in the knowledge of these parasites could lead to an increasing number of molecular methods at different levels of specificity (i.e. genus, species and strain). These could include techniques such as restriction fragment lengh polymorphism (RFLP) and reverse blot PCR. At a national and regional level, reference laboratories will provide sequences of primers and oligoprobes to be used. The role of these laboratories in the validation of molecular reagents as diagnostic tools is obvious. Furthermore, these laboratories will have a growing responsibility in providing standardized protocols including positive and negative controls. Proficiency evaluations such as ring tests should also be organized for these diagnostic procedures in order to avoid misinterpretation of results. It should be said here that some of the DNA-based methods presented in this paper were aiming the study of life-cycle of Marteilia refringens which may include intermediate hosts or free-living stages (Berthe et al., 1998). With similar goals, a number of research laboratories are already engaged in developing DNA-based diagnostic techniques for mollusc pathogens. Therefore, several new diagnostic tools for mollusc diseases should be available in the future. Another potential use of molecular diagnostic tools is for detection of Haplosporidium nelsoni, one of the causative agent of haplosporidiosis - a disease of eastern oyster (Crassostrea virginica). A parasite morphologically similar to H. nelsoni was described in the Pacific oyster (Crassostrea gigas) on the west coast of the USA (Friedman et al., 1991). This parasite was identified as H. nelsoni by the use a specific DNA probe (Stokes and Burreson, 1995). Furthermore, some of the C. gigas stocks were traced back to Japan where the examination of native C. gigas, demonstrated infection by a Haplosporidium sp. indistinguishable from H. nelsoni described in C. gigas from the USA (Friedman, 1996). A Minchinia sp. (Haplosporidium-like organism) has also been known to occur in C. gigas in Korea since the mid-1970s (Kern, 1976). In France, several authors have reported the occurance Haplosporidium spp. in several species of molluscs (Bonami et al., 1985; Chagot et al., 1987; Comps and Pichot, 1991). There is some confusion in the taxonomic relationship of the two pathogens H. nelsoni and H. costale. This should be investigated in the near future. This example illustrates an unforseen consequence of the use of DNA probes and reveals how taxonomy is an underestimated key point in mollusc pathology.
Acknowledgements Work reported in this paper was partly conducted as a cooperative project - MARS - funded by the FAIR programme, and by the Community Reference Laboratory for mollusc diseases (Ifremer La Tremblade) with the financial assistance of the EU DG VI.
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References Azevedo C. (1989). Fine structure of Perkinsus atlanticus n. sp. (Apicomplexa, Perkinsea) parasite of clams, Ruditapes decussatus, from Portugal. J. Parasitol., 75, 627-635. Auffret M. and Poder M. (1985). Recherches sur Marteilia maurini, parasite de Mytilus edulis sur les côtes de Bretagne nord. Rev. Trav. Inst. Pêches marit. 47 :105-109. Bannister, C. and Key D. (1982). Bonamia a new threat to the native oyster fishery. Fish. Nat. MAFF Direct . Fish. Res. 71,7. Barber, B.J. and Davis C. (1994). Prevalence of Bonamia ostreae in Ostrea edulis populations in Maine. J. Shellfish Res. 13, 298. Berthe, F.C.J., Pernas, M., Zerabib, M., Haffner, P., Thébault, A. and Figueras, A.J. (1998). Experimental transmission of Marteilia refringens with special consideration of the life cycle. Diseases of Aquatic Organisms 34, 135-144. Berthe, F.C.J., Le Roux F., Peyretaillade E., Peyret P., Rodriguez D., Gouy M. and Vivarès C.P. (1999). The existence of the phylum Paramyxea Desportes and Perkins, 1990 is validated by the phylogenetic analysis of the Marteilia refringens small subunit ribosomal RNA. Submitted in Molecular and Biochemical Parasitology. Bonami, J.R., C.P. Vivares and M. Brehelin. (1985). Étude d'une nouvelle haplosporidie parasite de l'huître plate Ostrea edulis L.: morphologie et cytologie de différents stades. Protistologica 21: 161-173. Boudry, P., Heurtebise S., Collet B., Cornette F and A. Gérard (1998). Differentiation between populations of the Portuguese oyster Crassostrea angulata (Lamark) and the Pacific oyster, Crassostrea gigas (Thunberg), revealed by mtDNA RFLP analysis. J. Exp. Mar. Biol. Ecol. 226, 279-291. Chagot, D., Bachère, E., Ruano, F., Comps, M. and Grizel, H. (1987). Ultrastructural study of sporulated instars of a haplosporidian parasitizing the clam Ruditapes decussatus. Aquaculture 67: 262-263. Cigarria, J. and Elston, R. (1997). Independent introduction of Bonamia ostreae, a parasite of Ostrea edulis, to Spain. Dis. aquat. Org. 29, 157-158. Claver-Derqui A. (1990). Datos sobre Marteiliosis en la provincia de Cadiz. Acta III Congreso Nac. Acuicult. 885. Comps M., Grizel H. and Papayanni Y. (1982) Infection parasitaire causée par Marteilia maurini sp. nov. chez la moule Mytilus galloprovincialis. Cons. Int. Explor. Mer, F:1-3. Comps, M. and Y. Pichot. (1991). Fine spore structure of a haplosporidan parasitizing Crassostrea gigas: taxonomic implications. Diseases of Aquatic Organisms 11: 73-77. Elston, R.A., Farley, C.A., and Kent, M.L. (1986). Occurrence and significance of bonamiasis in European flat oyster Ostrea edulis in North Amerinca. Dis. aquat. Org. 2, 49-54. Friedman C.S. (1996). Haplosporidian infections of the Pacific oyster, Crassostrea gigas (Thunberg), in California and Japan. Journal of Shellfish Research 15 : 597-600. Friedman C.S., Cloney D.F., Manzer D. and Hedrick R.P. (1991). Haplosporidiosis of the Pacific oyster, Crassostrea gigas. J. Invertebr. Pathol., 58, 367-372. Friedman, C.S., and Perkins, F.O. (1994). Range extension of Bonamia ostreae to Maine, U.S.A. J. Invertebr. Pathol. 64, 179-181. Grizel H., Comps M., Bonami J.R., Cousserans F., Duthoit J.L., and Le Pennec M.A. (1974). Recherche sur l'agent de la maladie de la glande digestive de Ostrea edulis Linne. Sci. Pêche. Bull. Inst. Pêches marit. 240:7-29. Goggin, C.L. (1994). Variation in the two internal transcribed spacers and 5.8S ribosomal RNA from five isolates of the marine parasite Perkinsus (Protista, Apicomplexa). Molecular and Biochemical Parasitology 65: 179-182. Hamaguchi, M., Suzuki, N., Usuki, H. and Ishioka, H. (1998). Perkinsus protozoan infection in shortnecked clam Tapes (=Ruditapes) philippinarum in Japan. Fish Pathology 33, 473-480. Hine P.M. (1996). Southern hemisphere mollusc diseases and an overview of associated risk assessment problems. Rev. Sci. Tech. Off. Int. Epiz., 15(2) 563-577.
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Kern F.G. (1976). Sporulation of Minchinia sp. (Haplosporida, Haplosporidiidae) in the Pacific oyster Crassostrea gigas (Thunberg) from the Republic of Korea. J. Protozool., 23, 498-500. Le Roux, F., Audemard, C., Barnaud, A., and Berthe, F. (1999). DNA probes as potential tools for the detection of Marteilia refringens. Submitted in Mol. Mar. Biol. Biotechnol Marteil L. (1976). La conchyliculture française. 2. Biologie de l’huître et de la moule. Rev. Trav. Inst. Pêches marit., 40, 149-345. Mc Ardle, J.F., McKiernan, F., Foley, H., and Jones D.H. (1991). The current status of Bonamia disease in Ireland. Aquaculture 93, 273-278. Mialhe, E., Boulo, V., Elston, R., Hill, B., Hine, M., Montes, J., Van Banning, P., and Grizel, H. (1988). Serological analysis of Bonamia in Ostrea edulis and Tiostrea lutaria using polyclonal and monoclonal antibodies. Aquat. Living Resour. 1, 67-69. Pichot, Y., Comps, M., Tigé, G., Grizel, H., and Rabouin M.A. (1979). Recherche sur Bonamia ostreae gen. n., sp. n., parasite nouveau de l’huître plate Ostrea edulis L. Revue Trav. Inst. scient. tech. Pêch. marit. 43, 131-140. Stokes, N. A., and Burreson, E. M. (1995). A sensitive and specific DNA probe for the oyster pathogen Haplosporidium nelsoni. J. Euk. Microbiol., 42 (4) : 350-357. Tigé G., and Rabouin, M.A. (1976). Etude d'un lot de moules transférées dans un centre touché par l'épizootie affectant l'huitre plate. Cons. Int. Explor. Mer, K:1-10. Van Banning, P. (1982). The life cycle of the oyster pathogen Bonamia ostreae with a presumtive phase in the ovarian tissue of the European flat oyster, Ostrea edulis. Aquaculture 84, 189192. Villalba A., Mourelle S. G., López M. C., Carballal M. J., and Azevedo C. (1993). Marteiliasis effecting cultured mussels Mytilus galloprovincialis of Galicia (NW. Spain). I. Etiology, phases of the infection, and temporal and spatial variability in prevalence. Dis. aquat. Org. 16:61-72.
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Molecular diagnostics for the oyster pathogens Haplosporidium nelsoni (MSX Disease) and Perkinsus marinus (Dermo Disease) in Chesapeake Bay, Virginia, USA Eugene M. Burreson Virginia Institute of Marine Science, College of William and Mary, Gloucester Point, Virginia 23062, USA
__________________________________________________________ Background on Haplosporidium nelsoni (MSX) and Perkinsus marinus (Dermo) Haplosporidium nelsoni The protozoan oyster pathogen Haplosporidium nelsoni, commonly called MSX, has been responsible for periodic, large–scale oyster mortality in estuaries of the middle Atlantic coast of the United States. The pathogen appeared in Delaware Bay in 1957 and killed 90–95% of all oysters on planted grounds within three years. It appeared in the lower Chesapeake Bay in 1959 and was responsible for the death of about 95% of the oysters in that region by 1962 (Haskin and Andrews, 1988). There have been additional epizootics during periods of extreme drought, especially 1980–81 and 1987–88. The origin of H. nelsoni is uncertain, but there has been considerable speculation that the parasite was introduced with exotic oysters (Andrews, 1984). Although the annual distribution and abundance of H. nelsoni vary with salinity, its general prevalence and intensity have not decreased since the original epizootics and the parasite has now spread along most of the Atlantic coast of the United States. The widespread distribution and high virulence of H. nelsoni have greatly reduced traditional onbottom culture and have limited the development of off-bottom oyster aquaculture in high salinity areas. Early diagnosis of H. nelsoni infection is an essential management tool (Ford and Haskin, 1988). Thus, rapid, sensitive diagnostic assays may be critical in avoiding losses to the pathogen. Paraffin-based histological examination, the most commonly used diagnostic technique for H. nelsoni, involves finding parasite cells in one or two 5 µm thick sections through an oyster’s visceral mass, gills and mantle. There is a period of several weeks between the time early-summer H. nelsoni infections are heavy enough to be detected histologically and when mortality begins. Therefore, the earlier infections can be detected, the more time is available for oyster growers to decide whether to harvest to limit mortality, or whether to move oysters to low salinity to eliminate the parasite (Ford, 1985). Infections of H. nelsoni acquired in late summer or fall remain nonlethal until spring; early detection would also allow harvest to avoid losses from these late season infections. Perkinsus marinus Perkinsus marinus has been a significant cause of mortality of the eastern oyster Crassostrea virginica along the east coast of the United States since the 1950s (Andrews, 1988; Burreson and Ragone Calvo, 1996). The origin of P. marinus is obscure, but it probably has always been an associate of oysters. Along the east coast prior to the late 1980s, P. marinus was restricted to high salinity portions of coastal bays and estuaries south of Delaware Bay, although it apparently was absent from the seaside bays of the eastern shore of Virginia and Maryland (Andrews, 1988). In the Chesapeake Bay, P. marinus was prevalent in the lower bay, but was restricted to the mouths of the major tributaries in Virginia and southern Maryland. There were a few localized concentrations of P. marinus in Maryland, primarily in Fishing Bay and Eastern Bay. The pathogen was observed sporadically in Delaware Bay in the mid-1950s, as a result of importing infected oysters from Chesapeake Bay, but it never caused significant mortality in oysters and appeared to die out as importations stopped in the late 1950s (Ford, 1992). North of Delaware Bay the parasite was absent or at least undetectable. In
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endemic areas P. marinus has always been responsible for some oyster mortality, but it did not significantly affect harvest most years. The four consecutive drought years in the middle Atlantic region from 1985 through 1988 caused elevated salinities throughout the Chesapeake Bay and allowed P. marinus to spread naturally into areas where oysters were abundant and highly susceptible. Unusually warm winters during the period also contributed to the spread by allowing higher overwintering survival of the pathogen and movement of infected oysters into areas where P. marinus was historically absent also contributed to the problem. Although drought conditions have abated and rainfall patterns have returned to more or less typical conditions, with wet winters and springs, P. marinus continues to persist tenaciously in most areas of the Chesapeake Bay. Unlike H. nelsoni, which is very susceptible to salinity below 10 ppt, P. marinus can survive in oysters even in areas where salinity drops below 3 ppt for periods of weeks. Thus, even though P. marinus is not causing oyster mortality in many areas, it is present on all beds and if drought conditions return and salinity becomes favorable for development, it will be able to quickly multiply and cause mortality. Investigators generally rely on Ray’s fluid thioglycollate medium (FTM) culture of Perkinsus marinus cells from oyster tissue (rectum, gill, and mantle) for the diagnosis of infected oysters (Ray, 1952). An oyster hemolymph FTM assay was developed to allow monitoring of P. marinus infections without sacrificing the host and to improve quantitation of systemic infections (Gauthier and Fisher, 1990). Using these FTM methods, quantitation relies on accurate counting and the use of a subjective scale developed by Mackin (1962) and modified by Craig et al. (1989). Diagnosis based on FTM methods makes several assumptions. It is assumed that all life stages of P. marinus found in the host are retrieved and that the number of parasites remains constant during incubation in FTM. Furthermore, it is assumed that the distribution of P. marinus in the assayed tissues is representative of the distribution of the parasite throughout the oyster. To overcome this latter assumption, a total body burden assay was developed (Bushek et al., 1994) which employs a procedure using sodium hydroxide to digest oyster tissues after incubation in FTM (Choi et al., 1989). The body burden assay is quantitative and allows enumeration of the total number of parasites in whole oyster homogenates. FTM culture diagnostic methods are relatively simple to perform; however, insensitivity of the assays often causes very light infections to be overlooked.
Molecular diagnostics There has been increasing interest in developing DNA probes and/or PCR primers as tools for detection of disease agents in aquatic animals. These detection methods have been shown to be more sensitive and specific than previously developed procedures, such as histological examination or immunoassays. DNA probe/PCR technology is ideal for specifically recognizing target DNA regardless of the life history stage present, for identifying cryptic life cycle stages in any potential host organism and for detecting pathogens early in the infection cycle. The ability to rapidly and accurately detect parasites of oysters has broad implications for both research and industry. As rapid diagnostic methods become more sensitive, early detection of disease agents permits the management of oysters or other cultured species in a manner that is more responsive to patterns of naturally occurring diseases. However, caution must be exercised in the interpretation of molecular diagnostic results, especially from the polymerase chain reaction (PCR). The extreme sensitivity of this technique may allow amplification of DNA from non-viable or non-pathogenic organisms. A PCR-positive result does not necessarily mean that viable, pathogenic organisms are present or that mortality will eventually occur. More research and experience is needed to fully understand the meaning of PCR-positive results for aquaculture or management. Nonetheless, molecular diagnostic techniques hold great promise for early diagnosis of disease outbreaks in aquaculture and natural populations.
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Development of molecular diagnostics for Haplosporidium nelsoni A sensitive and specific DNA probe and PCR primers have been developed for Haplosporidium nelsoni at the Virginia Institute of Marine Science (Stokes and Burreson, 1995; Stokes et al., 1995). One difficulty in developing molecular diagnostics for molluscan pathogens in the phylum Haplosporidia is that in vitro culture of these organisms has not been accomplished. Thus it is very difficult to obtain purified organisms for DNA extraction. For our work, plasmodia were concentrated and purified from hemolymph of infected oysters by panning in petri dishes. This technique involves settling of hemocytes, which adhere to the petri dish, allowing the H. nelsoni plasmodia to be decanted. Repeated panning results in an enrichment of plasmodia, although not all hemocytes are excluded. DNA was extracted from the H. nelsoni plasmodia and the small subunit (SSU) rDNA was amplified from the H. nelsoni genomic DNA using the universal primers for eukaryotic 16S-like rDNA. PCR amplification products were cloned into pCRII and INV F' using the TA Cloning system (Invitrogen) and subclones were cloned into pUC8 and DH5 . Clones with rDNA inserts were sequenced via the dideoxy chain termination method using the Sequenase kit; the collections of putative H. nelsoni sequence data were assembled into a composite gene sequence using the Gene Jockey software package (GenBank acession number U19538). The variable regions of the SSU rDNA sequence were examined for a section that could be used as a species–specific probe. A putative H. nelsoni–specific oligonucleotide probe, designated MSX1347 (5'– ATGTGTTGGTGACGCTAACCG–3'), was chosen from sequence alignment with the SSU rDNA of Minchinia teredinis, C. virginica, Perkinsus marinus and various protists from GenBank. The probe was synthesized with Figure 1. In situ hybridization with the incorporation of digoxigenin at the 5' end. In situ H. nelsoni DNA probe on histological hybridizations of this H. nelsoni probe react strongly with section of an oyster infected with H. H. nelsoni plasmodia (Figure 1) and immature spores, but nelsoni. weakly with mature spores. The probe does not react with oyster tissue, Perkinsus marinus or any of the other haplosporidians present in the middle Atlantic region, including Haplosporidium costale, H. louisiana, or Minchinia teredinis. The variable regions of the H. nelsoni SSU rRNA gene were examined for areas which appeared to be species specific and would be appropriate for use as PCR primers. Following identification of suitable priming regions, the putative oligonucleotide sequences were sent as queries to the BLAST electronic mail server (
[email protected]) to determine whether the primers would anneal to non-target genes. Two oligonucleotides, designated MSX-A (5’-GCATTAGGTTTCAGACC-3’) and MSX-B (5’-ATGTGTTGGTGACGCTA-ACCG-3’), were selected and commercially synthesized. The sequence of the MSX-B primer is the same as the H. nelsoni DNA probe MSX1347 described above. These primers amplify a 564 base pair region of the SSU rRNA gene. Primer sensitivity was tested in ten-fold serial dilutions from 1 ng to 1 fg of cloned H. nelsoni SSU rDNA. A PCR product was easily detectable by agarose gel electrophoresis from a single amplification of 100 fg of cloned H. nelsoni SSU rDNA. Specificity was tested in PCR reactions using genomic DNA from oysters infected and not infected with H. nelsoni, and cloned SSU rDNA of three haplosporidians, M. teredinis, H. costale and H. louisiana. The PCR primers amplified H. nelsoni SSU rDNA from genomic DNA of an infected oyster, but no product was obtained from uninfected oysters or from the other haplosporidians tested.
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Validation of PCR diagnosis for Haplosporidium nelsoni During May 1995, 400 oysters uninfected with H. nelsoni were imported to the lower York River estuary, Virginia. These oysters were examined monthly through December for H. nelsoni using three diagnostic techniques—paraffin histology, the routine diagnostic technique (i.e. gold standard), PCR of oyster hemolymph and PCR of gill and mantle tissue. A sample of 25 oysters was collected each month. The oyster shells were notched and 0.5 ml of hemolymph was withdrawn from the adductor muscle sinus with a needle and syringe for DNA extraction. Each oyster was then shucked and a 0.25 g piece of gill and mantle tissue was removed for DNA extraction. The remaining oyster was preserved in Davidson’s AFA fixative for histological analysis. Results of the analyses are shown in Figure 2. PCR of hemolymph detected H. nelsoni infections two months earlier (May) than either PCR of tissue or histology (July). PCR of both hemolymph and tissue always detected a higher prevalence of infection that histology, although during November and December, when all infections are typically systemic, there was little difference in prevalence among the three techniques. Overall, infections were detected in 129 oysters by one of the three methods. Fifty five infections were detected by one of the two PCR techniques that were negative by histological examination and two infections were detected by histology that were negative by PCR. These results demonstrate that PCR diagnosis is much more sensitive than histology and allowed infections to be detected much earlier in the natural infection cycle (For another interpretation from a different perspective see the paper by Dan Fegan in this report). This early detection can be very important for mitigating mortality from the disease. The decline in prevalence during August is the result of H. nelsoni-induced mortality that typically occurs during late July as infections intensify.
Development of molecular diagnostics for Perkinsus marinus One of the problems with normal PCR is that it is not quantitative and there is much interest in quantification of infections and in detection and quantification of P. marinus in natural water samples. A semi-quantitative PCR assay for P. marinus was developed by Marsh et al. (1995), so our interest was in developing a quantitative analysis (Yarnall et al., 1999) called quantitative competitive PCR (QCPCR).
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Figure 2. Prevalence ofHaplosporidium nelsoni in oysters through a natural infection cycle in Chesaspeake Bay, Virginia as determined by three different diagnostic techniques.
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Limitations of precise quantitation using PCR techniques stem from the lack of consistent initial exponential increase in product during the amplification reaction. Replicate samples can be subjected to variations in reaction conditions such as inhibitor concentrations, polymerase integrity, and even thermal cycler block positions. These differences can result in variation in amplification efficiencies that may obscure differences in the amounts of DNA or RNA that are being measured and thus preclude accurate quantitation. In competitive PCR, the standard and target DNA are present in the same reaction tube. The target DNA and internal standard DNA compete for the same DNA polymerase and deoxynucleotide triphosphates. In addition, the two DNA templates are equally affected by tube to tube variations in the PCR conditions including inhibitor effects. Furthermore, competitive PCR overcomes the need to perform quantitation in the exponential phase since both the standard and the target are equally affected by the changes in amplification parameters that occur as the reactions enter the plateau phase. Temperature cycling into the plateau phase, therefore, does not interfere with quantitation and even increases the sensitivity of the assay. Ultimately, quantitation by competitive PCR involves a set of reactions that include a constant volume aliquot of unknown DNA concentration and a dilution series of known concentrations of the standard competitor DNA. At the point where the molar amounts of the two products are equivalent, the amount of original target DNA present in the sample is equivalent to the amount of standard initially added. Thus, quantitation of the unknown DNA is based on the attainment of an equivalence point at a known concentration of standard competitor DNA. Internal competitive standards for quantitative PCR typically have sequences that are homologous to the target nucleic acids and amplify with the same or slightly modified primers. The target and the competitor, therefore, amplify with the same kinetics. Altering the size of the competitor molecule relative to the target sequence has been shown to be an excellent method for competitor. This method eliminates the need to use restriction enzymes with variable digestion efficiencies to cut competitor molecules engineered with unique sites. Using automated sequencers, a one base pair size difference between PCR products can be detected, allowing extremely similar competitor molecules to be utilized. Primers were derived from the published P. marinus DNA sequences for the ribosomal RNA gene (Fong et al., 1993) and the adjacent internal transcribed spacer (ITS-1) region (Goggin, 1994). The primers, designated PER-18S (5’CCTA-CGGGATTGGTTGTATCAG3’) and PER-ITS (5’CATCTCGCAACTCTCTAACA-AAAG3’) specifically amplified a 1210 base pair fragment of DNA from within the SSU rRNA gene to within the ITS-1 of the ribosomal DNA region. The primers were tested for sensitivity with cloned SSU rDNA derived from pure cultures of P. marinus. As little as 0.005 fg of P. marinus DNA was detected. They were tested for specificity with DNA from cultured cells of P. atlanticus, from cultured cells of a variety of P. marinus isolates from along the east coast of the United States, and from a variety of cultured dinoflagellates. The primers amplified DNA from all isolates of P. marinus, but not from P. atlanticus or any of the dinoflagellates. A competitor DNA molecule was constructed using the PCR primers PER-18S and ITS-MUT (5’CATCTCGCAACTCTCTAACAAAAGagcaagagagagcGAGACCGC-TG3’). The latter (ITSMUT) contained the entire sequence of the PER-ITS primer; however, the ITS-MUT primer created a gap of thirteen nucleotides in the sequence to be amplified by linking to a region of sequence several bases upstream, within the ITS-1 region. The PER-18S and ITS-MUT PCR product was and cloned. The QCPCR assay involved a two-phase approach. Titration spanning a broad range of competitor dilutions was performed to obtain a rough estimate of the amount of DNA and then a second titration over a narrower range was performed for precise quantitation. A standard curve was prepared by extracting DNA from a quantified number of cultured P. marinus cells (102 to 106 cells). The standard curve related the number of cells to the amount of DNA as determined by QCPCR.
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Validation of PCR diagnostics for Perkinsus marinus Target DNA was obtained from oysters collected from the lower Chesapeake Bay, Virginia. Oyster shells were notched and hemolymph samples were withdrawn with needle and syringe. Approximately 0.6 ml of hemolymph was divided equally for DNA extraction and for hemolymph Fluid Thioglycollate (FTM) analysis. Oysters were then shucked and 0.25 g of gill/mantle tissue was removed for DNA extraction and an equal amount for standard FTM analysis. The remaining oyster tissue was processed for the total body burden FTM analysis (Fisher and Oliver, 1996). Results are shown in Table 1. For the QCPCR results, any number >0 but
