Post 10: Glugeosis

Glugeosis is a marine and freshwater parasitic disease and is well documented in Japanese aquaculture and reported outbreaks in North America. The main causative agent of glugeosis disease is microsporidian parasites in the Glugea genus (Koesharyani et al 2005).

In 1998, a private hatchery in Indonesia culturing juvenile barramundi found that in the early stages of infection, fish exhibited lethargic swimming and a disinterest in food (Koesharyani et al 2005). The longer a fish goes infected, the easier it is to externally diagnose, with heavily infected fish exhibiting a cripplingly swollen abdomen, the cause of which are white cysts ranging in diameter from 0.5-1.5 mm, growing inside the fish (Zafran et al 1998). If left untreated, the cysts may multiply until the abdomen bursts or the fish starves to death (Zafran et al 1998).

An effective treatment for glugeosis disease is administering copper sulfate into the water body. Prevention of glugeosis in an aquaculture setting is to reduce the stocking density and to maintain good water quality, however research shows that infections have occurred mostly due to live shrimp being fed to fish. This suggests oral transmission of the parasite (Koesharyani et al), therefore decent food supervision should decrease infection rates.

numerous white cysts in the abdomen of fish infected with glugeosis

References:

Koesharyani, I. Roza, D. Mahardika, K. Johnny, F. Zafran, A. Yuasa, K 2005, ‘Manual for Fish Disease Diagnosis – II’, ‘Japan International Cooperation Agency’

Zafran, A. Rosa, D. Koesharyani, I. Johnny, F. Youssa, K 1998, ‘Manual for Fish Diseases Diagnosis’, ‘Japan International Cooperation Agency’

Post 9: Marine Velvet Disease

Marine Velvet disease (Amyloodinium ocellatum) is a microscopic, parasitic protista in the phylum dinoflagellata (Brown & Hovasse 1946). Mass mortalities resulting from A. ocellatum have been documented since the 1930's with the parasite infecting fish in both fresh and salt water; tolerating salinity levels from 3-45 ppt (Paperna 1980).

A. ocellatum has a 3 stage life-cycle, beginning as a trophont (The feeding stage of its life cycle). The trophont then matures into a reproductive tomont which produces the third stage in the life cycle; the free swimming dinospores-which can survive up to a week whilst looking for a host organism (Paperna 1980).

A fish with A. occeltatum can be visually diagnosed in an aquarium or aquaculture environment by the infected fish 'flashing' against the substrate and/or gasping for air at the surface as it suffers from anoxia due to A. ocellatum occuring on the gills of the host fish (Lawler 1980). In the later stages, the skin of infected fish may appear powdery (appearing as if it has been rolled in flour) as the Marine Velvet disease can cause signficant damage to the hosts skin and fins (Lawler 1980).

Copper is an effective treatment to aquatic parasites and fungi and is popular for treatment of A. ocellatum, in the form of copper sulfate. A knowledge of the water chemistry such as pH levels in the hosts surrounding water body should be known to avoid potential copper toxicity (Cardeilhac & Whitaker 1988). Therefore knowing the concentrations of copper in the administering treatment and the volume of water in the tank is recommended for this treatment process.

Amyloodinium ocellatum can be diagnosed from observed powedery skin

  
     

References:

Brown, E. Hovasse, R 1946, 'Amyloodinium ocellatum (Brown), A Peridinian Parasitic on Marine Fishes. A Complementary Study', 'Journal of Zoology', vol. 116, pp. 33-46

Cardeilhac, P. Whitaker, B 1988, 'Copper Treatments: Uses and Precautions', 'Veterinary Clinics of North America: Small Animal Practice', vol. 18, pp. 435-448

Lawler, A 1980, 'Studies on Amyloodinium ocellatum (Dinoflagellata) in Mississippi Sound: Natural and Experimental Hosts', 'Gulf and Carribean Research', vol. 6, pp. 403-413

Paperna, I 1980, 'Amyloodinium ocellatum (Brown, 1931) (Dinoflagellida) Infestations in Cultured Marine Fish at Eilat, Red Sea: Epizootiology and Pathology', 'Journal of Fish Diseases', vol. 3, pp. 363-372

Post 8: Gill Flukes

Gill flukes are highly infectious, 1-2 mm long, flatworm parasites commonly found in fresh and salt water aquariums (Klinger & Floyd 1998). This post will focus on the monogenean trematodes-trematodes being a common name for the flukes, and monogenean which describes the host-site specific life cycle of the parasite. Monogenean trematodes affect the gills, skin and fins of the host fish (Klinger & Floyd 1998).

Gill flukes can enter an aquarium water body through water exchanges, transplants of live rocks or be brought in with introduced fish. Gill fluke parasites are equipped with hooks which they use to latch onto their host-with the parasites life span being roughly 7-9 days (Ogawa 2015).

Directly diagnosing flukes through visual observation of the parasite is unlikely until the later stages of infection. Fish that are infected can almost always be seen participating in 'flashing', where the fish brushes the infected area on the sides or bottom of the tank due to physical irritation (Kiskaroly 1977). In the later stages, flukes growing on the fishes gills will interfere with respiration, thus the host will either be seen gasping for air (generally at the surface where oxygen diffusion is greatest), or lethargically sitting at the bottom as it dies.

Preventative steps can be taken to avoid outbreaks of gill flukes, such as fresh water bathing new fish and live rocks before introducing them into a saltwater aquarium; although doing this to fish that have suffered a prolonged fluke infection can cause further damage to the skin and fins (Ogawa 2015). Praziquantel has been shown to be effective in treating gill flukes and other aquatic worm parasites as it generally only requires 1-2 treatments and is non toxic to plants (Ishimaru et al 2013).

Gill fluke vs Skin fluke in fish 

References:

Ishimaru, K. Mine, R. Shirakashi, S. Kaneko, E. Kubono, K. Okada, T. Sawada, Y. Ogawa, K 2013, '

Praziquantel Treatment against Cardicola Blood Flukes: Determination of the Minimal Effective Dose and Pharmacokinetics in Juvenile Pacific Bluefin Tuna', 'Aquaculture', vol. 402-403, pp. 24-27

Kiskaroly, M 1977, 'Study of the Parasite Fauna of Freshwater Fishes from Fish Ponds of Bosnia and Herzegovina. A. Monogenean Trematodes 1. I. Cyprinid fish ponds', 'Veterinaria', vol. 26, pp. 195-208

Klinger, R. Floyd, R 1998, 'Introduction to Freshwater Fish Parasites', 'University of Florida-Fredrick, Aldridge & Jerome' 

Ogawa, K 2015, 'Diseases of Cultured Marine Fishes caused by Platyhelminthes (Monogenea Digenea, Cestoda)', 'Parasitology', vol. 142, pp. 178-195




  

Post 7: Taura Syndrome in American Shrimp Farming

Shrimp farming is occasionally reliant on capturing wild shrimp in a post-larvae stage for grow out or for broodstock prposes (Argue et al 2002). Unfortunately this stocking technique means potentially introducing diseased shrimp into a facility. One such virus that has caused billions of damge to shrimp stocks since the 1990's within the American region alone is a disease now known as Taura Syndrome in the Aparavirus genus (Lightner 2011).

Taura Syndrome emerged in 1992 in Ecuador and since then, certain species have shown great susceptibility-specifically the Whiteleg Shrimp (Litopenaeus vannamei) where outbreaks have occuered throughout the entirity of the species life cycle (Lightner et al 1995). Diagnosis of the disease in a farm setting occurs generally after noticing significant mortalities of shrimp in ponds/the presence of birds feeding on deceased shrimp (Brock 1997). further visual diagnosis of Taura Syndrome are the presence of multifocal, melanized spots on the body and tail of those effected (Lightner at al 1995).

Taura Syndrome induced melanized spots

    
 No current cure for Taura Syndrome currently exists, however resistance to the virus has been discovered in individuals of L. vannamei, which are undergoing selective breeding to pass on this resistance to future stocks (Argue at al 2002).


References:

Argue, B. Arce, S. Lotz, J. Moss, S 2002, 'Selective Breeding of Pacific White Shrimp (Litopenaeus vannamei) for Growth and Resistance to Taura Syndrome Virus', 'Aquaculture', vol. 204, pp. 447-460

Brock, J 1997, 'Taura Syndrome, a Disease Important to Shrimp Farms in the Americas', '
World Journal of Microbiology and Biotechnology', vol. 13, pp. 415-418


Lightner, D 2011, 'Virus Diseases of Farmed Shrimp in the Western Hemisphere (the Americas): A Review', 'Journal of Verterbrate Pathology', vol. 106, pp. 110-130

Lightner, D. Redman, R. Hasson, K. Pantoja, C 1995, 'Taura Syndrome in Penaeus vannamei (Crustacea: Decapoda): Gorss signs, Histopathology and Ultrastructure', 'Diseases of Aquatic Organisms', vol. 21, pp. 53-59

Post 6: Lymphocystis

Lymphocystis is a common viral disease, occuring in more than 100 marine and freshwater fish species (Yan et al 2011).
Lymphocystis is reported widely in home aquariums and is associated with fish facing high levels of stress; which is usually attibuted to poor water quality (Lawler & Donnes 1977) or the stress of moving fish from one environment to another e.g. moving fish from the pet shop to the home aquarium.
Lymphocystis causes hypertrophy (increase in volume) of the tissue cells of infected fish, which leads to visible white, pinprick nodules forming usually on the fins and body (Lawler & Donnes 1977). Unfortunately the formation of these nodules often lead to a mis-diagnosis of Ichthyophthirius multifillis (Fresh Water White Spot Disease).

The diagnosis of Lymphocystis is difficult in the early stages due to its similar resemblance to Fresh Water Whitespot Disease, however the white nodules in Lymphocystis will clump together to form large clusters, usually within two weeks of exhibiting symptoms (Paperna et al 1982). Infected fish also exhibit swimming difficulty and may also be inactive due to a breathing difficulty associated with nodules growing over the gills (Paperna et al 1982).

Viruses can evolve rapidly because they lack many factors that correct impurities in their DNA or RNA that exist in humans. Therfore mutations in viruses are a common occurence (Bamford et al 2002). Because of this, there is no existing cure for Lymphocystis. Therefore treatment should be focused on decreasing the stress on infected fish to allow the immune system to fight the virus and to decrease the risk of secondary infections.  

    

Early stage Lymphocystis

Late stage Lymphocystis

References:

Bamford, D. Bernett, R. Stuart, D 2002, 'Evolution of Viral Sructure', 'Theoretical Population Biology', vol. 61, pp. 461-470

Lawler, A. Donnes, J 1977, 'New Hosts for Lymphocystis, and a List of Recent Hosts', 'Journal of Wildlife Disease' vol. 13, pp. 307-312

Paperna, I. Sabnai, H. Colorni, A 1982, 'An Outbreak of Lymphocystis in Sparas aurata L. in the Gulf of Aqaba, Red Sea', 'Journal of Fish Diseases', vol. 5, pp. 433-437

Yan, X. Wu, Z. Jian, J 2011, 'Analysis of the Genetic Diversity of the Lymphocystis Virus and its Evolutionary Relationship with its Host', 'Virus Genes, vol. 43, pp. 358-366

Post 5: Vibriosis-An Examination of one of the most Recurring Bacterial Diseases in the Marine Environment

In 1997, Japanese researchers captured both juvenile and adult individuals of Estuary Cod, Mangrove Grouper and Napoleon Fish and transported these individuals back to Gondol Research station. In less than a week after transportation, all captured fish had died (Zafran et al 1998).
The cause of which came to be regarded as the most common bacterial disease in marine fishes; Vibriosis-from the genus Vibrio (Zafran et al 1998).

Bacteria in the Vibrio genus have evolved to become highly versatile, as strains of the disease have been documented to infect fish in marine, brackish and freshwater environments (Bullock 1977).
A study on Vibrio cholerea even found significant genotypical and phenotypical trait variations within the individual bacterial species (Cho et al 2010).

Signs of Vibriosis in juvenile fish simply appear simply as having a darker body coloration and a loss of appetite (Munn 2008), with older fish displaying haemorrhaging from hemorrhagic septicaemia and/or ulcerative lesions occurring on the fishes skin. Fish that are infected in the eyes also exhibit eye bulging (Zafran et al 1998).

A control of Vibriosis in an aquaculture or aquarium setting involve maintaining high water quality and a low density of fish within the tank in order to avoid any further stress on the fishes immune system. Administration of antibiotics is also a valid treatment to stop the spread of the disease, usually the use of the broad-spectrum antibiotic; OxyTetra Cycline over an 8-10 day period (Zafran et al 1998).

A snapper with the whole Vibrosis package; darker skin, lesions and haemorrhaging.

Stay tuned for more exciting pathogens and parasites.

References:

Bullock, G 1977, 'Vibriosis in Fish', 'U.S. Fish and Wildlife Service'.

Cho, Y. Yi, H. Lee, J. Kim, D. Chun, J 2010, 'Genomic Evolution of Vibrio cholerae', 'Current Opinion in Microbiology', vol. 13, pp. 646-651

Munn, C 2008, 'Vibriosis in Fish and its Control', 'Aquaculture Research', vol. 8, pp. 11-15

Zafran, A. Rosa, D. Koesharyuni, I. Johnny, F. Yuasa, K 1998, 'Manual for Fish Diseases Diagnosis', 'Japan International Cooperation Agency'.

Post 4: Freshwater White Spot Disease in Home Aquariums

Home aquariums have become a popular way of having pets and and decorating the home in the 21st century. Due to the costs associated with saltwater aquariums, plenty of people are opting for freshwater.
Unfortunately there are diseases that have evolved to thrive in these conditions that pose risks to freshwater fish.

Ichthyophthirius multifiliis (Freshwater White Spot) is a parasite that exists on the outside of a fish that can cause 100% of mortalities in an aquarium or aquaculture setting (Jiravanichpaisal et al 2004). It gets its name from the 1mm long white cysts that typically grow on the gills and body of the fish (Wu et al 2002), and each spot is a cyst comparable to grains of sugar that encloses a parasite (Nigrelli et al 1976).

I. multifiliis can become introduced into an aquarium usually by introducing contaminated fish, exchanging untreated and contaminated water into the system or using contaminated instruments in the same water body as the fish (Dickerson 2006).
As the cysts grow on the individuals gills and body, respiration becomes increasingly difficult as the cysts block water flow through the gills and because fish excrete excess mucous due to irritation from the cysts (Dickerson 2006). Wounds left by parasites also affect the fishes swimming ability making it harder to feed (Dickerson 2006).

A visual diagnosis of the fish is the fastest way to detect an I. multifiliis infection, with the stand out symptoms being the presence of white cysts, rapid operculum movements as the fish struggles to breath and anorexia due to a lack of feeding (Dickerson 2006).
Treatment is the quarantine of infected individuals into clean water until the symptoms subside-or the individual dies and/or the addition of salt into the system to a maximum of 4 parts per thousand, as freshwater fish can tolerate higher salinity than the parasite (Dickerson 2006).

I. multifiliis targets the gills and body

References:

Dickerson, H 2006, 'Ichthyophthirius multifiliis and Cryptocaryon irritans (Phylum Ciliophora)', 'Fish Diseases and Disorders' pp. 116-118

Jiravanichpaisal, P. Soderhall, K. Soderhall, I 2004, 'Effect of Water Temperature on the Immune Response and Infectivity Pattern of White Spot Syndrome Virus (WSSV) in Freshwater Crayfish', 'Fish and Shellfish Immunology' vo. 17, pp. 265-275

Nigrelli, R. Pokorny, K. Ruggieri, G 1976, 'Notes on Ichthyophthirius multifiliis, a Ciliate Parasitic on Fresh-Water Fishes, with Some Remarks on Possible Physiological Races and Species', 'Transactions of the American Microscopical Society', vol. 95, pp. 607-613

Wu, J. Nishioka, T. Mori, K. Nishizawa, T. Muroga, K 2002, '

A Time-Course Study on the Resistance of Penaeus japonicus Induced by Artificial Infection with White Spot Syndrome Virus', 'Fish and Shellfish Immunology', vol. 13, pp. 391-403

   

Post 3: Streptococcus: the Flesh Eating Bacteria in the Far North & the Evolution of an Epidemic

Between the years 2007 to 2011, over 90 Queensland Grouper (Epinephelus lanceolatus) and a variety of wild fish and estuarine stingrays were found dead in the Cairns inlet in Far North Queensland (Bowater et al 2012). The symptoms were black patches of necrotic tissue and visible signs of both internal and external bleeding. The Department of Northern Fisheries diagnosed the problem as a disease called Streptococcus agalactiae, a flesh eating bacterium, which lead to septicaemia (Presence of bacterium in the blood) in the infected (Bowater et al 2012). This was the first recorded outbreak of S. agalactiae in Australia (Bowater et al 2012) and because it occurred in the wild, there was very little that responders could do besides removing the infected and deceased individuals from the environment. 

In the last 30 years, Streptococcus has been realised as an epidemic, with S. agalactiae being the leading cause of fast spreading infections in new born mammals (Brochet et al 2006). 
Other symptoms aside from skin necrosis and internal bleeding, infected fish can be seen swimming erratically and their eyes appear bulging (Abuseliana et al 2011). 

In an aquaculture or aquarium setting, antibiotics are the only realistic treatment for S. agalactiae when septicaemia is present. S. agalactiae has however evolved a number of resistances to antibiotics with 9 similtaneous antibiotic resistances being seen using PCR tests (Zeng et al 2006).
The first treatment should be a ten day course of the antibiotic Oxy-tetra Cycline (OTC) by either incorporating this into food or dosing the water body. If this treatment doesn't work, it becomes a trial and error process with other antibiotics to see what can irradicate the bacterium from the system.

A deceased Queensland Grouper that died during the Streptococcus outbreak in Cairns

References:

Abuseliana, A. Daud, H. Aziz, S. Bejo, S. Alsaid, M 2011, 'Pathogenicity of Streptococcus agalactiae Isolated from a Fish Farm in Selangor to Juvenile Red Tilapia (Oreochromis sp.)', 'Journal of Animal and Veterinary Advances', vol.10, pp. 914-919

Bowater, R. Forbes-Faulkner, J. Anderson, I. Condon, K. Robinson, B. Kong, F. Gilbert, G. Reynolds, A. Hyland, S. McPherson, G. O'Brien, J. Blyde, D 2012, 'Natural Outbreak of Streptococcus agalactiae (GBS) Infection in Wild Giant Queensland Grouper, Epinephelus lanceolatus (Bloch), and Other Wild Fish in Northern Queensland, Australia', 'Journal of Fish Dieseases', vol. 35, pp. 173-186

Brochet, M. Couve, E. Zouine, M. Vallaeys, T. Rusniok, C. Lamy, M. Buchrieser, C. Trieu-Cuot, P. Kunst, F. Poyart, C. Glaser, P 2006, 'Genomic Diversity and Evolution within the Species Streptococcus agalactiae', 'Microbes and Infection', vol. 8, pp. 1227-1243

Zeng, X. Kong, F. Wang, H. Darbar, A. Gilbert, G 2006, 'Simultaneous Detection of Nine Antibiotic Resistance-Related Genes in Streptococcus agalactiae Using Multiplex PCR and Reverse Line Blot Hybridization Assay', 'Antimicrobial Agents and Chemotherapy', vol. 50, pp. 204-209

Post 2: Bacterial Fin Rot in Aquaculturally Reared Queensland Grouper and Issues Surrounding the Frequent use of Antibiotics

The Queensland Grouper (Epinephelus lanceolatus) is the largest bony fish to reside around coral reefs in the world (Jennings & Lock, 1996). Aqua cultural facilities and aquariums that work in the breeding and rearing of Queensland Grouper for research and/or other purposes sometimes lose fish due to pathogens and parasites. One such pathogen is a disease called Fin Rot, which gets its name from the way it slowly destroys mainly the tail fin (caudal fin) of Grouper and many other fish species and can be caused by either a fungal infection or by a bacterium (Nagasawa & Cruz-Lacierda 2004). In this post, I will be discussing the bacterial fin rot disease which results from a Pseudomonas Fluorescens infection.

Bacterial fin rot affecting the Caudal, Elongated Dorsal and Anal fin

This disease is often associated with poor water quality and dense populations. Poor water quality can increase the stress in cultured grouper which in turn lowers their immune system which makes them more vulnerable to pseodomonas fluorescens (Nagasawa & Cruz-Lacierda, 2004). Queensland Grouper are also cannibalistic and so in dense populations, the larger fish attack the smaller fish and the injuries incurred allow for opportunistic fin rot infections.
Treatment for bacterial fin rot is the simple use of antibiotics, the most commonly used is Oxy-Tetra Cycline (OTC) over a 10 day treatment period (Austin & Austin, 2007). Issues surrounding the frequent use of this however include the bacterium developing an immunity to this. Bacterial resistance for antibiotics has been observed since the 1950's and the two theories surrounding Fin Rot is whether it is the result of bacteria evolving in response to the use of OTC or whether the bacterium already possesses genes which give it immunity (Groisman & Ochman, 1996).
Hopefully with more research, more effective treatments can be found.

References:

Austin, B. Austin, D. 2007, 'Bacterial Fish Pathogens: Disease of Farmed and Wild Fish', 'Springer Science'

Groisman, E. Ochman, H. 1996, 'Pathogenicity Island: Bacterial Evolution in Quantum Leaps', 'Cell', vol. 87, pp. 791-794

Jennings, S. Lock, J. 1996, 'Population and Ecosystem Effects of Reef Fishing', 'Reef Fisheries', vol. 20, pp. 193-218

Nagasawa, K. Cruz-Lacierda, E. 2004, 'Diseases of Cultured Groupers', 'South-East Asian Fisheries Development Centre'

Somga. J, Somga, S. 2002, 'Impacts of Disease on Small-Scale Grouper Culture in the Philippines', 'Primary Aquatic Animal Healthcare', vol. 14, pp. 248-261

Post 1: Betanodavirus-An examination of potential management strategies.

Betanodavirus is a genus of virus that causes a specific strain of a disease called Viral Nervous Necrosis (VNN) in a variety of marine organisms including fin-fish and shellfish (Chi et al, 2008). VNN is a single stranded, RNA virus and is one of the main problems aquaculture facilities face as it can cause mortalities of up to 100% of reared populations and it effects 157 known species worldwide (Chi et al, 2003). VNN is regarded as a very hardy virus, affecting fish in both tropical and temperate environments and  has been known to be able to survive for up to 10 years in soil (Munday et al, 2002).
Because viruses have a high mutation rate and exist within the hosts cells (Gojobori et at, 1990), they are very hard to treat and thus there is no current cure for VNN.
VNN symptoms can be witnessed as the infected organisms exhibit very lethargic behaviour, show a disinterest for food which leads to anorexia and in fin-fish, spiral swimming is a symptom due to a loss of motor function (Arimoto et al, 1996).
VNN outbreaks can be linked to environmental factors such as large salinity and temperature fluctuations and high sediment loads in the water. To reduce outbreaks, especially in an Aquaculture setting, temperature and salinity should be maintained at a level that minimises stress to the fish.
Prevention can also include incorporating ozone into a circulating water system before the water is added into the tank containing fish, as ozone is toxic to pathogens. Also being an unstable molecule, ozone quickly dissipates out of the system and therefore if incorporated in correct doses will not be harmful to the fish.
VNN outbreaks are most common in fish aged between 4-70 days old (Fakuda et al, 2005) and hopefully through further research, Vaccines may be found to prevent this disease when organisms are most susceptible.

References:

Chi, S. Shieh, J. 2003, 'Genetic and Antigenic Analysis of Betanodaviruses Isolated from Aquatic Organisms in Taiwan', 'Diseases of Aquatic Organisms', vol. 7, pp. 238-249

1. Fenner, B. 
2. Thiagarajan, R.
3.  Chua. H, Kwang, J. 2005, '
Betanodavirus B2 Is an RNA Interference Antagonist That Facilitates Intracellular Viral RNA Accumulation', 'Journal of Virology', vol. 80, pp. 85-94

Arimoto, M. Sato, J. Maruyama, K. Mimura, G. Furusawa, I. 1996, 'Effect of Chemical and Physical Treatments on the Inactivation of Striped Jack Nervous Necrosis Virus (SJNNV)', 'Aquaculture', vol 143, pp. 15-22

Chi, C. Lo, B. Lin, S. 2008, 'Characterization of Grouper Nervous Necrosis Virus (GNNV)', 'Fish Diseases', vol. 24, pp. 3-13

Fakuda, Y. Nguyen, H. Furuyashi, M. Nakai, T. 1996, 'Mass Mortality of CUltured Sevenbanded Grouper, Epinephelus Septemifasciatus, Associated with Viral Nervous Necrosis', 'Fish Pathology, vol. 31, pp. 165-170

Munday, B. Kwang, J. Moody, N. 2002, 'Betanodavirus Infections of Teleost Fish', 'Journal of Fish Diseases', vol. 25, pp. 127-142

Gojobori, T. Moriyama, E. Kimura, M. 1990, ' Molecular Clockwork of Viral Evolution and the Neutral Theory', 'Current Issue' vol. 87, pp. 24-32