© 2019 Nelmarie Landrau Giovannetti

DETECTION AND CHARACTERIZATION OF EMERGING PATHOGENS IN STRANDED

CETACEANS

By

NELMARIE LANDRAU GIOVANNETTI

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL

OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2019

To my Mom, Dad, Sisters and Niece and Nephew

ACKNOWLEDGMENTS

I would like to acknowledge the following people, institutions, and organizations because

without their assistance my dissertation research would not have been possible. I would like to

thank my advisor Dr. Thomas Waltzek for all his guidance and patience throughout my

dissertation. I would also like to sincerely thank my other committee members: Dr.

Kuttichantran Subramaniam, Dr. Kristi West, Dr. John Lednicky, and Dr. Salvatore Frasca Jr. for

their guidance and many contributions to my dissertation research.

I want to thank the institutions and organizations that provided financial assistance during

my PhD studies including the Aquatic Animal Health Program, University of Florida (UF)

College of Veterinary Medicine (CVM), and the Florida Fish and Wildlife Conservation

Commission (FWC Grant #15145). I am also indebted to the UF Veterinary Graduate Student

Association and Graduate Student Council for providing travel grant funding, which allowed me

to present my research at several conferences.

I thank all the people who helped me during my time at the University of Florida,

Gainesville, starting with the staff and students at UF Wildlife Aquatic Veterinary and Disease

Laboratory (WAVDL) including: Patrick Thompson, Linda Archer, Allison Cauvin, Dr. Abigail

Clark, Dr. Galaxia Cortes-Hinojosa, Jared Freitas, Rachel Henríquez, Dr. Thaís Rodrigues,

Samantha Koda, Elizabeth Scherbatskoy, Dr. Preeyanan Sriwanayos, Kamonchai Imnoi, Dr.

Natalie Stilwell, Jaime Haggard, Dr. Alissa Deming, and Dr. María José Robles-Malagamba.

WAVDL Warriors, your words of encouragement, our lunch escapades and daily fun adventures

made graduate life more pleasurable than I ever thought possible. I am also grateful to the faculty

and staff within the UF CVM, including Dr. Iskande Larkin, Nina Thompson, Kia Hendrix,

Melissa Ann Brown and Sally O’Connell. I am thankful to various people across the globe for

their expertise and for providing samples and/or sequences: Dr. David Rotstein, Dr. Katia Groch,

Dr. Jerry Saliki, Ole Nielsen, Dr. Nahiid Stephens, and Dr. Pádraig Duignan. Lastly, I would like

to thank the Hawaii Pacific University Marine Mammal Response Team for making my three

month stay such an amazing and unforgettable experience! A special thanks to Dr. Kristi West,

Erin Hanahoe, Sarah Donahue, Nick Hoffman, Jackie Ton, Stacia Marcoux, Kyanna Tamborini,

and Clare Wolf. Finally, my sincerest thanks to several lifelong friends and mentors (Michelle

De Jesús, Marinelly Rodríguez, Antonio A. Mignucci-Giannoni) as well as, my family for their

love, encouragement, and patience as I pursued my dreams.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...............................................................................................................4

LIST OF TABLES ...........................................................................................................................8

LIST OF FIGURES .........................................................................................................................9

ABSTRACT ...................................................................................................................................12

CHAPTER

1 LITERATURE REVIEW .......................................................................................................14

Introduction to Marine Mammal Strandings ..........................................................................14 Marine Mammal Regulations .................................................................................................16 Florida Marine Mammal Strandings .......................................................................................18

Hawaii Marine Mammal Strandings .......................................................................................20 Conclusion ..............................................................................................................................22

2 MOLECULAR DETECTION OF PATHOGENS IN STRANDED CETACEANS .............26

Introduction .............................................................................................................................26 Materials and Methods ...........................................................................................................27

Samples ............................................................................................................................27

Nested Universal Adenovirus Endpoint PCR ..................................................................28 Semi-Nested Universal Respovirus/Morbillivirus/Henipavirus Endpoint RT-PCR .......28 Nested Universal Herpesvirus Endpoint PCR .................................................................29

Nested Hammondia heydorni/Toxoplasma gondii/Neospora caninum Endpoint

PCR ..............................................................................................................................29

Next-Generation Sequencing (NGS) ...............................................................................30 Results.....................................................................................................................................31

Florida Molecular Diagnostics ........................................................................................31 Hawaii Molecular Diagnostics ........................................................................................31

Discussion ...............................................................................................................................32

3 PHYLOGENOMIC DIVERSITY OF CETACEAN MORBILLIVIRUSES ............................51

Introduction .............................................................................................................................51

Materials and Methods ...........................................................................................................53 Samples and RNA Extractions ........................................................................................53 RT-PCR Conditions and Sanger Sequencing of PCR Amplicons ...................................53 Next-Generation Sequencing and Genome Annotation ..................................................54 Phylogenetic Analysis: Alignment and Model Selection ................................................55 Sequence Identity Matrix ................................................................................................55

Results.....................................................................................................................................56

Genome Organization ......................................................................................................56

Phylogenetic Analysis .....................................................................................................56 Sequence Identity Matrix ................................................................................................57

Discussion ...............................................................................................................................57

4 CHARACTERIZATION OF A NOVEL CIRCOVIRUS FROM A STRANDED

LONGMAN’S BEAKED WHALE (INDOPACETUS PACIFICUS) ....................................86

Introduction .............................................................................................................................86 Materials and Methods ...........................................................................................................89

Samples and Genome Sequencing ...................................................................................89 Phylogenetic and Genetic Analyses ................................................................................90 PCR Detection of BWCV in Longman’s Beaked Whale Tissues ...................................90 In Situ Hybridization (ISH) .............................................................................................91

Results.....................................................................................................................................92 Genomic Sequence Annotation .......................................................................................92

Phylogenetic and Genetic Analyses ................................................................................93 Detection of BWCV in Longman’s Beaked Whale Tissues ...........................................93 In Situ Hybridization (ISH) .............................................................................................93

Discussion ...............................................................................................................................94

5 TOXOPLASMA GONDII IN STRANDED HAWAIIAN CETACEANS ............................108

Introduction ...........................................................................................................................108 Methods and Materials .........................................................................................................113

Postmortem Examination and PCR Screening for Toxoplasma gondii .........................113

Histopathology and Immunohistochemistry ..................................................................113

Multilocus Sequence Typing, Genetic, and Phylogenetic Analysis ..............................114 Results...................................................................................................................................114

PCR Screening for Toxoplasma gondii .........................................................................114

Postmortem Examination, Histopathology, and Immunohistochemistry ......................115 Multilocus Sequence Typing, Genetic and Phylogenetic Analysis ...............................117

Discussion .............................................................................................................................117

6 CONCLUDNG STATEMENTS ..........................................................................................129

LIST OF REFERENCES .............................................................................................................131

BIOGRAPHICAL SKETCH .......................................................................................................147

LIST OF TABLES

Table page

2-1 PCR test results for Florida marine mammal samples submitted to the UF Wildlife

and Aquatic Veterinary Diagnostic Laboratory from 2014 – 2018 ...................................36

2-2 PCR test results for Hawaiian marine mammal samples submitted to the UF Wildlife

and Aquatic Veterinary Diagnostic Laboratory from 2014 – 2018 ...................................37

3-1 Description of the CeMV samples sequenced in this chapter 3 .........................................61

3-2 List of the primers used in PCR reactions to sequence the genomes of seven CeMV ......62

3-3 Virus names, GenBank accession numbers and references for the morbilliviruses

used in genetic and phylogenetic analyses.........................................................................72

4-2 Circoviruses used in the phylogenetic and genetic analyses .............................................99

LIST OF FIGURES

Figure page

1-1 Sperm whale stranding near Egmond, the Netherlands, 15 February 1764. The

picture reveals that measurements were not always taken accurately ...............................24

1-2 In 1577, Flemish artist Jan Wierix engraved Three Beached Whales, which depicts

three stranded sperm whales ..............................................................................................25

2-1 Schematic of the location of the primers used in the nested PCR assay targeting the

adenovirus DNA polymerase gene ....................................................................................47

2-2 Schematic of the location of the primers used in semi-nested RT-PCR assay targeting

the RNA polymerase gene of the genera Morbillivirus, Respovirus, Henipavirus ...........48


herpesvirus DNA polymerase gene ...................................................................................49


Toxoplasmatinae internal transcribed spacer 1 (ITS1) of the rDNA .................................50

3-1 Genome organization of the CeMVs sequenced in this Chapter 3 ....................................75

3-2 Cladogram depicting the relationship of the CeMV strains within the genus

Morbillivirus, based on the concatenated nucelotide sequence of the 6 genes ..................76

3-3 Cladogram depicting the relationship of the CeMV strains within the genus

Morbillivirus, based on the nucleotide sequence of the partial P gene .............................77

3-4 Sequence identity matrix illustrating the genetic distance of the CeMVs to other

members of the genus Morbillivirus based on the concatenated nucleotide sequence

of the 6 genes .....................................................................................................................78


members of the genus Morbillivirus based on the nucleotide sequence of the partial P

gene. ...................................................................................................................................79


members of the genus Morbillivirus based on the nucleotide sequence of the N gene .....80


members of the genus Morbillivirus based on the nucleotide sequence of the P gene .....81


members of the genus Morbillivirus based on the nucleotide sequence of the M gene .....82


members of the genus Morbillivirus based on the nucleotide sequence of the F gene .....83


members of the genus Morbillivirus based on the nucleotide sequence of the H gene .....84


members of the genus Morbillivirus based on the nucleotide sequence of the L gene ......85

4-1 Genome schematics of the beaked whale circovirus (BWCV) ........................................100

4-2 Phylogram depicting the relationship of the BWCV from Longman’s beaked whale

to species of the genus Circovirus based on their capsid amino acid sequences .............101

4-3 Phylogram depicting the relationship of the BWCV from Longman’s beaked whale

to species of the genus Circovirus based on their replication-associated amino acid

sequences .........................................................................................................................102

4-4 Genetic comparison of the full genome of the novel BWCV from Longman’s beaked

whale to 29 accepted type species in the genus Circovirus .............................................103

4-5 RNAscope® in situ hybridization (ISH) results from lymph node (LN) tissue of the

Longman’s beaked whale ................................................................................................104

4-6 RNAscope® in situ hybridization (ISH) results from the diaphragm tissue of the


4-7 RNAscope® in situ hybridization (ISH) results from the liver tissue of the


4-8 RNAscope® in situ hybridization (ISH) results from the liver tissue of the


5-1 Gross examination of an adult male spinner dolphin infected with T. gondii .................122

5-2 Microscopic examination of internal organs of a spinner dolphin infected with T.

gondii in the pulmonary interstitium................................................................................123

5-3. Microscopic examination of the heart of a spinner dolphin infected with T. gondii in

the heart ............................................................................................................................124

5-4 Microscopic examination of the brain of a spinner dolphin infected with T. gondii in

the brain ...........................................................................................................................125

5-5 Microscopic examination of the adrenal gland of a spinner dolphin infected with T.

gondii in the adrenal gland ...............................................................................................126

5-6 Immunohistochemistry (IHC) performed on internal organ tissue sections of a

spinner dolphin infected with T. gondii in the brain, adrenal gland, and liver ................127

5-7 Cladogram depicting the relationship of the Toxoplasma gondii linages 1-12, based

on the concatenated nucelotide sequences of GRA6, SAG1, MIC1 & 2, and BTUB1 ...128

Abstract of Dissertation Presented to the Graduate School

of the University of Florida in Partial Fulfillment of the

Requirements for the Degree of Doctor of Philosophy

DETECTION AND CHARACTERIZATION OF EMERGING PATHOGENS IN STRANDED

CETACEANS

By

Nelmarie Landrau Giovannetti

May 2019

Chair: Thomas B. Waltzek

Major: Veterinary Medical Sciences

Stranding events provide scientists with important information on the biology and health

status of marine mammal populations. They provide baseline information on marine mammal

ecology including anatomy/physiology, dietary habits, genetics, distribution, and occurrence of

disease within populations. In recent years, characterization of infectious agents in samples from

stranded marine mammal has increased because of the development of improved molecular

diagnostic tools including universal PCR assays, in situ hybridization (ISH) assays, and next-

generation sequencing (NGS) approaches.

This dissertation outlines several research projects in which these molecular diagnostic

tools assisted in the discovery of previously unknown pathogens in stranded marine mammals

from Florida and Hawaii. Chapter 1 provides an overview of marine mammal unusual mortality

events and single animal strandings, while Chapter 2 describes different molecular diagnostic

techniques (e.g., PCR and NGS) used to detect and characterize the genomes of the pathogens

that can cause these disease episodes. Genomic sequencing and phylogenomic analyses were

used to characterize globally emerging cetacean morbilliviruses (Chapter 3), the first ever

cetacean circovirus detected in a stranded Hawaiian Longman’s beaked whale (Indopacetus

pacificus) (Chapter 4), and disseminated toxoplasmosis in a stranded Hawaiian spinner dolphin

(Stenella longirostris) (Chapter 5). The characterized infectious agents represent pathogens

responsible for unexplained mortality events in cetaceans (e.g., Cetacean morbillivirus) and

pathogens responsible for single strandings (e.g., Toxoplasma gondii) in cetaceans. Two new

virus species were discovered from Hawaiian cetaceans using NGS approach, a Fraser’s dolphin

(Lagenodelphis hosei) morbillivirus (Chapter 3) and a beaked whale circovirus (Chapter 4). An

ISH assay using RNAscope® technology was used to confirm the beaked whale circovirus was

present in the tissues of the stranded Longman’s beaked whale (Chapter 4). In recent years, T.

gondii has become a pathogen of increasing concern to Hawaiian wildlife. PCR screening of

tissue samples from 16 stranded cetacean species from Hawaii (302 tissue samples) resulted in

one stranded spinner dolphin testing positive for T. gondii in all nine tissues tested (Chapter 5).

Using a multilocus sequence typing approach we determined the spinner dolphin’s T. gondii

strain to be most closely related to strains reported from human toxoplasmosis cases in North

America and Europe.

14

LITERATURE REVIEW

Introduction to Marine Mammal Strandings

In past times, stranded marine mammals were used as a source of food or functioned as

religious or spiritual symbols in communities living near the coast. One of the earliest recorded

mass stranding events occurred in the Bay of Biscay in the 7th century, according to Saint

Philibert’s of Jumièges biography, the vita Filiberti (i.e., Life of St. Philibert) (Arnold, 2016).

The event had a religious context as Saint Philibert prayed for sustenance and was met with a

great number of “fish” as food. The following is the full account of the event:

And at another time, when a great famine had begun to constrain the territory of

Poitiers, and the man of God [St. Philibert], mindful of the needs of the brothers,

lay himself down, praying. And when he rose, a great multitude of the fish that

are called marsuppas (harbor porpoises) found themselves in the river. When the

sea receded, two hundred and thirty-seven of them remained on the dry banks.

From that point and for the space of an entire year, the brothers had a huge

increase in their comfort, and many monasteries and paupers had food aid.

A 16th century author, Olaus Magnus, described how a single stranded whale provided resources

(Arnold, 2016):

When sea-monsters or whales have been hauled out of the sea…the people of the

neighborhood divide the booty…in such a way that with the meat, blubber, and

bones of a single whale or monster they can fill between 250 and 300 carts. After

they have put the meat and fat into vast numbers of large barrels, they preserve it

in salt, as they do other huge sea-fish.

Smeenk (1997) compiled strandings of sperm whales in the North Sea from 1560-1995. These

strandings were infrequent and in small numbers in that region and of which the details indicate

males ranging from 12-18 m in size were involved (Smeenk, 1997). Some of these records

include fairly accurate descriptive engravings of the animals (Figure 1-1, 1-2).

Modern stranding events capture public attention and are of great scientific interest

versus serving as a food source or holding religious meaning as in the past. Prior to necropsy, the

condition of the marine mammal carcass is assessed to determine the sampling scheme that will

15

be of greatest value (Pugliares et al., 2007). The Woods Hole Oceanographic Institution technical

report has been used to classify carcasses into five code categories based on the level of

decomposition: code 1 is alive, code 2 is fresh carcass, code 3 is moderate decomposition, code 4

is advanced decomposition and code 5 is mummified or skeletal remains (Pugliares et al., 2007).

Post-mortem examinations may provide valuable information about the ecology and

health status of marine mammal populations including baseline data on their

anatomy/physiology, dietary habits, genetics, distribution, and occurrence of diseases. For

example, studying the skull and/or skeleton of the animal provides clues about how the animal is

adapted for survival in its environment. The teeth are good indicators on the feeding habitats,

phylogenetic relationship, and can be used to estimate age. The condition of teeth and bones can

provide information about the animal’s history. For example, heavily worn teeth may indicate

that an animal is old or similarly, missing, chipped or broken teeth, or broken bones may indicate

aggressive interactions.

The biology and trophic ecology of some marine mammal species is known mostly from

stranded specimens (e.g., elusive pelagic beaked whales of the family Ziphiidae). A review of

stomach content from stranded beaked whales of the genera Hyperoodon, Mesoplodon, Ziphius,

revealed they primarily feed on cephalopods, fish, and to a lesser extent crustaceans (MacLeod et

al., 2003). Mesoplodon spp. were found to be the most piscivorous, while the southern bottlenose

whale (Hyperoodon planifrons) and Cuvier's beaked whale (Ziphius cavirostris) seldom

consumed fish (MacLeod et al., 2003).

Data accumulated over the years for some well-studied marine mammal species has led

to a better understanding of their growth rates, longevity, reproductive season, and age at

maturity (Bowen and Siniff, 1999). Data collected during stranding events can be used to

16

understand the effects of climatic change on the health of marine mammal populations. Truchon

et al. (2013) used general linear models to analyze a 15-year database documenting marine

mammal strandings and environmental parameters known to affect marine mammal survival,

from regional (sea ice) to continental scales (North Atlantic Oscillation). These analyses can

provide insights into the effects of environmental contaminants, algal toxins, anthropomorphic

factors, and pathogens on the health of marine mammal populations.

Marine Mammal Regulations

The federal Marine Mammal Protection Act (MMPA) was passed in 1972 to prevent any

live marine mammal within Unites States waters from being harmed, harassed, and harvested for

body parts. It mandated the creation of stranding networks to respond to marine mammal

strandings, with volunteering networks established all along the coasts of the United States. It

protects all marine mammals, which include cetaceans (dolphins, porpoises, and whales), polar

bears, sea otters, pinnipeds (sea lions and seals), and sirenians (manatees) within the waters of

the United States. A year later, the Endangered Species Act (ESA) was passed. The ESA

mandated protection for species that are considered endangered or threatened in all or some

portion of their range. The two agencies that were assigned to administrate these two acts are the

National Oceanic and Atmospheric Administration (NOAA) and the US Fish and Wildlife

Service (FWS). The FWS is responsible for walruses, manatee and dugongs, sea otters, and polar

bears and NOAA manages whales, dolphins, porpoises, seals, and sea lions.

Although marine mammals have been washing ashore for centuries, major marine

mammal die-offs started in the 1980s and have been argued to be indicators of the deteriorating

health of our oceans (Gulland and Hall, 2007). Two significant stranding events lead the US

Congress to pass the Marine Mammal Health and Stranding Response Act (MMHSRA) in 1992.

One of the events was the 1987-1988 Cetacean morbillivirus (CeMV) epizootic that occurred

17

along the Mid-Atlantic coast of the United States in which approximately 750 bottlenose

dolphins (Tursiops truncatus) perished. The other event was in the winter months of 1987, were

14 humpback whales were found dead along the coast of Cape Cod Bay, MA. Post-mortem

analyses revealed the 14 humpback whales had consumed prey contaminated with brevetoxin or

saxitoxin (Geraci et al., 1989). The MMHSRA coordinates emergency responses to sick, injured,

distressed, or dead pinnipeds and cetaceans. The network currently consists of five US regions,

representing the Southeast coast, West coast, Pacific Islands, Mid-Atlantic/New England, and

Alaska.

A single stranding, the most common stranding event, refers to when only one marine

mammal comes ashore dead or in need of intervention. Mass-stranding events generally refer to

the simultaneous stranding of two or more marine mammals of the same species, other than a

female and calf. Unusual mortality events (UMEs) can involve a few animals or large-scale

epizootics (i.e., die-off) (Geraci and Lounsbury, 2005). According to NOAA, UMEs are

unexpected strandings, involving significant die-offs of any marine mammal population and

demand immediate response (NOAA Fisheries, 2018a). In 1991, NOAA established the Working

Group on Marine Mammal Unusual Mortality Events (WGMMUME). Members of the

WGMMUME include experts from conservation organizations, state and federal agencies, and

scientific and academic institutions who work closely with stranding networks and have a wide

variety of experience in epidemiology, pathology, toxicology, biology, and ecology (NOAA

Fisheries, 2018a). They establish that an event is an UME if it meets one or more of the

following seven criteria:

1. A temporal change in morbidity, mortality, or strandings is occurring.

2. Morbidity is observed concurrent with or as part of an unexplained continual decline of a

marine mammal population, stock, or species.

18

3. Potentially significant morbidity, mortality, or stranding is observed in species, stocks, or

populations that are particularly vulnerable (e.g., listed as depleted, threatened, or

endangered or declining).

4. The species, age, or sex composition of the affected animals is different than that of

animals that are normally affected.

5. A marked increase in the magnitude or a marked change in the nature of morbidity,

mortality, or strandings when compared with prior records.

6. A spatial change in morbidity, mortality, or strandings is occurring.

7. Affected animals exhibit similar or unusual pathologic findings, behavior patterns,

clinical signs, or general physical condition (e.g., blubber thickness).

From 1991 to 2018, 67 UMEs have been formally recognized in the USA and the

etiologies were classified as undetermined/pending, biotoxins, ecological factors, human

interactions, and infectious diseases (NOAA Fisheries, 2018b). Three infectious agents have

been recognized as causes of UMEs: CeMV, avian influenza subtype H3N8, and Phocine

distemper virus (Karlsson et al., 2014; NOAA Fisheries, 2018c; Van Bressem et al., 2014).

Phocine distemper virus is believed to be responsible for an ongoing pinniped UME along the

Northeast US coastline (NOAA Fisheries, 2018c).

Florida Marine Mammal Strandings

There have been a total of 21 marine mammal UMEs in Florida, some solely

concentrated in Florida, while others spanned additional states. Thus, Florida has been involved

in 31% of the total UMEs and the causes have been human interaction, biotoxin, infectious

diseases, ecological factors, and undetermined. Manatees and bottlenose dolphins have been the

most commonly affected species in Florida UMEs and biotoxins are the most common cause

(10/21 events) (NOAA Fisheries, 2018b).

The Indian River Lagoon System (IRL), Florida, has an estimated population of 211

bottlenose dolphins and from January to May of 1982, 43 carcasses were recovered in the IRL

19

(Hersh et al., 1990). Serological data supported CeMV as the likely culprit of the disease episode

(Duignan et al., 1996). In addition, serological studies using samples collected from 2003-2007

indicated that IRL dolphins born after the 1982 mortality had antibodies to CeMV, indicating

exposure and infection (Bossart et al., 2010).

CeMV was confirmed as the cause of two large UMEs involving stranded bottlenose

dolphins along the Mid-Atlantic coastline from New York to Florida in 1987-1988 (Lipscomb et

al., 1994) and 2013-2015 (NOAA Fisheries, 2018d). A total of 742 and 1,650 bottlenose

dolphins were confirmed dead in the 1987-1988 and 2013-2015 UMEs, respectively. In both

events, all ages of bottlenose dolphins were affected with gross lesions observed on their skin,

mouth, joints, and lungs (NOAA Fisheries, 2018d). Diagnostics used to confirm the CeMV

diagnosis in these two events included histopathology, immunohistochemistry, serology, and

endpoint RT-PCR assays (Krafft et al., 1995; Lipscomb et al., 1994; NOAA Fisheries, 2018d;

Taubenberger et al., 1996).

CeMV UMEs involving large numbers of stranded bottlenose dolphin along the US Gulf

of Mexico (GOM) are thought to have occurred as far back as 1990 (Litz et al., 2014). Prior to

start of the formal UME program, a large bottlenose dolphin die-off (n = 342) occurred in the

GOM from Florida to Texas in 1990 (Hansen, 1992). Although never officially confirmed,

serological and histopathological evidence supported CeMV as the cause of this epizootic (Litz

et al., 2014). A confirmed CeMV UME involving more than 240 bottlenose dolphins occurred in

the GOM coastline from Alabama to Texas in 1993-1994 (Litz et al., 2014). Histopathology,

immunohistochemistry, and an endpoint RT-PCR assay supported CeMV as the cause of the

UME (Lipscomb et al., 1996). In contrast to mass stranding events caused by CeMV, strandings

involving smaller numbers or even single animals have been reported. In 1993 (Lipscomb et al.,

20

1994) and 2012 (Cassle et al., 2016) single bottlenose dolphins stranded along the Florida GOM

coastline and were found to be co-infected with CeMV and an ascomycete fungus (Aspergillus

sp.). The observed gross and microscopic lesions in these cases supported the co-infection and

the CeMV diagnoses were confirmed by immunohistochemistry and endpoint RT-PCR assays.

Infection by Toxoplasma gondii, an obligate coccidian parasite within the family

Sarcocystidae that infects a range of warm-blooded animals, has also resulted in marine mammal

strandings in Florida (Buergelt and Bonde, 1983; Inskeep II et al., 1990; Smith et al., 2016). The

first report of toxoplasmosis in cetaceans occurred in a stranded female bottlenose dolphin and

her calf from Tampa, Florida in 1987 (Inskeep II et al., 1990). The pathogen was identified by

light and electron microscopy and by immunohistochemistry in tissues of both animals. T. gondii

was associated with interstitial pneumonia, necrotizing adrenalitis, and cardiac myonecrosis in

the mother and with lymphoid necrosis in both dolphins (Inskeep II et al., 1990).

Hawaii Marine Mammal Strandings

The Pacific Island Region Marine Mammal Response Network responds to strandings of

cetaceans and Hawaiian monk seals (Neomonachus schauinslandi) in Guam, the Northern

Mariana Islands, American Samoa, the main Hawaiian Islands, and the Northwest Hawaiian

Islands. The Hawaiian monk seal is one of the most endangered seal species in the world and

they are only found in the Hawaiian archipelago.

There have been two UMEs involving Hawaiian monk seal, one was prior to the formal

UME program and involved >50 monk seals that died in Laysan Island, Hawaii in 1978 (Johnson

and Johnson, 1981). This mortality event was believed to be the result of gastrointestinal

parasitism and/or ciguatoxin produced by a dinoflagellate (Johnson et al., 1982). The second

UME occurred in 2001 in the Laysan Islands and involved 11 emaciated monk seals. In this

event, the necropsy results revealed no signs of infectious disease or toxicosis linking (Yochem

21

et al., 2004). Antonelis et al. (2001) reported that the unusually high mortality in yearling seals

appeared to be a result of their inability to forage successfully during the post-weaning transition

to nutritional independence.

Analyzed data from 65 years (1937-2002) of odontocete stranding data around the main

Hawaiian Islands showed that pygmy and dwarf sperm whales (Kogia spp.) (18%), spinner

dolphins (Stenella longirostris) (15%), striped dolphins (S. coeruleoalba) (11%), and sperm

whales (Physeter macrocephalus) (10%) were the four most commonly involved species

(Maldini et al., 2005). Mazzuca et al. (1999) compiled stranding data from 1957-1998 in the

Hawaiian Islands and identified nine mass stranding events, involving four species and 96

animals. The largest event occurred in 1959 when 28 short-finned pilot whales (Globicephala

macrorhynchus) became trapped and died behind a reef in Kauai, Hawaii. Since 1997, there have

been >150 cetacean strandings in Hawaii (West, unpublished data). It is common to find stranded

cetaceans in an advanced state of decomposition along the shorelines of the Hawaiian Islands

because of the tropical climate and remote locations. In 2011, a neonate female sperm whale

stranded alive and died at Oahu, Hawaii (West et al., 2015). West and colleagues (2015) found

that the neonate was co-infected CeMV and a Brucella sp. Microscopic lesions included a

chronic meningitis, lymphoid depletion, and pneumonia. The Brucella sp. was considered

responsible for the observed multisystemic pathology because it was isolated from all organs

tested (lung, cerebrum, lymph nodes, and umbilicus) as compared to the limited detection of

CeMV by RT-PCR and IHC (West et al., 2015).

The Longman’s beaked whale (Indopacetus pacificus) is one of the least studied cetacean

species with <10 confirmed recorded strandings worldwide. In 2010, a juvenile male was

stranded in Oahu, Hawaii (West et al., 2013). A necropsy was performed, and morphometric data

22

was collected, and samples were preserved for histopathology, genetics, and pathogen discovery

using molecular assays (West et al., 2013). Histopathological findings included

lympoplasmacytic periglomerulitis, mild lymphoid depletion, encephalitis, pulmonary edema,

thyroid gland atrophy, and bilateral hypertrophy of the adrenal cortex (West et al., 2013).

Diagnostic testing using conventional PCR assays detected a novel CeMV and a novel

alphaherpesvirus. This case was the first report of CeMV in a marine mammal from the central

Pacific.

The first reported case of disseminated toxoplasmosis in a Hawaiian marine mammal was

in a wild male spinner dolphin (Stenella longirostris) (Migaki et al., 1990). Additional cases of

fatal toxoplasmosis in Hawaiian marine mammals include reports in Hawaiian monk seals

(Barbieri et al., 2016; Honnold et al. 2005). Diagnostics used to confirm the T. gondii infection

in stranded marine mammals have included histopathology, immunohistochemistry, serology,

and endpoint PCR assays.

Conclusion

Although marine mammals have been stranding for centuries, major marine mammal die-

offs started in the 1980s and have been argued to be indicators of the deteriorating health of our

oceans. Modern stranding events capture public attention and are of great scientific interest

versus serving as a food source or holding religious meaning as in the past. Post-mortem

examinations of stranded marine mammals may provide valuable information about the ecology

and health status of marine mammal populations including baseline data on their

anatomy/physiology, dietary habits, genetics, distribution, and occurrence of diseases. Marine

mammal unusual mortality events have five etiologies including: undetermined/pending,

biotoxins, ecological factors, human interactions, and infectious diseases. Within the infectious

disease category, CeMV has been responsible for the greatest number of UMEs as well as being

23

responsible for single animal strandings in both the Hawaiian Islands and Gulf and Atlantic

coastlines of Florida. Similarly, the parasite Toxoplasma gondii has resulted in a number of

marine mammal strandings along the coastlines of Florida and Hawaii. The objectives of my

dissertation were to molecularly characterize pathogens within the tissues of stranded cetaceans

in Florida and Hawaii and to better understand their potential role in disease.

24

Figure 1-1. Sperm whale stranding near Egmond, the Netherlands, 15 February 1764. The

picture reveals that measurements were not always taken accurately. Anonymous

drawing in East Indian ink, collection Municipal Archive, Haarlem (Smeenk, 1997).

25

Figure 1-2. In 1577, Flemish artist Jan Wierix engraved Three Beached Whales, which depicts

three stranded sperm whales.

26

CHAPTER 2

MOLECULAR DETECTION OF PATHOGENS IN STRANDED CETACEANS

Introduction

In recent years, the characterization of infectious agents in tissue samples from stranded

marine mammal has increased because of the development of improved molecular diagnostic

tools. For example, the use of universal PCR assays have become routine diagnostic tools for

screening samples derived from stranded marine mammals. Using endpoint PCR, a single copy

of a DNA sequence is exponentially amplified to generate millions of copies of that particular

DNA target and this amplification allows visualization on an agarose gel (Green and Sambrook,

2012). PCR has several advantages such as it can be used on samples from code 1-4 animals and

even formalin-fixed tissues, can be performed using small tissue quantities, is relatively cheap,

can be designed to generate quantitative results, and has rapid turnaround. The amplification of

pathogen nucleic acid using universal PCR assays coupled with Sanger sequencing has proven to

be a useful approach in detecting marine mammal herpesviruses (Maness et al., 2011;

VanDevanter et al., 1996), adenoviruses (Rubio-Guerri et al., 2015; Wellehan et al., 2004),

paramyxoviruses (Cassle et al., 2016; Tong et al., 2008), and strains of Toxoplasma gondii (Silva

et al., 2009; Smith et al. 2016).

Additionally, viral enrichment strategies paired with high-throughput next-generation

sequencing (NGS) approaches have revolutionized the discovery and genomic characterization

of novel viruses. Virus discovery using NGS approaches provides several advantages over PCR

by providing an unbiased snapshot of all RNA and DNA viruses in a sample and recovering full

genomes given the large amount of sequence data generated. Recently, there has been an

increase in published papers describing novel viral pathogens from wild marine mammal

populations using NGS approaches (Bellehumeur et al., 2016; Davison et al., 2017a, b; Kluge et

27

al., 2016; Malmberg et al., 2017; Ng et al., 2011, 2015; Phan et al., 2015; Russo et al., 2018;

Siqueira et al., 2017). For example, the first full genomes of a marine mammal alphaherpesvirus,

gammaherpesvirus, and poxvirus have recently been reported from a wild beluga whale, a wild

common bottlenose dolphin, and a wild sea otter, respectively (Davison et al., 2017a, b; Jacob et

al., 2018).

In this chapter, we utilized universal PCR assays to detect RNA viruses (e.g. Cetacean

morbillivirus; CeMV), DNA viruses (e.g. herpesvirus), and Toxoplasma gondii in tissue samples

collected from Florida and Hawaiian stranded marine mammals. Additionally, a NGS approach

was used to detect novel viral pathogens in the tissue samples that the conventional PCR assays

failed to detect and to obtain complete viral genomes. This NGS approach facilitated the

genomic characterization of novel CeMVs and the first marine mammal circovirus.

Materials and Methods

Samples

A total of 344 tissue samples from stranded cetaceans from Florida or Hawaii between

2013 and 2017 were sent to the Wildlife and Aquatic Veterinary Disease Laboratory (WAVDL),

Gainesville, Florida for diagnostic screening (Table 2-1, 2-2). Tissues received by WAVDL

included brain, liver, spleen, lung, heart, kidney, adrenal gland, and lymph nodes. Tissue samples

were archived at -80˚C until used. Nucleic acid extractions (RNA, DNA, or both) were

performed on frozen tissue samples and an isolate using the following commercial kits: Maxwell

16 LEV simplyRNA Tissue Kit (ProMega Corp., Madison, WI), Qiagen DNeasy Kit (QIAGEN,

Valencia, CA), and AllPrep DNA/RNA Mini Kit (QIAGEN, Valencia, CA) according to the

manufacturer’s instructions. In addition, RNA extraction from a formalin-fixed paraffin-

embedded (FFPE) lung tissue block was extracted using a RNeasy FFPE Kit (QIAGEN,

28

Valencia, CA) (Table 2-1). This was from a bottlenose dolphin stranded in Alabama in 1993.

The universal conventional PCR assays and NGS approach are described below.

Nested Universal Adenovirus Endpoint PCR

The DNA samples were screened for the adenovirus DNA polymerase using two pairs of

primers, polFouter/polRouter and polFinner/polRinner (Figure 2-1; Wellehan et al., 2004).

Briefly, the following steps were used for the first round of amplification: one cycle of 94°C for

5 min for denaturing, followed by 40 amplification cycles of 94°C for 30 sec, annealing step at

46°C (polFouter/polRouter) for 1 min, elongation step at 72°C for 1 min, and a final elongation

step at 72°C for 7 min. For the second round of amplification, the following steps were: one

cycle of 94°C for 5 min for denaturing, followed by 40 amplification cycles of 94°C for 30 sec,

annealing step at 46°C (polFinner/polRinner) for 1 min, elongation step at 72°C for 1 min, and a

final elongation step at 72°C for 7 min. The PCR products from the second round were then

subjected to electrophoresis in 1% agarose gel stained with ethidium bromide. Fragments of the

expected size were purified using a QIAquick Gel Extraction Kit (QIAGEN, Valencia, CA) and

was sequenced in both directions using an ABI 3130 DNA sequencer (Life Technologies,

Carlsbad, CA).

Semi-Nested Universal Respovirus/Morbillivirus/Henipavirus Endpoint RT-PCR

The samples were screened for the paramyxoviruses RNA dependent RNA polymerase

(i.e., members of the genera Respovirus, Morbillivirus, and Henipavirus) using primers designed

for a semi-nested RT-PCR (Figure 2-2; Tong et al., 2008). Briefly, the following steps were used

for the first round of amplification: one cycle of 50°C for 30 min for cDNA synthesis and one

cycle of 95°C for 15 min for denaturing, followed by 40 amplification cycles of 94°C for 1 min,

annealing step at 45°C (ResMorHen-F1/ResMorHen-R) for 1 min, elongation step at 72°C for 1

min, and a final elongation step at 72°C for 10 min. For the second round of amplification, the

29

following steps were used: one cycle of 94°C for 5 min for denaturing, followed by 40

amplification cycles of 94°C for 1 min, annealing step at 45°C (ResMorHen-F2/ResMorHen-R)

for 1 min, elongation step at 72°C for 1 min, and a final elongation step at 72°C for 10 min. The

PCR products from the second round were then subjected to electrophoresis in 1% agarose gel

stained with ethidium bromide. Fragments of the expected size were purified using a QIAquick

Gel Extraction Kit (QIAGEN, Valencia, CA) and was sequenced in both directions using an ABI

3130 DNA sequencer (Life Technologies, Carlsbad, CA).

Nested Universal Herpesvirus Endpoint PCR

The samples were screened for the herpesvirus DNA polymerase using two pairs of

primers (nested PCR), DFA/ILK/KG1 and TGV/IYG (Figure 2-3; VanDevanter et al., 1996).

Briefly, the following steps were used for the conventional PCR for the first round of

amplification steps amplification steps: one cycle of 94°C for 5 min for denaturing, followed by

40 amplification cycles of 94°C for 30 sec, annealing step at 46°C (DFA/ILK/KG1) for 1 min,

elongation step at 72°C for 1 min, and a final elongation step at 72°C for 7 min. For the second

amplification, the following steps were used for conventional PCR: one cycle of 94°C for 5 min

for denaturing, followed by 40 amplification cycles of 94°C for 30 sec, annealing step at 46°C

(TGV/IYG) for 1 min, elongation step at 72°C for 1 min, and a final elongation step at 72°C for

7 min. The PCR products from the second round were then subjected to electrophoresis in 1%

agarose gel stained with ethidium bromide. Fragments of the expected size were purified using a

QIAquick Gel extraction kit (QIAGEN, Valencia, CA) and was sequenced in both directions

using an ABI 3130 DNA sequencer (Life Technologies, Carlsbad, CA).

Nested Hammondia heydorni/Toxoplasma gondii/Neospora caninum Endpoint PCR

The samples were screened for T. gondii using a nested PCR with external (JS4/CT2b)

and internal (CT1/CT2) primer pairs as previously described by Silva et al. (2009). The PCR

30

target is specific to Toxoplasmatinae internal transcribed spacer 1 (ITS1) of the rDNA (Figure 2-

4). The thermocycling conditions for the nested PCR included one cycle of 94°C for 5 min for

denaturation, followed by 40 amplification cycles of 94°C for 1 min, annealing step at 60°C

(JS4/CT2b) or 55°C (CT1/CT2) for 1 min, elongation step at 72°C for 1 min, and a final

elongation step at 72°C for 7 min. The PCR products from the second round were then subjected

to electrophoresis in 1% agarose gel stained with ethidium bromide. Fragments of the expected

size were purified using a QIAquick Gel extraction kit (QIAGEN, Valencia, CA) and was

sequenced in both directions using an ABI 3130 DNA sequencer (Life Technologies, Carlsbad,

CA).

Next-Generation Sequencing (NGS)

RNA extracts from the tissues of a Fraser’s dolphin that stranded in Hawaii in 2018

(DMV-Lh-18-Hawaii) and from formalin-fixed paraffin-embedded tissues (PMV-Tt-93-

Alabama) of a bottlenose dolphin that stranded along the Gulf coastline in 1993 were prepared

for NGS. A DNA extract from the tissues of a Longman’s beaked whale tissue sample (lung)

was also prepared for NGS. DNA and cDNA libraries were created and sequenced using a 600-

cycle version 3 MiSeq Reagent Kit on an Illumina MiSeq (Illumina, San Diego, CA) sequencer.

After trimming the resulting NGS data for quality in CLC Genomics Workbench 10.1.1

software, the reads were assembled de novo using SPades 3.10.1. (Bankevich et al., 2012). The

assembled contigs were subjected to BLASTX searches against a custom viral database, created

from virus protein sequences retrieved from UniProt Knowledgebase (UniProt Consortium,

2017), in CLC Genomics Workbench 10.1.1 software.

31

Results

Florida Molecular Diagnostics

A total of 9 cetacean individuals from two species (i.e., Clymene dolphins; Stenella

clymene and bottlenose dolphins) with 18 tissues were tested by PCR or RT-PCR for CeMV,

herpesvirus, and Toxoplasma gondii (Table 2-1). 2/18 tissues were positive for CeMV, which

were from two bottlenose dolphin samples collected in 2012 and 2013. After primer removal, the

bottlenose dolphin sample from 2012 (TT12047) produced a 559 bp amplicon with highest

nucleotide sequence identity of 100% (559/559) to dolphin morbillivirus strain BCF20110815-

LA001 (GenBank accession no. KU720624; Cassle et al., 2016). The bottlenose dolphin sample

from 2013 (TT13198) produced a 559 bp amplicon with highest nucleotide sequence identity of

99% (558/559) nucleotide sequence identity to six CeMV sequences in GenBank. Tissues tested

for T. gondii, herpesvirus, and adenovirus were negative (Table 2-1). The NGS approach

generated the complete genome sequence for a CeMV from a stranded bottlenose dolphin sample

(FFPE tissue; PMV-Tt-93-Alabama) derived from the CeMV UME that occurred along the US

Gulf coastline from 1993-1994 (Chapter 3).

Hawaii Molecular Diagnostics

A total of 330 tissue samples, from 55 individual cetaceans were tested by PCR or RT-

PCR for CeMV, herpesvirus, adenovirus, and/or Toxoplasma gondii (Table 2-2). 13/44 tissue

samples (IP13001-05, WVL-18068-01A-01G, and WVL-18068-01A) were positive for CeMV in

a Longman’s beaked whale (Indopacetus pacificus) and a Fraser’s dolphin (Lagenodelphis hosei)

(Chapter 3). The CeMV sequence from the Longman’s beaked whale showed the highest

nucleotide sequence identity of 86% (479/559) to a CeMV isolated from a stranded harbor

porpoise (McCullough et al., 1991). The sequence of the Fraser’s dolphin CeMV displayed

highest nucleotide identity (78%) to a CeMV characterized from a white-beaked dolphin

32

stranded in the Netherlands in 2011 (Jo et al., 2018). 11/115 tissue samples were positive for

herpesviruses in a sperm whale (Physeter macrocephalus), a spinner dolphin (Stenella

longirostris), a Longman’s beaked whale, and a Cuvier’s beaked whale (Ziphius cavirostris).

After primer removal, the 173 bp amplicon from the sperm whale sample (PM13001-03) showed

highest nucleotide sequence identity of 95% (165/173) to a sperm whale gammaherpesvirus

(GenBank accession no. AB510475). The spinner dolphin tissue sample (SL13010-11) showed

the highest nucleotide sequence identity of 92% (166/181) to a melon-headed whale

alphaherpesvirus (GenBank accession no. AB510474). The Longman’s beaked whale and

Cuvier’s beaked whale tissue samples (IP13001-05; ZCA13005) showed the highest nucleotide

sequence identities of 65% (106/163) and 99% (179/181) to Ziphius cavirostris alphaherpesvirus

(GenBank accession no. KP995682), respectively. All 88 tissue samples tested were negative for

adenovirus. 9/302 tissues samples (WVL-17010-23A-23C; SL15001A-01F) were positive for T.

gondii in a spinner dolphin (Chapter 5) (Table 2-2). The next-generation sequencing data

recovered the complete genome sequences of a CeMV recovered from a Fraser's dolphin tissue

sample (WVL-18068-01A) and the complete genome sequence of a novel circovirus recovered

from a Longman’s beaked whale tissue sample (IP13001) (Chapters 3 and 4, respectively).

Discussion

This study demonstrated the utility of universal endpoint PCR and RT-PCR assays in

screening marine mammal tissue samples for known pathogens and the usefulness of next-

generation sequencing (NGS) approaches in recovering the full genomes of marine mammal

pathogens that the universal PCR assays failed to detect. A universal endpoint RT-PCR detected

Toxoplasma gondii DNA in tissue samples from a stranded Hawaiian spinner dolphin. A

universal endpoint RT-PCR assay detected CeMV RNA in tissue samples from stranded

bottlenose dolphins, a Longman’s beaked whale, and a Fraser’s dolphin. A universal endpoint

33

PCR detected gammaherpesvirus DNA in tissue samples from a stranded Hawaiian sperm whale

and alphaherpesvirus DNA in tissue samples from a stranded Hawaiian spinner dolphin, a

Longman’s beaked whale, and a Cuvier’s beaked whale. The NGS data produced the complete

CeMV genomes from cDNA libraries generated from a formalin-fixed paraffin-embedded

bottlenose dolphin tissue sample and a Fraser’s dolphin frozen tissue sample. The NGS approach

also recovered the full genome of the first circovirus detected in a marine mammal (i.e., from a

DNA library generated from a Longman’s beaked whale frozen tissue sample).

In this study, an endpoint RT-PCR assay (Tong et al. 2008) detected CeMV RNA in

Florida bottlenose dolphin samples taken from strandings involving a single animal that occurred

along the Gulf of Mexico (GOM) in 2012 (Cassle et al., 2016) and an animal in 2013 that was

part of the 2013-2015 Mid-Atlantic CeMV UME (Fauquier et al., 2014). The partial RNA

dependent RNA polymerase sequence of the 2012 GOM CeMV was identical to the sequence of

a previously reported CeMV strain reported from a bottlenose dolphin that stranded along the

Louisiana coastline in 2011 (Fauquier et al., 2017). This suggests that a strain of CeMV may be

circulating in GOM bottlenose dolphins (Chapter 3). Additionally, our NGS data recovered the

full genome of a CeMV in a bottlenose dolphin that stranded in Alabama as part of a confirmed

CeMV UME that occurred along the GOM coastline from 1993-1994 (Litz et al., 2014).

Interestingly, this GOM CeMV was found to be a strain of porpoise morbillivirus (PMV) that

typically circulates in harbor porpoises within the North Sea (van de Bildt et al., 2005). These

data confirm the findings of a previous study, based on partial sequences of the CeMV

phosphoprotein, that PMV was likely responsible for the CeMV UME in GOM bottlenose

dolphins from 1993-1994 (Taubenberger et al., 1996). To our knowledge, this study represents

the first time a full morbillivirus genome has been determined from a formalin-fixed paraffin-

34

embedded tissue block using an NGS approach. We believe these techniques may be used in

future studies to unravel the evolution and epidemiology of viruses circulating in wildlife

populations.

An endpoint RT-PCR assay detected CeMV RNA in a Longman’s beaked whale and a

Fraser’s dolphin that stranded in Hawaii in 2010 (West et al., 2013) and 2018 (Rotstein et al.,

2018), respectively (Tong et al., 2008). The partial RNA dependent RNA polymerase sequence

of the Longman’s beaked CeMV displayed highest nucleotide identity (86%) to a PMV isolated

from a harbor porpoise that stranded on the coast of Northern Ireland in 1988 (McCullough et al.,

1991). The sequence of the Fraser’s dolphin CeMV displayed highest nucleotide identity (78%)

to CeMV characterized from a white-beaked dolphin that stranded in the Netherlands in 2011 (Jo

et al., 2018). The relatively high sequence divergence of Hawaiian CeMVs as compared to other

CeMV was the rationale for sequencing their full genomes and exploring their evolutionary

relationships to other morbilliviruses (Chapter 3). Finally, a DNA extract from the tissues of the

same Longman’s beaked whale was used to construct a DNA library and then sequenced on a

next-generation sequencer (NGS). The resulting NGS data recovered the full genome of a

Longman’s beaked whale circovirus, expanding the known host range of circoviruses into

marine mammals (Chapter 4).

A universal endpoint RT-PCR assay (Silva et al., 2009) detected Toxoplasma gondii

DNA in 4% (1/25 animals) of code 1, 2, and 3 stranded spinner dolphins analyzed in this study

from the Hawaii Pacific University stranding archive. All tissues from this spinner dolphin tested

positive for T. gondii by PCR and BLASTN searches of the resulting sequences against the

NCBI non-redundant database revealed 25 T. gondii sequences with 100% coverage and 100%

sequence identity. After 25 years, this is the only the second report of toxoplasmosis in a

35

stranded Hawaiian spinner dolphin (Migaki et al., 1990). In chapter 5, we performed a multilocus

sequencing approach, histopathology, and immunohistochemistry to compare the 2015 spinner

dolphin toxoplasmosis case to the aforementioned case of disseminated toxoplasmosis in a

spinner dolphin.

36

Table 2-1. PCR test results for Florida marine mammal samples submitted to the University of Florida Wildlife and Aquatic

Veterinary Diagnostic Laboratory from 2014 – 2018. WAVDL: Wildlife and Aquatic Veterinary Diagnostic Laboratory.

LN: lymph node. FFPE: formalin-fixed-paraffin embedded. CeMV: Cetacean morbillivirus. Herpes: herpesvirus. Adeno:

adenovirus. Toxo: Toxoplasma gondii. NGS: next-generation sequencing. +: positive. –: negative. n.d.: not done.

WAVDL ID Species Tissue Type CeMV Herpes Adeno Toxo NGS

TT13198 Bottlenose dolphin Lung + n.d. n.d. n.d. n.d.

TT13199 Bottlenose dolphin Pulmonary LN – n.d. n.d. n.d. n.d.

TT13200 Bottlenose dolphin Blowhole Swab – n.d. n.d. n.d. n.d.

TT12047 Bottlenose dolphin Lung + – n.d. – n.d.

PMV-Tt-93-Alabama* Bottlenose dolphin FFPE lung n.d. n.d. n.d. n.d. +

WVL-17377-1A Clymene dolphin Lung – n.d. – – n.d.

WVL-17377-1B Clymene dolphin Pulmonary Marginal LN – n.d. – – n.d.


WVL-17377-2B Clymene dolphin Lung LN – n.d. – – n.d.




WVL-17377-4B Clymene dolphin Pulmonary Marginal LN – n.d. – – n.d.






WVL-17377-7B Clymene dolphin Lung LN – n.d. – – n.d. *This sample is from Alabama, USA

37

Table 2-2. PCR test results for Hawaiian marine mammal samples submitted to the University of Florida Wildlife and Aquatic

Veterinary Diagnostic Laboratory from 2014 – 2018. WAVDL: Wildlife and Aquatic Veterinary Diagnostic Laboratory.

LN: lymph node. FFPE: formalin-fixed-paraffin-embedded. L: left. R: right. TB: tracheobronchial. CeMV: Cetacean

morbillivirus. Herpes: herpesvirus. Adeno: adenovirus. Toxo: Toxoplasma gondii. NGS: next-generation sequencing. +:

positive. –: negative. n.d.: not done. WAVDL ID Species Tissue Type CeMV Herpes Adeno Toxo NGS Comment

WVL-17379-1A Melon-headed whale Cerebrum – n.d. n.d. – n.d.

WVL-17379-1B Melon-headed whale Cerebellum – n.d. n.d. – n.d.

WVL-17379-1C Melon-headed whale Liver – n.d. n.d. – n.d.

WVL-17379-1D Melon-headed whale Spleen – n.d. n.d. – n.d.

WVL-17379-1E Melon-headed whale R Lung – n.d. n.d. – n.d.

WVL-17379-1F Melon-headed whale L Ventricle – n.d. n.d. – n.d.

WVL-17379-1G Melon-headed whale R Mediastinal LN – n.d. n.d. – n.d.

WVL-17379-1H Melon-headed whale L Mediastinal LN – n.d. n.d. – n.d.

WVL-17379-1I Melon-headed whale L Marginal LN – n.d. n.d. – n.d.

WVL-17379-1J Melon-headed whale R Marginal LN – n.d. n.d. – n.d.

WVL-17379-1K Melon-headed whale Mesenteric LN – n.d. n.d. – n.d.

WVL-17379-1L Melon-headed whale Anal LN – n.d. n.d. – n.d.

WVL-17349-01A Cuvier's beaked whale Anal LN – n.d. n.d. – n.d.

WVL-17349-01B Cuvier's beaked whale Spleen n.d. n.d. n.d. – n.d.

WVL-17349-01C Cuvier's beaked whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-01D Cuvier's beaked whale Mesenteric LN n.d. n.d. n.d. – n.d.

ZCA13001 Cuvier's beaked whale R Lung n.d. – – – n.d. Same as WVL-17349-01

ZCA13002 Cuvier's beaked whale L Lung n.d. – – – n.d. Same as WVL-17349-01

ZCA13003 Cuvier's beaked whale TB Ln n.d. – – – n.d. Same as WVL-17349-01

WVL-17349-02A Spotted dolphin R Lung n.d. n.d. n.d. – n.d.

WVL-17349-02B Spotted dolphin Liver n.d. n.d. n.d. – n.d.

WVL-17349-02C Spotted dolphin Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-02D Spotted dolphin R Ventricle n.d. n.d. n.d. – n.d.

STA13001 Spotted dolphin R Lung n.d. – – n.d. n.d. Same as WVL-17349-02

STA13002 Spotted dolphin L Lung n.d. – – n.d. n.d. Same as WVL-17349-02

WVL-17349-03A False killer whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-03B False killer whale L Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-03C False killer whale Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-03D False killer whale Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-04A Blainville's beaked whale Spleen n.d. n.d. n.d. – n.d.

38

Table 2-2. Continued WAVDL ID Species Tissue Type CeMV Herpes Adeno Toxo NGS Comment

WVL-17349-04B Blainville's beaked whale Mesenteric LN n.d. n.d. n.d. – n.d.

WVL-17349-04C Blainville's beaked whale Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-04D Blainville's beaked whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-04E Blainville's beaked whale R Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-04F Blainville's beaked whale Cerebrum n.d. n.d. n.d. – n.d.

MD13001 Blainville's beaked whale R Lung n.d. – – – n.d. Same as WVL-17349-04

MD13002 Blainville's beaked whale Mediastinal LN n.d. – – – n.d. Same as WVL-17349-04

MD13003 Blainville's beaked whale L Lung n.d. – – – n.d. Same as WVL-17349-04

WVL-17349-05A Rough-toothed dolphin Spleen n.d. n.d. n.d. – n.d.

WVL-17349-05B Rough-toothed dolphin Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-05C Rough-toothed dolphin Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-05D Rough-toothed dolphin Liver n.d. n.d. n.d. – n.d.

WVL-17349-05E Rough-toothed dolphin L Lung n.d. n.d. n.d. – n.d.

WVL-17349-06A Sperm whale Mesenteric LN n.d. n.d. n.d. – n.d.

WVL-17349-06B Sperm whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-06C Sperm whale Spleen n.d. n.d. n.d. – n.d.

WVL-17349-06D Sperm whale Brain n.d. n.d. n.d. – n.d.

WVL-17349-06E Sperm whale R Ventricle n.d. n.d. n.d. – n.d.

PM13001 Sperm whale R Lung n.d. + – – n.d. Same as WVL-17349-06

PM13002 Sperm whale L Lung n.d. + – – n.d. Same as WVL-17349-06

PM13003 Sperm whale TB LN n.d. + – – n.d. Same as WVL-17349-06

WVL-17349-07A Cuvier's beaked whale Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-07B Cuvier's beaked whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-07C Cuvier's beaked whale Spleen n.d. n.d. n.d. – n.d.

WVL-17349-07D Cuvier's beaked whale R Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-07E Cuvier's beaked whale Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-07F Cuvier's beaked whale R Lung n.d. n.d. n.d. – n.d.

WVL-17349-08A Longman's beaked whale Spleen n.d. n.d. n.d. – n.d.

WVL-17349-08B Longman's beaked whale Muscle n.d. n.d. n.d. – n.d.

WVL-17349-08C Longman's beaked whale L Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-08D Longman's beaked whale Mesenteric LN n.d. n.d. n.d. – n.d.

WVL-17349-08E Longman's beaked whale L Adrenal n.d. n.d. n.d. – n.d.

WVL-17349-08F Longman's beaked whale Liver n.d. n.d. n.d. – n.d.

IP13001 Longman's beaked whale Lung + + – – + Same as WVL-17349-08

39


IP13002 Longman's beaked whale Cerebrum + + – – n.d. Same as WVL-17349-08

IP13003 Longman's beaked whale Cerebellum + + – – n.d. Same as WVL-17349-08

IP13004 Longman's beaked whale Scapular LN + + – – n.d. Same as WVL-17349-08

IP13005 Longman's beaked whale Mediastinal LN + + – – n.d. Same as WVL-17349-08

WVL-17349-09A Melon-headed whale L Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-09B Melon-headed whale Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-09C Melon-headed whale Spleen n.d. n.d. n.d. – n.d.

WVL-17349-09D Melon-headed whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-09E Melon-headed whale Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-09F Melon-headed whale Mesenteric LN n.d. n.d. n.d. – n.d.

PEPE13002 Melon-headed whale R Lung n.d. – – – n.d. Same as WVL-17349-09

PEPE13003 Melon-headed whale L Lung n.d. – – – n.d. Same as WVL-17349-09

PEPE13004 Melon-headed whale R Pulmonary LN n.d. – – – n.d. Same as WVL-17349-09

PEPE13005 Melon-headed whale Mediastinal LN n.d. – – – n.d. Same as WVL-17349-09

PEPE13006 Melon-headed whale Hylar LN n.d. – – – n.d. Same as WVL-17349-09

WVL-17349-10A Risso's dolphin L Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-10B Risso's dolphin Liver n.d. n.d. n.d. – n.d.

WVL-17349-10C Risso's dolphin Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-10D Risso's dolphin Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-10E Risso's dolphin Spleen n.d. n.d. n.d. – n.d.

WVL-17349-10F Risso's dolphin Mesenteric LN n.d. n.d. n.d. – n.d.

GG13001 Risso's dolphin R Lung n.d. n.d. – – n.d. Same as WVL-17349-10

GG13002 Risso's dolphin L Lung n.d. n.d. – – n.d. Same as WVL-17349-10

GG13003 Risso's dolphin Prescapular LN n.d. n.d. – – n.d. Same as WVL-17349-10

GG13004 Risso's dolphin TB LN n.d. n.d. – – n.d. Same as WVL-17349-10

GG13005 Risso's dolphin Lung LN n.d. n.d. – – n.d. Same as WVL-17349-10

WVL-17349-11A Short-finned pilot whale Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-11B Short-finned pilot whale Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-11C Short-finned pilot whale Liver n.d. n.d. n.d. – n.d.

WVL-17349-11D Short-finned pilot whale Gastric LN n.d. n.d. n.d. – n.d.

WVL-17349-11E Short-finned pilot whale Spleen n.d. n.d. n.d. – n.d.

WVL-17349-11F Short-finned pilot whale R Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-11G Short-finned pilot whale R Lung n.d. n.d. n.d. – n.d.

WVL-17349-12A Bottlenose dolphin Spleen n.d. n.d. n.d. – n.d.

40


WVL-17349-12B Bottlenose dolphin Liver n.d. n.d. n.d. – n.d.

WVL-17349-12C Bottlenose dolphin Cerebellum n.d. n.d. n.d. – n.d.

WVL-17349-12D Bottlenose dolphin Cerebrum n.d. n.d. n.d. – n.d.

WVL-17349-12E Bottlenose dolphin L Ventricle n.d. n.d. n.d. – n.d.

WVL-17349-12F Bottlenose dolphin Anal LN n.d. n.d. n.d. – n.d.

WVL-17349-12G Bottlenose dolphin R Lung n.d. n.d. n.d. – n.d.

WVL-17349-12H Bottlenose dolphin Mesenteric LN n.d. n.d. n.d. – n.d.

ZCA13004 Cuvier's beaked whale R Lung n.d. – – – n.d.

ZCA13005 Cuvier's beaked whale L Lung n.d. – – – n.d.

ZCA13006 Cuvier's beaked whale Pulmonary LN n.d. – – – n.d.

ZCA13007 Cuvier's beaked whale TB LN n.d. – – – n.d.

PEPE13001 Melon-headed whale Right Lung n.d. – – – n.d.

PEPE13007 Melon-headed whale Right Lung n.d. + – – n.d.

PEPE13008 Melon-headed whale Left Lung n.d. – – – n.d.

PEPE13009 Melon-headed whale R Pulmonary LN n.d. – – – n.d.

PEPE13010 Melon-headed whale Mediastinal LN n.d. – – – n.d.

PEPE13011 Melon-headed whale Right Lung n.d. – – – n.d.

PEPE13012 Melon-headed whale Left Lung n.d. – – – n.d.

PEPE13013 Melon-headed whale TB Ln n.d. – – – n.d.

PEPE13014 Melon-headed whale Mediastinal LN n.d. – – – n.d.

WVL17010-01A Spinner dolphin Brain n.d. – – – n.d.

WVL17010-01B Spinner dolphin Liver n.d. n.d. n.d. – n.d.

WVL17010-01C Spinner dolphin Lung n.d. n.d. n.d. – n.d.

WVL17010-02A Spinner dolphin Cerebellum n.d. n.d. n.d. – n.d.


WVL17010-02C Spinner dolphin Lung n.d. n.d. n.d. – n.d.

WVL17010-02D Spinner dolphin Spleen n.d. n.d. n.d. – n.d.

WVL17010-03A Spinner dolphin Brain n.d. n.d. n.d. – n.d.

WVL17010-03B Spinner dolphin Lung n.d. n.d. n.d. – n.d.

WVL17010-03C Spinner dolphin Spleen n.d. n.d. n.d. – n.d.

WVL17010-04A Spinner dolphin Liver n.d. n.d. n.d. – n.d.

WVL17010-04B Spinner dolphin Spleen n.d. n.d. n.d. – n.d.

WVL17010-05A Spinner dolphin Brain n.d. n.d. n.d. – n.d.


41



WVL17010-06A Spinner dolphin Cerebellum n.d. n.d. n.d. – n.d.



WVL17010-07B Spinner dolphin R Lung n.d. n.d. n.d. – n.d.


WVL17010-08B Spinner dolphin L Lung n.d. n.d. n.d. – n.d.


WVL17010-08D Spinner dolphin L Adrenal n.d. n.d. n.d. – n.d.

WVL17010-08E Spinner dolphin Anal LN n.d. n.d. n.d. – n.d.

WVL17010-08F Spinner dolphin R Ventricle n.d. n.d. n.d. – n.d.

WVL17010-09A Spinner dolphin Cerebrum n.d. n.d. n.d. – n.d.


WVL17010-09C Spinner dolphin L Lung n.d. n.d. n.d. – n.d.

SL13001 Spinner dolphin R Lung n.d. – – n.d. n.d. Same as WVL-17010-09

SL13002 Spinner dolphin L Lung n.d. – – n.d. n.d. Same as WVL-17010-09



WVL17010-10C Spinner dolphin R Lung n.d. n.d. n.d. – n.d.


WVL17010-10E Spinner dolphin L Ventricle n.d. n.d. n.d. – n.d.

WVL17010-10F Spinner dolphin Anal LN n.d. n.d. n.d. – n.d.



SL13005 Spinner dolphin TB LN n.d. – – n.d. n.d. Same as WVL-17010-10

SL13006 Spinner dolphin Scapular LN n.d. – – n.d. n.d. Same as WVL-17010-10


WVL17010-11B Spinner dolphin Cerebellum n.d. n.d. n.d. – n.d.

WVL17010-11C Spinner dolphin Liver n.d. n.d. n.d. – n.d.

WVL17010-11D Spinner dolphin L Lung n.d. n.d. n.d. – n.d.

WVL17010-11E Spinner dolphin Spleen n.d. n.d. n.d. – n.d.

WVL17010-11F Spinner dolphin L Ventricle n.d. n.d. n.d. – n.d.

WVL17010-11G Spinner dolphin Colonic LN n.d. n.d. n.d. – n.d.

WVL17010-11H Spinner dolphin L Adrenal n.d. n.d. n.d. – n.d.

42








WVL17010-12G Spinner dolphin Mesenteric LN n.d. n.d. n.d. – n.d.






WVL17010-13F Spinner dolphin Mesenteric LN n.d. n.d. n.d. – n.d.

WVL17010-13G Spinner dolphin R Adrenal n.d. n.d. n.d. – n.d.

WVL17010-13H Spinner dolphin L Ventricle n.d. n.d. n.d. – n.d.






WVL17010-14F Spinner dolphin Anal LN n.d. n.d. n.d. – n.d.





WVL17010-15D Spinner dolphin R Adrenal n.d. n.d. n.d. – n.d.









43





SL13009 Spinner dolphin L lung LN n.d. – – n.d. n.d. Same as WVL-17010-16







SL13010 Spinner dolphin R Lung n.d. + – n.d. n.d. Same as WVL-17010-17

SL13011 Spinner dolphin L Lung n.d. + – n.d. n.d. Same as WVL-17010-17

SL13012 Spinner dolphin Prescapular LN n.d. – – n.d. n.d. Same as WVL-17010-17

SL13013 Spinner dolphin TB LN n.d. – – n.d. n.d. Same as WVL-17010-17

SL13014 Spinner dolphin L Marginal LN n.d. – – n.d. n.d. Same as WVL-17010-17

SL13015 Spinner dolphin R Marginal LN n.d. – – n.d. n.d. Same as WVL-17010-17

WVL17010-18A Spinner dolphin L Lung n.d. n.d. n.d. – n.d.

WVL17010-18B Spinner dolphin R Lung n.d. n.d. n.d. – n.d.

WVL17010-18C Spinner dolphin Mesenteric LN n.d. n.d. n.d. – n.d.




WVL17010-19D Spinner dolphin R Lung n.d. n.d. n.d. – n.d.


WVL17010-19F Spinner dolphin R Adrenal n.d. n.d. n.d. – n.d.

WVL17010-19G Spinner dolphin L Ventricle n.d. n.d. n.d. – n.d.

WVL17010-19H Spinner dolphin Mesenteric LN n.d. n.d. n.d. – n.d.






WVL17010-20F Spinner dolphin L Ventricle n.d. n.d. n.d. – n.d.


44





WVL17010-21E Spinner dolphin L Adrenal n.d. n.d. n.d. – n.d.


WVL17010-21G Spinner dolphin Anal LN n.d. n.d. n.d. – n.d.







WVL17010-22F Spinner dolphin L Adrenal n.d. n.d. n.d. – n.d.

WVL17010-22G Spinner dolphin L Ventricle n.d. n.d. n.d. – n.d.

WVL17010-23A Spinner dolphin Liver n.d. n.d. n.d. + n.d.

WVL17010-23B Spinner dolphin R Adrenal n.d. n.d. n.d. + n.d.

WVL17010-23C Spinner dolphin L Ventricle n.d. n.d. n.d. + n.d.

SL15001A Spinner dolphin Brain – – – + n.d. Same as WVL-17010-23

SL15001B Spinner dolphin Spleen – – – + n.d. Same as WVL-17010-23

SL15001C Spinner dolphin L Lung – – – + n.d. Same as WVL-17010-23

SL15001D Spinner dolphin L Marginal LN – – – + n.d. Same as WVL-17010-23

SL15001E Spinner dolphin L Prescapular LN – – – + n.d. Same as WVL-17010-23

SL15001F Spinner dolphin Mesenteric LN – – – + n.d. Same as WVL-17010-23












45



WVL17010-25G Spinner dolphin Anal LN n.d. n.d. n.d. – n.d.


PC15001A False killer whale L Adrenal n.d. – n.d. – n.d.

PC15001B False killer whale R Adrenal n.d. – n.d. – n.d.

PC15001C False killer whale L Lung n.d. – n.d. – n.d.

PC15001D False killer whale R Lung n.d. – n.d. – n.d.

PC15001E False killer whale L Orbital LN n.d. – n.d. – n.d.

PC15001F False killer whale R Orbital LN n.d. – n.d. – n.d.

PC15001G False killer whale Submandibular LN n.d. – n.d. – n.d.

PC15002A False killer whale L Atrium n.d. – n.d. – n.d.

PC15002B False killer whale R Atrium n.d. – n.d. – n.d.

PC15002C False killer whale L Lung n.d. – n.d. – n.d.

PC15002D False killer whale R Lung n.d. – n.d. – n.d.

PC15002E False killer whale L Ventricle n.d. – n.d. – n.d.

PC15002F False killer whale R Ventricle n.d. – n.d. – n.d.

PC15002G False killer whale L Marginal LN n.d. – n.d. – n.d.

PC15002H False killer whale Mediastinal LN n.d. – n.d. – n.d.

PC15003A False killer whale Spleen – – – – n.d.

PC15003B False killer whale Cerebrum – – – – n.d.

PC15003C False killer whale Left Lung – – – – n.d.

PC15003D False killer whale Submandibular LN – – – – n.d.

PC15003E False killer whale Prescapular LN – – – – n.d.

PEPE15001A Melon-headed whale Brain – – – – n.d.

PEPE15001B Melon-headed whale Spleen – – – – n.d.

PEPE15001C Melon-headed whale L Lung – – – – n.d.

PEPE15001D Melon-headed whale L Marginal LN – – – – n.d.

PEPE15001E Melon-headed whale Prescapular LN – – – – n.d.

PEPE15001F Melon-headed whale Sublumbar LN – – – – n.d.

FA13001 Pygmy killer whale R Lung n.d. – – – n.d.

FA13002 Pygmy killer whale L Lung n.d. – – – n.d.

FA13003 Pygmy killer whale Prescapular LN n.d. – – – n.d.

MN13001 Humpback whale R Lung n.d. – – – n.d.

MN13002 Humpback whale L Lung n.d. – – – n.d.

46


MN13003 Humpback whale Hylar LN n.d. – – – n.d.

MN13004 Humpback whale Marginal LN n.d. – – – n.d.

MN13005 Humpback whale TB LN n.d. – – – n.d.







MN13012 Humpback whale Hylar LN n.d. – – – n.d.

OO13001 Killer whale R Lung – – – – n.d.

OO13002 Killer whale L Lung – – – – n.d.

SCO13001 Striped dolphin R Lung n.d. – n.d. n.d. n.d.

SCO13002 Striped dolphin L Lung n.d. – n.d. n.d. n.d.

SCO13003 Striped dolphin Pulmonary LN n.d. – n.d. n.d. n.d.

SCO13004 Striped dolphin Hylar LN n.d. – n.d. n.d. n.d.

SCO13005 Striped dolphin R Lung n.d. – n.d. – n.d.

SCO13006 Striped dolphin L Lung n.d. – n.d. – n.d.

SCO13007 Striped dolphin Bronchial LN n.d. – n.d. – n.d.

SCO13008 Striped dolphin TB LN n.d. – n.d. – n.d.

SCO13009 Striped dolphin L Lung n.d. – n.d. – n.d.

WVL-18068-01A Fraser’s dolphin L Lung + – n.d. n.d. +

WVL-18068-01B Fraser’s dolphin Cerebellum + – n.d. n.d. n.d.

WVL-18068-01C Fraser’s dolphin Liver + – n.d. n.d. n.d.

WVL-18068-01D Fraser’s dolphin L Hylar LN + – n.d. n.d. n.d.

WVL-18068-01E Fraser’s dolphin Spleen + – n.d. n.d. n.d.

WVL-18068-01F Fraser’s dolphin Mediastinal LN + – n.d. n.d. n.d.

WVL-18068-01G Fraser’s dolphin R Prescapular LN + – n.d. n.d. n.d.

47

Figure 2-1. Schematic of the location of the primers used in the nested PCR assay targeting the

adenovirus DNA polymerase gene. R1 = round one and R2 = round two.

48

Figure 2-2. Schematic of the location of the primers used in semi-nested RT-PCR assay targeting

the RNA polymerase gene of the genera Morbillivirus, Respovirus, Henipavirus

within the subfamily Paramyxovirinae. R1 = round one and R2 = round two.

49


herpesvirus DNA polymerase gene. R1 = round one and R2 = round two.

50


Toxoplasmatinae internal transcribed spacer 1 (ITS1) of the rDNA. R1 = round one

and R2 = round two.

51

CHAPTER 3

PHYLOGENOMIC DIVERSITY OF CETACEAN MORBILLIVIRUSES

Introduction

The genus Morbillivirus, family Paramyxoviridae, includes negative-sense single-

stranded, enveloped RNA viruses of importance in both human and veterinary medicine.

Morbilliviruses infect a range of mammalian hosts including: felids, canids, marine mammals,

nonhuman primates, and humans (Appel and Gillespie, 1972; Carpenter et al., 1998; Osterhaus et

al., 1995; Qiu et al., 2011; Viana et al., 2015). Over the last three decades, morbilliviruses have

emerged as among the most important causative agents of infectious diseases in wild pinnipeds

and odontocete populations (Duignan et al., 2014; Van Bressem et al., 2014).

In 1987-1988, 742 bottlenose dolphins (Tursiops truncatus) died along the Atlantic coast

of the United States in an epizootic event caused by a morbillivirus (Lipscomb et al., 1994). A

dolphin morbillivirus (DMV) was identified from a 1990 outbreak in the Mediterranean Sea in

striped dolphins (Stenella coeruleoalba) (Domingo et al., 1990). The DMV was later designated

Cetacean morbillivirus (CeMV) (Bolt et al., 1994) and accepted as a species in the genus

Morbillivirus. The species CeMV include strains genetically distinct from DMV that have

emerged in a variety of odontocetes and mysticetes including: porpoise morbillivirus (PMV),

pilot whale morbillivirus (PWMV), and beaked whale morbillivirus (BWMV) (West et al.,

2013). The PMV was originally isolated from a harbor porpoise (Phocoena phocoena) that

stranded in Northern Ireland (McCullough et al., 1991) and the PWMV was detected from a

long-finned pilot whale (Globicephala melas) that stranded in New Jersey, USA (Taubenberger

et al., 2000). Another CeMV strain was first identified from a stranded Hawaiian Longman’s

beaked whale (Indopacetus pacificus) (West et al., 2013) and later from several Hawaiian

cetaceans (Jacob et al., 2016). Partial sequencing of morbilliviruses from Southern Hemisphere

52

odontocetes in Brazil and Western Australian suggest that they might represent a new strain or

even a new species in the genus Morbillivirus (Groch et al., 2014, 2018; Stephens et al., 2014).

In recent years, CeMV has infected numerous cetaceans including: short-finned

(Globicephala macrorhynchus) and long-finned pilot whales, striped and bottlenose dolphins, fin

whale (Balaenoptera physalus), pygmy sperm whale (Kogia breviceps), harbor porpoise, Guiana

dolphin (Sotalia guianensis), Indo-Pacific bottlenose dolphins (T. aduncus), and white-beaked

dolphins (Lagenorhynchus albirostris) (Bellière et al., 2011a; Groch et al., 2014; Lipscomb et

al., 1994; Mazzariol et al., 2012; McCullough et al., 1991; Stephens et al., 2014; Van Elk et al.,

2014; Wierucka et al., 2014; Yang et al., 2006). In addition to cetaceans, CeMV has also been

detected in harbor seals (Phoca vitulina) and manatees (Trichechus manatus) (Duignan et al.,

1995; Mazzariol et al., 2013).

The availability of CeMV genomic sequences has led to the development of molecular

diagnostic assays capable of rapidly detecting these viruses in stranded cetacean tissues. An

endpoint reverse transcription (RT) PCR assay was developed to detect morbilliviruses based on

a highly conserved region of the phosphoprotein gene (P gene) and has been used in several

studies to detect CeMV (Barrett et al., 1993). In this assay, universal primers generate a 429 bp

amplicon with the upstream primer binding to positions 400-419 of the measles virus P gene and

the downstream primer binding to positions 808-828 (Barrett et al., 1993; Bellini et al., 1985).

Other assays to detect CeMV have been designed using PCR primers that bind to other

morbillivirus genes, such as the nucleocapsid protein (N gene), fusion protein (F gene), and the

hemagglutinin protein (H gene) (Raga et al., 2008; Rubio-Guerri et al., 2013b; van de Bildt et al.,

2005). Besides endpoint RT-PCR assays, real-time (rt) quantitative reverse transcription PCR (rt-

qRT-PCR) assays have been utilized for the detection and quantification of CeMV in marine

53

mammals. Grant and colleagues (2009) developed an rt-qRT-PCR assay for detecting CMV and

PMV by targeting a hypervariable region in the N gene. Rubio-Guerri et al. (2013b) developed a

rt-qRT-PCR which amplifies a highly conserved region within the F gene of three strains: one

primer set detects both DMV and PMV RNAs and the other primer set detects PWMV RNA.

To date, four CeMV genome sequences have been determined sequenced. Rima and

colleagues (2005) sequenced an isolate from a Mediterranean striped dolphin. Fauquier and

colleagues (2017) sequenced the genome of viruses infecting two bottlenose dolphins and a

striped dolphin in the Gulf of Mexico. In the present study, we sequenced and described seven

CeMV genomes from stranded odontocetes from the Gulf and Pacific coasts of North America

and the North and Mediterranean Seas of Europe to elucidate the evolution, epidemiology, and

taxonomy of CeMVs.


Samples and RNA Extractions

CeMV-infected tissue samples and a CeMV isolate were collected during seven cetacean

mortality events in the USA, Spain, and Ireland between 1993 and 2018 (summarized in Table 3-

1). RNA was extracted from frozen samples or cell culture supernatant using either a Maxwell 16

LEV simplyRNA Tissue Kit (ProMega Corp., Madison, WI) or a RNeasy Mini Kit (QIAGEN,

Valencia, CA) according to the manufacturer’s instructions. RNA extraction from a formalin-

fixed paraffin-embedded (FFPE) lung tissue was extracted using a FFPE RNeasy Kit (QIAGEN,

Valencia, CA).

RT-PCR Conditions and Sanger Sequencing of PCR Amplicons

The genomes of PMV-Tt-93-Alabama, PMV-Pp-88-North Ireland, DMV-Tt-12-

Florida(GOM), DMV-Sc-02-Canary Islands, DMV-Sc-11-Canary Islands, DMV-Ip-10-Hawaii,

and DMV-Lh-18-Hawaii were sequenced by designing overlapping pairs of primers (Table 3-2).

54

The primer sequences were based on the complete genome sequence of a CeMV (DMV-Sc-90-

92-Meditteranean; GenBank accession no. AJ608288) previously determined from a

Mediterranean striped dolphin. To acquire the complete CeMV genome sequences, RNA

samples were subjected to a series of overlapping one-step RT-PCR reactions using a OneStep

RT-PCR Kit (QIAGEN, Valencia, CA) as recommended by the manufacturer. The following

steps were used for the one-step RT-PCR reactions: one cycle of 50°C for 30 min for cDNA

synthesis and one cycle of 95°C for 15 min for denaturing, followed by 40 amplification cycles

of 94°C for 1 min for denaturing, annealing (temperature dependent upon the primer pair used)

for 1 min, elongation step at 72°C for 1 min, and a final elongation step at 72°C for 10 min. The

PCR products were then subjected to electrophoresis in 1% agarose gel stained with ethidium

bromide. DNA fragments of the expected sizes were purified using a QIAquick Gel Extraction

Kit (QIAGEN, Valencia, CA) and was sequenced in both directions using an ABI 3130 DNA

sequencer (Life Technologies, Carlsbad, CA).

Next-Generation Sequencing and Genome Annotation

RNA from the FFPE lung tissue of a bottlenose dolphin that stranded along the coast of

Alabama in 1993 (PMV-Tt-93-Alabama) and from the frozen lung tissue of a Fraser’s dolphin

that stranded in Hawaii in 2018 (DMV-Lh-18-Hawaii) were used to generate cDNA libraries

using a Nextera XT DNA Library Prep Kit (Illumina, San Diego, CA) according to the

manufacturer’s protocol. The libraries were sequenced using a 600-cycles version 3 Kit on an

Illumina MiSeq sequencer (Illumina, San Diego, CA). After trimming the resulting NGS data for

quality in CLC Genomics Workbench 10.1.1 software, the reads were assembled de novo using

SPAdes 3.10.1. (Bankevich et al., 2012). The assembled contigs were subjected to BLASTX

searches against a custom viral database, created from virus protein sequences retrieved from

UniProt Knowledgebase (UniProt Consortium, 2017), using CLC Genomics Workbench 10.1.1

55

software. Annotation of viral genomes was completed in CLC Genomics Workbench 10.1.1

software and gene function was predicted using BLASTP searches against the National Center

for Biotechnology Information (NCBI) GenBank non-redundant protein sequence database.

Phylogenetic Analysis: Alignment and Model Selection

A total of 18 fully sequenced morbillivirus genomes were selected (Table 3-3) to produce

a concatenated six gene tree. In addition, all available CeMV partial P gene sequences were

compared with the homologous sequences of five other morbillivirus species to produce a

separate gene tree (Table 3-3). Sequences were trimmed and edited in BioEdit (Hall, 1999) and

aligned in MAFFT 7.0 using default settings. MEGA 7.0 was used to determine best-fitting

model of evolution for the nucleotide (nt) concatenated six gene and partial P gene alignments.

Maximum Likelihood phylogenetic analyses were performed using the IQ-TREE server (Nguyen

et al., 2014) with 1000 non-parametric bootstraps to test clade robustness. The trees were edited

using FigTree v1.4.2 (Rambaut, 2014).

Sequence Identity Matrix

Identity matrices were generated for the nt sequences of the six paramyxovirus gene

alignments separately, the concatenated six gene alignment (14,057 characters), and partial P

gene alignment (389 characters) using the Sequence Demarcation Tool v1.2 (Muhire et al.,

2014). The genetic distance between Canine distemper virus (CDV) and Phocine distemper virus

(PDV), which are classified as different species, was used as a reference measure to define a new

species. CDV and PDV were selected because these two species are the two most closely related

species in the genus Morbillivirus. The nt sequence identity between these two species is 77.7%

for the concatenated 6 gene alignment (Figure 3-4) and 81.5% for the partial P gene alignment

(Figure 3-5). The nt sequence identity between CDV and PDV ranged between 71.5-80.5% for

the six separate gene alignments (Figures 3-6, 3-7, 3-8, 3-9, 3-10, 3-11).

56

Results

Genome Organization

Seven complete CeMV genomes were sequenced either by PCR and Sanger sequencing

or NGS. The genome length of the seven CeMV ranged from 15,647 to 15,702 bp. For each

CeMV genome, six open reading frames were predicted encoding the nucleoprotein,

phosphoprotein, matrix protein, fusion protein, hemagglutinin protein, and RNA polymerase

protein (Figure 3-1). The upstream sequence of the co-transcriptional editing V protein in DMV-

Lh-18-Hawaii was found to be identical to that of all other CeMVs

(AAAUCCAUUAAAAAGG), except for the last four amino acids of the protein (-DIPE) that

were found to be identical to Rinderpest virus (Kabete O strain, GenBank accession no.

X98291). The last four amino acids of all other CeMVs were -QWPF.

Phylogenetic Analysis

Both ML analyses produced well-supported trees (Figures 3-2, 3-3). DMV-Tt-12-Florida

(GOM) was found to branch together with the other CeMVs from the Gulf of Mexico (Figure 3-

2). The GOM CeMVs grouped together with CeMVs from the Mediterranean (DMV-Sc-90-92-

Meditterranean) including those from the Canary Islands (DMV-Sc-02-Canary Islands and

DMV-Sc-11-Canary Islands). All DMV strains of CeMV clustered together as a clade,

independent of geographic distribution or host species (Figure 3-3). The newly sequenced PMV

strains of CeMV (PMV-Tt-93-Alabama and PMV-Pp-88-North Ireland) formed a clade with a

PMV from the Netherlands (Figures 3-2, 3-3). The PWMV strains of CeMV formed a well-

supported clade (Figure 3-3). The ML analysis of the partial P gene sequences illustrated that

most Hawaiian strains (DMV-Ip-10-Hawaii et al.) clustered together at the base of the CeMV

tree (Figure 3-3). The DMV-Lh-18-Hawaii strain was supported as the most divergent CeMV in

57

the tree, branching between the clade formed by the Southern Hemisphere CeMV strains from

Brazil and Western Australia and the other morbillivirus species (Figures 3-2, 3-3).

Sequence Identity Matrix

Genetic analyses of the concatenated six genes alignment revealed nt identities ranging

from 71.5-99.9% between CeMV strains and 57.1-66.5% of CeMV strains to the other

morbillivirus species (Figure 3-4). Based on partial P gene sequences, the nt identities ranged

from 61.3-99.6% between CeMV strains and 45.2-59.2% between CeMVs and the other

morbillivirus species (Figure 3-5). The nt identity of the concatenated six gene analysis of DMV-

Lh-18-Hawaii to the other CeMV strains ranged from 71.5-72.0 %, while the nt identity between

the other CeMV strains to each other ranged from 84.0-100.0% (Figures 3-6, 3-7, 3-8, 3-9, 3-10,

3-11).

Discussion

Cetacean morbillivirus infections have emerged around the world as important causes of

mortality in a variety of cetacean species. Characterization of the complete genomes of seven

new CeMVs in this study contributes to a better understanding of the molecular epidemiology of

these viruses. The genome sequences of CeMV strains from the Canary Islands and of DMV-Tt-

12-Florida (GOM) that were sequenced in this study grouped together with those of other DMV

strains, despite their different host and geographic ranges. Our genetic analyses indicate that

similar DMV strains of CeMV circulate in the Mediterranean Sea and Gulf of Mexico, adding to

the known genetic diversity of CeMVs circulating in these disparate regions. CeMV epizootics

have been reported in the Gulf of Mexico since 1990s (Lipscomb et al., 1994, 1996; Litz et al.,

2014).) and other locations such as the Mid-Atlantic coastline of the United States, different

regions of the Mediterranean Sea, and Brazil (Aguilar et al., 1993; Duignan et al., 1996;

Fernández et al., 2008; Groch et al., 2018; Raga et al, 2008; Taubenberger et al., 1996; Van

58

Bressem et al., 1991). DMV is believed to be endemic in the Mediterranean basin and bottlenose

dolphins and striped dolphins are considered typical hosts (Centelleghe et al., 2017; Mazzariol et

al., 2013). The genetic relatedness between the two Canary Island DMV strains of CeMV

sequenced in this study, collected from animals that stranded nine years apart, supports the

hypothesis of prolonged DMV circulation in the region.

Both PMV strains sequenced in this study (PMV-Tt-93-Alabama and PMV-Pp-88-North

Ireland) formed a clade with PMV-Pp-90-Netherlands (GenBank accession no. KF650727).

PMV was previously reported in the North Sea, mainly in the United Kingdom and the

Netherlands (Kennedy et al., 1988, 1991; McCullough et al., 1991; Visser et al., 1993). The

PMV-Tt-93-Alabama sequence was obtained from a bottlenose dolphin that died in the 1993

Gulf of Mexico epizootic, and both DMV and PMV partial sequences were reported from

bottlenose dolphin samples in the 1987-1988 Mid-Atlantic epizootic (Taubenberger et al., 1996).

The complete genome sequencing of PMV from a bottlenose dolphin stranded in Alabama in the

1993 mortality event, determined in this study, confirms the PMV strain of CeMV as the likely

causative agent of this epizootic and confirms PMV circulates outside of the North Sea in species

other than harbor porpoises. Our findings may also support the hypothesis that certain cetaceans

(e.g., harbor porpoise and pilot whales) may spread strains of CeMV between disparate

geographic regions (e.g., Europe and North America). To our knowledge, this study represents

the first time a complete CeMV genome has been determined from a FFPE tissue using NGS.

We believe these techniques may be used in future studies to unravel the evolution and

epidemiology of other viruses circulating in wildlife populations.

The genomes sequenced in this study from animals that died in Hawaii were the most

divergent when compared to other CeMVs. The first CeMV reported in Hawaii was identified

59

from a Longman’s beaked whale that stranded in 2010, using a RT-PCR assay (West et al.,

2013). Jacob et al. (2016) determined the partial CeMV N and P gene sequences from 12

cetacean species and 15 individuals that stranded in Hawaii and analyzed their heterogenicity

compared to other CeMVs. These authors suggested the beaked whale morbillivirus (BWMV)

represents a novel Hawaiian CeMV strain. Using the nt identity between the CDV and PDV as

reference for species demarcation, our genetic analyses confirmed that most Hawaiian CeMVs

including DMV-Ip-10-Hawaii represent related strains. In contrast, the Hawaiian CeMV from a

Fraser’s dolphin (DMV-Lh-18-Hawaii) likely represents a new species within the genus

Morbillivirus. The discovery of a divergent CeMV raises important questions involving the

taxonomy and evolution of morbilliviruses infecting cetaceans around the globe as well as

questions related the pathogenicity, epidemiology, and the impact of this novel morbillivirus to

the health of Hawaiian cetaceans.

Based on the partial P gene analysis, the Brazilian and Western Australian CeMV

sequences clustered together forming a Southern Hemisphere clade (Figure 3-3). The Guiana

dolphin CeMV (GDMV) is the only CeMV described in the South Atlantic Ocean and was

recently associated with a mass cetacean die-off in Rio de Janeiro, Brazil (Groch et al., 2018).

The Western Australian CeMV (WAMV) was recently described from an Indo-Pacific

bottlenose dolphin (Tursiops aduncus) (Stephens et al., 2014). Interestingly, despite the

geographical distance, GDMV-Sg-17a-Brazil and WAMV-Ta-09a-Western Australia displayed

99.6% nt identity for the partial P gene (Figure 3-5). The mechanism of spread of these very

CeMV strains into different oceans is unknown; however, Duignan and colleagues (1995)

speculated that the long-finned pilot whale could play a role in the interspecies transmission of

CeMV strains. Our partial P gene analysis revealed these Southern Hemisphere CeMVs form a

60

phylogenetically distinct clade from other CeMVs (Figure 3-5). Further genomic-scale analyses

are needed to determine whether these Southern Hemisphere CeMVs represents a distinct species

in the genus Morbillivirus.

This study provides a much-needed update to CeMV genetics, provides a foundation for

future efforts aimed at developing improved CeMV molecular diagnostics, and a better

understanding of the temporospatial dynamics of these emerging cetacean viruses. We sequenced

the complete genomes of seven CeMVs associated with mortality events in the Gulf of Mexico,

Pacific Ocean (Hawaii), and the Mediterranean and North Seas. The complete genome of a

divergent and potentially novel species within the genus Morbillivirus from Hawaii suggests

much is still to be learned about the evolution, epidemiology, pathogenicity of morbilliviruses

circulating around the globe in cetaceans.

61

Table 3-1. Description of the CeMV samples sequenced in this chapter including: host species,

year and location sample was obtained, sample type, and associated reference. FFPE:

formalin-fixed paraffin embedded. GOM: Gulf of Mexico.

Species Year Location Sample Reference

Bottlenose dolphin

(Tursiops truncatus)

1993 Alabama, USA FFPE lung tissue This study

Harbor porpoise

(Phocoena phocoena)

1995 Coast of

Northern

Ireland

Viral isolate McCullough et al.,

1991

Striped dolphin

(Stenella

coeruleoalba)

2002 Canary Islands,

Spain

Frozen lung tissue This study

Striped dolphin 2011 Canary Islands,

Spain

Frozen lung tissue This study

Bottlenose dolphin 2012 Florida (GOM),

USA

Frozen lung tissue Cassle et al, 2016

Longman’s beaked

whale (Indopacetus

pacificus)

2013 Hawaii, USA Frozen lung tissue West et al., 2013

Fraser’s dolphin

(Lagenodelphis hosei)

2018 Hawaii, USA Frozen lung tissue This study

62

Table 3-2. List of the primers used in PCR reactions to sequence the genomes of seven CeMV including: DMV-Tt-12-Florida(GOM),

DMV-Ip-10-Hawaii, DMV-Sc-02-Canary Islands, DMV-Sc-11-Canary Islands, PMV-Pp-88-North Ireland. Primer Sequence (5’-3’) Combined with Amplicon size (bp) Genome sequenced

DMVP2490F

GGGCACAGGAGAGAGATCAG

DMVP3248R

758

DMV-Sc-02-Canary Islands


DMV-Tt-12-Florida(GOM)

DMV-Ip-10-Hawaii

PMV-Pp-88-North Ireland

DMVP3248R CCTATCATGCTTCGTTTGTGTTC

DMVP3139F

AGCTCTGCTGTGGGATTTGT

DMVM3967R

828




DMV-Ip-10-Hawaii


LBWMV2701F CCAGAAATCCCAGAGCAATG DMVM3967R 1266 DMV-Ip-10-Hawaii

DMVM3967R

TCCAGCATCTTTCTGGGAAC

DMVM3876F

CTTGACACTCCGCAAAGGTT

DMVM4505R

629




DMVM4505R

GCTCTGTTGATTCTGCTGGA

DMVF5343F

ACTGAAGGGCAAATCCATTG

DMVF6166R

823




DMV-Ip-10-Hawaii


DMVF6166R ATCACCCCCTTGACCTCTGA

63

Table 3-2. Continued Primer Sequence (5’-3’) Combined with Amplicon size (bp) Genome sequenced

DMVF6064F

AGAGTCGAGGGATAAAAGCAAA

DMVF6773R

709




DMV-Ip-10-Hawaii


DMVF6773R

GTACATTGCTCCTCCAAAGG

PMV6485F

GGCACCATAATTAGCCAGGA PMV7427R

PMV7904R

942

1419



PMV7427R

TGTTAACTTCTGGGGCATCC

PMV7904R TTCGTCCTCATCAATCACCA

PMV5endF ACCAGACAAAGCTGGCTAGGG PMV546R 525 PMV-Pp-88-North Ireland

PMV546R

ACCAGTTGGTTCCTCTGGTG

PMV2404F GAATGGATTCGGAGGAGACA PMV3420R 1016 PMV-Pp-88-North Ireland

PMV3420R TTGGCTCAATCCTGACTCCT

PMV14586F

CAACGGGAAACCAGAGGTAA

PMV15302R

716


PMV15302R

CAGCTGATTAAGCGCTCCTT

PMV15465F

ATAGGGTATGGCGCCCTTAT

PMV3endR

134 PMV-Pp-88-North Ireland

PMV3endR

ACCAGACAAAGCTGGGTATAG

PMV3517F

GGAAGATTGATCCCCCAAGT

PMV5659R

2142


PMV5659R CCACTCCCAATGCTACACCT

64


PMV6755F

CCTTTGGAGGAGCCAATGTA

PMV8381R

1626


PMV8381R

TATCAAGGGCCCGTAGTTTG

PMV10366F

TTTGCTGGGGTTAAGTTTGG

PMV12546R

2180


PMV12546R

GGGACAAAGAACCAGCCATA

PMV12752F

CTGCTGTTAGGATTGCCACA

PMV13518R

766


PMV13518R

AGCTGCACATTGCCCTAAGT

PMV13994F

CTCCTGGAGGATTGAATTGG PMV14176R

182


PMV14176R

TAATCCCCTTGGCCTTACCT

DMVP1808F

AGGAGCAGGCCTATCATGTC

DMVP2603R

795




DMV-Ip-10-Hawaii


DMVP2603R

GGTGCACTTGACTCTGACGA

DMVL13419F

GCTGCTTTAATAGGAGACGATGA

DMVL14117R

698




DMV-Ip-10-Hawaii


DMVL14117R CGATACGAACCCAGGATCAAC

65


MV5end

ACCAGACAAAGCTGGSTARGG

DMVN1123R

1102 DMV-Tt-12-Florida(GOM)

DMV-Ip-10-Hawaii


MV5end ACCAGACAAAGCTGGSTARGG DMVN520R 500 DMV-Sc-02-Canary Islands



DMVN520R

TCCTCCCCTGCTTGAATAGA

DMVL10176F

GTTATCAACTACGAGACTATGATGAA

DMVL10867R

691





DMVL10867R

CCACCTTCACCTCTATGGTAATC

DMV10700F

TGCCAAGTCATTGCAGAGAA

DMV11500R

800





DMV11500R

TTTCAAATGATGCCCTATATCGT

DMVL12147F

GCGGATAGCCATGAGGTAGA

DMVL12895R

748


DMVL12895R

TTGGTGGATGTCGATATTGG

DMVL15338F

CGCTAGGAAGTATTGGGGGTA

PMV3end

556 DMV-Sc-02-Canary Islands



DMV-Ip-10-Hawaii


DMVH8225F CAAGCATTGTGCAAGAGTAGACC LBWMV9237R 1012 DMV-Ip-10-Hawaii

66


DMVH8225F

CAAGCATTGTGCAAGAGTAGACC

DMVL9263R 1038 DMV-Sc-02-Canary Islands




DMVL9263R

TAGATTTACCACATTCCCAATTTC

DMVH7375F

GTTTAAGATCATCGGAGACGAAGT

DMVH8284R

909 DMV-Sc-02-Canary Islands




DMVH8284R

CAATGGCTCCCAATCAGAACT

DMVH7375F

GTTTAAGATCATCGGAGACGAAGT

LBWMV9237R 1862 DMV-Ip-10-Hawaii

LBWMV9237R

ACCACATTCCCAACTTCCAG

DMVN465F

GGCGCTAATATGGAGGATGA

DMVN1123R

658




DMV-Ip-10-Hawaii


DMVN1123R

ACTCCCATTGCATAGCTCCA

DMVN1014F TACCAACAGATGGGCGAGAC

DMVP1875R DMV-Sc-02-Canary Islands



DMV-Ip-10-Hawaii


DMVP1875R ATCGGGCGGATTTTCTCT

67


DMVM4350F

GCTGTTCTTCAGCCATCAGTC

DMVF5405R

1055




DMV-Ip-10-Hawaii


DMVF5405R GGCACTTCCTGTCCCAACTAT

DMVL9044F

GGAGTCCATCTCAATCAACCA

DMVL9751R

707




DMV-Ip-10-Hawaii


DMVL9751R

CCTTCCACTACATCGCAGTACAT

DMVL9618F

CCTTCATTTGTGCATGGTGA

DMVL10252R

634





DMVL10252R

TGCCTATCACGGTACCCATT

DMVL9618F

CCTTCATTTGTGCATGGTGA

LBWMV10387R

769

DMV-Ip-10-Hawaii

LBWMV10387R

ACTCAAGGGCATGAAACACC

DMVL10802F

ACTCCACACACTAGCGGTTTC

DMVL11581R

779




DMV-Ip-10-Hawaii


DMVL11581R AGTGATTGAGAGATCAGCATTCC

68


DMVL11466F

CAGCGATTGCACGATATAGG

DMVL12285R

819





DMVL12285R

TAGTATCGAGCATCCCTGCAA

DMVL14067F

AGAGGGGCGGTTAAACAGAT

DMVL14824R

757





DMVL14824R

CCTTGAACTAAGTCCCCGCT

DMV14450F

TCGAGTCTAGATCTGGGCAAA

DMV15450R

1000


DMV15450R

AACCATTCTCTCCGTTGGAT

DMV14700F

ATAAGGACCTAGTTGAAAAGCTGGA

DMV15500R

800


DMV14595F

CCAGAGGTGACATGGGTAGG

DMV15500R

905


DMV15500R

AAGAGTCCGTATTAAGACCCTTTC

DMV4412F

TGATCAAGGGTTATTCAAGATTC

DMV5037R

650


DMV5037R

CCAATGGATGACTGGTGGTC

DMVL12828F

GCGAGGCAGAGAGCTAACAT

DMVL13529R

701





DMVL13529R GCCCAATTAATAGCTGCACA

69


DMVL14705F

GGACCTAGTTGAAAAGCTGGA

DMVL15487R

782





DMVL15487R

AAGACCCTTTCGGATTTGGA

DMV5000F CAGGCTCCATCTCCAGGAC DMV5600R 600 DMV-Sc-02-Canary Islands



DMV-Ip-10-Hawaii

DMV5600R

AGGCTTAATATTCTTTGTTATCACTGT

DMV1600F

CAGCGTCAGACTTTGTATTTTCA

DMV2580R

980


DMV2580R

GCCCATCTTGATTCTTGAGC

DMV6610F

TCTCGGCCCAGCTATATCAC

DMV7580R

970


DMV7580R

CTGCGATGTGGTTGCAGTA

DMV8110F

CGATTGATTCTTCTATCGAGAAGTT

DMV8395R

285


DMV8395R

ATGGATTATCAAGGGTCCGTA

LBWMV3876F

CTCGACACTCCGCAAAGATT

LBWMV4557R

681

DMV-Ip-10-Hawaii

LBWMV4557R

CGTTACAAGCCTGAAATGAGAAG

LBWMV5847F

TCCATGCACCAGTTGTCTTG LBWMV6234R

387

DMV-Ip-10-Hawaii

LBWMV6234R TAGCCGTGGGTTGCTACATA

70


LBWMV6672F

GGGAGTGCTCTCACGAAACT

DMVH7426R

754


DMV-Ip-10-Hawaii

DMVH7426R

TGTTAACTTCTGGGGCATCC

LBWMV6672F

GGGAGTGCTCTCACGAAACT LBWMV9237R

2565

DMV-Ip-10-Hawaii

LBWMV9237R ACCACATTCCCAACTTCCAG

LBWMV9578F TCAAATCAGCGACCCATACA

LBWMV10932R 1354 DMV-Ip-10-Hawaii

LBWMV10932R AGTCTGTCTGCGTCGCCTAT

LBWMV14000F

AGGGCTGAATTGGAATGATG

LBWMV15580R

1580

DMV-Ip-10-Hawaii

LBWMV15580R CAGGGGTAGGAATCCGTTTT

LBWMV4864F

GGAATCAAAGCTCCAACCAA

LBWMV5209R

345

DMV-Ip-10-Hawaii

LBWMV5209R

GCTCGATTGCAGGTTTTCTC

LBWMV7401F

GATTACGAATGCCCCAGAAA

LBWMV8354R

953

DMV-Ip-10-Hawaii

LBWMV8354R

CAAGCTCTGGACCTGAATCC

LBWMV13324F

CAACGGGGCAGTTATACCAT

LMBWMV14224R

900

DMV-Ip-10-Hawaii

LMBWMV14224R

AATCGTGCAGCATCTTCCTT

LBWMV11483F AGGGCATCATTTGAAAGCAA LBWMV13419R 1936 DMV-Ip-10-Hawaii

LBWMV13419R TGTCATCATCTCCTATTAAAGCAA

71


LBWMV12146F

TTAGATTGGGCCAGTGATCC

LBWMN13931R

1785

DMV-Ip-10-Hawaii

LBWMN13931R

TTCTCTTATCGACGGGCAGT

LBWMV12146F

TTAGATTGGGCCAGTGATCC

LBWMV12982R

836

DMV-Ip-10-Hawaii

LBWMV12982R CTCCTCCAAGGTCACATTAGC

72

Table 3-3. Virus names, GenBank accession numbers and references for the morbilliviruses used

in genetic and phylogenetic analyses.

Species name (virus abbreviation) Accession

number

Reference

1,2,3,4,5Mediterranean Dolphin morbillivirus

(DMV-Sc-90-92-Meditteranean)

AJ608288 Rima et al., 2005

1,2,3,5Gulf Dolphin morbillivirus (DMV-Sc-10-

Florida(GOM))

KU720623 Fauquier et al., 2017

1,2,3,5Gulf Dolphin morbillivirus (DMV-Tt-11-

Louisiana)


1,2,3,5Gulf Dolphin morbillivirus (DMV-Tt-11-

Mississippi)


1,2,3,5Gulf Dolphin morbillivirus (PMV-Tt-93-

Alabama)

This study This study

1,2,3,4,5Gulf Dolphin morbillivirus (DMV-Tt-12-

Florida(GOM))

This study Cassle et al., 2016

1,2,3,4,5Dolphin morbillivirus (DMV-Sc-02-

Canary Islands)


1,2,3,5Dolphin morbillivirus (DMV-Sc-11-

Canary Islands)


1,2,3,4,5Porpoise morbillivirus (PMV-Pp-88-

North Ireland)

This study McCullough et al., 1991

1,2,3,4,5Hawaii Dolphin Morbillivirus (DMV-Ip-

10-Hawaii)

This study West et al., 2013

1,2,3,4,5Hawaii Dolphin Morbillivirus (DMV-Lh-

18-Hawaii)


3(DMV-Sc-12-Portugal) KT878658 Bento et al., unpublished 3(DMV-Sc-12-Spain) KP835987 Bento et al., unpublished 3(DMV-Sc-13-Portugal) KT878656 Bento et al., unpublished 3(DMV-Sc-07-Portugal) KT878657 Bento et al., unpublished 3(DMV-Sc-09- Spain) KT878660 Bento et al., unpublished 3(DMV-Bp-16-Denmark) KY681807 Jo et al., unpublished 3(DMV-Dd-13-Portugal) KP836003 Bento et al., unpublished 3(DMV-Sc-11-Canary Islands) KJ139454 Sierra et al., 2014a 3,4(DMV-Sc-02-Canary Islands) KJ139451 Sierra et al., 2014a 3 (DMV-Sc-12-Spain) KT878661 Bento et al., unpublished 3,4(DMV-Dd-12-Portugal) KP835999 Bento et al., unpublished 3(DMV-Tt-13-Florida(GOM)) KU720622 Fauquier et al., 2017 3(DMV-Sc-14-Portugal) KT878659 Bento et al., unpublished 3,4(DMV-Sc-11-Portugal) KP835983 Bento et al., unpublished

Strains used in: 1Genome annotation; 2Concatenated six gene tree; 3Partial P gene tree; 4Sequence identity matrix

(partial P gene); 5Sequence identity matrix (6 concatenated gene and individual gene analyses)

73

Table 3-3. Continued


number

Reference

3(DMV-Sc-12-Spain) KC572861 Rubio-Guerri et al.

unpublished 3,4(DMV-Sc-12-Spain) KP835991 Bento et al., unpublished 3(DMV-Tt-05-Canary Islands) KF695110 Sierra et al., 2014b 3,4(DMV-Sc-09-Canary Islands) KJ139453 Sierra et al., 2014a 3,4(DMV-Sc-07-Portugal) KP835995 Bento et al., unpublished 3,4(DMV-Sc-07-Canary Islands) KJ139452 Sierra et al., 2014a 3,4(DMV-Bp-13-Italy) KR337460 Mazzariol et al., 2012 3(DMV-Zc-15-Italy) KX237511 Centelleghe et al., 2017

3(DMV-Gme-07-Spain) EU039963 Esperon et al., unpublished 3(DMV-Bp-13-Italy) KU977450 Mazzariol et al., 2016 3(DMV-Sc-07-Spain) HQ829973 Bellière et al., 2011b 3(DMV-Gme-07-Spain) HQ829972 Bellière et al., 2011b 3(DMV-Sc-11-Spain) JN210891 Rubio-Guerri et al., 2013a 3,4(DMV-Kb-Taiwan) AF333347 Yang et al., 2006 3(DMV-Sc-90-92-Mediterranean) Z47758 Bolt et al., 1995 3,4(DMV-La-11-Netherlands) KC888945 van Elk et al., unpublished 3,4(DMV-La-11-Netherlands) KC888946 van Elk et al., unpublished 3,4(DMV-Gg-08-Canary Islands) KX512307 Sierra et al., unpublished 3,4(PWMV-Gme-00-New Jersey) AF200817 Taubenberger et al., 2000 3,4(PWMV-Gma-96-Canary Islands) FJ842381 Bellière et al., 2011b 3(PWMV-Gma-15-Canary Islands) KT006289 Sierra et al., 2016 3(PWMV-Gma-15-Canary Islands) KT006290 Sierra et al., 2016 3,4(PWMV-Gma-15-Canary Islands) KT006291 Sierra et al., 2016 3(PMV-Pp-90-Netherlands) KF650727 van de Bildt et al.,

unpublished 3,4(DMV-Sl-01-Hawaii) KM460048 Jacob et al., 2016 3(DMV-Sc-14-Hawaii) KM460055 Jacob et al., 2016 3(DMV-Sl-13-Hawaii) KM975648 Jacob et al., 2016 3(DMV-Md-10-Hawaii) KM460052 Jacob et al., 2016 3(DMV-Zc-08-Hawaii) KM460049 Jacob et al., 2016 3(DMV-Gg-13-Hawaii) KM460054 Jacob et al., 2016 3(DMV-Sb-11-Hawaii) KM460053 Jacob et al., 2016


(partial P gene); 5Sequence identity matrix (Concatenated six gene and individual gene analyses)

74

Table 3-3. Continued


number

Reference

3(DMV-Kb-00-Hawaii) KM460047 Jacob et al., 2016 3(DMV-Mn-98-Hawaii) KM460046 Jacob et al., 2016 3(DMV-Sc-10-Hawaii) KM460051 Jacob et al., 2016 3(DMV-Pm-11-Hawaii) KJ482570 West et al., 2015 3,4(DMV-Sl-09-Hawaii) KM460050 Jacob et al., 2016 3(GDMV-Sg-10-Brazil) KF711855 Groch et al., 2014 3,4(GDMV-Sg-17a-Brazil) MG845551 Groch et al., 2018 3(GDMV-Sg-17b-Brazil) MG845552 Groch et al., 2018

3,4(WAMV-Ta-09a-Western Australia) unpublished Stephens et al., 2014 3(WAMV-Ta-09b-Western Australia) unpublished Stephens et al., 2014 1,2,3,4,5Measles virus (MeV) AF266287 Parks et al., 2001 1,2,3,4,5Rinderpest virus (RPV) JN234010 Jeoung et al., 2012 1,2,3,4,5Small ruminants virus (PPRV) KJ867542 Muniraju & Parida,

unpublished 1,2,3,4,5Phocine distemper morbillivirus (PDV) KC802221 de Vries et al., 2013 1,2,3,4,5Canine distemper morbillivirus (CDV) AF164967 Wiederkehr et al.,

unpublished 1,2,3,4,5Feline morbillivirus (FeMV) JQ411016 Woo et al., 2012 1,2,3,4,5Feline morbillivirus (FeMV) AB924120 Park et al., 2014


(partial P gene); 5Sequence identity matrix (Concatenated six gene and individual gene analyses)

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Figure 3-1. Genome organization of the CeMVs sequenced in this study. The genes are shown as

colored arrows that are drawn to scale. For the P gene, the smaller green arrows

represent the region of C protein (leaky scanning) and V protein (RNA editing). See

Table 3-2 for the virus abbreviations.

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Figure 3-2. Cladogram depicting the relationship of the CeMV strains within the genus

Morbillivirus, based on the concatenated nucelotide sequence alignment of the six

genes. Numbers at each node represent the bootstrap values from the Maximum

Likelihood analysis. Branch lengths are based on the number of inferred substitutions,

as indicated by the scale. See Table 3-2 for the virus abbreviations.

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Figure 3-3. Cladogram depicting the relationship of the CeMV strains within the genus

Morbillivirus, based on the nucleotide sequence alignemnt of the partial P gene.

Numbers at each node represent the bootstrap values from the Maximum Likelihood

analysis. Branch lengths are based on the number of inferred substitutions, as

indicated by the scale. Bootstrap values <70% were removed. Bolded strains indicate

they are new sequences generated in this study. See Table 3-2 for the virus

abbreviations.

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Figure 3-4. Sequence identity matrix illustrating the genetic distance of the CeMVs to other members of the genus Morbillivirus based

on the concatenated nucleotide sequence alignment of the six genes: nucleocapsid protein, phosphoprotein, matrix protein,

fusion protein, hemagglutinin protein, and RNA polymerase. Values are expressed as a percentage of identity. See Table 3-

2 for the virus abbreviations.

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on the nucleotide sequence alignment of the partial phosphoprotein gene. Values are expressed as a percentage of identity.

See Table 3-2 for the virus abbreviations.

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on the nucleotide sequence alignment of the nucleocapsid gene. Values are expressed as a percentage of identity. See Table

3-2 for the virus abbreviations.

81


on the nucleotide sequence alignment of the phosphoprotein gene. Values are expressed as a percentage of identity. See

Table 3-2 for the virus abbreviations.

82


on the nucleotide sequence alignment of the matrix gene. Values are expressed as a percentage of identity. See Table 3-2

for the virus abbreviations.

83


on the nucleotide sequence alignment of the fusion gene. Values are expressed as a percentage of identity. See Table 3-2

for the virus abbreviations.

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Figure 3-10. Sequence identity matrix illustrating the genetic distance of the CeMVs to other members of the

genus Morbillivirus based on the nucleotide sequence alignment of the hemagglutinin gene. Values are expressed as a

percentage of identity. See Table 3-2 for the virus abbreviations.

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Figure 3-11. Sequence identity matrix illustrating the genetic distance of the CeMVs to other members of the

genus Morbillivirus based on the nucleotide sequence alignment of the RNA polymerase gene. Values are expressed as a

percentage of identity. See Table 3-2 for the virus abbreviations.

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CHAPTER 4

CHARACTERIZATION OF A NOVEL CIRCOVIRUS FROM A STRANDED LONGMAN’S

BEAKED WHALE (INDOPACETUS PACIFICUS)

Introduction

Members of the family Circoviridae are small non-enveloped viruses with icosahedral

nucleocapsids of approximately 17 nm in diameter that enclose a single-stranded circular DNA

(ssDNA) genome ranging in size from 1.7-2.3 kilobases (Herbst and Willems, 2017). The

ambisense genome contains two open reading frames (ORFs) that encode the capsid and

replication-associated proteins (Biagini et al., 2012). The intergenic region between these genes

contains a conserved nonamer at the apex of a stem-loop motif where replication of the ssDNA

genome is initiated and then completed by rolling circle mechanism (Biagini et al., 2012).

According to a recent taxonomic revision of the family Circoviridae, the family is divided into

the genera Circovirus and Cyclovirus (Breitbart et al., 2017). The genus Circovirus currently

includes 29 recognized species from fish (2), bats (9), birds (12), human (1), pigs (3), dog (1),

and chimpanzee (1) (Breitbart et al., 2017). The genus Cyclovirus currently includes 45

recognized species from bats (16), invertebrates (9), mammals (8), and humans (12) (Breitbart et

al., 2017).

The genus Circovirus includes agents responsible for diseases of veterinary significance

such as the porcine circoviruses. Porcine circovirus 1 (PCV-1) was first identified in 1974 in a

persistently infected permanent pig kidney cell line (PK15) and considered nonpathogenic to

pigs (Tischer et al., 1974). Porcine circovirus 2 (PCV-2) was first isolated in 1997 from pigs

with chronic wasting disease syndrome and reported to cause various disease syndromes in pigs

(e.g., post weaning multisystemic wasting syndrome, porcine dermatitis and nephropathy

syndrome, proliferative and necrotizing pneumonia, reproductive failure, and enteritis),

collectively known as porcine circovirus-associated disease (PCVAD). PCVAD is one of the

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most economically important diseases in the swine industry and occurs most commonly in

piglets of 5-18 weeks of age (Tischer et al., 1974). The most common microscopic lesions

observed with PCVAD include mild to severe lymphocyte depletion with an accompanying

granulomatous inflammation within lymphoid tissues (Segalés, 2012). Thus, it is believed that

PCV-2 is immunosuppressive, which predisposes infected pigs to secondary infections. Other

microscopic lesions include necrotizing bronchiolitis, interstitial nephritis, non-suppurative to

necrotizing or fibrosing myocarditis of fetuses, and hepatic congestion in fetuses (Segalés, 2012).

PCV-2 infected cells may present with cytoplasmic and basophilic botryoid (grape-like)

inclusion bodies.

Lorincz and colleagues (2011) characterized the first circovirus in an aquatic vertebrate,

e.g. diseased barbel fry (Barbus barbus). Their metagenomic analyses of the moribund fish

recovered two complete genomes, barbel circovirus 1 (BaCV1) and barbel circovirus 2 (BaCV2),

but linkage of these viruses to the clinical disease was not established (Lorincz et al., 2011). A

second fish circovirus, the European catfish circovirus (EcatfishCV), was detected in spawning

sheetfish (Silurus glanis) experiencing high mortality in Lake Balaton, Hungary in 2011 (Lorincz

et al., 2012). Circoviruses are generally considered to be immune-suppressive; so, it was

speculated that the EcatfishCV plays a role in the development of diseases, especially during the

stress of spawning (Lorincz et al., 2012).

A number of studies have sought to understand the mechanism by which circoviruses

induce host immunosuppression. Shibahara et al. (2000) reported PCV induces apoptosis in B

lymphocytes, which leads to B lymphocyte depletion and therefore suppression of humoral

immunity. Abadie and colleagues (2001) demonstrated a significantly higher percentage of

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apoptotic lymphocytes in the bursa of Fabricius of birds infected with Pigeon circovirus than in

uninfected birds.

In situ hybridization (ISH) is a molecular method used to localize specific nucleic acid

sequences, DNA or RNA, within tissue sections. Older ISH methods have a high degree of

technical complexity, but low sensitivity and specificity (Wang et al., 2012). However,

RNAscope® technology is a novel in situ hybridization approach for detection of target RNA

within tissue sections. This approach is a major advance in RNA ISH technology and hinges on a

proprietary probe design to amplify target-specific signals and reduced background signal from

non-specific hybridization. In recent years, ISH has been used to detect circoviruses, including

porcine circoviruses 1-3 with the improvement of its technology. Positive staining of PCV3 has

been detected in lymphocytes infiltrating the alveolar spaces of the lungs, in macrophages in the

alveolar wall, and in histiocytes of the lymphoid follicles in lymph nodes (Kedkovid et al., 2018;

Kim et al., 2018).

In 2010, a juvenile male Longman’s beaked whale (Indopacetus pacificus) stranded in

Maui, Hawaii. Histopathological findings included lymphoplasmacytic periglomerulitis, mild

lymphoid depletion, encephalitis, pulmonary edema, thyroid gland atrophy, and bilateral

hypertrophy of the adrenal cortex (West et al., 2013). Diagnostic testing using conventional PCR

assays detected a novel cetacean morbillivirus and a novel alphaherpesvirus (West et al., 2013).

Herein, we report the genetic characterization of a novel circovirus also detected in the tissues of

this Longman’s beaked whale using a high-throughput sequencing approach, followed by

detection of this novel circovirus in tissues by ISH using RNAscope® technology.

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Samples and Genome Sequencing

Tissue samples of spleen, muscle, left ventricle, mesenteric lymph node (LN), left

adrenal, liver, lung, cerebrum, cerebellum, scapular LN, and mediastinal LN from a juvenile

Longman’s beaked whale that stranded in Hawaii (West et al., 2013) were sent frozen on dry ice

from the Hawaii Pacific University to the Wildlife and Aquatic Veterinary Disease Laboratory

(Gainesville, FL). DNA from lung was extracted using a Qiagen DNeasy Kit (Qiagen, Valencia,

CA) following the manufacturer’s instructions. A DNA library was prepared by using an

Illumina Nextera XT DNA Library Preparation Kit and sequenced by using a 600-cycle V3 kit

on an Illumina MiSeq sequencer (Illumina, San Diego, CA). The paired-end reads were quality

trimmed and assembled de novo in CLC Genomic Workbench V7 using default settings.

BLASTX analysis of the assembled contigs was conducted against the GenBank non-redundant

(nr) protein sequence database provided by the National Center for Biotechnology Information

(NCBI). A 1,434 bp contig was identified with highest identity (91/174 amino acid (aa); 52%) to

a replication-associated protein of canine circovirus (GenBank accession no. AML03152)

previously sequenced from the mesenteric lymph node of a dog (Canis lupus familiaris).

Conventional and inverse PCRs, with primers designed from the known replication-associated

gene sequence (Table 4-1), were implemented to amplify the complete genome sequence of the

beaked whale circovirus (BWCV). The Sanger sequence reads were then assembled into a single

contiguous sequence and open reading frames (ORFs) were predicted using CLC Genomic

Workbench V7. Gene functions were determined based on BLASTP searches against the NCBI

GenBank nr protein sequence database.

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Phylogenetic and Genetic Analyses

The phylogenetic analysis was performed on the full-length predicted aa sequences of the

replication-associated and capsid genes for 30 circoviruses, including BWCV. The aa sequences

were aligned for each gene in MAFFT 7 using default parameters (Katoh and Standley, 2013).

The final dataset contained 379 and 415 aa characters (including gaps) for the replication-

associated and capsid genes, respectively. Maximum Likelihood phylogenetic analysis was

performed in IQ-TREE (Nguyen et al., 2014) with the Bayesian information criterion to

determine the best model fit and 1000 non-parametric bootstraps to test the robustness of the

clades (Nguyen et al., 2014). The phylogenetic trees were then edited using FigTree v1.4.2

(Rambaut, 2014). For the genetic analysis, complete genome sequence of the BWCV was

compared to 29 circoviruses in the Sequence Demarcation Tool v1.2 (Muhire et al., 2014) with

the MAFFT 7 alignment option implemented.

PCR Detection of BWCV in Longman’s Beaked Whale Tissues

A primer pair specific to BWCV (Table 4-1) was designed using the Primer3 program

(Rozen and Skaletsky, 2000). Conventional PCR assays were conducted on all 11 available

frozen tissues (i.e., spleen, muscle, left ventricle, mesenteric LN, left adrenal, liver, lung,

cerebrum, cerebellum, scapular lymph node, mediastinal lymph node) retrospectively to

determine the distribution of BWCV in Longman’s beaked whale tissues. The 30 µl BWCV PCR

cocktail consisted of 0.15 µl of Platinum Taq DNA Polymerase (Invitrogen), 3.0 µl of 10× PCR

Buffer, 1.2 µl of 50 mM MgCl2, 0.6 µl of 10 mM dNTPs, 1.5 µl of 20 µM forward and reverse

primers, 17.55 µl of molecular grade water, and 4.5 µl of DNA template. The following thermal

cycling conditions were used for the BWCV conventional PCR assay: one cycle of 94°C for 5

min for initial denaturation, followed by 40 amplification cycles of 94°C for 30 sec for

denaturation, 56°C for 30 sec for annealing, 72°C for 30 sec for elongation, and a final

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elongation cycle of 72°C for 5 min. PCR products were then subjected to electrophoresis in 1%

agarose gel stained with ethidium bromide. Fragments of the expected size (400 bp) were

purified using a QIAquick PCR Purification Kit. The concentration of purified DNA was

quantified fluorometrically using a Qubit® 3.0 Fluorometer and dsDNA BR Assay Kit. Purified

DNA was sequenced in both directions on an ABI 3130 platform (Applied Biosystems).

In Situ Hybridization (ISH)

Tissue sections were fixed in 10% neutral buffered formalin for 24 hours for RNAscope®

ISH. RNAscope® target probes for the BWCV and the Longman’s beaked whale cytochrome B

(positive control) were designed and provided by Advanced Cell Diagnostics, and the negative

control probe was designed to a sequence of the bacterial gene DapB. Formalin-fixed paraffin-

embedded tissue sections were deparaffinized in xylene, followed by dehydration in an ethanol

series. Chromogenic in situ hybridization was performed on a Leica BOND RX Fully Automated

Research Stainer (Leica Biosystems, Buffalo Grove, Illinois) using RNAscope® technology

(Wang et al., 2012) and probes designed by Advanced Cell Diagnostics (Newark, California).

Blocks of formalin-fixed paraffin-embedded tissues were sectioned at 4um and mounted on

Fisherbrand SuperFrost Plus glass slides (Fisher Scientific, Pittsburgh, PA) for use in a single-

plex automated RNAscope® assay (Anderson et al., 2016). The entire assay was performed on

the Leica BOND and consisted of four steps, i.e. pretreatment, hybridization, signal

amplification, and detection, with pretreatment, amplification, and detection divided into a series

of sequential reactions (Advanced Cell Diagnostics, Inc., 2018), with slight modifications to

standard conditions (e.g., Anderson et al., 2016). Pretreatment was carried out by sequential

deparaffinization, target retrieval (i.e., 15 min incubation at 95°C using Leica Epitope Retrieval

Buffer 2), protease digestion (20 min at 40°C), and endogenous enzyme block. This was

followed by target probe hybridization (120 min at 42°C). After hybridization, signal

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amplification was performed through a series of reactions, which was then followed by fast red

chromogenic development and hematoxylin staining. The resultant histologic sections were

interpreted using an Olympus BX53 light microscope.

The following tissues were tested using the RNAscope® technique: diaphragm, liver,

five lymph nodes (not specified which lymph nodes), lung, pericardium, oral mucosa and tongue,

adrenal gland, testis, aorta, intestine, stomach, spleen, heart, and skeletal muscle. Histologic

sections from each paraffin block were tested in a three-slide experimental design wherein one

slide received the negative control (DapB) probe, a second slide received the positive internal

control (LBW cytochrome B) probe, and a third slide received the test (BWCV) probe.

Results

Genomic Sequence Annotation

The complete circular BWCV genome is 1,894 bp with a G+C content of 46.5%. Two

ORFs were identified, on complementary strands in opposite orientation, which encode the

capsid and replication-associated proteins (Figure 4-1a). The BWCV contains a stem-loop motif

between the intergenic region of the two ORFs which consists of palindromic 15 bp stem, a 11

bp open loop for the initiation of rolling-circle replication, and a 9 bp long conserved nonamer

(5’-TAGTATTAC-3’) on the apex of the open loop (Figure 4-1b). The replication-associated

protein contains 3 rolling circle replication motifs at the N-terminus including, motif I [FTVNN;

aa 10-14], motif II [PHLQG; aa 45-49], and motif III [YCSK; aa 84-87]. The superfamily 3

helicase motifs located at the C-terminus of the BWCV replication-associated protein include a

Walker-A motif [GPPGVGKS; aa 163-170], a Walker-B motif [IIDDF; aa 200-204], and motif

C [ITSN; aa 240-243]. The N-terminus of BWCV capsid protein contains a 41-aa-long arginine

(R) rich stretch (RRYRRHRPYFRRRRRYHGYRKRFNRRRRFRKPRLFHFRFER) starting at

residue 2.

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Phylogenetic and Genetic Analyses

The ML analyses based on the aligned aa sequences of replication-associated and capsid

genes, using the best fit models LG+I+G4 and PMB+I+G4, respectively, produced well-

supported trees with the BWCV grouping as the closest relative to the canine circovirus isolate

214 (Figure 4-2, 4-3; GenBank accession no. JQ821392). The nucleotide identity ranged from

50.9-56.7% to other circoviruses and the highest identity was observed with bat associated

circovirus 2 isolate XOR7 (Figure 4-4; GenBank accession no. KC339249).

Detection of BWCV in Longman’s Beaked Whale Tissues

A single band of 400 bp was observed for all 11 frozen tissues samples (i.e., spleen,

muscle, left ventricle, mesenteric LN, left adrenal, liver, lung, cerebrum, cerebellum, scapular

LN, mediastinal LN) and the resulting edited Sanger sequences were identical to the BWCV

genome determined through NGS. This confirms all tissue samples from the Longman’s beaked

whale were positive for BWCV.

In Situ Hybridization (ISH)

No reactivity was observed when the negative control probe (DapB) was applied to tissue

sections of diaphragm, liver, five lymph nodes (not specified which lymph nodes), lung,

pericardium, oral mucosa and tongue, adrenal gland, testis, aorta, intestine, stomach, spleen,

heart, and skeletal muscle from the Longman’s beaked whale (Figure 4-5a, 4-6a, 4-7a, 4-8a).

Host cells in histologic sections of all tissues except spleen and skeletal muscle were labeled by

the LBW cytochrome B probe; histologic sections could not be interpreted further because of an

absence of labeling by the internal control probe (i.e. LBW cytochrome B probe), which may

have been a result of the autolysis of those tissues. RNAscope® technology revealed circovirus

nucleic acid labeling of cells in the remaining tissues. The heart and stomach had labeling of

individual cells in the endothelium and vascular smooth muscle, occasionally in the myocardial

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fibers, and in the glandular epithelium. The oral mucosa and tongue had BWCV nucleic acid

labeling in cells in the subepithelial connective tissue, possibly macrophages within the

submucosa and white blood cells in the tongue vasculature. There was intense labeling in some

areas of the lymph nodes within individual lymphocytes and monocytes (Figure 4-5e, f). In the

diaphragm the presence of BWCV nucleic acid was seen in cells in the interstitium of the muscle

and occasionally cells of the blood vessels (Figure 4-6e, f). BWCV nucleic acid was detected in

the cells of the zona glomerulosa and blood vessels of the capsule of the adrenal gland. The liver

had intense staining in the cytoplasm of many hepatocytes (Figure 4-7e, f). BWCV was detected

in the cytoplasm of biliary epithelium (Figure 4-8e, f). Lung was positive for BWCV, with probe

labeling of macrophages in the lumen of alveoli and also cells in blood vessels. Lastly, in the

testis labeling for BWCV was observed in cells of the interstitium, in the spermatogenetic

epithelium of seminiferous tubules, and smooth muscle cells in blood vessels.

Discussion

The current study provides the first complete genome sequence of a circovirus from a

marine mammal. In addition, the presence of BWCV nucleic acids in the Longman’s beaked

whale tissues was verified retrospectively by conventional PCR and ISH. The BWCV displayed

a typical circovirus genome organization including two proteins (i.e., replication-associated and

capsid) and a conserved nonamer (5’-TAGTATTAC-3’) within a stem-loop motif. The BWCV

replication-associated protein, containing rolling cycle replication motifs at the N-terminus and

superfamily 3 helicase motifs at the C-terminus, mediates the initiation and termination of rolling

cycle replication (Ilyina & Koonin, 1992). The BWCV capsid protein displayed an arginine (R)-

rich N-terminus which is believed to involve DNA binding and nuclear localization (Sarker et

al., 2016).

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A methionine (ATG) start codon was not identified for the BWCV capsid protein and we

postulate the use of alternative start codon (GTT; valine) at nucleotide position 1843 in the

BWCV genome. Future transcriptomic analyses are needed to verify the start codon used by

BWCV. The use of alternative start codons is commonplace in avian circoviruses (Mankertz et

al., 2000; Phenix et al., 2001; Stewart et al., 2006; Todd et al., 2001, 2007). Porcine circovirus 3

also utilize an alternative start codon (valine), which may be similar to the situation in BWCV

(Palinski et al., 2017). Although the significance of the usage of an alternative start codon in

circoviruses remains to be determined, in eukaryotes and RNA viruses they act as a regulatory

mechanism for proteins with key cellular functions (Kearse and Wilusz, 2017) and translation of

multiple proteins from alternative reading frames (Firth and Brierley, 2012), respectively.

The BWCV genome sequence reported herein permitted phylogenetic and genetic

analyses that support its inclusion as a novel species within the family Circoviridae.

Phylogenetic analyses based on aa sequences of replication-associated and capsid genes revealed

the BWCV branches most closely to canine circovirus isolate 214 (GenBank accession no.

JQ821392). One criterion for the species demarcation of circoviruses is the genome-wide

pairwise nucleotide identity of <80% represent circoviruses from different species and >80%

represent different isolates within a species (Rosario et al., 2017). The genome-wide pairwise

identity of BWCV as compared to 29 other circoviruses revealed highest nucleotide identity of

56.7% identity. Given the unique phylogenetic position and sequence divergence of the BWCV,

we propose the formal species designation of beaked whale circovirus (BWCV) to be considered

for approval by the International Committee on Taxonomy of Viruses.

The clinical significance and pathogenicity of BWCV is unknown, but its genetic

proximity to canine circovirus isolate 214 (GenBank accession no. JQ821392) and the

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association of canine circoviruses in dogs with vasculitis and hemorrhage (Li et al., 2013),

hemorrhagic diarrhea (Anderson et al., 2017), and hemorrhagic enteritis (Decaro et al., 2014)

highlight the importance of further investigation into the potential role of BWCV in the stranding

of the juvenile Longman’s beaked whale in Maui, Hawaii in 2010. Circoviruses are known to

infect lymphoid tissues and cause persistent immunosuppression in hosts (Maclachlan and

Dubovi, 2010). Likewise, histopathological analysis of the stranded Longman’s beaked whale

revealed mild lymphoid depletion (West et al., 2013), all 11 tissues tested positive for BWCV by

conventional PCR, and ISH confirmed the presence of BWCV nucleic acids in all of those

tissues, with the exception of two which could not be interpreted (skeletal muscle and spleen). A

consistent feature of the labeling with the BWCV probe between tissues was labeling of the cells

of blood vessels. Taking into consideration that all the tissues tested positive by PCR, this

systemic infection could be explained by this ISH observation. BWCV could be an opportunistic

agent replicating in lymphoid tissues without clinical significance, or it alone may have

contributed to the disease, or it may have worked in concert with beaked whale morbillivirus

(Chapter 3) and the alphaherpesvirus previously reported in this stranded Longman’s beaked

whale juvenile (West et al., 2013).

The BWCV is the first circovirus to be detected in a cetacean, expanding the known host

range of circoviruses into marine mammals. The role of the BWCV in disease and the stranding

of the juvenile Longman’s beaked whale could not be determined in this study due to the co-

infection with two other viruses, a morbillivirus and an alphaherpesvirus (West et al., 2013).

Additional techniques, such as transmission electron microscopy and qPCR can help elucidate

and correlate the intense staining with visual evidence and quantitative numbers. Hawaii is home

for many resident marine mammal populations including some that are critically endangered.

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Further research to investigate the prevalence and health impacts of circoviruses in marine

mammals in Hawaii is warranted.

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Table 4-1. Primers used to amplify the complete genome of the beaked whale circovirus

(BWCV) and screen Longman’s beaked whale tissue samples for BWCV.

Primer name Sequence Comments

BWCVInv-F1 AGACTCTCCTTTCTCTCGGC Amplification

of BWCV

BWCVInv-R1 CGGCATCATCGGGGATTTTC

BWCVConv-F2 CCCGACAGAAGCAGATGAAG

BWCVConv-R2 TTTAGACTTGCCCACCCCAG

BWCV-F CTTCAGATTCCCCGTCAAGA Tissue

screening BWCV-R GTCTCCCCACAATGGTTCAC

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Table 4-2. Circoviruses used in the phylogenetic and genetic analyses (adapted from Rosario et

al., 2017).

Species name Abbreviation Accession #

Barbel circovirus BarCV GU799606

Bat associated circovirus 1 BatACV-1 JX863737

Bat associated circovirus 2 BataCV-2 KC339249

Bat associated circovirus 3 BatACV-3 JQ814849

Bat associated circovirus 4 BatACV-4 KT783484

Bat associated circovirus 5 BatACV-5 KJ641727

Bat associated circovirus 6 BataCV-6 KJ641724



Beak and feather disease

virus BFDV AF31129

Canary circovirus CaCV AJ301633

Canine circovirus CanineCV JQ821392

Chimpanzee associated

circovirus ChimpACV

GQ404851

Duck circovirus DuCV AY228555

European catfish

circovirus EcatfishCV JQ011377

Finch circovirus FiCV DQ845075

Goose circovirus GoCV AF418552

Gull circovirus GuCV DQ845074

Human associated

circovirus 1 HuACV GQ404856

Mink circovirus MiCV KJ020099

Pigeon circovirus PiCV AF252610

Porcine circovirus 1 PCV-1 AF012107 Porcine circovirus 2 PCV-2 AF027217

Porcine circovirus 3 PCV-3 KT869077

Raven circovirus RaCV DQ146997

Starling circovirus StCV DQ172906

Swan circovirus SwCV EU056309

Zebra finch circovirus ZfiCV KP793918

Beaked whale circovirus BWCV This study

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Figure 4-1. Genome schematics of the beaked whale circovirus (BWCV). A) Two open reading

frames (ORFs) were annotated including the capsid protein (Cap) and the replication-

associated protein (Rep). B) The stem-loop motif of the BWCV consist of

palindromic 15 bp stem and a 11 bp open loop that includes a 9 bp long conserved

nonamer (5’-TAGTATTAC-3’) highlighted in yellow.

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Figure 4-2. Phylogram depicting the relationship of the beaked whale circovirus (black arrow)

from Longman’s beaked whale (Indopacetus pacificus) to representatives of the

genus Circovirus based on their aligned capsid amino acid sequences (415 characters

including gaps). Bootstrap values of >70% were indicated at the nodes and the branch

lengths represent the number of inferred substitutions as indicated by the scale. See

Table 4-2 for circovirus abbreviations.

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Figure 4-3. Phylogram depicting the relationship of the novel beaked whale circovirus (black

arrow) from Longman’s beaked whale (Indopacetus pacificus) to representatives of

the genus Circovirus based on their aligned replication-associated amino acid

sequences (379 characters including gaps). Bootstrap values of >70% were indicated

at the nodes and the branch lengths represent the number of inferred substitutions as

indicated by the scale. See Table 4-2 for circovirus abbreviations.

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Figure 4-4. Genetic comparison of the full genome of the novel BWCV from Longman’s beaked whale (Indopacetus pacificus) to 29

accepted type species in the genus Circovirus. Values are expressed as a percentage of nucleotide identity. See Table 4-2

for circovirus abbreviations.

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Figure 4-5. RNAscope® in situ hybridization (ISH) results from lymph node (LN). A) LN DapB

negative control (10x objective). B) LN DapB negative control (60x objective). C)

Longman’s beaked whale (LBW) cytochrome B positive control (red) within the LN

(10x objective). D) LBW cytochrome B positive control (red) (60x objective). E) LN

ISH indicates circovirus nucleic acid labeling (red) within mononuclear cells (10x

objective). F) LN ISH indicates circovirus nucleic acid labeling (red) in mononuclear

cells (60x objective).

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Figure 4-6. RNAscope® in situ hybridization (ISH) results from the diaphragm. A) DapB

negative control probe (10x objective). B) Blood vessel within the interstitium with

DapB negative control probe (60x objective). C) Longman’s beaked whale (LBW)

cytochrome B positive control within muscle cells(red) (10x objective). D) LBW

cytochrome B positive control probe labeling of cells of a blood vessel. Staining

occurs in the endothelium of the blood vessel (red) (60x objective). E) Diaphragm

ISH indicates circovirus nucleic acid labeling (red) (10x objective). F) ISH indicates

circovirus nucleic acid labeling (red) in the endothelium and smooth muscle fibers of

the blood vessel (red) (60x objective).

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Figure 4-7. RNAscope® in situ hybridization (ISH) results from the liver. A) DapB negative

control probe (10x objective). B) Hepatocytes are not labeled by the DapB negative

control probe (60x objective). C) Longman’s beaked whale (LBW) cytochrome B

positive control probe (red) (10x objective). D) LBW cytochrome B positive control

probe gives mild staining (red) of the cytoplasm of hepatocytes (60x objective). E)

Liver ISH indicates circovirus nucleic acid labeling (red) (10x objective). F) ISH

indicates circovirus nucleic acid labeling (red) in hepatocytes (60x objective).

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Figure 4-8. RNAscope® in situ hybridization (ISH) results from the liver. A) DapB negative

control probe (10x objective). B) Bile ductule without labeling by the DapB negative

control probe (60x objective). C) Longman’s beaked whale (LBW) cytochrome B

positive control probe within the liver (red) (10x objective). D) LBW cytochrome B

positive control probe gives punctate stain precipitate in the cytoplasm of biliary

epithelial cells (red) (60x objective). E) ISH indicates circovirus nucleic acid labeling

(red) of hepatocytes (10x objective). F) ISH indicates circovirus nucleic acid labeling

(red) of the biliary epithelium and cytoplasm of hepatocytes (red) (60x objective).

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CHAPTER 5

TOXOPLASMA GONDII IN STRANDED HAWAIIAN CETACEANS

Introduction

Toxoplasma gondii is an obligate coccidian parasite within the family Sarcocystidae that

infects a range of warm-blooded animals including domesticated species (e.g., cats, cattle, sheep,

pigs), wildlife (e.g., birds, rodents, marine mammals), and humans. Members of the family

Felidae are the definitive host in which T. gondii undergoes sexual reproduction within the

enterocytes of infected cats resulting in formation of oocysts that sporulate and are released into

the feces where they contaminate the environment and can infect other cats or the

aforementioned intermediate hosts. Intermediate hosts typically become infected by ingestion of

sporulated oocysts carrying infectious sporozoites that invade enterocytes and develop into

tachyzoites that multiply asexually and expand the population of the parasite within the host

through the bloodstream. Tachyzoites can produce disease if the immune system of the host

becomes weak and they are also known to infect the fetus across the placenta resulting in

congenital toxoplasmosis (Tenter et al., 2000). After the initial tachyzoite phase which may or

may not result in disease, T. gondii persists in the intermediate host as cysts within the

musculature or nervous tissues known as bradyzoites. The definitive host (i.e., felid) can then

become infected if it consumes prey (i.e., intermediate host) carrying tissue cysts (i.e.,

bradyzoites). Felid feces can carry large numbers of oocysts that sporulate within 1-5 days and

can remain viable for several months resulting in environmental contamination of marine coastal

waterways via sewage systems, storm water drainage, and freshwater runoff (Fayer et al., 2004;

Lindsay and Dubey, 2009; Miller et al., 2002a).

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Given there are no pathognomonic clinical signs associated with T. gondii infections, a

diagnosis is typically made using bioassays, serological tests, and molecular methods (reviewed

in Liu et al., 2015). Varying molecular methods for detecting T. gondii in infected tissue samples

have been developed including conventional PCR, real-time PCR, loop-mediated isothermal

amplification. Determination of T. gondii the genetic lineage has been accomplished by

microsatellite analysis, random amplification of polymorphic DNA (PCR-RADP), restriction

fragment length polymorphism (PCR-RFLP), high-resolution melting PCR, and multilocus

sequence typing (MLST) (Liu et al., 2015). Of these methods, MLST has the greatest resolution

of genetic lineages and can be used in downstream phylogenetic analyses.

Comprehensive phylogenetic analyses based on MLST data including coding (exons) and

non-coding (introns) loci have resulted in the discrimination of 12 T. gondii lineages in Europe,

North America, and South America (Khan et al., 2007, 2011). Clonal lineages 1, 2, and 3 are

present in Europe and North America (Howe and Sibley, 1995). Toxoplasma gondii lineages 4,

5, 6, 8, 9, and 10 have primarily been characterized from cases involving humans, domestic cats,

and pigs in South American (Khan et al., 2007, 2011). Toxoplasma gondii lineage 7 includes two

human cases and lineage 11 includes two cougar (Puma concolor) cases from the Pacific

Northwest of North America. Genotyping of T. gondii isolates from California sea otters

revealed two additional novel genotypes, designated as Type X and Type A (Miller et al., 2004;

Sundar et al., 2008), that were later grouped together as lineage 12 (Khan et al., 2011).

Marine mammal species inhabiting coastal marine ecosystems serve as sentinels for both

public and aquatic ecosystem health. Disease resulting from T. gondii infection has been reported

in a wide range of marine mammals worldwide including sirenians (Attademo et al., 2016;

Bossart et al., 2012; Buergelt and Bonde, 1983; Greenland et al., 2005; Smith et al., 2016),

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pinnipeds (Measures et al., 2004; Migaki et al., 1977; Rengifo-Herrera et al., 2012; Roe et al.,

2017), cetaceans (Di Guardo et al., 2009; Dubey et al., 2008; Gonzales-Viera et al., 2013; Jardine

and Dubey, 2002; Mazzariol et al., 2012; Pretti et al, 2010; Resendes et al, 2002) and Southern

sea otters (Hanni et al., 2003; Kreuder et al., 2003; Miller et al., 2002b; Shapiro et al., 2015;

Thomas and Cole, 1996). The Southern sea otter appears especially prone to T. gondii infection

and it is a significant cause of mortality that has impeded the recovery of this endangered

species. The seroprevalence of T. gondii infected Southern sea otters was approximately 38% in

live animals and 52% in freshly dead, beach cast animals along the California coast from 1998-

2004 (Conrad et al., 2005).

Buergelt and Bonde (1983) reported the first case of systemic toxoplasmosis in a Florida

manatee (Trichechus manatus latirostris). A second case of disseminated toxoplasmosis was

later reported by Smith and colleagues (2016). However, the same authors reported a low T.

gondii seroprevalence in two Florida manatee populations. In Puerto Rico, toxoplasmosis may be

an emerging disease in the West Indian manatee (Trichechus manatus manatus). Gross,

microscopic, and ultrastructural examination of four West Indian manatee carcasses from August

2010 to August 2011 supported toxoplasmosis as the cause of the strandings (Bossart et al.,

2012). In Australia, disseminated toxoplasmosis has been reported in dugongs (Dugong dugon)

(Greenland et al., 2005; Owen et al., 2012). The dugongs displayed granulomas in several organs

with numerous bradyzoites observed in the liver and mesenteric lymph nodes that tested positive

for T. gondii by immunohistochemistry (Greenland et al., 2005; Owen et al., 2012).

A systemic toxoplasmosis case was recently reported in a free-ranging New Zealand sea

lion (Phocarctos hookeri) (Roe et al., 2017). The female was found dead and had T. gondii-

associated lesions in the pelvic muscles, caudal spinal cord, and spinal nerves. These

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neuromuscular lesions were believed to have resulted in hindlimb paresis thereby reducing the

animal’s mobility (Roe et al., 2017). Although the cause of death was attributed to crushing and

asphyxiation by a male sea lion, the T. gondii infection was believed to have played a significant

role as a result of the decreased mobility of the young female. Other cases of T. gondii in

stranded and managed marine mammals have been genetically characterized as lineage 2

including cases involving striped dolphin (Stenella coeruleoalba), bottlenose dolphin (Tursiops

truncatus), Hector’s dolphin (Cephalorhynchus hectori), walrus (Odobenus rosmarus), and New

Zealand fur seal (Arctocephalus forsteri) by PCR-RFLP analysis (Donahoe et al., 2014; Dubey et

al., 2007, 2008, 2009; Roe et al., 2013).

The first reported case of toxoplasmosis in a Hawaiian marine mammal was in a wild

male spinner dolphin (Stenella longirostris), with lesions observed in the liver, brain, and

adrenals (Migaki et al., 1990). The adrenal glands were severely affected, with numerous large,

discrete areas of coagulative necrosis in the cortex, a diffuse encephalitis was observed with

numerous small foci of gliosis associated with tissue cysts, and the liver displayed numerous

small randomly distributed necrotic foci containing mononuclear leukocytic infiltrates (Migaki et

al., 1990). Tachyzoites and bradyzoites were observed within parenchymal cells.

Additional cases of fatal toxoplasmosis in Hawaiian marine mammals include reports in

Hawaiian monk seals (Neomonachus schauinslandi), a critically endangered species (Barbieri et

al., 2016; Honnold et al. 2005). To characterize the T. gondii genetic lineage infecting monk

seals, Honnold and colleagues (2005) used an PCR-RFLP approach targeting the SAG2 gene and

determined it as lineage 3. Barbieri et al. (2016) studied protozoal-related mortalities in the

Hawaiian monk seal between 2001 and 2015. They found eight cases of protozoal-related

mortalities, seven of these were attributed to toxoplasmosis and the eighth case was attributed to

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vertical transmission of T. gondii in which co-infection with Sarcocystis neurona was suspected

(Barbieri et al., 2016). The T. gondii lineage(s) involving these monk seal cases as well as the

aforementioned single spinner dolphin case were not determined.

Toxoplasmosis is an emerging as a significant threat to endangered Hawaiian wildlife

including birds. The Hawaiian Islands have the highest per capita number of endangered birds in

the USA (Dobson et al., 1997). Fatal disseminated toxoplasmosis has been described from the

‘Alala (Corvus hawaiiensis), nene (Branta sandvicensis), red-footed booby (Sula sula), and

Erckel’s francolin (Francolinus erckelii) (Work et al., 2000, 2002). Recently, Work et al. (2015)

examined 300 carcasses of the endangered nene between 1992-2013 to better understand causes

of death to aid in its management and recovery. Infectious/inflammatory diseases were the third

most common cause of death with many cases resulting from T. gondii infections (Work et al.,

2015). Four nene that succumbed to T. gondii were determined to be harboring two novel genetic

lineages (ToxDB #261 and #262), based on the results of multlocus PCR-RFLP analysis, that

have not been detected in any other region (Work et al., 2016).

In the fall 2015, a suspected case of disseminated toxoplasmosis in a stranded Hawaiiain

spinner dolphin was suspected based on microscopic lesions observed in various internal tissues.

Histological lesions included a myocarditis with extracellular and intracellular protozoal cysts

and tachyzooites, adrenalitis with intralesional protozoal cysts and tachyzooites, and

bronchointerstitial pneumonia with intra-endothelial trachyzooites. In this chapter, we confirmed

the T. gondii infection based on histopathology, immunohistochemistry, and conventional nested

PCR assay. The genetic lineage was then determined for positive samples using a previously

described MLST approach.

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Methods and Materials

Postmortem Examination and PCR Screening for Toxoplasma gondii

A total of 25 Code 1, 2, and 3 stranded spinner dolphin tissue samples, regardless of

gross pathology or histopathology findings, were retrieved from the Hawaii Pacific University

stranding archive to obtain an unbiased estimate of the prevalence of T. gondii from 1997-2016.

In addition, 29 stranded individuals representing 15 species, which included: Blainville’s beaked

whale (Mesoplodon densirostris), Cuvier’s beaked whale (Ziphius cavirostris), false killer whale

(Pseudorca crassidens), Longman’s beaked whale (Indopacetus pacificus), spotted dolphin

(Stenella frontalis), rough-toothed dolphin (Steno bredanensis), Risso’s dolphin (Grampus

griseus), sperm whale (Physeter macrocephalus), pygmy sperm whale (Kogia breviceps), short-

finned pilot whale (Globicephala macrorhynchus), bottlenose dolphin, killer whale (Orcinus

orca), striped dolphin, melon-headed whale (Peponocephala electra), and humpback whale

(Megaptera novaeangliae) were also included in this retrospective study.

A suite of ten frozen tissue types per individual were selected for the detection of T.

gondii by nested PCR, although not all individuals had each of the ten tissue type. The tissues

screened by nested PCR were brain (cerebrum, cerebellum), left or right ventricle, left or right

lung, lymph nodes (mesenteric, anal and, colonic), spleen, liver, and left or right adrenal glands.

DNA was extracted from frozen tissues using a Qiagen DNeasy Kit (Qiagen, Valencia, CA)

according to the manufacturer’s instructions. Each individual was processed on separate days to

reduce the risk of cross contamination. The samples were screened for T. gondii by nested PCR

as described in Chapter 2.

Histopathology and Immunohistochemistry

PCR positive tissue samples that was previously fixed in 10% neutral buffered formalin,

were then routinely processed, embedded in paraffin, sectioned at 5 µm, and stained with

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hematoxylin and eosin for histological evaluation. Formalin-fixed paraffin-embedded (FFPE)

tissue sections of the brain, liver, lung, adrenal gland, lymph nodes and heart were processed for

IHC alongside appropriate positive and negative controls at the California Animal Health and

Food Safety Laboratory Systems (Davis, CA). Immunohistochemistry (IHC) using a rabbit

primary polyclonal antiserum against T. gondii and Neospora caninum was performed according

to Miller and colleagues (2001).

Multilocus Sequence Typing, Genetic, and Phylogenetic Analysis

Cetacean tissue DNA determined to be positive by PCR were then used to determine the

genetic lineage through additional PCR and Sanger sequencing of coding (exons) and non-

coding (introns) regions of the T. gondii genome using a previously described multilocus

sequence typing method (Khan et al., 2007, 2011). The T. gondii exon sequences including dense

granule protein 6 (GRA6) and surface antigen 1 (SAG1) and the intron sequences including

microneme protein 1 and 2 (MIC1 and 2), and beta-tubulin 1 (BTUB1). Nucleotide sequences

from 66 T. gondii strains, representing 12 known genetic lineages originating from animal and

human samples from Europe, North America, and South America were retrieved from the

National Center for Biotechnology Information GenBank sequence database to be included in the

phylogenetic analyses as previously described (Khan et al., 2011). These outgroup sequences

were aligned with the ingroup sequences (i.e., T. gondii nucleotide sequences from Hawaiian

cetaceans) in MAFFT 7.0 using default settings. Maximum Likelihood analyses were performed

in MEGA 7.0 with 1000 non-parametric bootstraps to test the robustness of the clades.

Results

PCR Screening for Toxoplasma gondii

Spinner dolphin tissue samples screened by nested PCR assay resulted in nine positives

(out of 138 samples), all from the same individual (KW2015013). Samples from the other 15

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Hawaiian cetacean species tested negative (0/164). Following the removal of primer sequences,

the round one and two reactions resulted in 514 bp and 392 bp consensus sequences,

respectively. BLASTN searches of the resulting spinner dolphin T. gondii sequences (329 bp)

against the NCBI non-redundant database revealed 25 T. gondii sequences with 100% coverage

and 100% sequence identity.

Postmortem Examination, Histopathology, and Immunohistochemistry

The carcass of a 184 cm, 60.7 kg, fresh (code 2), adult male spinner dolphin

(KW2015013) was recovered from Waimea, Hawaii on 19 October 2015 and brought to Hawaii

Pacific University in Kaneohe, Hawaii for necropsy. The carcass was examined and determined

to be in poor body condition (i.e., thin). The animal had multiple healed partial cookie cutter

shark bite wounds with necrotic centers and irregular healing patterns. Mild bruising was noted

on the right mandible and blood was observed in the oral cavity (Figure 5-1a). A 6 cm x 5 cm,

healed scar and fresh bruising were noted just below the left eye and above the oral commissure

(Figure 5-1b). Musculature along the right rib cage was pale, edematous, and gelatinous. No

signs of skin trauma or broken ribs were noted. The main stomach was empty, the pyloric

chamber had a small ulcer (3 mm), and no fecal matter was present in the colon. The left lung

appeared consolidated, there was prominent lymph nodes associated with the left lung and

sublumbar region. The liver was small and firm and displayed an abnormal bluish-silver

appearance on the serosal surface. The serosal surface of the kidney appeared bluish-silver as

compared to the pink parenchyma and pale-yellow foci were noted in the center of many

reniculi. The spleen was irregular with protruding masses (5 mm-2 cm diameter) noted (Figure

5-1d). A probable urinary stone was observed in the bladder, a 2 x 1 cm area of mucosal

hemorrhage was noted, and the urine was a dark amber color.

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Histological examination of the respiratory system revealed diffuse, moderate

submucosal vascular congestion in the trachea. A bronchointerstitial pneumonia was noted with

moderate, diffuse histiocytic and lymphoplasmacytic infiltrates (Figure 5-2a). Within the

pulmonary interstitium, occasional epithelial cells appeared clumped (syncytial cells), scattered

bacterial cocci were observed, and a few vessels displayed suspect tachyzoites within endothelial

cells (Figure 5-2b). Multiple lymph nodes exhibited multifocal lymphocytolysis with associated

lymphadenitis, hemorrhage, fibrin exudation, and occasional tachyzoites within affected regions.

Multifocally, there was splenic capsular expansion by basophilic mineral and hemosiderophages

(Figure 5-2c). The liver had patchy hepatocellular necrosis characterized by hepatocellular

swelling, nuclear pyknosis, and hypereosinophilia with associated necrotic cellular debris,

clumps of golden pigment (bilirubin), and fibrin (Figure 5-2d). The ventricle displayed a few foci

of cardiomyocyte necrosis with associated fibrin exudation. A few cardiomyocytes contained 15-

20 µm protozoal cysts containing numerous tachyzoites (Figure 5-3a & b). In the cerebrum and

cerebellum, there was patchy effacement of the neuropil by clusters of macrophages, neutrophils,

and elongate microglial cells and occasional astrocytic aggregates (astrocytic scars) were present

(Figure 5-4a, b, c). There was multifocal adrenal cortical necrosis. Necrotic corticocytes

displayed eosinophilic cytoplasms, pyknotic nuclei, and were present within pools of

erythrocytes, macrophages, neutrophils, fibrin, and cellular debris (Figure 5-5a). There were free

tachyzoites and intracellular protozoal cysts present (Figure 5-5b).

Discrete areas of positive IHC staining for T. gondii was observed in all six tissues,

especially the brain, liver, and adrenal gland (Figure 5-6a, b, c). Although, the amount of T.

gondii was not heavy, they were observed at the edges of necrotic zones. N. caninum slides were

negative, but with some non-specific staining was observed.

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Multilocus Sequence Typing, Genetic and Phylogenetic Analysis

The Maximum Likelihood analysis of the concatenated 5 loci data set (2,835 aligned

nucleotide characters) produced a tree (Figure 5-7) separating a T. gondii clade involving strains

from North American and Europe from a T. gondii Brazilian clade as previously described (Khan

et al. 2011). The Hawaiian spinner dolphin T. gondii strain (KW2015013) was well supported as

the sister group to a clade of North American and European T. gondii strains, derived primarily

from human cases, including lineages 1 (e.g., ENT, GT1, MOR, VEL, RH) and 7 (e.g., JGM,

CAST). T. gondii strains from lineages 1 and 7 exhibited between 4-7 differences to the spinner

dolphin strain as compared to the other strains from other lineages that exhibited between 10-50

differences across the multilocus alignment. The clade formed by the Hawaiian spinner dolphin,

lineage 1 and lineage 7 strains displayed a cytosine at position 1996 in the alignment as

compared to all other T. gondii strains displaying a guanine. The Hawaiian spinner dolphin strain

displayed a cytosine at position 1573 in the alignment as compared to all other T. gondii strains

that displayed a thymine.

Discussion

The molecular assays, histopathology, and immunohistochemistry findings are consistent

with disseminated toxoplasmosis in 4% (1/25 animals) of code 1, 2, and 3 stranded spinner

dolphins analyzed in this study from the Hawaii Pacific University stranding archive. After 25

years, this is the second report of disseminated toxoplasmosis in a stranded Hawaiian spinner

dolphin. Similar to our case, the first report involved an adult male spinner dolphin that stranded

in Haleiwa, Oahu and was diagnosed with fatal disseminated toxoplasmosis (Migaki et al.,

1990). Both cases involved dolphins in poor body condition with little or no ingesta or feces

found in their digestive tracts. Microscopic lesions in the 1990 case attributed to T. gondii

included the adrenals, brain, and liver (other tissues were not available for microscopic

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examination in Migaki et al. (1990). In our case, the disseminated T. gondii infection was

confirmed in the adrenals, brain, heart, liver, lung, lymph nodes, and spleen. Both cases involved

bronchopneumonias with bacteria observed upon microscopic examination (our case) or cultured

(Aeromonas sp. and Pseudomonas putrefaciens) from the lungs (Migaki et al., 1990).

Monk seals are the only other marine mammal species in Hawaii in which T. gondii has

been confirmed (Barbieri et al. 2016; Honnold et al., 2005). The first monk seal case reported in

2005 found prominent lesions in multiple tissues (adrenals, brain, diaphragm, lymph nodes, and

spleen), and was characterized by necrosis with variable numbers of extracellular and

intracellular protozoal tachyzoites (Honnold et al., 2005). Based on the characteristics of the

lesions, Honnold and colleagues (2005) suggested that the monk seal had recently acquired the T.

gondii infection that had then spread systemically and overwhelmed the adult male seal that was

otherwise in good body condition. During the acute phase of infection, there was the formation

of localized, small tissue cysts within the brain (Honnold et al., 2005). Similar cases of

disseminated toxoplasmosis occurring from 2001-2015 have been reported in adult and aborted

Hawaiian monk seals (Barbieri et al., 2016).

Native and introduced Hawaiian bird species are another group negatively impacted by T.

gondii in the Hawaiian Islands. Postmortem examination, histopathology,

immunohistochemistry, bioassay, serological assays, and PCR-RFLP assays have been utilized

to diagnose T. gondii in ‘Alala (Work et al., 2000), the red-footed booby, Erckel’s francolin

(Work et al., 2002), and nene (Work et al., 2016). These avian species exhibited microscopic

findings consistent with T. gondii infection including tissue cysts and tachyzoites in different

organs (e.g., brain, spleen, liver, adrenals, skeletal muscle), nonsuppurative inflammation,

multifocal necrosis in different organs, and consequently several of them died from disseminated

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toxoplasmosis. Work and colleagues (2016) argued that the high rate of exposure (based on

serosurveys) and the low rate of mortality (4%) in nene suggests T. gondii does not have severe

population impacts in nene. Similarly, this prevalence is identical to what we observed in

stranded spinner dolphins analyzed in this study from the Hawaii Pacific University stranding

archive. In contrast to nene, the highly endangered ‘Alala appears highly sensitive to T. gondii

and it has been argued to be a significant threat to reintroduction programs in Hawaii (Work et

al., 2000). Work and colleagues (2016) also point out that a low prevalence of clinically

significant disease may underestimate other indirect impacts such as when T. gondii infected

animals are at greater risk of anthropogenic induced trauma (e.g., road-killed; Hollings et al.,

2013). To date, the prevalence of T. gondii induced clinical disease in wild spinner dolphins

appears low (2/26 = 7.7%); however, the seroprevalence and any indirect effects of exposure in

wild spinner dolphin populations remain unknown.

The source(s) of the T. gondii infections in Hawaiian mammals and birds is unknown.

Work and colleagues (2000, 2002, 2016) suggested the source of infection in Hawaiian birds was

through ingestion of oocysts from cat feces or through ingestion of a transport host (e.g., birds

ingesting insects harboring oocysts after grazing on contaminated cat feces). However, in a

marine environment, the source of infection can be acquired from ingestion of land-based water

runoff and sewage dispersal contaminated with cat feces. Cetaceans lead an aquatic life that

allows little opportunity for contact with terrestrial animals. On the other hand, pinnipeds, such

as monk seals, have an amphibious lifestyle in which they have more opportunities to acquire T.

gondii through contact with infected intermediate hosts and felines. The spinner dolphin

populations in Hawaii have a marked difference in the use of their marine environment. They

commonly use inshore habitat for daytime rest and social interactions, while at night they spend

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their time foraging in deep waters. Migaki et al. (1990) postulated that although the route of

infection in the spinner dolphin was unknown, it is possible it became infected with T. gondii

after eating garbage containing feline fecal material or eating dead or crippled birds infected with

the parasite.

Toxoplasma gondii genotypes in Hawaiian wildlife have been determined by PCR-RFLP

analysis using between 1 and 10 genetic markers including: SAG1, (SAG2 (5’3’ SAG2 and

alt. SAG2), SAG3, BTUB, GRA6, c22-8, c29-2, L358, PK1, and Apico (Dubey et al., 2011;

Honnold et al., 2005; Verma et al., 2015; Work et al., 2016). Honnold and colleagues (2005)

used the SAG2 RFLP profile to determine the disseminated toxoplasmosis in an adult monk seal

was caused by a genotype III (i.e., lineage 3) T. gondii. Work and colleagues (2015)

characterized two new T. gondii genotypes (ToxoDB PCR genotypes #261 and #262) from four

nene carcasses. Finally, Verma and colleagues (2015) characterized a genotype III (i.e., lineage

3) and a new T. gondii genotype (ToxoDB PCR genotype #249) from two Hawaiian mouflon

sheep (Ovis ammoni). The disseminated toxoplasmosis cases involving the first reported stranded

spinner dolphin and more recent cases in monk seals have not been genetically characterized

(Barbieri et al., 2016; Migaki et al., 1990).

Our MLST approach provided greater resolution of the genetic lineages as compared to

the PCR-RFLP approach and was used in downstream phylogenetic analyses in which the

spinner dolphin T. gondii strain formed a distinct branch most closely related to strains from

human toxoplasmosis cases from North America and Europe. Feral cats in Hawaii are believed

to have originated from Hawaii, but appear to have diverged from the original source population

and perhaps the T. gondii parasites they harbor have as well (Hansen et al., 2007; Work et al.,

2016) as indicated by the novel genotypes characterized in Hawaiian birds, sheep, and spinner

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dolphin. Future research is needed to determine whether the spinner dolphin PCR-RFLP profile

matches any of those previously determined from Hawaiian wildlife (birds, mouflon sheep, and

monk seal). Additionally, we hope to determine the lineage of other T. gondii from Hawaiian

wildlife using the greater discriminating power of the MLST approach we used in this study.

Understanding the genetic lineage(s) circulating in Hawaiian wildlife should facilitate a better

understanding of host-parasite dynamics.

The markers used in the phylogenetic analysis included GRA6, SAG1, BTUB1, MIC2

intron 1 and 2, while the other loci had poor phylogenetic signal and were not useful for

phylogenetic analysis (data not shown). Our phylogenetic analysis revealed that spinner dolphin

T. gondii is a North American strain and groups together with Clade 1 and 7. The intermediate

hosts for T. gondii in clade 1 and 7 are human and goat.

Over the last two decades, there appears to be a rise in T. gondii infections in Hawaiian

wildlife. The increase of infections may be due to an increase in surveillance efforts to the

Hawaiian wildlife, improved diagnostic approaches, or an actual increase in disease prevalence.

Many of the Hawaiian marine mammal species exhibit unique habitat structure and are

considered separate populations than more cosmopolitan species. Nearshore species, like

Hawaiian spinner dolphins, may be at increased risk from environmental contamination of

marine coastal waterways via sewage systems, storm water drainage, and freshwater runoff.

Future research is needed to understand the transmission dynamics of T. gondii from terrestrial to

marine environments in the Hawaiian Islands and the impact (including indirect effects) this

zoonotic parasite has on its marine mammals.

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Figure 5-1. Gross examination of an adult male spinner dolphin infected with T. gondii. A)

Inflammation and ulceration observed in the oral cavity. B) Mild bruising on right

mandible with a 6 cm x 5 cm healed scar and fresh bruising just below the left eye

and gape of mouth. C) The kidney parenchyma was pale and irregular. D) The spleen

was irregular with protruding masses (5 mm – 2 cm diameter) observed.

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Figure 5-2. Microscopic examination of internal organs of a spinner dolphin infected with T.

gondii. Hematoxylin and eosin stain. A) A few protozoal cysts are present within

endothelial cells (arrow). B) Expansion of the pulmonary interstitium by lymphocytes

and plasma cells. Alveoli are flooded with macrophages. C) The splenic capsule has

deposits of mineral, hemosiderin, and extravasated erythrocytes. D) There are foci of

hepatocellular necrosis filled with fibrin and cellular debris (arrow).

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Figure 5-3. Microscopic examination of the heart of a spinner dolphin infected with T. gondii.

Hematoxylin and eosin stain. Adult male spinner dolphin tested positive for T. gondii.

A) There is multifocal cardiomyocyte necrosis and inflammation in the heart (arrow).

B) Protozoal cysts are present within myofibers (circle).

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Figure 5-4. Microscopic examination of the brain of a spinner dolphin infected with T. gondii.

Hematoxylin and eosin stain. A, B & C) Brain. Patchy effacement of the neuropil by

clusters of macrophages, neutrophils, and elongate microglial cells and occasional

astrocytic aggregates (astrocytic scars) were present.

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Figure 5-5. Microscopic examination of the adrenal gland of a spinner dolphin infected with T.

gondii. Hematoxylin and eosin stain. A) Necrosis of the adrenal gland and within the

necrotic foci (circle). B) Free tachyzoites and intracellular protozoal cysts present

(arrow).

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Figure 5-6. Immunohistochemistry (IHC) performed on internal organ tissue sections of a

spinner dolphin infected with T. gondii. A) Brain. Discrete areas of positive IHC

staining observed at the edges of necrotic zones (arrows). B) Adrenal gland. Discrete

areas of positive IHC staining observed at the edges of necrotic zones (arrows). C)

Liver. Discrete areas of positive IHC staining observed at the edges of necrotic zones

(arrows).

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Figure 5-7. Cladogram depicting the relationship of the Toxoplasma gondii linages 1-12, based

on the concatenated nucelotide sequences of GRA6, SAG1, MIC1 and 2, and

BTUB1. Numbers at each node represent the bootstrap values of the Maximum

Likelihood analysis. Branch lengths are based on the number of inferred substitutions,

as indicated by the scale. Red represents South American strains. Blue represents

North American and Europe strains. Black arrow denotes the position of the spinner

dolphin T. gondii.

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CHAPTER 6

CONCLUDNG STATEMENTS

Marine mammals are sentinel species for ocean health because they have long-life spans,

are near-shore residents, and feed at high trophic levels. However, marine mammals are exposed

to a variety of environmental stressors including: contaminants, anthropogenic noise pollution,

harmful algal biotoxins, decreasing prey availability, and emerging pathogens. Stranded marine

mammals offer important information about the ecology and health status of marine mammal

populations including baseline data on their anatomy/physiology, dietary habits, genetics,

distribution, and occurrence of diseases. Data collected from stranded animals represents a

biased snapshot of marine mammal populations; however, stranding events can provide

invaluable clues to the problems marine mammal populations are experiencing (e.g., UMEs

caused by infectious agents such as CeMV).

Molecular diagnostics, including PCR and next-generation sequencing approaches, have

proven to be invaluable tools in determining the infectious agents responsible for marine

mammal stranding events. Comparative genomic and phylogenetic analyses as part of this

dissertation, led to the characterization of seven CeMVs from around the globe, the first

description of a marine mammal circovirus from a Longman’s beaked whale (BWCV) that

stranded in Hawaii, and a novel Toxoplasma gondii strain that resulted in disseminated

toxoplasmosis in a Hawaiian spinner dolphin. Methods based on the alignment of the complete

coding sequences for CeMVs and the BWCV produced well resolved and supported trees

providing new insights into the evolution and taxonomy of these respective viral families. The

results suggest that BWCV represents a new species in the genus Circovirus and is most closely

related to a canine circovirus. Our results illustrated that six of the sequenced CeMVs represent

130

novel strains and the Fraser’s dolphin morbillivirus (DMV-Lh-18-Hawaii) represents a new

species in the genus Morbillivirus.

PCR screening of Toxoplasma gondii in stranded Hawaiian marine cetaceans resulted in

9/187 positives, all from the same spinner dolphin. This is the second case of disseminated T.

gondii infection in a stranded spinner dolphin in the last 25 years. Nearshore species, like

Hawaiian spinner dolphins, may be at increased risk from environmental contamination of

marine coastal waterways via sewage systems, storm water drainage, and freshwater runoff.

Further research is needed to understand the transmission dynamics of T. gondii from terrestrial

to marine environments in the Hawaiian Islands and the impact (including indirect effects) this

zoonotic parasite has on wildlife including its marine mammal populations.

131

LIST OF REFERENCES

Abadie, J., Nguyen, F., Groizeleau, C., Amenna, N., Fernandez, B., Guereaud, C., ... & Wyers,

M. (2001). Pigeon circovirus infection: pathological observations and suggested

pathogenesis. Avian Pathology, 30(2), 149-158.

Advanced Cell Diagnostics, Inc. (2018). The Definitive ISH Guide. Retrieved December 26,

2018, from https://acdbio.com/referenceguide.

Aguilar, A., & Raga, J. A. (1993). The striped dolphin epizootic in the Mediterranean

Sea. Ambio, 22, 524-528.

Anderson, A., Hartmann, K., Leutenegger, C. M., Proksch, A. L., Mueller, R. S., & Unterer, S.

(2017). Role of canine circovirus in dogs with acute haemorrhagic diarrhoea. Veterinary

Record, 180(22), 542-542.

Anderson, C. M., Zhang, B., Miller, M., Butko, E., Wu, X., Laver, T., ... & Park, E. (2016). Fully

automated RNAscope in situ hybridization assays for formalin‐ fixed paraffin‐embedded cells and tissues. Journal of Cellular Biochemistry, 117(10), 2201-2208.

Antonelis, G., Ryon, B., Braun, R., Spraker, T., Baker, J., & Rowles, T. (2001). Juvenile

Hawaiian monk seal (Monachus schauinslandi) unusual mortality event in the

Northwestern Hawaiian Islands. In 14th Biennial Conference on the Biology of Marine

Mammals (pp. 7-8 abstract). Vancouver, CA.

Appel, M. J. G. & Gillespie, J.H. (1972). Canine distemper virus. Virology Monographs, 11, 1-

96.

Arnold, E. F. (2016, March 9). Washed ashore: Marine mammals from medieval times to today

origins: Current events in historical perspective. Retrieved September 17, 2018, from

https://origins.osu.edu/connecting-history/washed-ashore-marine-mammals-medieval-

times-today

Attademo, F.L., Ribeiro, V.O., Soares, H.S., Luna, F.O., Sousa, G.P., Freire, A.C., . . . & Silva,

J.C. (2016). Seroprevalence of Toxoplasma gondii in captive Antillean manatee

(Trichechus manatus manatus) in Brazil. Journal of Zoo and Wildlife Medicine, 47(2),

423-426.

Bankevich, A., Nurk, S., Antipov, D., Gurevich, A. A., Dvorkin, M., Kulikov, A. S., ... &

Pyshkin, A. V. (2012). SPAdes: A new genome assembly algorithm and its applications

to single-cell sequencing. Journal of Computational Biology, 19(5), 455-477.

Barbieri, M. M., Kashinsky, L., Rotstein, D. S., Colegrove, K. M., Haman, K. H., Magargal, S.

L., ... & Littnan, C. L. (2016). Protozoal-related mortalities in endangered Hawaiian

monk seals Neomonachus schauinslandi. Diseases of Aquatic Organisms, 121(2), 85-95.

https://acdbio.com/referenceguide

132

Barrett, T., Visser, I. K. G., Mamaev, L., Goatley, L., Van Bressem, M. F., & Osterhaus, A. D.

M. E. (1993). Dolphin and porpoise morbilliviruses are genetically distinct from phocine

distemper virus. Virology, 193(2), 1010-1012.

Bellehumeur, C., Nielsen, O., Measures, L., Harwood, L., Goldstein, T., Boyle, B., & Gagnon,

C. A. (2016). Herpesviruses including novel gammaherpesviruses are widespread among

phocid seal species in Canada. Journal of Wildlife Diseases, 52(1), 70-81.

Bellière, E. N., Esperón, F., Fernández, A., Arbelo, M., Muñoz, M. J., & Sánchez-Vizcaíno, J.

M. (2011a). Phylogenetic analysis of a new Cetacean morbillivirus from a short-finned

pilot whale stranded in the Canary Islands. Research in Veterinary Science, 90(2), 324-

328.

Bellière, E. N., Esperón, F., & Sánchez-Vizcaíno, J. M. (2011b). Genetic comparison among

dolphin morbillivirus in the 1990–1992 and 2006–2008 Mediterranean

outbreaks. Infection, Genetics and Evolution, 11(8), 1913-1920.

Bellini, W. J., Englund, G., Rozenblatt, S., Arnheiter, H., & Richardson, C. D. (1985). Measles

virus P gene codes for two proteins. Journal of Virology, 53(3), 908-919.

Biagini, P., Bendinelli, M., Hino, S., Kakkola, L., Mankertz, A., Niel, C., … & Todd, D. (2012)

Family Circoviridae. In: King A, Adams MJ, Carstens EB, Lefkowitz EJ (eds) Virus

taxonomy: Classification and nomenclature of viruses: Ninth report of the International

Committee on Taxonomy of Viruses. Academic Press, London, pp 343–349.

Bolt, G., Alexandersen, S., & Blixenkrone-Møller, M. (1995). The phosphoprotein gene of a

dolphin morbillivirus isolate exhibits genomic variation at the editing site. Journal of

General Virology, 76(12), 3051-3058.

Bolt, G., Blixenkrone-Møller, M., Gottschalck, E., Wishaupt, R. G., Welsh, M. J., Earle, J. A. P.,

& Rima, B. K. (1994). Nucleotide and deduced amino acid sequences of the matrix (M)

and fusion (F) protein genes of cetacean morbilliviruses isolated from a porpoise and a

dolphin. Virus research, 34(3), 291-304.

Bossart, G., Mignucci-Giannoni, A.A., Rivera-Guzman, A., Jimenez-Marrero, N., Camus, A.,

Bonde, R., . . . Reif, J. (2012). Disseminated toxoplasmosis in Antillean manatees

Trichechus manatus manatus from Puerto Rico. Diseases of Aquatic Organisms, 101(2),

139-144.

Bossart, G. D., Reif, J. S., Schaefer, A. M., Goldstein, J., Fair, P. A., & Saliki, J. T. (2010).

Morbillivirus infection in free-ranging Atlantic bottlenose dolphins (Tursiops truncatus)

from the Southeastern United States: Seroepidemiologic and pathologic evidence of

subclinical infection. Veterinary Microbiology, 143(2-4), 160-166.

Bowen, W.D., & Siniff, D.B. (1999). Distribution, population biology, and feeding ecology of

marine mammals. In: Reynolds III, J.E., & Rommel, S.A. (eds) Biology of Marine

Mammals. Smithsonian Institution, Washington DC, pp 423-484.

133

Breitbart, M., Delwart, E., Rosario, K., Segalés, J., Varsani, A., and ICTV Report

Consortium. (2017). ICTV Virus Taxonomy: Circoviridae. Journal of General Virology,

98(8), 1997–1998.

Buergelt, C.D., & Bonde, R.K. (1983). Toxoplasmic meningoencephalitis in a West Indian

manatee. Journal of the American Veterinary Medical Association, 183, 1294-1296.

Carpenter, M. A., Appel, M. J., Roelke-Parker, M. E., Munson, L., Hofer, H., East, M., &

O'Brien, S. J. (1998). Genetic characterization of Canine distemper virus in Serengeti

carnivores. Veterinary Immunology and Immunopathology, 65(2-4), 259-266. Cassle, S. E., Landrau-Giovannetti, N., Farina, L. L., Leone, A., Wellehan Jr, J. F., Stacy, N. I.,

... & Saliki, J. T. (2016). Coinfection by Cetacean morbillivirus and Aspergillus

fumigatus in a juvenile bottlenose dolphin (Tursiops truncatus) in the Gulf of

Mexico. Journal of Veterinary Diagnostic Investigation, 28(6), 729-734.

Centelleghe, C., Beffagna, G., Palmisano, G., Franzo, G., Casalone, C., Pautasso, A., ... & Di

Guardo, G. (2017). Dolphin morbillivirus in a Cuvier’s beaked whale (Ziphius

cavirostris), Italy. Frontiers in Microbiology, 8, 111.

Conrad, P., Miller, M., Kreuder, C., James, E., Mazet, J., Dabritz, H., . . . Grigg, M. (2005).

Transmission of toxoplasma: Clues from the study of sea otters as sentinels of

Toxoplasma gondii flow into the marine environment. International Journal for

Parasitology, 35(11-12), 1155-1168.

Davison, A. J., Nielsen, O., Subramaniam, K., Jacob, J. M., Romero, C. H., Burek-Huntington,

K. A., & Waltzek, T. B. (2017a). Genome sequence of an alphaherpesvirus from a beluga

whale (Delphinapterus leucas). Genome Announcements, 5(42), e01100-17.

Davison, A. J., Subramaniam, K., Kerr, K., Jacob, J. M., Landrau-Giovannetti, N., Walsh, M. T.,

… Waltzek, T. B. (2017b). Genome sequence of a gammaherpesvirus from a common

bottlenose dolphin (Tursiops truncatus). Genome Announcements, 5(31), e00777-17.

Decaro, N., Martella, V., Desario, C., Lanave, G., Circella, E., Cavalli, A., ... & Buonavoglia, C.

(2014). Genomic characterization of a circovirus associated with fatal hemorrhagic

enteritis in dog, Italy. PloS One, 9(8), e105909.

de Vries, R. D., Verburgh, R. J., van de Bildt, M. W., Osterhaus, A. D., & de Swart, R. L.

(2013). Complete genome sequence of phocine distemper virus isolated from a harbor

seal (Phoca vitulina) during the 1988 North Sea epidemic. Genome Announcements, 1(3),

e00291-13.

Di Guardo, G., Proietto, U., Francesco, C.E., Marsilio, F., Zaccaroni, A., Scaravelli, D., . . .

Casalone, C. (2009). Cerebral toxoplasmosis in striped dolphins (Stenella coeruleoalba)

stranded along the Ligurian Sea Coast of Italy. Veterinary Pathology, 47(2), 245-253.

Dobson, A.P., Rodriguez, J.P., Roberts, W.M., & Wilcove, D.S. (1997). Geographic distribution

of endangered species in the United States. Science, 275(5299), 550-553.

134

Domingo, M., Ferrer, L., Pumarola, M., Marco, A., Plana, J., Kennedy, S., ... & Rima, B. K.

(1990). Morbillivirus in dolphins. Nature (London), 348(6296).

Donahoe, S.L., Rose, K., & Šlapeta, J. (2014). Multisystemic toxoplasmosis associated with a

type II-like Toxoplasma gondii strain in a New Zealand fur seal (Arctocephalus forsteri)

from New South Wales, Australia. Veterinary Parasitology, 205(1-2), 347-353.

Dubey, J.P., Fair, P.A., Sundar, N., Velmurugan, G., Kwok, O.C., Mcfee, W.E., . . . Su, C.

(2008). Isolation of Toxoplasma gondii from bottlenose dolphins (Tursiops truncatus).

Journal of Parasitology, 94(4), 821-823.

Dubey, J.P., Mergl, J., Gehring, E., Sundar, N., Velmurugan, G.V., Kwok, O.C., . . . Martineau,

D. (2009). Toxoplasmosis in captive dolphins (Tursiops truncatus) and walrus (Odobenus

rosmarus). Journal of Parasitology, 95(1), 82-85.

Dubey, J.P., Morales, J.A., Sundar, N., Velmurugan, G.V., González-Barrientos, C.R.,

Hernández-Mora, G., & Su, C. (2007). Isolation and genetic characterization of

Toxoplasma gondii from striped dolphin (Stenella coeruleoalba) from Costa Rica.

Journal of Parasitology, 93(3), 710-711.

Dubey, J. P., Velmurugan, G. V., Rajendran, C., Yabsley, M. J., Thomas, N. J., Beckmen, K. B.,

... & McFee, W. E. (2011). Genetic characterisation of Toxoplasma gondii in wildlife

from North America revealed widespread and high prevalence of the fourth clonal

type. International journal for parasitology, 41(11), 1139-1147.

Duignan, P. J., House, C., Odell, D. K., Wells, R. S., Hansen, L. J., Walsh, M. T., ... & Geraci, J.

R. (1996). Morbillivirus infection in bottlenose dolphins: Evidence for recurrent

epizootics in the western Atlantic and Gulf of Mexico. Marine Mammal Science, 12(4),

499-515.

Duignan, P. J., House, C., Walsh, M. T., Campbell, T., Bossart, G. D., Duffy, N., ... & Geraci, J.

R. (1995). Morbillivirus infection in manatees. Marine Mammal Science, 11(4), 441-451.

Duignan, P., Van Bressem, M. F., Baker, J., Barbieri, M., Colegrove, K., De Guise, S., ... &

Early, G. (2014). Phocine distemper virus: current knowledge and future

directions. Viruses, 6(12), 5093-5134.

Fauquier, D., Goldstein, T., Colegrove, K., Rotstein, D., Di Giovanni, R. A. J. R., & McLellan,

W. (2014). Update on the dolphin morbillivirus outbreak and the 2013–2014 US mid-

Atlantic bottlenose dolphin (Tursiops truncatus) unusual mortality event. Rep.

International Whal. Commission. See https://events. iwc. int/index.

php/scientific/SC65B/paper/viewFile/759/757/SC-65b-E03. pdf.

Fauquier, D. A., Litz, J., Sanchez, S., Colegrove, K., Schwacke, L. H., Hart, L., ... & McFee, W.

(2017). Evaluation of morbillivirus exposure in cetaceans from the northern Gulf of

Mexico 2010-2014. Endangered Species Research, 33, 211-220.

135

Fayer, R., Dubey, J.P., & Lindsay, D.S. (2004). Zoonotic protozoa: From land to sea. Trends in

Parasitology, 20(11), 531-536.

Fernández, A., Esperón, F., Herraéz, P., de los Monteros, A. E., Clavel, C., Bernabé, A., ... &

Bayón, A. (2008). Morbillivirus and pilot whale deaths, Mediterranean Sea. Emerging

Infectious Diseases, 14(5), 792.

Firth, A. E., & Brierley, I. (2012). Non-canonical translation in RNA viruses. Journal of General

Virology, 93(7), 1385-1409.

Geraci, J. R., Anderson, D. M., Timperi, R. J., St. Aubin, D. J., Early, G. A., Prescott, J. H., &

Mayo, C. A. (1989). Humpback whales (Megaptera novaeangliae) fatally poisoned by

dinoflagellate toxin. Canadian Journal of Fisheries and Aquatic Sciences, 46(11), 1895-

1898.

Geraci, J. R., & Lounsbury, V. J. (2005). Marine mammals ashore: A field guide for strandings

(2nd ed.). Baltimore, MD: National Aquarium in Baltimore.

Gonzales-Viera, O., Marigo, J., Ruoppolo, V., Rosas, F., Kanamura, C., Takakura, C., . . . Catão-

Dias, J. (2013). Toxoplasmosis in a Guiana dolphin (Sotalia guianensis) from Paraná,

Brazil. Veterinary Parasitology, 191(3-4), 358-362.

Grant, R. J., Banyard, A. C., Barrett, T., Saliki, J. T., & Romero, C. H. (2009). Real-time RT-

PCR assays for the rapid and differential detection of dolphin and porpoise

morbilliviruses. Journal of Virological Methods, 156(1-2), 117-123. Green, M. R., & Sambrook, J. (2012). Molecular Cloning: A Laboratory Manual (4th ed., Vol.

1). New York: Cold Spring Harbor Laboratory Press.

Greenland, J.A., Limpus, C.J. & Currie, K.J. (2005). Marine wildlife stranding and mortality

database annual report 2003. I Dugong. Conservation Technical and Data Report,

2003(1). Environmental Protection Agency.

Groch, K. R., Colosio, A. C., Marcondes, M. C., Zucca, D., Díaz-Delgado, J., Niemeyer, C., ... &

Catão-Dias, J. L. (2014). Novel Cetacean morbillivirus in Guiana dolphin,

Brazil. Emerging Infectious Diseases, 20(3), 511.

Groch, K. R., Santos-Neto, E. B., Díaz-Delgado, J., Ikeda, J. M., Carvalho, R. R., Oliveira, R. B.,

... & Catão-Dias, J. L. (2018). Guiana dolphin unusual mortality event and link to

Cetacean morbillivirus, Brazil. Emerging Infectious Diseases, 24(7), 1349-1354.

Gulland, F. M., & Hall, A. J. (2007). Is marine mammal health deteriorating? Trends in the

global reporting of marine mammal disease. EcoHealth, 4(2), 135-150.

Hall, T. A. (1999, January). BioEdit: A user-friendly biological sequence alignment editor and

analysis program for Windows 95/98/NT. In Nucleic acids symposium series (Vol. 41,

No. 41, pp. 95-98). [London]: Information Retrieval Ltd., c1979-c2000.

136

Hanni, K.D., Mazet, J.A., Gulland, F.M., Estes, J., Staedler, M., Murray, M.J., . . . Jessup, D.A.

(2003). Clinical pathology and assessment of pathogen exposure in Southern and Alaskan

sea otters. Journal of Wildlife Diseases, 39(4), 837-850.

Hansen, L. J. (Ed.). (1992). Report on investigation of 1990 Gulf of Mexico bottlenose dolphin

strandings. National Marine Fisheries Service, Southeast Fisheries Science Center.

Hansen, H., Hess, S. C., Cole, D., & Banko, P. C. (2007). Using population genetic tools to

develop a control strategy for feral cats (Felis catus) in Hawaii. Wildlife Research, 34(8),

587-596.

Herbst, W., & Willems, H. (2017). Detection of virus particles resembling circovirus and porcine

circovirus 2a (PCV2a) sequences in feces of dogs. Research in Veterinary Science, 115,

51-53.

Hersh, S. L., Odell, D. K., & Asper, E. D. (1990). Bottlenose dolphin mortality patterns in the

Indian/Banana River system of Florida. pp. 155-64. The Bottlenose Dolphin. Academic

Press, San Diego, CA., USA.

Hollings, T., Jones, M., Mooney, N., & McCallum, H. (2013). Wildlife disease ecology in

changing landscapes: Mesopredator release and toxoplasmosis. International Journal for

Parasitology: Parasites and Wildlife, 2, 110-118.

Honnold, S.P., Braun, R., Scott, D.P., Sreekumar, C., & Dubey, J.P. (2005). Toxoplasmosis in a

Hawaiian monk seal (Monachus schauinslandi). Journal of Parasitology, 91(3), 695-697.

Howe, D.K., & Sibley, L.D. (1995). Toxoplasma gondii comprises three clonal lineages:

Correlation of parasite genotype with human disease. Journal of Infectious Diseases,

172(6), 1561-1566.

Ilyina, T. V., & Koonin, E. V. (1992). Conserved sequence motifs in the initiator proteins for

rolling circle DNA replication encoded by diverse replicons from eubacteria, eucaryotes

and archaebacteria. Nucleic Acids Research, 20(13), 3279-3285.

Inskeep II, W., Gardiner, C. H., Harris, R. K., Dubey, J. P., & Goldston, R. T. (1990).

Toxoplasmosis in Atlantic bottle-nosed dolphins (Tursiops truncatus). Journal of Wildlife

Diseases, 26(3), 377-382.

Jacob, J. M., Subramaniam, K., Tu, S. L., Nielsen, O., Tuomi, P. A., Upton, C., & Waltzek, T. B.

(2018). Complete genome sequence of a novel sea otterpox virus. Virus Genes, 54(6),

756-767.

Jacob, J. M., West, K. L., Levine, G., Sanchez, S., & Jensen, B. (2016). Initial characterization of

novel beaked whale morbillivirus in Hawaiian cetaceans. Diseases of Aquatic

Organisms, 117, 215-227.

Jardine, J.E., & Dubey, J.P. (2002). Congenital toxoplasmosis in an Indo-Pacific bottlenose

dolphin (Tursiops aduncus). Journal of Parasitology, 88(1), 197-199.

137

Jeoung, H. Y., Lee, M. H., Yeh, J. Y., Lim, J. A., Lim, S. I., Oem, J. K., ... & An, D. J. (2012).

Complete genome analysis of three live attenuated Rinderpest virus vaccine strains

derived through serial passages in different culture systems. Journal of Virology, 86(23),

13115-13116.

Jo, W. K., Kruppa, J., Habierski, A., van de Bildt, M., Mazzariol, S., Di Guardo, G., ... &

Ludlow, M. (2018). Evolutionary evidence for multi-host transmission of Cetacean

morbillivirus. Emerging Microbes & Infections, 7(1), 201.

Johnson, A. M., Delong, R. L., Fiscus, C. H., & Kenyon, K. W. (1982). Population status of the

Hawaiian monk seal (Monachus schauinslandi), 1978. Journal of Mammalogy, 63(3),

415-421.

Johnson, B. W., & Johnson, P. A. (1981). The Hawaiian monk seal on Laysan Island: 1978. US

Department of Commerce, National Technical Information Service, Springfield, VA. PB-

285-428.

Karlsson, E. A., Ip, H. S., Hall, J. S., Yoon, S. W., Johnson, J., Beck, M. A., ... & Schultz-

Cherry, S. (2014). Respiratory transmission of an avian H3N8 influenza virus isolated

from a harbour seal. Nature Communications, 5, 4791.

Katoh, K., & Standley, D. M. (2013). MAFFT multiple sequence alignment software version 7:

improvements in performance and usability. Molecular biology and evolution, 30(4),

772-780.

Kearse, M. G., & Wilusz, J. E. (2017). Non-AUG translation: a new start for protein synthesis in

eukaryotes. Genes & Development, 31(17), 1717-1731.

Kedkovid, R., Woonwong, Y., Arunorat, J., Sirisereewan, C., Sangpratum, N., Lumyai, M., ... &

Thanawongnuwech, R. (2018). Porcine circovirus type 3 (PCV3) infection in grower pigs

from a Thai farm suffering from porcine respiratory disease complex (PRDC). Veterinary

Microbiology, 215, 71-76.

Kennedy, S., Smyth, J. A., Cush, P. F., McAliskey, M., McCullough, S. J., & Rima, B. K.

(1991). Histopathologic and immunocytochemical studies of distemper in harbor

porpoises. Veterinary Pathology, 28(1), 1-7.

Kennedy, S., Smyth, J. A., Cush, P. F., McCullough, S. J., Allan, G. M., & McQuaid, S. (1988).

Viral distemper now found in porpoises. Nature, UK, 336(6194).

Khan, A., Dubey, J.P., Su, C., Ajikoka, J.W., Rosenthal, B.M. & Sibley, L.D. (2011). Genetic

analyses of atypical Toxoplasma gondii strains reveal a fourth clonal lineage in North

America. International Journal for Parasitology, 41, 645-655.

Khan, A., Fux, B., Su, C., Dubey, J. P., Dardé, M. L., Ajioka, J. W., ... & Sibley, L. D. (2007).

Recent transcontinental sweep of Toxoplasma gondii driven by a single monomorphic

chromosome. Proceedings of the National Academy of Sciences, 104(37), 14872-14877.

138

Kim, S. H., Park, J. Y., Jung, J. Y., Kim, H. Y., Park, Y. R., Lee, K. K., ... & Park, C. K. (2018).

Detection and genetic characterization of porcine circovirus 3 from aborted fetuses and

pigs with respiratory disease in Korea. Journal of Veterinary Science, 19(5), 721-724.

Kluge, M., Campos, F. S., Tavares, M., de Amorim, D. B., Valdez, F. P., Giongo, A., ... &

Franco, A. C. (2016). Metagenomic survey of viral diversity obtained from feces of

subantarctic and south american fur seals. PLoS One, 11(3), e0151921.

Krafft, A., Lichy, J. H., Lipscomb, T. P., Klaunberg, B. A., Kennedy, S., & Taubenberger, J. K.

(1995). Postmortem diagnosis of morbillivirus infection in bottlenose dolphins (Tursiops

truncatus) in the Atlantic and Gulf of Mexico epizootics by polymerase chain reaction-

based assay. Journal of Wildlife Diseases, 31(3), 410-415.

Kreuder, C., Miller, M.A., Jessup, D.A., Lowenstine, L.J., Harris, M.D., Ames, J.A., . . . Mazet,

J.A. (2003). Patterns of mortality in Southern sea otters (Enhydra lutris nereis) from

1998–2001. Journal of Wildlife Diseases, 39(3), 495-509.

Li, L., Mcgraw, S., Zhu, K., Leutenegger, C. M., Marks, S. L., Kubiski, S., . . . Pesavento, P. A.

(2013). Circovirus in tissues of dogs with vasculitis and hemorrhage. Emerging Infectious

Diseases, 19(4), 534-541.

Lindsay, D.S., & Dubey, J.P. (2009). Long-term survival of Toxoplasma gondii sporulated

oocysts in seawater. Journal of Parasitology, 95(4), 1019-1020.

Lipscomb, T. P., Kennedy, S., Moffett, D., Krafft, A., Klaunberg, B. A., Lichy, J. H., ... &

Taubenberger, J. K. (1996). Morbilliviral epizootic in bottlenose dolphins of the Gulf of

Mexico. Journal of Veterinary Diagnostic Investigation, 8(3), 283-290.

Lipscomb, T. P., Schulman, F. Y., Moffett, D., & Kennedy, S. (1994). Morbilliviral disease in

Atlantic bottlenose dolphins (Tursiops truncatus) from the 1987-1988 epizootic. Journal

of Wildlife Diseases, 30(4), 567-571.

Litz, J. A., Baran, M. A., Bowen-Stevens, S. R., Carmichael, R. H., Colegrove, K. M., Garrison,

L. P., ... & Jones, W. (2014). Review of historical unusual mortality events (UMEs) in the

Gulf of Mexico (1990-2009): Providing context for the multi-year northern Gulf of

Mexico cetacean UME declared in 2010. Diseases of Aquatic Organisms, 112(2), 161-

175.

Liu, Q., Wang, Z. D., Huang, S. Y., & Zhu, X. Q. (2015). Diagnosis of toxoplasmosis and typing

of Toxoplasma gondii. Parasites & vectors, 8(1), 292.

Lorincz, M., Csagola, A., Farkas, S. L., Szekely, C., & Tuboly, T. (2011). First detection and

analysis of a fish circovirus. Journal of General Virology, 92(8), 1817-1821.

Lorincz, M., Dán, Á, Láng, M., Csaba, G., Tóth, Á G., Székely, C., . . . Tuboly, T. (2012). Novel

circovirus in European catfish (Silurus glanis). Archives of Virology, 157(6), 1173-1176.

139

Maclachlan, N. J., & Dubovi, E. J. (2010). Circoviridae and Anelloviridae. In Fenner's

Veterinary Virology (4th ed., pp. 259-268). London, UK: Academic Press.

MacLeod, C. D., Santos, M. B., & Pierce, G. J. (2003). Review of data on diets of beaked

whales: Evidence of niche separation and geographic segregation. Journal of the Marine

Biological Association of the United Kingdom, 83(3), 651-665.

Maldini, D., Mazzuca, L., & Atkinson, S. (2005). Odontocete stranding patterns in the main

Hawaiian Islands (1937–2002): How do they compare with live animal surveys?. Pacific

Science, 59(1), 55-67.

Malmberg, M., Rubio-Guerri, C., Hayer, J., Garcia-Parraga, D., Nieto-Pelegrin, E., Melero, M.,

... & Granberg, F. (2017). Phylogenomic analysis of the complete sequence of a

gastroenteritis-associated cetacean adenovirus (bottlenose dolphin adenovirus 1) reveals a

high degree of genetic divergence. Infection, Genetics and Evolution, 53, 47-55.

Maness, H. T., Nollens, H. H., Jensen, E. D., Goldstein, T., LaMere, S., Childress, A., ... &

Wellehan Jr, J. F. (2011). Phylogenetic analysis of marine mammal

herpesviruses. Veterinary Microbiology, 149(1-2), 23-29.

Mankertz, A., Hattermann, K., Ehlers, B., & Soike, D. (2000). Cloning and sequencing of

columbid circovirus (CoCV), a new circovirus from pigeons. Archives of

Virology, 145(12), 2469-2479.

Mazzariol, S., Centelleghe, C., Beffagna, G., Povinelli, M., Terracciano, G., Cocumelli, C., ... &

Di Francesco, C. E. (2016). Mediterranean fin whales (Balaenoptera physalus) threatened

by dolphin morbillivirus. Emerging Infectious Diseases, 22(2), 302.

Mazzariol, S., Marcer, F., Mignone, W., Serracca, L., Goria, M., Marsili, L., ... & Casalone, C.

(2012). Dolphin morbillivirus and Toxoplasma gondii coinfection in a Mediterranean fin

whale (Balaenoptera physalus). BMC Veterinary Research, 8(1), 20.

Mazzariol, S., Peletto, S., Mondin, A., Centelleghe, C., Di Guardo, G., Di Francesco, C. E., ... &

Acutis, P. L. (2013). Dolphin morbillivirus infection in a captive harbor seal (Phoca

vitulina). Journal of Clinical Microbiology, 51(2), 708-711.

Mazzuca, L., Atkinson, S., Keating, B., & Nitta, E. (1999). Cetacean mass strandings in the

Hawaiian Archipelago, 1957-1998. Aquatic Mammals, 25, 105-114.

McCullough, S. J., McNeilly, F., Allan, G. M., Kennedy, S., Smyth, J. A., Cosby, S. L., ... &

Rima, B. K. (1991). Isolation and characterisation of a porpoise morbillivirus. Archives of

Virology, 118(3-4), 247-252.

Measures, L.N., Dubey, J.P., Labelle, P., & Martineau, D. (2004). Seroprevalence of Toxoplasma

gondii in Canadian Pinnipeds. Journal of Wildlife Diseases, 40(2), 294-300.

Migaki, G., Allen, J.F., & Casey, H.W. (1977). Toxoplasmosis in California sea lion (Zalophus

californianus). American Journal of Veterinary Research, 38(1), 135-136.

140

Migaki, G., Sawa, T.R., & Dubey, J.P. (1990). Fatal disseminated toxoplasmosis in a spinner

dolphin (Stenella longirostris). Veterinary Pathology, 27(6), 463-464.

Miller, M., Gardner, I., Kreuder, C., Paradies, D., Worcester, K., Jessup, D., . . . Conrad, P.

(2002a). Coastal freshwater runoff is a risk factor for Toxoplasma gondii infection of

southern sea otters (Enhydra lutris nereis). International Journal for Parasitology, 32(8),

997-1006.

Miller, M., Gardner, I., Packham, A., Mazet, J. K., Hanni, K. D., Jessup, D., . . . Conrad, P.

(2002b). Evaluation of an indirect fluorescent antibody test (IFAT) for demonstration of

antibodies to Toxoplasma gondii in the sea otter (Enhydra lutris). Journal of


Miller, M., Grigg, M., Kreuder, C., James, E., Melli, A., Crosbie, P., . . . Conrad, P. (2004). An

unusual genotype of Toxoplasma gondii is common in California sea otters (Enhydra

lutris nereis) and is a cause of mortality. International Journal for Parasitology, 34(3),

275-284.

Miller, M. A., Sverlow, K., Crosbie, P. R., Barr, B. C., Lowenstine, L. J., Gulland, F. M., ... &

Conrad, P. A. (2001). Isolation and characterization of two parasitic protozoa from a

Pacific harbor seal (Phoca vitulina richardsi) with meningoencephalomyelitis. Journal of


Muhire, B. M., Varsani, A., & Martin, D. P. (2014). SDT: A virus classification tool based on

pairwise sequence alignment and identity calculation. PLoS ONE, 9(9).

Ng, T. F. F., Miller, M. A., Kondov, N. O., Dodd, E. M., Batac, F., Manzer, M., ... & Delwart, E.

(2015). Oral papillomatosis caused by Enhydra lutris papillomavirus 1 (ElPV-1) in

southern sea otters (Enhydra lutris nereis) in California, USA. Journal of Wildlife

Diseases, 51(2), 446-453.

Ng, T. F. F., Wheeler, E., Greig, D., Waltzek, T. B., Gulland, F., & Breitbart, M. (2011).

Metagenomic identification of a novel anellovirus in Pacific harbor seal (Phoca vitulina

richardsii) lung samples and its detection in samples from multiple years. Journal of

General Virology, 92(6), 1318-1323.

Nguyen, L., Schmidt, H. A., Haeseler, A. V., & Minh, B. Q. (2014). IQ-TREE: A fast and

effective stochastic algorithm for estimating maximum-likelihood

phylogenies. Molecular Biology and Evolution, 32(1), 268-274.

NOAA Fisheries. (2018a). Marine mammal unusual mortality events. Retrieved September 17,

2018, from https://www.fisheries.noaa.gov/national/marine-life-distress/marine-mammal-

unusual-mortality-events.

NOAA Fisheries. (2018b). Active and closed unusual mortality events. Retrieved September 17,

2018, from https://www.fisheries.noaa.gov/national/marine-life-distress/active-and-

closed-unusual-mortality-events.

https://www.fisheries.noaa.gov/national/marine-life-distress/active-and-closed-unusual-mortality-events

https://www.fisheries.noaa.gov/national/marine-life-distress/active-and-closed-unusual-mortality-events

141

NOAA Fisheries. (2018c). 2018 Pinniped unusual mortality event along the northeast coast,

Retrieved November 30, 2018, from https://www.fisheries.noaa.gov/new-england-mid-

atlantic/marine-life-distress/2018-pinniped-unusual-mortality-event-along-northeast.

NOAA Fisheries. (2018d). 2013 – 2015 bottlenose dolphin unusual mortality event in the Mid-

Atlantic (closed). Retrieved September 17, 2018, from

https://www.fisheries.noaa.gov/national/marine-life-distress/2013-2015-bottlenose-

dolphin-unusual-mortality-event-mid-atlantic.

Osterhaus, A. D., de Swart, R. L., Vos, H. W., Ross, P. S., Kenter, M. J., & Barrett, T. (1995).

Morbillivirus infections of aquatic mammals: newly identified members of the

genus. Veterinary Microbiology, 44(2-4), 219-227.

Owen, H., Gillespie, A. & Wilkie, I. (2012). Postmortem findings from dugong (Dugong dugon)

submissions to the University of Queensland: 1997–2010. Journal of Wildlife Diseases,

48(4), 962-970.

Palinski, R., Piñeyro, P., Shang, P., Yuan, F., Guo, R., Fang, Y., ... & Hause, B. M. (2017). A

novel porcine circovirus distantly related to known circoviruses is associated with

porcine dermatitis and nephropathy syndrome and reproductive failure. Journal of

Virology, 91(1), e01879-16.

Park, E. S., Suzuki, M., Kimura, M., Maruyama, K., Mizutani, H., Saito, R., ... & Morikawa, S.

(2014). Identification of a natural recombination in the F and H genes of Feline

morbillivirus. Virology, 468, 524-531.

Parks, C. L., Lerch, R. A., Walpita, P., Wang, H. P., Sidhu, M. S., & Udem, S. A. (2001).

Comparison of predicted amino acid sequences of measles virus strains in the Edmonston

vaccine lineage. Journal of Virology, 75(2), 910-920.

Phan, T. G., Gulland, F., Simeone, C., Deng, X., & Delwart, E. (2015). Sesavirus: Prototype of a

new parvovirus genus in feces of a sea lion. Virus Genes, 50(1), 134-136.

Phenix, K. V., Weston, J. H., Ypelaar, I., Lavazza, A., Smyth, J. A., Todd, D., ... & Raidal, S. R.

(2001). Nucleotide sequence analysis of a novel circovirus of canaries and its relationship

to other members of the genus Circovirus of the family Circoviridae. Journal of General

Virology, 82(11), 2805-2809.

Pretti, C., Mancianti, F., Nardoni, S., Ariti, G., Monni, G., Di Bello, D., … Papini, R. (2010).

Detection of Toxoplasmosis gondii infection in dolphins stranded along the Tuscan coast,

Italy. Revue de médecine vétérinaire, 161(10), 428-431.

Pugliares, K. R., Bogomolni, A., Touhey, K. M., Herzig, S. M., Harry, C. T., & Moore, M. J.

(2007). Marine Mammal Necropsy: An introductory guide for stranding responders and

field biologists (pp. 1-131, Tech. No. WHOI-2007-06). Woods Hole Oceanographic

Institution.

https://www.fisheries.noaa.gov/new-england-mid-atlantic/marine-life-distress/2018-pinniped-unusual-mortality-event-along-northeast

https://www.fisheries.noaa.gov/new-england-mid-atlantic/marine-life-distress/2018-pinniped-unusual-mortality-event-along-northeast

https://www.fisheries.noaa.gov/national/marine-life-distress/2013-2015-bottlenose-dolphin-unusual-mortality-event-mid-atlantic

https://www.fisheries.noaa.gov/national/marine-life-distress/2013-2015-bottlenose-dolphin-unusual-mortality-event-mid-atlantic

142

Qiu, W., Zheng, Y., Zhang, S., Fan, Q., Liu, H., Zhang, F., ... & Hu, R. (2011). Canine distemper

outbreak in rhesus monkeys, China. Emerging Infectious Diseases, 17(8), 1541.

Raga, J. A., Banyard, A., Domingo, M., Corteyn, M., Van Bressem, M. F., Fernández, M., ... &

Barrett, T. (2008). Dolphin morbillivirus epizootic resurgence, Mediterranean

Sea. Emerging Infectious Diseases, 14(3), 471.

Rambaut, A. (2014). FigTree v1. 2.2. Institute of Evolutionary Biology. University of Edinburgh.

Rengifo-Herrera, C., Ortega-Mora, L.M., Álvarez-García, G., Gómez-Bautista, M., García-

Párraga, D., García-Peña, F.J., & Pedraza-Díaz, S. (2012). Detection of Toxoplasma

gondii antibodies in Antarctic pinnipeds. Veterinary Parasitology, 190(1-2), 259-262.

Resendes, A.R., Almería, S., Dubey, J.P., Obón, E., Juan-Sallés, C., Degollada, E., . . . Domingo,

M. (2002). Disseminated toxoplasmosis in a Mediterranean pregnant Risso's dolphin

(Grampus griseus) with transplacental fetal infection. Journal of Parasitology, 88(5),

1029-1032.

Rima, B. K., Collin, A. M. J., & Earle, J. A. P. (2005). Completion of the sequence of a

Cetacean morbillivirus and comparative analysis of the complete genome sequences of

four morbilliviruses. Virus Genes, 30(1), 113-119.

Roe, W., Howe, L., Baker, E., Burrows, L., & Hunter, S. (2013). An atypical genotype of

Toxoplasma gondii as a cause of mortality in Hector's dolphins (Cephalorhynchus

hectori). Veterinary Parasitology, 192(1-3), 67-74.

Roe, W. D., Michael, S., Fyfe, J., Burrows, E., Hunter, S. A., & Howe, L. (2017). First report of

systemic toxoplasmosis in a New Zealand sea lion (Phocarctos hookeri). New Zealand

veterinary journal, 65(1), 46-50.

Rosario, K., Breitbart, M., Harrach, B., Segalés, J., Delwart, E., Biagini, P., & Varsani, A.

(2017). Revisiting the taxonomy of the family Circoviridae: Establishment of the genus

Cyclovirus and removal of the genus Gyrovirus. Archives of Virology, 162(5), 1447-

1463.

Rotstein, D., West, K., Landrau-Giovannetti, N., & Waltzek, T. (2018). Novel morbillivirus in a

Fraser’s dolphin (Lagenodelphis hosei) from Hawaii. In 2018 ACVP and ASVCP

Concurrent Annual Meeting. Washington, DC. Retrieved December 23, 2018, from

https://cdn.ymaws.com/www.acvp.org/resource/resmgr/meetings_&_events/2018/acvp_2

018-10-25_abstracts.pdf.

Rozen, S., & Skaletsky, H. (2000). Primer3 on the WWW for general users and for biologist

programmers. Bioinformatics Methods and Protocols, 365-386.

Rubio-Guerri, C., García-Párraga, D., Nieto-Pelegrín, E., Melero, M., Álvaro, T., Valls, M., ... &

Sánchez-Vizcaíno, J. M. (2015). Novel adenovirus detected in captive bottlenose

dolphins (Tursiops truncatus) suffering from self-limiting gastroenteritis. BMC

Veterinary Research, 11(1), 53.

143

Rubio-Guerri, C., Melero, M., Esperón, F., Bellière, E. N., Arbelo, M., Crespo, J. L., ... &

Sánchez-Vizcaíno, J. M. (2013a). Unusual striped dolphin mass mortality episode related

to Cetacean morbillivirus in the Spanish Mediterranean Sea. BMC Veterinary

Research, 9(1), 106.

Rubio-Guerri, C., Melero, M., Rivera-Arroyo, B., Bellière, E. N., Crespo, J. L., García-Párraga,

D., ... & Sánchez-Vizcaíno, J. M. (2013b). Simultaneous diagnosis of Cetacean

morbillivirus infection in dolphins stranded in the Spanish Mediterranean Sea in 2011

using a novel Universal Probe Library (UPL) RT-PCR assay. Veterinary

Microbiology, 165(1-2), 109-114.

Russo, C. D., Weller, D. W., Nelson, K. E., Chivers, S. J., Torralba, M., & Grimes, D. J. (2018).

Bacterial species identified on the skin of bottlenose dolphins off southern California via

next generation sequencing techniques. Microbial Ecology, 75(2), 303-309.

Sarker, S., Terrón, M. C., Khandokar, Y., Aragao, D., Hardy, J. M., Radjainia, M., ... & Raidal,

S. R. (2016). Structural insights into the assembly and regulation of distinct viral capsid

complexes. Nature Communications, 7, 13014.

Segalés, J. (2012). Porcine circovirus type 2 (PCV2) infections: Clinical signs, pathology and

laboratory diagnosis. Virus Research, 164(1-2), 10-19.

Shapiro, K., Miller, M.A., Packham, A.E., Aguilar, B., Conrad, P.A., Vanwormer, E., & Murray,

M.J. (2015). Dual congenital transmission of Toxoplasma gondii and Sarcocystis neurona

in a late-term aborted pup from a chronically infected southern sea otter (Enhydra lutris

nereis). Parasitology, 143(3), 276-288.

Shibahara, T., Sato, K., Ishikawa, Y., & Kadota, K. (2000). Porcine circovirus induces B

lymphocyte depletion in pigs with wasting disease syndrome. Journal of Veterinary

Medical Science, 62(11), 1125-1131.

Sierra, E., Fernández, A., Suárez-Santana, C., Xuriach, A., Zucca, D., de Quirós, Y. B., ... &

Díaz-Delgado, J. (2016). Morbillivirus and pilot whale deaths, Canary Islands, Spain,

2015. Emerging Infectious Diseases, 22(4), 740.

Sierra, E., Sánchez, S., Saliki, J. T., Blas-Machado, U., Arbelo, M., Zucca, D., & Fernández, A.

(2014a). Retrospective study of etiologic agents associated with nonsuppurative

meningoencephalitis in stranded cetaceans in the Canary Islands. Journal of Clinical

Microbiology, 52(7), 2390-2397.

Sierra, E., Zucca, D., Arbelo, M., García-Álvarez, N., Andrada, M., Déniz, S., & Fernández, A.

(2014b). Fatal systemic morbillivirus infection in bottlenose dolphin, Canary Islands,

Spain. Emerging Infectious Diseases, 20(2), 269.

Siqueira, J. D., Ng, T. F., Miller, M., Li, L., Deng, X., Dodd, E., ... & Delwart, E. (2017).

Endemic infection of stranded southern sea otters (Enhydra lutris nereis) with novel

parvovirus, polyomavirus, and adenovirus. Journal of Wildlife Diseases, 53(3), 532-542.

144

Silva, M.S.A., Uzeda, R.S., Costa, K.S., Santos, S.L., Macedo, A.C.C., Abe-Sandes, K. &

Gondim, L.F.P. (2009). Detection of Hammondia heydorni and related coccidia

(Neospora caninum and Toxoplasma gondii) in goats slaughtered in Bahia, Brazil.

Veterinary Parasitology, 162, 156-159.

Smeenk, C. (1997). Strandings of sperm whales Physeter macrocephalus in the North Sea:

History and patterns. Bull Inst R Sci Nat Belg Biol, 67, 15-28.

Smith, L.N., Waltzek, T.B., Rotstein, D.S., Francis-Floyd, R., Walsh, M.T., Wellehan, J.F.X. …

de Witt, M. (2016). Disseminated toxoplasmosis Toxoplasma gondii in a wild Florida

manatee Trichechus manatus latirostris and seroprevalence in two wild populations.

Diseases of Aquatic Organisms, 122, 77-83.

Stephens, N., Duignan, P. J., Wang, J., Bingham, J., Finn, H., Bejder, L., ... & Holyoake, C.

(2014). Cetacean morbillivirus in coastal Indo-Pacific bottlenose dolphin, Western

Australia. Emerging Infectious Diseases, 20(4), 666.

Stewart, M. E., Perry, R., & Raidal, S. R. (2006). Identification of a novel circovirus in

Australian ravens (Corvus coronoides) with feather disease. Avian Pathology, 35(02), 86-

92.

Sundar, N., Cole, R., Thomas, N., Majumdar, D., Dubey, J., & Su, C. (2008). Genetic diversity

among sea otter isolates of Toxoplasma gondii. Veterinary Parasitology, 151(2-4), 125-

132.

Taubenberger, J. K., Tsai, M. M., Atkin, T. J., Fanning, T. G., Krafft, A. E., Moeller, R. B., ... &

Lipscomb, T. P. (2000). Molecular genetic evidence of a novel morbillivirus in a long-

finned pilot whale (Globicephalus melas). Emerging Infectious Diseases, 6(1), 42.

Taubenberger, J. K., Tsai, M., Krafft, A. E., Lichy, J. H., Reid, A. H., Schulman, F. Y., &

Lipscomb, T. P. (1996). Two morbilliviruses implicated in bottlenose dolphin

epizootics. Emerging Infectious Diseases, 2(3), 213.

Tenter, A.M., Heckeroth, A.R., & Weiss, L.M. (2000). Toxoplasma gondii: From animals to

humans. International Journal for Parasitology, 30(12-13), 1217-1258.

Thomas, N.J., & Cole, R.A. (1996). The risk of disease and threats to the wild population.

Endangered Species Update Special Issue: Conservation and Management of the

Southern Sea Otter, 13(12), 23-27.

Tischer, I., Rasch, R., & Tochtermann, G. (1974). Characterization of papovavirus and

picornavirus-like particles in permanent pig kidney cell lines. Zentralbl Bakteriol Orig

A, 226, 153-67.

Todd, D., Scott, A. N. J., Fringuelli, E., Shivraprasad, H. L., Gavier-Widen, D., & Smyth, J. A.

(2007). Molecular characterization of novel circoviruses from finch and gull. Avian

Pathology, 36(1), 75-81.

145

Todd, D., Weston, J. H., Soike, D., & Smyth, J. A. (2001). Genome sequence determinations and

analyses of novel circoviruses from goose and pigeon. Virology, 286(2), 354-362.

Tong, S., Chern, S. W., Li, Y., Pallansch, M. A., & Anderson, L. J. (2008). Sensitive and broadly

reactive reverse transcription-PCR assays to detect novel Paramyxoviruses. Journal of

Clinical Microbiology, 46(8), 2652-2658.

Truchon, M.H., Measures, L., L’Hérault, V., Brêthes, J.C., Galbraith, P. S., Harvey, M., . . .

Lecomte, N. (2013). Marine mammal strandings and environmental changes: A 15-year

study in the St. Lawrence ecosystem. PLoS ONE,8(3).

UniProt Consortium. (2017). UniProt: the universal protein knowledgebase. Nucleic Acids

Research, 45, D158-D169.

van de Bildt, M. W. G., Kuiken, T., & Osterhaus, A. D. M. E. (2005). Cetacean morbilliviruses

are phylogenetically divergent. Archives of Virology, 150(3), 577-583.

Van Bressem, M. F., Duignan, P. J., Banyard, A., Barbieri, M., Colegrove, K. M., De Guise, S.,

... & Fernandez, A. (2014). Cetacean morbillivirus: Current knowledge and future

directions. Viruses, 6(12), 5145-5181.

Van Bressem, M. F., Visser, I. K., van de Bildt, M. W., Teppema, J. S., Raga, J. A., & Osterhaus,

A. D. (1991). Morbillivirus infection in Mediterranean striped dolphins (Stenella

coeruleoalba). The Veterinary Record, 129(21), 471.

VanDevanter, D. R., Warrener, P., Bennett, L., Schultz, E. R., Coulter, S., Garber, R. L., & Rose,

T. M. (1996). Detection and analysis of diverse herpesviral species by consensus primer

PCR. Journal of Clinical Microbiology, 34(7), 1666-1671.

Van Elk, C. E., van de Bildt, M. W. G., Jauniaux, T., Hiemstra, S., Van Run, P. R. W. A., Foster,

G., ... & Kuiken, T. (2014). Is dolphin morbillivirus virulent for white-beaked dolphins

(Lagenorhynchus albirostris)?. Veterinary Pathology, 51(6), 1174-1182.

Verma, S. K., Su, C., & Dubey, J. P. (2015). Toxoplasma gondii isolates from mouflon sheep

(Ovis ammon) from Hawaii, USA. Journal of Eukaryotic Microbiology, 62(1), 141-143.

Viana, M., Cleaveland, S., Matthiopoulos, J., Halliday, J., Packer, C., Craft, M. E., ... & Ernest,

E. (2015). Dynamics of a morbillivirus at the domestic–wildlife interface: Canine

distemper virus in domestic dogs and lions. Proceedings of the National Academy of

Sciences, 112(5), 1464-1469.

Visser, I. K., Van Bressem, M. F., de Swart, R. L., van de Bildt, M. W., Vos, H. W., van der

Heijden, R. W., ... & Barrett, T. (1993). Characterization of morbilliviruses isolated from

dolphins and porpoises in Europe. Journal of General Virology, 74(4), 631-641.

Wang, F., Flanagan, J., Su, N., Wang, L. C., Bui, S., Nielson, A., ... & Luo, Y. (2012).

RNAscope: a novel in situ RNA analysis platform for formalin-fixed, paraffin-embedded

tissues. The Journal of Molecular Diagnostics, 14(1), 22-29.

146

Wellehan, J. F., Johnson, A. J., Harrach, B., Benko, M., Pessier, A. P., Johnson, C. M., …

Jacobson, E. R. (2004). Detection and analysis of six lizard adenoviruses by consensus

primer PCR provides further evidence of a reptilian origin for the

Atadenoviruses. Journal of Virology, 78(23), 13366-13369.

West, K. L., Levine, G., Jacob, J., Jensen, B., Sanchez, S., Colegrove, K., & Rotstein, D. (2015).

Coinfection and vertical transmission of Brucella and Morbillivirus in a neonatal sperm

whale (Physeter macrocephalus) in Hawaii, USA. Journal of Wildlife Diseases, 51(1),

227-232.

West, K. L., Sanchez, S., Rotstein, D., Robertson, K. M., Dennison, S., Levine, G., ... & Jensen,

B. (2013). A Longman's beaked whale (Indopacetus pacificus) strands in Maui, Hawaii,

with first case of morbillivirus in the central Pacific. Marine Mammal Science, 29(4),

767-776.

Wierucka, K., Verborgh, P., Meade, R., Colmant, L., Gauffier, P., Esteban, R., de Stephanis, R.,

Cañadas, A. (2014). Effects of a morbillivirus epizootic on long-finned pilot whales

Globicephala melas in Spanish Mediterranean waters. Marine Ecology Progress Series,

502, 1-10.

Woo, P. C., Lau, S. K., Wong, B. H., Fan, R. Y., Wong, A. Y., Zhang, A. J., ... & Wang, M.

(2012). Feline morbillivirus, a previously undescribed paramyxovirus associated with

tubulointerstitial nephritis in domestic cats. Proceedings of the National Academy of

Sciences, 201119972.

Work, T.M., Dagenais, J., Rameyer, R., & Breeden, R. (2015). Mortality patterns in endangered

Hawaiian geese (nene; Branta sandvicensis). Journal of Wildlife Diseases, 51(3), 688-

695.

Work, T.M., Massey, J.G., Lindsay, D.S., & Dubey, J.P. (2002). Toxoplasmosis in three species

of native and introduced Hawaiian birds. Journal of Parasitology, 88(5), 1040-1042.

Work, T.M., Massey, J.G., Rideout, B.A., Gardiner, C.H., Ledig, D.B.,…Dubey, J.P. (2000).

Fatal toxoplasmosis in free-ranging endangered ‘Alala from Hawaii. Journal of Wildlife

Diseases, 36(2), 205-212.

Work, T.M., Verma, S.K., Su, C., Medeiros, J., Kaiakapu, T., Kwok, O.C., & Dubey, J.P. (2016).

Toxoplasma gondii antibody prevalence and two new genotypes of the parasite in

endangered Hawaiian geese (nene: Branta sandvicensis). Journal of Wildlife Diseases,

52(2), 253-257.

Yang, W.-C., Pang, V. F., Jeng, C.-R., Chou, L.-S., & Chueh, L.-L. (2006). Morbilliviral

infection in a pygmy sperm whale (Kogia breviceps) from Taiwanese waters. Veterinary

Microbiology, 116(1-3), 69–76.

Yochem, P. K., Braun, R. C., Ryon, B., Baker, J. D., & Antonelis, G. A. (2004). Contingency

plan for Hawaiian monk seal unusual mortality events. NOAA Tech Memo NMFS-PIFSC-

2, 1-27.

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BIOGRAPHICAL SKETCH

From an early age, Nelmarie Landrau Giovannetti devoted her life to the study of marine

mammals. At the age of 15, she began her marine mammal experience with the Caribbean

Stranding Network/Puerto Rico Manatee Conservation Center and she is still an active member

of the organization. In 2008, Nelmarie graduated with a Bachelor of Science in Coastal Marine

Biology from the University of Puerto Rico, Humacao Campus. She continued her passion for

marine mammals by completing a 5-month internship at the Whale Center of New England,

Gloucester, MA in 2012. In 2014, Nelmarie completed a Master of Science degree in

Environmental Science and Ecology under the mentorship of Dr. Antonio A. Mignucci-Giannoni

at the Inter American University of Puerto Rico, Bayamón campus. Her thesis project was

focused on the acoustical and anatomical determination of sound production and transmission in

West Indian (Trichechus manatus) and Amazonian (T. inunguis) manatees. She completed her

Doctor of Philosophy degree in Veterinary Medical Sciences at the University of Florida,

College of Veterinary Medicine under the mentorship of Dr. Thomas Waltzek in the Spring of

2019.

© 2019 Nelmarie Landrau Giovannetti

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Transcript of © 2019 Nelmarie Landrau Giovannetti