Anguillicoloides crassus: an invasive parasitic nematode infecting American eel (Anguilla rostratd) in Cape Breton, Nova Scotia

by

Cheryl Wall

A Thesis Submitted to Saint Mary's University, Halifax, Nova Scotia in Partial Fulfillment of the Requirements for the Degree of Master of Science in Applied Science

August 19, 2011, Halifax, Nova Scotia

©Cheryl Wall 2011

Approved: Dr. Katherine Jones Supervisor

Approved: Dr. David Cone Supervisor

Approved: Dr. Gary Conboy Examiner

Approved: Mr. John MacMillan Supervisory Committee

Approved: Dr. William Jones Supervisory Committee

Approved: Dr. Kevin Vessey Dean of Graduate Studies

Approved: Dr. Jeremy Lundholm Program Co-ordinator

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1*1 Canada Certification

Name: Cheryl Wall

Degree: Master of Science in Applied Science

Title of Thesis: Anguillicoloides crassus: an invasive parasitic nematode infecting

American eel (Anguilla rostrata) in Cape Breton, Nova Scotia

Examining Committee:

Dr. Kevin Vessey, Dean of Graduate Studies

Dr. Jeremy Lundholm, Program Co-ordinator

Dr. Gary Conboy, External Examiner

Dr. Katherine Jones, Supervisor

Dr. David Cone, Supervisor

Mr. John MacMillan, Supervisory Committee

Dr. William Jones, Supervisory Committee TABLE OF CONTENTS

ABSTRACT iv

ACKNOWLEDGEMENTS v

LIST OF FIGURES vi

LIST OF TABLES viii

CHAPTER 1 1

Ecology oi Anguillicoloides crassus infecting American

eel (Anguilla rostrata) in Cape Breton, Nova Scotia

CHAPTER 2 79

Inferring the origin of Anguillicoloides crassus infecting American

eel (Anguilla rostrata) in Cape Breton, Nova Scotia using the

cytochrome c oxidase I gene of the mitochondrial DNA

SUMMARY 103

FUTURE RESEARCH 104

iii Anguillicoloides crassus: an invasive parasitic nematode infecting American eel (Anguilla rostrata) in Cape Breton, Nova Scotia

Cheryl Wall

August 19,2011

Abstract

In 2007, Anguillicoloides crassus, an invasive, parasitic nematode found in the swimbladder of anguillid eels was documented for the first time in American eel in

Canadian waters, including two sites in Cape Breton (CB) Island, Nova Scotia. That discovery necessitated this study on the distribution, ecology and origin of the parasite. It was found that sites throughout CB contain infected eels. Comparing the COI gene of the mitochondrial DNA among nematodes from CB, USA, Europe, and published sequences, it was determined that the CB infection likely originated from a direct transmission of the parasite from the USA or Japan via natural eel, intermediate or paratenic host movement and/or within the ballast water of boats. Infection occurred in both freshwater and estuarine sites, with intensity of infection increasing with length of eel. Pathological damage to the swimbladder was evident; however condition factor, liver somatic and spleen somatic indices were not affected by infection.

iv Acknowledgements

As with any research, this project would not be possible without the knowledge, support, advice and assistance of many individuals. I would like to sincerely thank all of those helping to make this thesis possible:

Katherine Jones David Cone

John MacMillan William Jones

Timothy Frasier Jonathon Forest

Dollie Campbell Stanley King

Russell Easy Laurelle Saccary

Christa Davis Alanna Khattar

Carmen Cranley Heidi de Boer

Mark Sullivan Dewayne Fox

Lori Brown Francoise Daverat

Florian Nagal Janek Simon

Yu San Han Russell Poole

Ciar O'Toole Jennie Dahlberg

Hakan Wickstrom Susan Meek

Susan Dore

v LIST OF FIGURES

CHAPTER 1

Figure 1 Taxonomy of Anguillicoloides crassus 29

Figure 2 ArcGIS map of sampling sites in Cape Breton, Nova Scotia: 31 eels caught

Figure 3 ArcGIS map of Cape Breton, Nova Scotia: no eels caught 33

Figure 4 American eel 35

Figure 5 American eel length measurement 37

Figure 6 American eel weight measurement 39

Figure 7 American eel anal color 41

Figure 8 American eel swimbladder 43

Figure 9 Swimbladder yielding 50 Anguillicoloides crassus 45

Figure 10 Primary watersheds of Cape Breton, Nova Scotia 47 Figure 11 Scatter plot of intensity of infection vs. total length of 49 American eel in Cape Breton, Nova Scotia

Figure 12 Scatter plot of intensity of infection vs. weight of 51 American eel in Cape Breton, Nova Scotia

Figure 13 Scatter plot of condition factor vs. intensity for infected 53 American eels in Cape Breton, Nova Scotia

Figure 14 Scatter plot of liver somatic index vs. intensity in infected 55 American eels in Cape Breton, Nova Scotia

vi Figure 15 Scatter plot of spleen somatic index vs. intensity in infected 57 American eels in Cape Breton, Nova Scotia

Figure 16 Histological section of American eel swimbladder 59

Figure 17 Histological section of American eel swimbladder 61

Figure 18 Bar graph of mean intensity vs. anal color in infected 63 American eel in Cape Breton, Nova Scotia

CHAPTER 2

Figure 1 1.5% agarose gel showing serial dilutions of mtDNA used to 91 determine the optimal concentration for sequence analysis

Figure 2 Maximum-Likelihood phylogenetic tree of Anguillicoloides 93 crassus cytochrome c oxidase I haplotypes

vii LIST OF TABLES

CHAPTER 1

Table 1 Sampling sites for American eels in Cape Breton, Nova 65 Scotia

Table 2 Gross pathology of the swimbladder of American eels in 67 Cape Breton, Nova Scotia

Table 3 Prevalence and mean intensity for the 10 primary 69 watersheds of Cape Breton, Nova Scotia

Table 4 Prevalence and mean intensity for the 30 sites yielding 71 American eel in Cape Breton, Nova Scotia

Table 5 Prevalence and mean intensity based on distance from 73 sea for American eel in Cape Breton, Nova Scotia

Table 6 Prevalence and intensities of Anguillicoloides crassus 75 in European eels throughout Europe

Table 7 Prevalence and mean intensities of Anguillicoloides 11 crassus in American eels from North American locations

CHAPTER 2

Table 1 Sampling sites for American eels in Cape Breton, Nova 95 Scotia

Table 2 Donated Anguillicoloides crassus samples by location 97 and sampler

Table3 Location and number of samples of Anguillicoloides 99 crassus that underwent sequencing of the cytochrome c oxidase 1 gene

Table 4 Identical sequences 101

Vlll Chapter 1: Ecology of Anguillicoloides crassus infecting American eel (Anguilla

rostrata) in Cape Breton, Nova Scotia

1 Introduction

American eel (Anguilla rostrata) are common in freshwater river systems and estuaries spanning the east coast of the western hemisphere. They occur as far north as

Greenland, southward along the east coast of North America, and as far south as Guyana in South America (Scott & Crossman, 1973; Tesch, 1977, 2003). This anguillid species is catadromous, spawning in the Sargasso Sea, with its larvae drifting northward for approximately one year before entering estuaries and fresh water rivers of the east coast.

In freshwater it pigments, grows, and develops sexually before returning to the sea to spawn (Scott & Crossman, 1973; Tesch, 1977; COSEWIC, 2006).

American eel once supported valuable fisheries throughout much of their North

American range, with high success occurring along the St. Lawrence estuary into the

Maritime Provinces (Eales, 1968). Furthermore, these eels have high social and ceremonial significance for the First Nations of Eastern Canada, making American eel an important aspect of their culture (COSEWIC 2006; S. Denny, pers. comm.).

Unfortunately, American eel has been in decline throughout North America during the last several decades, resulting in a recent (2006) listing of 'Special Concern' by the

Committee on the Status of Endangered Wildlife in Canada (COSEWIC), and it is currently being considered for listing under the federal Species at Risk Act (COSEWIC

2006). It is believed that the North Atlantic Oscillation, as well as natural causes such as migrational barriers, habitat loss, reduction in available prey, and predation may be affecting recruitment and survival of this eel (Simon, 2007). In addition, it has been suggested that infection by the swimbladder parasite, Anguillicoloides crassus

2 (Kuwahara, Niimi et Itagaki, 1974), is playing a role in the decline of American eel

(Simon, 2007).

Anguillicoloides crassus (Superfamily Dracunculoidea, Family Anguillicolidae;

Figure 1 for taxonomy) is a parasitic nematode that is native to eastern Asia where it parasitizes the Japanese eel (Anguilla japonica) and introduced European eel (Anguilla anguilla) (Koops and Hartmann 1989). The importation of infected Japanese eels from

Taiwan into Germany may have led to the introduction of A. crassus into Europe in the early 1980's. It was first recorded in the Weser-Ems region of northern Germany in 1982, and has since spread to Berlin, France, Great Britain, Italy, Holland, Belgium, Denmark,

Sweden, Czech Republic, Slovakia, Austria, Hungary, Poland, and even Egypt, Morocco and Tunisia (Moravec, 1994; Kirk, 2003). In 1995, North America reported its first infection of A. crassus at an aquaculture facility in Texas (Fries et al., 1996). Between

1995 and 2006, infection with this nematode was reported in South Carolina, North

Carolina, Chesapeake Bay, Maryland, New York, Hudson River, Massachusetts, and

Maine (Fries et al., 1996; Barse and Secor, 1999; Moser et al., 2001). The first Canadian

A. crassus infection was reported in Nova Scotia in 2007 (Aieta & Oliveira, 2009;

Rockwell et al., 2009) and in subsequent years has also been reported in New Brunswick and in new locations on mainland Nova Scotia (D. Campbell, pers. comm.).

The rapid spread of the parasite throughout Europe and North America may be attributed to its life cycle, which has been documented in the European eel (De Charleroy et al. 1990; Thomas & Ollevier, 1992b; Aieta & Oliveira, 2009). It is an indirect life cycle involving intermediate and possibly paratenic hosts. Paratenic, or reservoir hosts

3 accumulate the larvae of A. crassus and contribute to their transfer to the definitive eel

host, however, they are not necessary for the completion of the life cycle (Thomas &

Ollevier, 1992a). It is likely that infection by crustacean intermediate hosts occur in

smaller eels, while larger eels acquire infection from paratenic fish hosts. Nematode eggs

are released passively into their surroundings via the pneumatic duct where they hatch in

water; some hatch in the swimbladder (De Charleroy et al., 1990; Kirk, 2003). Eggs can

hatch in a variety of salinities and under laboratory conditions they are able to retain their

infective capabilities for up to 80 days in freshwater, 21 days in estuarine water (50%

seawater) and 8 days in marine water; i.e. 100% seawater (Kirk et al., 2000).

Transmission of the parasite was also completed in both estuarine and marine water;

however, transmission at sea is unlikely due to the lack of intermediate hosts. The free-

living second stage larvae (L2) attach to the substratum and undulate in order to attract

the intermediate host (Thomas & Ollevier, 1993). Within the intermediate host, A.

crassus moult and become third stage infective larvae (L3). Eels, from the glass eel stage

on, can become infected by consuming an infected intermediate host (De Charleroy et al,

1990; Kennedy & Fitch, 1990; Nimeth et al, 2000). Once inside the eel, the third stage juvenile larvae penetrate the digestive tract and migrate across the body cavity where they enter the swimbladder wall and moult into fourth stage juveniles (Haenen et al.,

1989; De Charleroy et al., 1990; Kirk, 2003). The moult from fourth stage juvenile to pre-adult and adult nematodes occurs in the swimbladder lumen. These adult nematodes feed on eel blood and become sexually mature after acquired growth.

4 A variety of intermediate hosts of this parasite have been documented in the

European eel, with the majority being crustaceans, such as copepods and several ostracods (Moravec & Konecny 1994). Examples of cyclopoid copepod intermediate hosts include Paracyclops fimbriatus, Macrocyclops albidus, M. fuscus, Euclyclops serrulatus, E. macruroides, Cyclops strenuous, C. vicinus, Acanthocyclops robustus, A. vernalis and Diacyclops bicuspidatus, as well as Cypria ophthalmica and Notodromas monacha (Haenen et al., 1989; De Charleroy et al., 1990; Kennedy & Fitch, 1990;

Moravec et al., 1993; Moravec & Konecny, 1994). It has also been demonstrated that juvenile Gammarus, an amphipod, and a brackish water calanoid copepod, Eurytemora affinis, were able to acquire an infection under laboratory conditions (Kennedy & Fitch,

1990).

Eels can acquire infection directly from ingesting these intermediate hosts or from a second host. Intermediate hosts may also be consumed by paratenic hosts and then ingested by eels. It is likely that larger eels acquire infection from these paratenic hosts, as eels larger than 50cm feed almost entirely on fish rather than copepods (Tesch, 1977).

Not only are paratenic hosts an important food source, they are a probable vector in the rapid spread of the parasite in Europe and North America (Thomas & Ollevier, 1992; K.

Oliveira, pers. comm.).

Thus far, over 30 species offish have been found to act as paratenic hosts for the parasite in Europe, with benthic (living on or near the bottom) fish being the most heavily infected because they are most likely to feed on the epibenthic (living on the surface of the bottom) intermediate hosts (Hoglund & Thomas, 1992; Thomas and Ollevier, 1992;

5 Kirk, 2003). In addition to fish, experimental laboratory infections have reported that

snails (Moravec, 1996), frogs, newt tadpoles and aquatic insect larvae (Moravec &

Skorikova, 1998) have the ability to act as paratenic hosts; however, it is still unknown if this occurs in natural populations. Anguillicola crassus larvae do not reach sexual maturity within the paratenic host; however it has been shown to moult to stage 4 larvae inside perch (De Charleroy et al. 1990; Moravec 1996).

Intermediate and paratenic host species of A. crassus in East Asia and North

America have not been researched (Nagasawa et al, 1994; Bares et al., 2001). In order to fully understand the spread of the parasite through the American eel population in North

America, one must determine the possible intermediate and paratenic hosts. As in

Europe, these hosts may serve as important vectors. However, this research is beyond the scope of the current project and will not be examined at this time.

Extensive research on European eel reports that infection of the swimbladder by

A. crassus can cause thickening, disruption, or even rupture of the bladder wall, secondary bacterial infections, decreased host activity level, and possibly interfere with eel migration to spawning grounds in the Sargasso Sea (Thomas & Ollevier, 1992b;

Molnar et al., 1993, 1995; Haenan et al, 1996; Nimeth et al, 2000; Lefebvre & Crivelli,

2004; Palstra et al, 2007; Han et al., 2008). Other harmful effects of A. crassus include a red, swollen, protruding anus that was reported in infected European eels in Ireland

(Crean et al., 2003) and increased spleen masses, due to infection, have been documented in European eels in France (Lefebvre et al., 2004). Limited research on the pathology of

6 A. crassus in American eel has described similar effects as those reported for European eels (Barse & Secor, 1999).

The study described in this chapter aims to gain a better understanding of A. crassus infections in American eel in Cape Breton, Nova Scotia. In particular, it is an examination of the overall geographic distribution of A. crassus throughout Cape Breton, the prevalence and intensity of infection within watersheds on Cape Breton Island, as well as the pathology of infection for individual eels.

Methodology

Study Sites and Sampling Effort

Eel sampling took place June to September during 2008 and 2009 throughout

Cape Breton Island. A total of 730 eels were sampled from 30 sites, and an additional 10 sites were sampled with no eels captured (Table 1; Figure 2, Figure 3). Backpack electrofishing was used in shallow, freshwater rivers and brooks, with double-ended fyke nets, minnow traps and eel pots being used in lakes and estuaries. Fyke nets, eel pots and minnow traps were typically set in the morning or early afternoon, baited with a can of sardines or frozen mackerel, left overnight and checked the next morning. The catch was emptied into a large bucket and examined on shore. By-catch specimens from all methods of sampling (i.e. other species sampled but not included in this study) were identified, counted and returned to the water. Throughout the summer of 2008, a brook trout

(Salvelinus fontinalis) project was being conducted with the Nova Scotia Department of

Fisheries and Aquaculture, and any eels that were caught as by-catch were kept for this eel study. Eels were also purchased from several eel fishermen in Cape Breton (Table 1).

7 Eels were euthanized with an overdose/overexposure to a mixture of clove oil and water (50 mg/1; Kennedy et al., 2007), placed on ice and transported to the Estuary

Bioassessment Laboratory at Cape Breton University (CBU) or the Saint Mary's

University Taxonomy (SMUT) Laboratory where they were frozen until necropsy. All animal procedures used in this project were in accordance with guidelines established by the Canadian Council of Animal Care (CCAC).

Eel necropsies

Each eel was individually thawed and necropsied (Figure 4). Necropsies included recording the total length (mm; Figure 5) and weight (to the nearest O.lg; Figure 6) of the eel (after thawing and blotting dry with paper towel), the anal opening color (ranging from normal to light red; Figure 7), the liver and spleen weights (to the nearest 0.00 lg), and a coarse description of the stomach contents (diet) where possible. The swimbladder was opened and examined for gross pathology and the presence of A. crassus (Figure 8).

Parasites were removed, counted, and individually weighed. The nematodes were placed in ethanol (C2H5OH) to be identified and used in molecular analysis (Chapter 2). The swimbladder was further examined under a dissecting microscope for the presence of larvae or parasites within the swimbladder wall. A general description on the state of the swimbladder was also noted (i.e. healthy, slightly thickened, hemorrhaging, extremely thickened, etc.) and categorized from 0 to 2 based on the damage (Table 2). Random samples were also fixed in 10% buffered formalin (CH2O) and 6 (2 representing each category - 0, 1, and 2) samples were sent to the Veterinary Diagnostic Laboratory in

Truro, Nova Scotia for histological sectioning and staining. Each sample was cut 3 times,

8 one at each end of the swimbladder and a mid-section and stained with hematoxylin and eosin.

Statistical Analysis

Some sites had low sample sizes of eels (Table 1), so sites were categorized and pooled by watersheds prior to some analyses (Table 3). Data analyses are presented for individual sites (Table 4), pooled by watershed (Table 3) and pooled for all of Cape

Breton (Table 4). Prevalence and mean intensity were calculated according to Bush et al.

(1997). Prevalence, in parasitology, is the percentage offish infected with at least one parasite divided by the number offish examined. The mean intensity is the mean total number of parasites per infected fish from one site (Bush et al. 1997). Prevalence and intensity in freshwater localities were compared to estuarine sites, as well as among freshwater sites according to distance to sea (nearest salt water). Condition factor, an indicator of the health of a fish, was calculated using K = (W(100))IL where Wis the weight of a fish in grams and L is the length of a fish in centimeters (Moyle and Cech,

2004). The mean liver somatic index (LSI = liver weight/total body weight) and mean spleen somatic index (SSI = spleen weight/total body weight) were calculated for both infected and uninfected eels. All necropsy data were recorded and stored in Microsoft

Access or Excel, and analyses were completed using a combination of SPSS Statistics and Minitab 14.

Parasite prevalence in eels in either freshwater or estuarine water were compared using a contingency table Chi-square test while intensity was compared using a Mann-

9 Whitney test. Distance from sea and intensity were tested with a Kruskal-Wallis (Zar,

1996).

Spearman's rank correlation was used to examine the relationships between total

length and intensity of infection, weight and intensity, condition factor and intensity, LSI

and intensity, and SSI and intensity. A Mann-Whitney test was used to determine if there

was a difference between condition factor and total length of eel for both infected and

uninfected eels. Mann-Whitney was also used to compare the condition factor vs. the

total length class for infected and uninfected eels. The length classes of eel were in 100

mm increments and compared within each class. Comparisons of LSI and SSI for

infected and uninfected eels were conducted using a Mann-Whitney. For all statistics,

confidence level was at p = 0.05. And finally, a Kruskal-Wallis test was used to

determine if anal color was related to intensity of infection

Results

A total of 40 sites was sampled throughout Cape Breton, with 30 of these yielding

eels (Table 1). Of the sites with eels, 25 had eels infected with A. crassus. The smallest

eel caught was from the Mira River and was 55 mm in total length. The largest eel was from Bell Lake and measured 852 mm in total length. The smallest eel parasitized by A. crassus was from Mira River and measured 70 mm long, and contained 1 adult nematode in the swimbladder. The largest infected eel was 821 mm long (Margaree Harbour) and had 4 swimbladder nematodes. Sampling bias for infected eels in passive (netting and trapping) versus active (electrofishing) collection methods was not examined in this study as no site had large enough sample size with both methods.

10 Prevalences ranged from 0 to 100% (Table 4), with an overall prevalence of 42% for Cape Breton. Mean intensity (± S.D.) of infection for the sites ranged from 1.00 ±

0.00 to 7.67 ± 9.52, and averaged 4.58 ± 5.60 overall for Cape Breton (Table 4). The highest intensity was in an eel from Margaree Harbour with 50 individual A. crassus within the swimbladder (Figure 9). All intensities exclude eels with swimbladders containing hundreds of thousands of larvae that occurred due to bursting pregnant female nematodes that likely ruptured during necropsy.

Prevalence of infection differed significantly between all sites (with samples n >

10; x2 = 49.901, DF = 14, p < 0.05; Table 5). Prevalence of A. crassus in eels from freshwater localities was 43% (n=T50), while prevalence in eels in estuarine habitats was

42% (n=578). Mean intensity for freshwater habitat was 3.63 ± 3.84, and 4.82 ± 5.90 in estuarine habitat. There was no significant difference in the prevalence (x2 = 0.081 df=1; p = 0.775) or intensity (U=6671.00; p=0.620) for freshwater and estuary localities. There was no significant difference in the prevalence based on the distance to sea (x2 ~ 1.372; df = 2; p=0.504). However, the distance to sea was significantly different between distances when comparing intensities (x2 = 6.802; df = 2; p = 0.033).

There were some differences among the 10 primary watersheds (Figure 10) with respect to prevalences and intensities (Table 3). Prevalences range from 0% for Indian

(n=5) and Wreck (n=3) watersheds to 67% for the River Denys watershed (n=21). Mean intensity ranges from 1.25 ± 0.5 in the Isle Madame watershed (n=10) to 4.88 ± 5.08 in the Grand watershed (n=46).

11 Intensity increased significantly with eel length (Spearman's rho = 0.124; p =

0.031; Figure 11). Parasite intensity increased significantly with eel weight (Spearman's rho = 0.098; p = 0.024; Figure 12). Condition factor was not significantly related to intensity of infection (Spearman's rho = 0.089; p = 0.089; Figure 13). There was no significant difference between the condition factor for uninfected and infected eels when lengths were pooled (U = 59249.500; p = 0.062). However, when compared among length classes, a significant difference in condition was reported for lengths 201-300 mm

(U = 813.00; p = 0.031).

Mean liver somatic index for infected eels was 0.0132 ± 0.0040 and 0.0134 ±

0.0037 for uninfected eels, and not significantly different (U = 91791.00; p = 0.170).

Similarly, mean spleen somatic index for infected eels was 0.0022 ± 0.0012 and 0.0020 ±

0.0012 for uninfected eels, again, with no significant difference (U = 45482.00; p =

0.017). There was no correlation between intensity and LSI (Spearman's rho = 0.018; p

= 0.762; Figure 14) and intensity and SSI (Spearman's rho = 0.010; p = 0.870; Figure

15).

Gross pathology of the eel swimbladder showed a range of characteristics. All categories were present within Cape Breton eel samples. The categories ranged from 0 to

2, with 0 being healthy and 2 being severely damaged. Any swimbladder that wasn't healthy but also wasn't severely damaged was put in category 1 (Table 2). Histology of the swimbladder samples were relative to the gross pathology with the 0 category

(healthy) showing no signs of damage, category 1 showing thickening of the connective tissue of the swimbladder and the category 2 showing severe multifocal melanizing

12 necrosis (Figure 16). Nematode larvae were also present in the category 2 samples

(Figure 17). Due to varying methods in the characterization of the state of the swimbladder by different observers, the pathologies were not compared to other health indices.

Both infected and uninfected eels showed varying shades of anal color from normal to red. There was no significant difference in intensity based on anal color (p =

0.992; Figure 18).

Discussion

Anguillicoloides crassus was first recorded in North America, in the United States

(US), in Texas in 1995 (Fries et al., 1996). Subsequent dispersal of the parasite northward along the east coast of the US reported infection in South Carolina (Fries et al.,

1996), Chesapeake Bay and Hudson River (Barse & Secor, 1999), North Carolina

(Moser et al., 2001), Rhode Island, Massachusetts and Maine (Aieta & Oliveira, 2009).

This dispersal into Canadian waters was not expected to occur, as the parasite's development and viability are temperature - dependent with the low temperature of

Canadian winters, preventing transmission from occurring (Knopf et al., 1998). Parasite surveys conducted throughout mainland Nova Scotia in the 1990's did not report the presence of A. crassus (Cone et al., 1993; Barker et al., 1996; Marcogliese & Cone,

1996). It was not until 2007 that two independent records were made of the parasite in

Canada (Aieta & Oliveira, 2009; Rockwell et al., 2009). Rockwell et al. (2009) first reported the presence of the parasite in Sydney Harbour and Mira River, Cape Breton,

Nova Scotia. Aieta and Oliveira (2009) were quick to follow with reports of infection in

13 commercially fished eels from Lochabere Lake, Margaree Harbour and Bras d'Or Lakes in Nova Scotia, as well as the Saint John River and Silver Lake in New Brunswick. Thus,

A. crassus is considered an alien parasite that arrived sometime between 5 and 10 years ago. It would appear that cold Canadian winters have not discouraged the establishment of A. crassus in American eel.

Cape Breton Distribution

Although its invasion was felt to be unlikely (Knopf et al. 1998), the reality now is that the nematode occurs throughout Cape Breton Island with nearly half of the eels sampled being infected. Additionally, several new sites in Nova Scotia and New

Brunswick have shown infection with the parasite (D. Campbell, pers. comm.) so it is starting to spread throughout the mainland and other parts of the Atlantic provinces.

All eel life stages were represented in this study with the exception of glass eels and marine leptocephala. Through laboratory experiments, De Charleroy et al. (1990),

Thomas and Ollevier (1992b) and Nimeth et al. (2000) report that all stages of the life cycle, from glass eel to silver eel, can be infected with A. crassus. The present field study also showed that elvers to silver eels carry the parasite in the wild. Elvers can become infected once they enter the estuary and either remain there, or move up into freshwater, where they would carry the infection and allow for localized upstream spread of the parasite. This indicates a widespread transmission of A. crassus within the freshwater systems themselves, possibly due to their complex life cycle which involves an intermediate, and sometimes paratenic, host (De Charleroy et al., 1990). In Europe, studies have shown that intermediate hosts include a variety of copepods and ostracods

14 (Moravec & Konecny, 1994; Kirk 2003), while paratenic hosts include fish species

(Hoglund & Thomas, 1992; Thomas and Ollevier, 1992a; Kirk, 2003), as well as aquatic insect larvae, frogs, and snails (Moravec, 1996; Moravec & Skorikova, 1998). It is suggested that such paratenic hosts allow for such rapid dispersal of the parasite as seen in Europe and North America (Thomas & Ollevier, 1992a; Aieta & Oliveira, 2009).

Eels in estuaries and freshwater localities had similar levels of infection. Although

A. crassus is known as a freshwater parasite, laboratory experiments have shown the parasite's ability to survive and maintain infectivity in full seawater (De Charleroy et al.

1989; Kennedy & Fitch, 1990; Reimer et al., 1994), and within wild eels, A. crassus has been found in brackish water, estuaries, and saline lagoons (Hoglund et al., 1992; Pilcher

& Moore, 1993; Lefebvre et al., 2002) but the infection levels are lower than in freshwater (Kirk et al., 2000; Kirk, 2003). The mean intensity for estuaries in the present study was slightly higher (4.82 ± 5.90) than freshwater (3.63 ± 3.84) with an intensity range of 1-50 parasites versus 1-18 respectively. Intermediate and paratenic hosts of A. crassus have not been identified in North America (Nagasawa et al, 1994; Barse et al.,

2001) and the Cape Breton estuaries may be providing a greater number of hosts for completing the life cycle than found in the Cape Breton freshwater. Alternatively, the eels in the freshwater may have been originally infected in the estuary, before moving into freshwater, and their swimbladder has been too damaged to allow further infection.

Accumulating histopathological changes in the swimbladder due to multiple infections may eventually render it inhospitable for further A. crassus infection (Molnar et al., 1993,

1995; Haenen et al., 1989; Wurtz & Taraschewski, 2000; Lefebvre & Crivelli, 2004;

15 Knopf et al, 2008; Abdelmonem et al., 2010). Recent studies on American eel have also found that some eels may never leave the estuary, choosing to live and complete their life cycle within salt water (Lamson et al., 2006; Jessop et al., 2008). If this is the case, infected eels that choose to permanently inhabit estuaries may be continuously exposed to the parasite throughout their life cycle. The observed difference in abundance of the parasite between the two habitats could simply reflect the early phase in its dispersal or the plasticity in the eels selection in habitat.

Prevalence and mean intensity within Cape Breton varied with freshwater distance from sea (Table 5). Sites that were located 15-30km from sea (or nearest salt water) had the highest prevalence and mean intensity. Eels within this range are significantly more infected than the more inland freshwater samples. Research has shown some eels tend to stay within a certain home range area, moving nocturnally to feed

(Lamothe at al., 2000; Thibault et al., 2007; Hedger et al., 2010) and as mentioned above, some eels are permanent residents of salt water, or freshwater; never leaving (Lamson et al., 2006; Jessop et al., 2008). Eels within this range (i.e. 15-30km from sea) may become infected, and because there is not a lot of movement in or out, re-infection continues to occur within the population located within this distance from sea.

Alternatively, Thibault et al. (2007) showed that during the summer months, some eels moved from the freshwater down to the estuaries for better foraging and remained there until the fall months. In the present study, estuaries showed a high prevalence and intensity of infection, which may be due to the time of sampling. The majority of samples were obtained in the summer and early fall, suggesting that eels may have been caught

16 during such seasonal forays. Eels returning from the highly infected estuary into freshwater for winter may be carrying the infection back into the upper reaches of the freshwater, spreading eggs along the way. It appears that the overall vagility of eels throughout aquatic habitats, coupled with the diversity of paratenic/intermediate hosts, fuels this nematode's aggressive dispersal.

Taking a closer look at differences among watersheds (Figure 9; Table 3), River

Denys, River Inhabitants and Grand had the highest prevalences (67%, 52% and 52% respectively). These watersheds are located near bodies of water that are connected to major shipping ports and boating traffic, suggesting ballast water as the route of arrival into Cape Breton. Distributional data suggests the nematode may have arrived first in the south west part of the island (in the Canso Strait area) and spread north and east, as indicated by the decreasing prevalence across the island and ending with the north western Indian and Wreck watersheds devoid of eels. River Denys, having the highest prevalence, is also connected to the Bras d'Or Lakes, which hosts many recreational vessels, as well as ballast-exchanging ships from the United States (ICES, 2006).

In comparison with the vast amount of research done in Europe, prevalences and intensities within the European eel are higher (Table 6) than in Cape Breton. The parasite has been present in Europe much longer than it has been in Canada, or even North

America, with almost 15 years between the first reports from either continent. Within the

European eel, the parasite has been reported to reach 100%) prevalence in one year

(Kennedy & Fitch, 1990; Thomas & Ollevier, 1992b). It is unknown if this is also the case for the American eel, but prevalences and intensities reported throughout North

17 America and specifically Cape Breton, thus far, have been lower than that of Europe

(Table 7) supporting the idea of recent colonization in Cape Breton.

Pathology in Cape Breton American eels

There was an increase in intensity with eel length and weight, a trend also documented in infected American (Moser et al., 2001) and European eel (Moller et al.,

1991; Thomas & Ollevier, 1992b; Molnar et al., 1994; Lefebvre et al., 2002), indicating that all sizes can be infected and that host size is likely not a determinate of A. crassus levels of infection (Thomas & Ollevier, 1992b; Barse et al., 2001; Moser et al., 2001;

Lefebvre et al., 2002). Several reasons for this trend are possible, including a longer exposure to the parasite over time, a larger body size (swimbladder) for the parasite to infect, and a higher consumption of infected intermediate/paratenic hosts with increasing host size (Lefebvre et al., 2002).

Eel condition factor (K), a measure of nutritional status, was not related to intensity of infection, nor was there a significant difference between K for infected and uninfected eels when samples from different localities were pooled. The results are consistent with findings from studies of infected European eel, which suggest that despite the damage of A. crassus to the swimbladder, infected eels still appear to be in good condition (Moller et al., 1991; Moser et al., 2001; Morrissey & McCarthy, 2007; Han et al., 2008). This contrasts studies on infected farmed eels which exhibit a loss of appetite and emaciation (Nagasawa et al., 1994; Kirk, 2003). It has been suggested that loss of body weight in wild eels has not been demonstrated because their K shows greater variation due to more diverse feeding, such as better nourishment from fish paratenic

18 hosts, or a more diverse infection history (Moller et al., 1991; Molnar et al., 1993; Kirk

2003). Interestingly, when K was compared among length classes for infected and

uninfected eels, a significant difference was found within the 201-3 00mm length class. It

is unclear as to why this is the case solely for this size class, but it could simply reflect

variability or susceptibility to infection by the eel host. Overall one can say that in spite

of this difference, for the most part, K is not affected.

Variations within the liver mass, an organ indicating energy reserves in the fish, may reflect the cost of A. crassus on the eel condition (Lefebvre et al., 2004); whereas

variations in the spleen mass, an organ responsible for immune defense and blood cell

synthesis, may reflect the response of the eel to fighting off infection (Lefebvre et al.,

2004). In the present study, there was no significant difference of LSI or SSI between

infected and uninfected eels, which is consistent with findings in infected European eel

(Moller et al., 1991; Han et al., 2008) Furthermore, Moller et al. (1991), Lefebvre et al.

(2004) and Han et al. (2008), found no relationship between LSI and intensity of infection, a result that was also found herein. Interestingly, Lefebvre and colleagues

(2004) determined that the spleen showed variation in mass in relation to parasite intensity, whereas the present study did not find such a relationship. However, Lefebvre et al. (2004) controlled for eel somatic mass and age when comparing spleen mass and parasite pressure and the present study did not. The two studies are questionably comparable.

Swimbladder function may be altered by infection with A. crassus (Kirk, 2003).

Gross pathology in European eels report fibrosis, which causes the swimbladder to

19 thicken and become opaque, resulting in a reduced lumen which becomes filled with

digested blood. Dark clots of dead and decaying nematode form, and the swimbladder

wall ends up with encapsulated adults and juveniles (Thomas & Ollevier, 1992b; Molnar

et al., 1993,1995; Wiirtz & Taraschweski, 2000; Abdelmonem et al., 2010). Fibrosis of

the swimbladder is caused by the blood sucking activity of the adult nematode and

migration of the larvae (Wiirtz & Taraschweski, 2000; Abdelmonem et al., 2010). The

breaking down and encapsulation of A. crassus is an immune response by the eel host

(for more in depth review see Kirk, 2003). Similar pathology was observed in the

American eel in the present study. Although qualitative observations made in this study

were basic as compared with other research, observations such as "thickening, dark

spotting, blood clots/clumps, hemorrhaging, etc." are on par with the mentioned pathology research on European eel.

Although in European eel A. crassus does not affect the overall condition of the eel host, the pathological damages to the swimbladder organ itself may be detrimental to the eel's ability to not only control its buoyancy, but its overall migration capabilities

(Palstra et al., 2007; Sjoberg et al., 2009). Palstra et al. (2007) assessed the effects of A. crassus on the host energy and swimbladder damage through experiments with infected eel in swim tunnels. They determined that infected eels and eels with damaged swimbladders were less efficient swimmers with reduced endurance that have a higher energy budget with a concluded overall reduced swimming performance. Palstra et al.

(2007), suggest that these effects would lengthen migration, making eels spend more energy, leaving less fat for egg production if they do spawn (Palstra et al. 2007). In contrast, Sjorberg et al. (2009) found that migration speed and short term swimming activity were not influenced by the parasite, but the study does not give detailed information about the eel behavior as it was done in the wild through mark-recapture studies. Additionally, they caution not to compare the two studies because experiments in the swim-tunnel do not allow the eel to be exposed to natural physical factors (such as current or lunar cycles), or predators, and it does not allow for rest or vertical movements.

They report that migration speed is unaffected, but the damaged swimbladder could result in impaired migration behavior (particularly vertical migration), causing eels to stay closer to shore and become exposed to nets, which was a result of their recapture experiment, as more heavily infected eels were more vulnerable to recapture in pound nets. Not only that, but this may make the eels more vulnerable to predators and lead to navigational problems because vertical migration may be required to derive directional cues (Sjorberg et al., 2009).

In order to detect the presence of A. crassus in the European eel using non­ invasive methods, Crean et al. (2003) determined that a red anus indicated a high probability of infection and high parasite abundance, and suggested that any divergence from normal coloration could be used as a diagnostic tool. The present study showed variation in anal color, from normal to red, in both infected and uninfected eels, indicating that anal color that varies from the norm is not necessarily indicative of A. crassus infection. Furthermore, intensity was not related to differences in anal color, suggesting that, for American eels in the present study, levels of intensity cannot be determined by anal color.

21 Literature Cited

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24 Lamson, H.M., Shiao, J-C, Iizuka, Y., Tzeng, W-N., and Cairns, D.K. 2006. Movement patterns of American eels (Anguilla rostrata) between salt-and freshwater in a coastal watershed, based on otolith microchemistry. Marine Biology. 149: 1567- 1576.

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27 Zar, J.H., 1996. Biostatistical Analysis, 3rd Edition. Prentice-Hall, Englewood Cliffs, NJ, 662 p. Figure 1. Taxonomy of Anguillicoloides crassus.

29 Phylum: Nematoda

Class: Secernentea

Order: Spirurida

Suborder: Camallanina

Superfamily: Dracunculoidea

Family: AnguiUcolidae

Genus: Anguillicola

Subgenus: Anguillicoloides

Species: Anguillicoloides crassus

30 Figure 2. ArcGIS map of Cape Breton (courtesy of Dollie Campbell), Nova Scotia. Each point represents a site that yielded American eels.

31 32 Figure 3. ArcGIS map of Cape Breton, Nova Scotia (courtesy of Dollie Campbell) depicting 10 sites that did not yield American eels while sampling.

33 »

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34 Figure 4. American eel ready to be necropsied.

35 36 Figure 5. Length measurement of an American eel. Length was recorded to the nearest millimetre (mm).

37 38 Figure 6. Weight measurement of an American eel. Weight was measured to the nearest O.lg. 40 Figure 7. Examining the anal color of an American eel. Image: a. Eel sample AR190710- 2; b. Close up of the same eel; arrow indicates pink color surrounding anal opening.

41 42 Figure 8. Swimbladder from American eel sample ARl 90710-2. Image a. intact swimbladder; b. opened swimbladder.

43 mmmm

44 Figure 9. The swimbladder from an American eel from Margaree Harbour that had the greatest parasite intensity with 50 parasites. Image: a. swimbladder after initial opening cut; b. swimbladder completely opened and A. crassus removed.

45 46 Figure 10. A map showing the 10 primary watersheds in Cape Breton, Nova Scotia.

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48 Figure 11. A scatter plot showing the relationship between intensity of infection and total length for American eels in Cape Breton, Nova Scotia.

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50 Figure 12. A scatter plot of intensity versus weight for American eels from Cape Breton, Nova Scotia.

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52 Figure 13. A scatter plot of condition factor versus intensity for infected American eels from Cape Breton, Nova Scotia.

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54 Figure 14. A scatter plot of liver somatic index versus intensity in infected American eels from Cape Breton, Nova Scotia.

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56 Figure 15. A scatter plot of spleen somatic index versus intensity in infected American eels in Cape Breton, Nova Scotia.

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58 Figure 16. Histological section of a category 2 American eel swimbladder showing severe multifocal melanizing necrosis (arrows).

59 60 Figure 17. Histological section of a category 2 American eel swimbladder from Cape Breton, Nova Scotia. The arrow indicates an encapsulated A. crassus larva.

61 7**

62 Figure 18. The mean intensity versus anal color of infected American eels in Cape Breton, Nova Scotia.

63 20.00-

15.00-

BROWN NORMAL ORANGE PINK RED UNKNOWN YELLOW Anal Color

Error Bars: +/-1 SE

64 Table 1. Cape Breton sites sampled for American eel during 2008 and 2009. CW = Cheryl Wall; DC = Dollie Campbell; DFO = Department of Fisheries and Oceans; KJ = Katherine Jones; NSDFA = Nova Scotia Department of Fisheries and Aquaculture

65 Site Latitude Longitude Sampled By Sampling Method Water Type n Baddeck Bay 46.112200 -60.722984 CW, NSDFA Fyke nets Estuarine 21 Bell Lake 46.290717 -60.735950 CW, NSDFA Fyke nets, Eel Pots Freshwater 4 Big Lorraine 45.934481 -59.931222 KJ Eel Pots Estuarine 26 Brec Brook 45.913668 -60.528288 NSDFA Electrofished Freshwater 1 Cheticamp Harbour 46.599825 -61.023835 Dale Larade Weir/Commercial Estuarine 52 Effies Brook 46.862809 -60.465397 Parks Canada Electrofished Freshwater 1 False Bay Brook 45.636133 -61.010917 CW, DC Electrofished Freshwater 24 Glace Bay 46.154778 -59.954672 KJ Eel Pots Estuarine 11 Lake Ainslie 46.162510 -61.252217 CW, NSDFA Fyke nets, Eel Pots Freshwater 22 Leitches Creek 46.142210 -60.351770 Leo Bungay Weir Estuarine 68 Little River 46.438799 -60.471148 Parks Canada Electrofished Freshwater 4 MacLennan Brook 46.288120 -60.641893 CW, NSDFA Electrofished Freshwater 1 Margree Harbour 46.441313 -61.103522 Gordon MacKay Weir Estuarine 128 Mill Brook 45.669469 -61.421902 DFO Electrofished Freshwater 11 Mira River 46.047467 -60.021133 KJ Eel Pots Estuarine 242 Morrisons Beach 45.763158 -60.297483 KJ Eel Pots Estuarine 9 Morrisons Brook 45.687567 -61.248283 CW, DC Electrofished Freshwater 11 Neils Brook 46.811072 -60.333459 CW, NSDFA Electrofished Freshwater 2 Northwest Brook 46.196060 -60.117550 Maureen C.-M., AM EC Electrofished Freshwater 4 NW Arm River Inhabitants 45.680450 -61.325633 CW, DC Electrofished Freshwater 24 NW Branch Baddeck River 46.292244 -60.785751 CW, NSDFA Electrofished Freshwater 6 Orangedale 45.902556 -61.091755 CW, NSDFA Fyke Nets Estuarine 6 Pondville Brook 45.540467 -60.992383 CW, DC Electrofished Freshwater 10 River Denys 45.853824 -61.215696 CW, NSDFA Fyke Nets Freshwater 8 River Ryan 46.214605 -60.081276 KJ Eel Pots Estuarine 10 Skye River 46.001610 -61.150352 CW, NSDFA Electrofished Freshwater 4 Sydney Harbour 46.140310 -60.200455 KJ Eel Pots Estuarine 1 Upper Marie Joseph Brook 45.703933 -60.424267 CW, DC Electrofished Freshwater 13 Wentworth Park 46.132588 -60.194617 KJ Eel Pots Estuarine 1 Whycocomagh Bay 45.967813 -61.126068 CW, NSDFA Fyke Nets Estuarine 3 66 Table 2. Gross pathology of the swimbladder of American eels in Cape Breton, Nova Scotia. Category range is 0-2 and is based on the observations at the time of necropsy.

67 Category Gross Pathology 0 Normal, healthy-looking Slightly thickened, partially blackened, or 1 slightly lumpy Extremely thickened, completely hardened, 2 hemorrhaging, or many clumps of dry blood

68 Table 3. Prevalence and mean intensity ±SD for the 10 primary watersheds of Cape Breton, Nova Scotia.

69 Watershed n Prevalence(%) Mean Intensity ± SD

Cheticamp 52 7.7 3.25 ±4.50

Grand 46 52.2 4.88 ± 5.08

Indian 5 0 0 Isle Madame 10 40 1.25 ±0.5

Margaree 150 46.7 4.41 ± 6.60

Mira/Salmon 364 43.1 4.77 ± 5.53

North Baddeck 31 35.5 4.91 ± 6.72 River Denys 21 66.7 4.86 ± 3.88

River Inhabitants 46 52.2 3.75 ± 3.60

Wreck 3 0 0

70 Table 4. Prevalence and Mean Intensity (±Standard Deviation) for 30 sites yielding American eels in Cape Breton, Nova Scotia. Sample size and number of eels infected are also indicated. Bolded names indicate the watershed with sites within listed below.

71 Watershed with Sites n Infected Prevalence Mean Intensity Cheticamp Cheticamp Harbour 52 4 7.69 3.25 (4.50) Grand Morrison Beach 9 8 88.89 7.25(6.56) Marie Joseph Brook 13 7 53.85 5.71 (5.28) False Bay Brook 24 9 37.50 2.11(0.93) Indian Little River 4 0 0.00 0.00(0.00) Maclennan Brook 1 0 0.00 0.00 (0.00) Isle Madame Pondville Brook 10 4 40.00 1.25 (0.50) Margaree Margaree Harbour 128 62 48.44 4.40 (6.74) Lake Ainslie 22 8 36.36 4.50(5.63) Mira/Salmon Big Lorraine 26 17 65.38 5.00 (6.82) Brec Brook 1 1 100.00 1.00(0.00) Glace Bay 11 6 54.55 7.67 (9.52) Leitches Creek 68 20 29.41 5.55 (5.67) Mira River 242 104 42.98 4.67(5.19) Northwest Brook 4 4 100.00 1.00(0.00) River Ryan 10 3 30.00 4.00 (4.36) Sydney Harbour 1 1 100.00 1.00 (0.00) Wentworth Park 1 1 100.00 3.00 (0.00) North Baddeck Baddeck Bay 21 10 47.62 5.30 (6.96) Bell Lake 4 0 0.00 0.00 (0.00) NW Branch of Baddeck Riv. 6 1 16.67 1.00(0.00) River Denys Orangedale 6 4 66.67 5.25 (4.99) River Denys 8 6 75.00 6.17(4.17) Skye River 4 1 25.00 3.00(0.00) Whycocomagh Bay 3 3 100.00 2.33 (0.58) River Inhabitants Mill Brook 11 3 27.27 1.33 (0.58) Morrison Brook 11 4 36.36 1.75 (0.50) NW Arm of Riv. Inhab. 24 17 70.83 4.65 (3.95) Wreck Effies Brook 1 0 0.00 0.00 (0.00) Neils Brook 2 0 0.00 0.00 (0.00) Unknown 2 1 50.00 1.00(0.00) Overall 730 309 42.33 4.58 (5.60)

72 Table 5. Prevalence and mean intensity (±Standard Deviation) based on distance fromse a for infected American eel found in freshwatersite s in Cape Breton, Nova Scotia.

73 Distance From Sea (km) n Prevalence (%) Mean Intensity ± SD

0-15 75 39 2.62 ±3.11

15-30 65 54 4.54 ± 4.22

30-45 10 10 1.00 ±0.00

74 Table 6. Prevalence and intensities of A. crassus in European eels throughout Europe from selected publications MI = mean intensity; *L.A. - Leimersheimer Allrhein; *Germ = Germersheim.

75 Location n Prevalence (%) Intensity Author Germany (Elbe) 1778 57.7 Ml = 7.5 Moller etal., 1991 Belgium (Albert Canal) 345 90.2 Ml = 17 Thomas & Ollevier, 1992) England (River Thames) 436 12-32 1-5 Pilcher & Moore, 1993 Germany (L.A.)* 61 83.6 Ml = 5.3 (Germ.)* 60 76.7 Ml = 5.0 Sures et al., 1999 Ireland (Erne System) 432 9.9 Ml = 6.7 Evans et al., 2001 Czech Republic (Korycany water reservoir) 143 64.4 Ml = 6.6 Palfkova & Navratil, 2001 Scotland 2 100 2-7 Lyndon & Pieters, 2005 Ireland Morrissey & McCarthy, (Camus Bay) 301 0-63 0-4.22 2007 Wickstrom unpub data in Sweden 15-94 Sjoberg et al., 2009 Egypt (East Delta) 65 10.7 Ml = 1.85 Abdelmonem et al., 2010 Netherlands (Rhine) 56 71.4 4.4 (Ijsseleer) 36 78.8 6.1 Haenen etal., 2010

76 Table 7. Prevalence and mean intensities of A. crassus in American eels from North American locations for American eels from selected publications. Nova Scotia is highlighted, showing one site from the mainland (Antigonish) and two Cape Breton sites (Margaree Harbour and Bras d'Or Lakes)

77 Mean Location n Prevalence (%) Intensity Author Chesapeake Bay 329 10-29 3.1-6.2 Barse & Secor, 1999 Hudson River 160 0-12 1.0-1.7 Barse & Secor, 1999 Chesapeake Bay 423 13-82 3.1-6.2 Barse et al., 2001 North Carolina 1111 52 3.9 Moser etal., 2001 Rhode Island 129 28-69 2.2-3.5 Aieta & Oliveira, 2009 Massachusetts 394 7-76 1.8-5.3 Aieta & Oliveira, 2009 Maine 313 26-65 1.4-4.5 Aieta & Oliveira, 2009 New Brunswick 60 3.1-3.6 1 Aieta & Oliveira, 2009

Nova Scotia Antigonish 27 14.8 2.5 Margaree Hbr. 26 30.1 6.1 Bras d'Or Lake 28 14.3 4.0 Aieta & Oliveira, 2009

78 Chapter 2: Inferring the origin of Anguillicoloides crassus infecting American eel

(Anguilla rostrata) in Cape Breton, Nova Scotia using the cytochrome c oxidase I

gene of the mitochondrial DNA

79 Introduction

The loss of biodiversity and species extinctions due to the introduction and spread of non-native species has become a global ecological and conservation crisis (Gurevitch

& Padilla, 2004). Among the most heavily invaded systems in the world, are coastal estuarine and marine habitats (Vignon & Sasal, 2010). Human-mediated transportation, such as in the ballast water of ships, has transferred thousands of freshwater, estuarine and marine species around the world and the rate of biological invasions in these systems has increased substantially in recent years (Carlton, 1996; Cohen & Carlton, 1998; Ruiz et al., 2000; Roman & Darling, 2007; Vignon & Sasal, 2010). Ballast water is used for stability on ships when not carrying cargo. Water is typically loaded into the ship's ballast tank or cargo hold at one port, and discharged en route or while loading cargo at the next port (Lavoie et al., 1999; Wonham et al., 2001). Additional sources of invasion include the aquarium industry, aquaculture, movement of shellfish and bait, the opening of new shipping channels, and intentional release by humans (Ruiz et al., 1997; Roman,

2006; Vignon & Sasal, 2010).

In 2007, the invasive parasitic nematode (Anguillicoloides crassus (Kuwahara,

Niimi & Itagaki, 1974)) was recorded for the first time in American eel (Anguilla rostrata) in Canadian waters (Aieta & Oliveira, 2009; Rockwell et al., 2009). Where it occurs, Angullicoloides crassus is found in the swimbladder of anguillid eels (Anderson,

2000). Originating from East Asia, A. crassus is widespread among its native host, the

Japanese eel (Anguilla japonica) (Koops and Hartmann 1989). However, the parasite is invasive in its infected, non-native hosts, including both the European eel (Anguilla 80 anguilla) and the American eel (Nagasawa et al., 1994; Kirk, 2003; Han et al., 2008). It is thought that international trade of parasitized Japanese eels from Taiwan into Germany in

1982 was responsible for the infection of European eels (Moravec, 1994; Kirk, 2003). In

1995, the parasite was reported in North America in a Texas aquaculture facility which had received infected elvers from South Carolina (Fries et al., 1996; Aieta & Oliveira,

2009). Since its introduction into North America, A. crassus has spread rapidly northward along the east coast (Barse & Secor, 1999; Moser et al, 2001; Aieta &

Oliveira, 2009). It is unknown as to how the parasite has spread so quickly, however, it is speculated that the parasite may be transported in the ballast water of ships, in live transport of eels as recreational bait, and in the natural movement of intermediate, paratenic and final hosts (Aieta & Oliveira, 2009).

In order to gain a better understanding of species invasions and develop strategies against their introduction and dispersal, molecular markers are now being used to study transmission, host-specificity and patterns of speciation (Lee, 2002; Criscione et al.,

2005; Taraschewski, 2006). Currently, molecular research on parasites is in its youth

(Criscione et al., 2005), and very little molecular research has been conducted on A. crassus. Wielgoss et al. (2007) developed seven microsatellite primers for A. crassus and, in 2008, they furthered their research by examining the seven microsatellites, as well as a universal mitochondrial marker (cytochrome c oxidase I - COI) in parasites from eels sampled in eleven European locations, three East Asian sites and one North

American river (the Saint Jones) (Wielgoss et al., 2007, 2008). Looking at the North

American, Saint Jones River samples, they found that these nematodes shared a unique 81 haplotype with a Japanese eel population, indicating these samples likely originated directly from Japan, as opposed to a primary introduction from Taiwan or secondary introduction from Europe. Using the Wielgoss study as a platform, this chapter aims to examine the COI gene of A. crassus from Cape Breton in order to better understand its origin. Additionally, examination of samples from the United States (USA), Europe,

Africa and Taiwan will also be conducted and compared to that reported in the literature.

Methodology

American eels were collected through a distributional survey and donations from other researchers throughout the summer of 2008 and 2009 (Table 1) (Wall et al., in prep). Nematode donations (A. crassus preserved in ethanol) were also provided from locations in the USA, Europe, Africa and Taiwan (Table 2). The eels were necropsied at

Cape Breton University and Saint Mary's University and any A. crassus removed from eels were placed in ethanol (C2H5OH) for molecular research.

Prior to DNA extraction, the outer cuticle was removed as much as possible and discarded and approximately half of the nematode was used for DNA analysis. The sample was then placed in sterile distilled water for approximately 15 minutes to remove residual ethanol. DNA was extracted from each A.crassus by following the procedure outlined in the Qiagen QIAamp® DNA Minikit 50 tissue protocol. The DNA was then put in a Polymerase Chain Reaction (PCR). Each 30 ul PCR reaction consisted of 2 ul

DNA template, 3 ul 10 x Taq PCR buffer, 3 ul of dNTP (10 mM), 3 ul of MgCl2

(50mM), 10 pmol of each universal primer, 0.2 ul Taq DNA polymerase. PCR samples were run in a thermalcycler (BioRad) according to Wielgoss et al. (2008) and consisted of 82 an initial 3 minutes at 94°C followed by 35 cycles of denaturation for 35 seconds at 95°C, annealing for 1 minute at 40°C, and elongation for 1 minute 30 seconds at 72°C, and then ending with a final elongation step of 15 minutes at 72°C. The PCR products were run out on a 1.5% agarose gel against alOO bp Ladder Sharp DNA Marker (United

Bioinformatica Inc.) containing a concentration gradient ranging from 30 ng to 80 ng, thus allowing for comparison and estimation of sample DNA concentrations. Following the run, gels were stained with ethidium bromide (EtBr) for approximately 20 minutes and then visualized and digitally photographed using UV transluminescence in the

Syngene ChemiGemus .

To determine the appropriate DNA concentration for sequencing, 2 PCR samples underwent a series of serial dilutions before being run against the concentration gradient

100 bp DNA ladder. The gel lanes were loaded with 10 ul of diluted PCR product in 2 ul of 6 x loading buffer (0.25% (weight/volume) bromophenol blue and 30%

(volume/volume) glycerol in water). The banding brightness of the sample was compared with that of the DNA ladder in order to obtain 2 similar bands of 30 ng/10 p.1 (Figure 1).

A dilution of 1:40 was selected for both samples and the approximate concentration of

DNA in each sample was calculated by CiVi = C2V2, where Ci and C2 are the initial and final concentrations and Vi and V2 are the initial and final volumes respectively. These concentrations were used to determine the appropriate concentrations for sequencing, as suggested by GENEWIZ Inc., New Jersey, U.S.A., the company used for sequencing products throughout this project. Samples of 40 ng/10 ul, 50 ng/10 ul, 60 ng/10 ul, 70 ng/10 ul and 80 ng/10 ul were sent in order to determine the best concentration for the A. 83 crassus sequence. It was determined that 70 ng/10 ul was appropriate for both samples that were sequenced and this was used as the concentration for all remaining samples.

Sequence Editing

The sequences were edited individually using CodonCode and BioEdit sequence editors. Once sequences were edited, and the primers trimmed off, consensus sequences were made. All sequences were aligned in MEGA 5 (Tamura et al., 2011) with all A. crassus COI samples present in GenBank (Wielgoss et al., 2008). All of the sequences were then aligned in Clustal X vers. 2 (Larkin et al., 2007) and saved. Tree-Puzzle version 5.2 (Schmidt et al., 2002) was then used to obtain the transition and transversion ratio and the rate of heterogeneity alpha (a) value. The species Hysterothylacium aduncum (Nematoda: Anisakidae) was selected from GenBank as the outgroup; a group more distantly related to any taxa in the ingroup than the ingroup taxa are related to each other (Hall, 2004). All sequences, including the outgroup, were re-aligned in MEGA 5, trimmed, and aligned using Clustal X version 2 (Larkin et al., 2007). Using PHYLIP version 3.69 (Felsenstein, 1989), a bootstrap of 1000 was performed and a distance matrix was created using the transition/transversion ratio and a acquired earlier. A Fitch tree was built using multiple data sets (of 1000) and from that a consensus tree was constructed. The consensus tree was viewed in the program FigTree version 1.3 (Morariu,

2008). A maximum likelihood tree was constructed in Tree-Puzzle with the quartet puzzling software, after which the consensus tree was built using 1000 bootstrap trees.

The HKY85/F84 model of evolution was used. The resulting tree was viewed in FigTree

84 (Figure 2). Both trees were rooted using the H. aduncum outgroup and bootstrap percentages are presented on the branch nodes.

Results

Sequence Analysis

A total of 115 samples were sent for sequencing, however, not all sequence results were optimal for editing (due to poor quality sequence results). A final total resulted in 65 nematodes from Cape Breton, 7 nematodes from the USA, 3 from Morocco and 2 from

Germany (Table 3; for Cape Breton coordinates see Table 1). These samples were aligned with all samples from Wielgoss et al. (2008).

All Cape Breton sequences were identical and put into a single consensus sequence. This was also the case for the USA samples. When calculating the transition/transversion ratio for all sequences, identical sequences were identified and all but one sequence was removed for each group and the group received a new name. The identical sequences and new group names are provided in Table 4. Cape Breton and USA samples were identified as being identical to all samples found in SEQ 9, however, they were kept separate from the grouped sequences (SEQ 9) for the purpose of identifying them geographically in the tree. The sequence from Uecker River (Germany) was grouped into SEQ 3, while the sequence from the Elbe River (Germany) was grouped into SEQ 10. Sequences from the Sebou River (Morocco) and Loukos River (Morocco) were both grouped into SEQ 11, while the third Moroccan sequence (from the Sebou

River) remained on its own.

85 Phylogenetic Tree

The phylogenetic tree constructed for this study can be seen in Figure 2. The Cape

Breton and USA samples fell into the same clade as SEQ 9 with 800 of 1000 bootstrap repeats. Uecker River, Germany (SEQ 3) fell into a clade with SEQ 4, SEQ 5 and with a sample from the Kao-Ping, Taiwan. The sample from the Elbe River, Germany fell out with a sample from Aland Islands, Finland and a sample from Lake Neagh, Great Britain.

The two Morrocan sequences in SEQ 11 fell into the same clade as SEQ 12, which consists of two samples from Angers, Loire, France. The last Moroccan sequence was its own haplotype.

Discussion

In 2008, Wielgoss et al. used microsatellites and the cytochrome c oxidase I (COI) gene to examine the genetic diversity of A. crassus in Japanese, European and American eel. The only North American site included in their study was the St. Jones River, USA.

They determined that both the nuclear and mtDNA from North American samples shared distinct signatures with the Japanese samples. They suggest that it is likely that the USA samples were of direct Japanese eel origin, as opposed to a primary colonization from

Taiwan or secondary movement from Europe (Wielgoss et al., 2008). In the present study, only the COI gene was used and both the Cape Breton and USA samples were found to be the same as that of the unique haplotype identified by Wielgoss et. al. (2008)

(Figure 2). This provides further evidence of a direct transfer from the Japanese eel to the

American eel in the USA.

86 In Cape Breton, firstly, it was hypothesized that the invasion may have occurred via the spread of the parasite from the USA, up the east coast and into Canada, through intermediate, paratenic and/or final host movement or in the ballast water of ships (Aieta

& Oliveira, 2009; Rockwell et al., 2009). Cape Breton appears to have populations of eels that were infected before those in mainland Nova Scotia, which seemed unusual as one would expect the mainland to be infected first if it was a direct spread of the parasite northward along the east coast. Secondly, it was suggested that the nematode may have been introduced from Europe, Japan, or other countries via ballast water from container ships. Thirdly, it was possible that both of the scenarios could be occurring and there were separate introductions from parasitic spread within hosts from the US and through ballast water introduction from the USA, Europe, Japan and/or various countries. Lastly, it is conceivable that due to the Sargasso Sea being the spawning ground for both

American and European eel, it is possible that European eel may stop along the North

American coast during their migration, thus carrying the parasite into North American waters. The results of the current study suggest that movement of the parasite northward along the east coast of North America via intermediate, paratenic or final host, or introduction via ballast water from USA or Japanese ships was responsible for the invasion of the parasite into Canada, as no European signatures were found. This suggests that introduction from the European eel is not occurring and, furthermore, that European eel are not entering North American rivers on their way to the Sargasso Sea.

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89 Ruiz, G. M., Fofonoff, P. W., Carlton, J. T., Wonham, M. J. & Hines, A. H. 2000. Invasion of coastal marine communities in North America: apparent patterns, processes, and biases. Annual Review of Ecology, Evolution and Systematics, 31, 481-53

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90 Figure 1. 1.5% agarose gel showing the concentration gradient of two Anguillicoloides crassus samples (53 and 54). These samples were used to determine the optimum concentration for sequencing. M= Molecular Marker (lOObp DNA Ladder) - numbers next to ladder indicate the concentration in ng/10 ul; ratios above lanes indicate dilution factor of the sample; red circles indicate the dilution selected for sequencing (~30 ng/10 ul).

91 53 M 1/5 1/10 1/20 1/40 1/80

54

92 Figure 2. Maximum-Likelihood phylogenetic tree of Anguillicoloides crassus cytochrome c oxidase I haplotypes. The outgroup was defined as H. aduncum (Hysterothylacium aduncum). Values on the nodes represent the percentage of bootstrap replicates out of 1000. The bolded names indicate a haplotype or group that contains haplotype(s) that were examined in the present study. The box indicates the clade of interest in this study containing the Cape Breton, United States (USA) and SEQ9 haplotype.

93 -OER09 -SEQ19 -M1K09 -SEQ 18 -KA054 -Cs! -MTK05 -SEQ3 -KA219A -SEQ4 -SEQ5

SFCAPE -BRETON SEQ9 -YA223C

0.07 94 Table 1. Cape Breton sites sampled for American eel during 2008 and 2009. CW = Cheryl Wall; DC = Dollie Campbell; DFO = Department of Fisheries and Oceans; KJ = Katherine Jones; NSDFA = Nova Scotia Department of Fisheries and Aquaculture

95 Site Latitude Longitude Sampled By Sampling Method Water Type n Baddeck Bay 46.112200 -60.722984 CW, NSDFA Fyke nets Estuarine 21 Bell Lake 46.290717 -60.735950 CW, NSDFA Fyke nets, Eel Pots Freshwater 4 Big Lorraine 45.934481 -59.931222 KJ Eel Pots Estuarine 26 Brec Brook 45.913668 -60.528288 NSDFA Electrofished Freshwater 1 Cheticamp Harbour 46.599825 -61.023835 Dale Larade Weir/Commercial Estuarine 52 Effies Brook 46.862809 -60.465397 Parks Canada Electrofished Freshwater 1 False Bay Brook 45.636133 -61.010917 CW, DC Electrofished Freshwater 24 Glace Bay 46.154778 -59.954672 KJ Eel Pots Estuarine 11 Lake Ainslie 46.162510 -61.252217 CW, NSDFA Fyke nets, Eel Pots Freshwater 22 Leitches Creek 46.142210 -60.351770 Leo Bungay Weir Estuarine 68 Little River 46.438799 -60.471148 Parks Canada Electrofished Freshwater 4 MacLennan Brook 46.288120 -60.641893 CW, NSDFA Electrofished Freshwater 1 Margree Harbour 46.441313 -61.103522 Gordon MacKay Weir Estuarine 128 Mill Brook 45.669469 -61.421902 DFO Electrofished Freshwater 11 Mira River 46.047467 -60.021133 KJ Eel Pots Estuarine 242 Morrisons Beach 45.763158 -60.297483 KJ Eel Pots Estuarine 9 Morrisons Brook 45.687567 -61.248283 CW, DC Electrofished Freshwater 11 Neils Brook 46.811072 -60.333459 CW, NSDFA Electrofished Freshwater 2 Northwest Brook 46.196060 -60.117550 Maureen C.-M., AMEC Electrofished Freshwater 4 NW Arm River Inhabitants 45.680450 -61.325633 CW, DC Electrofished Freshwater 24 NW Branch Baddeck River 46.292244 -60.785751 CW, NSDFA Electrofished Freshwater 6 Orangedale 45.902556 -61.091755 CW, NSDFA Fyke Nets Estuarine 6 Pondville Brook 45.540467 -60.992383 CW, DC Electrofished Freshwater 10 River Denys 45.853824 -61.215696 CW, NSDFA Fyke Nets Freshwater 8 River Ryan 46.214605 -60.081276 KJ Eel Pots Estuarine 10 Skye River 46.001610 -61.150352 CW, NSDFA Electrofished Freshwater 4 Sydney Harbour 46.140310 -60.200455 KJ Eel Pots Estuarine 1 Upper Marie Joseph Brook 45.703933 -60.424267 CW, DC Electrofished Freshwater 13 Wentworth Park 46.132588 -60.194617 KJ Eel Pots Estuarine 1 Whycocomagh Bay 45.967813 -61.126068 CW, NSDFA Fyke Nets Estuarine 3 Table 2. Donated Anguillicoloides crassus by location and who collected them.

97 Location Collected By France Francoise Daverat Germany Janek Simon, Florian Nagal Ireland Russell Poole, Ciar O'Toole Morocco Florian Nagal Sweden Hakan Wickstrom, Jennie Dahlberg Taiwan Yu San Han Mark Sullivan, Dewyane Fox, Lori United States Brown

98 Table 3. Location and number of samples of Anguillicoloides crassus that underwent sequencing of the cytochrome c oxidase 1 gene.

99 # of samples Site sequenced CAPE BRETON Baddeck Bay 6 Big Lorraine 2 Cheticamp Hbr. 3 False Bay Br. 2 Glace Bay 2 Lake Ainslie 5 Leitches Creek 4 Margaree Hbr. 10 Upper Marie Joseph Br. 2 Morrisons Br. 2 Mira River 9 Orangedale 2 River Denys 5 River Inhabitants 7 River Ryan 1 Pondville Br. 2 Skye River 1 USA Bargaintown, NJ 2 Delaware Bay, DE 1 St. Jones River, DE 3 Tuckerton, NJ 1 MOROCCO Loukos 1 Sebou 2 GERMANY Elbe 1 Uecker 1 Table 4. Identical sequences that were identified during sequence analysis. Each group name represents one single sequence from the list of identical sequence. Bolded sequences represent samples done in the present study. CB: Cape Breton, Nova Scotia, Canada; USA: United States of America; GerUEC: Uecker River Germany; GerELB: Elbe River Germany; M0I86S: Sebou River Morocco; Mol84L: Loukous River, Morocco; ALA: Aland Islands, Finland; OER: Kullen, Oresund/Kattegat, Sweden; SLP: Slapton Ley, Great Britain; NEA: Lake Neagh, Great Britain; SHA: Lough Dergh, Shannon, Ireland; FRE: Bois Jolie, Fremur, France; VIL: Brain-sur-Vilaine, France; LOI: Angers, Loire, France; ORI: Oria, Spain; RHO: Camargue, Rhone, France; TIB: Roma, Tiber, Italy; KAO: Tung-chiang, Kao-Ping, Taiwan; MIK: Mikawa Bay, Japan; YAM: Yamaguchi, Fushino, Japan; STJ: St. Jones River, USA.

101 Group Name Identical Sequences SEQ1 YA223A, YA223B SEQ2 KA226B, KA226C, KA216 GerUEC, ALA103, ORI34, ORI25, ORI23, KA015, KA038A, ORI 18, KA051, ORI 10, KA219A, SEQ3 STJ09, STJ17, STJ33, VIL36, VIL33, VIL30, MIK01, VIL28, MIK02, VIL26, VIL14, OER06, OER14, VIL03, TIB33, OER32, OER29, OER26, OER23 SEQ4 OER01,TIB07 SEQ5 SHA31, NEA15 SEQ6 KA064A, KA229A ORI41, KA235B, ORI40, ORI37, SHA02, ORI36, ORI32, ORI30,ORI29, SHAH, SHA17, SHA20, ORI28, LOIOl, ORI27, LOI06, LOI12, OER21, RHO01, ORI26, LOI13, LOI15, ORI24, ORI22, LOI17, LOI21, RHO07, RH021, TIB03, TIB04, TIB05, LOI28, ORI21, LOI34, ORI20, RHO10, LOI35, LOI37, SEQ7 LOI39, ORI17, ORI19, ORI01, VIL06, TIB49, TIB47, TIB46, TIB45, TIB44, TIB41, TIB36, TIB28, TIB16, TIB15,TIB14, TIB11, TIB08, RH049, RH048, RH047, RH046, RH043, RH042, RH041, RH038, RH037, RH032, RH031, RHO30, RH024, RH023, RH017, RH015, RH013, RH012, RHOll, LOI51, LOI50, LOI49, LOI46, LOI45, LOI40 SEQ8 KA229B, VIL29 CB, USA, STJ01, STJ02, STJ03, STJ04, STJ05, STJ06, STJ28, MIK22, STJ07, STJ29, MIK23, STJ08, STJ31, MIK24, STJ10, STJ34, MIK25, STJ11, STJ35, MIK26, STJ12, STJ37, MIK27, STJ14, STJ38, SEQ9 MIK28, STJ15, STJ16, MIK12, STJ18, MIK13, STJ20, MIK15, STJ21, MIK16, STJ24, MIK17, STJ25, MIK18, STJ26, MIK19, STJ27, MIK21 SEQ10 GerELB, TIB29, KA058, TIB27, TIB25, OER16, LOI55 SEQ11 M0I86S, Mol87L, ORI38 SEQ12 LOI42, LOI57 KA042C, ORI33, ORI31, KAO50, ORI35, ORI39, ORI09, ORI11, ORI14, KA062A, KA062B, KAO04A,KAO56, KA055, KAO04C, KA004D, KAO10, VIL38, VIL34, KA215, KA214, KA064B, VIL24, VIL23, VIL22, VIL27, VIL20, VIL19, VIL18, YA222A, KA226A, KA219B, KA239, KA228B, YA222C, VIL05, VIL04, VIL01, VIL10, VIL11, VIL07, RH016, NEA02, NEA04, NEA12, NEA16, NEA17, RHO04, NEA09, NEA08, NEA49, NEA30, NEA29, RH034, RH025, RH035, NEA26, SEQ13 TIB13, TIB12, RHO09, NEA23, FRE02, TIB19, TIB18, TIB09, TIB37, NEA20, TIB35, TIB30, FRE07, OER31, ALA114, ALA112, SHA05, ALA110, FRE22, ALA19, ALA18, ALA21, ALA105, ALA23, ALA12C, ALA12, ALA31, ALA24, FRE21, SHA09, FRE26, SHAM, FRE18, SHA18, FRE17, OER24, SHA29, FRE35, LOI10, SHA33, LOI02, FRE30, LOI48, FRE43,FRE48, COR21, OER17, FRE49, OER11, OER05, FRE52, FRE53, OER03, LOI33, FRE56, FRE60, OER08, SHA35, LOI53, LOI38, LOI47, SHA24, LOI25, SHA23, LOI09, LOI56 SEQ14 KA229C, KA235D, KA228A, KAO04B, KA038B, KAO40A, KA042A, KA042B, KA057, KA227A SEQ15 OER12, OER15, 0ER22 SEQ16 KA045, KA227B SHAOl, SHA12, SHA07, SHA15, SHA16, SHA19, SHA25, SHA27, SHA30, SHA34, SHA36, SHA37, SEQ17 NEA01, NEA05, NEA11, NEA13, NEA14, NEA18, NEA21, NEA22, NEA24, NEA27 SEQ18 KAO40B, KA049, NEA25 SEQ19 OER28, OER30, OER25, OER20, OER19, OER18, OER13, OER07, ALA109, STJ19 SEQ20 COR16, COR18, COR20, COR24, COR25, COR27, COR28, COR29, COR30, COR31, COR32,NEA28 SEQ21 COR22, COR26, COR19 FRE04, FRE05, FRE14, FRE15, FRE19, FRE24, FRE27, FRE36, FRE39, FRE45, FRE47, FRE54, SEQ22 FRE62, FRE64, FRE70, SHA04, SHA13, NEA06, NEA07, NEA10, NEA50, VIL12, VIL13, VUL16, VIL32, VIL37, VIL40 SEQ23 MIK29, MIK30 SEQ24 MIK11, MIK20 Summary

It is evident from the present study that throughout most of Cape Breton Island,

Nova Scotia, American eels are infected with Anguillicoloides crassus. This infection was likely introduced in the southwestern portion of the island as a result of movement of the nematode up the east coast of North America from the United States (Barse & Secor,

1999; Moser et al., 2001; Aieta & Oliveira, 2009). This could have occurred through eel host, intermediate and paratenic host movement, or from transport in the ballast water in either commercial or recreational vessels (Aieta & Oliveira, 2009). Infection is occurring all across Cape Breton Island, in freshwater lakes and streams and in estuaries, which is consistent with reports from American eel in the United States and European eel overseas

(Hoglund et al., 1992; Pilcher & Moore, 1993; Lefebvre et al, 2002; Kirk, 2003). All sizes of American eel in Cape Breton are infected, with the number of nematodes increasing as eel length increases (Moller et al., 1991; Thomas & Ollevier, 1992b;

Molnar et al., 1994; Moser et al., 2001; Lefebvre et al., 2002). There is no significant effect of the nematode on the liver or spleen as reported by Lefebvre et al. (2004) and, contrary to the study by Crean et al. (2003), prevalence and intensity cannot be determined based on the anal redness of the American eel in Cape Breton. While severe pathological damage can be seen in the swimbladder, the overall condition of the eel remains unaffected, at least for parameters measured in the present study, a conclusion consistent with research on European eel condition (Moller et al., 1991; Moser et al.,

2001; Morrissey & McCarthy, 2007; Han et al., 2008). This does not take into account

103 the eel's ability to control its buoyancy in the water column with a damaged swimbladder, a factor that could have detrimental effects on the eel's migration to the

Sargasso Sea. If the eel is unable to complete its migration to the spawning grounds, it can be argued that A. crassus may play a role in the decline of the whole panmictic

American eel population. Furthermore, the effects of the parasite on the eel in the wild are unknown, such that it may be causing the eel stress, making it more vulnerable to predation and other bacterial infections, or more likely to get caught in dams and traps; all scenarios that could result in eel mortality.

Future Research Considerations

Research on A. crassus in Canada is still in the early stages. The parasite has now been reported on mainland Nova Scotia, as well as New Brunswick (D. Campbell, pers. comm.). Further investigation into the remaining Atlantic Provinces (Newfoundland and

Labrador and Prince Edward Island) should be undertaken, as well as Quebec and

Ontario. Elvers from the east coast of Canada, including Nova Scotia, are transported to

Ontario for farming and eventual stocking of wild eel populations, thus introduction of the parasite into Ontario waters seems inevitable, if it has not already occurred. Further research should also be conducted throughout North America to determine the pathology caused by the parasite to the American eel. Where extensive research has been done in

Europe on the European eel, little is known about the effects of the parasite within the

American eel population.

104 The indirect life cycle of the parasite may be playing a large role in its rapid dispersal, thus research on the intermediate and paratenic hosts in North America is imperative. Sampling and necropsy offish species co-habiting rivers and lakes with

American eel may indicate possible paratenic or intermediate hosts. Examining similar fish species in North America that were found to act as hosts in Europe may aid in the process.

While the present genetic research indicates a direct spread of the parasite from the United States into Canada, further genetic research should be undertaken to examine the nematode's evolutionary history. Microsatellite primers, such as those developed by

Wielgoss et al. (2007) should be used to determine genetic diversity within the A. crassus population in North America.

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