Assessment of the Immune Response to Posthodiplostomum Minimum Infection in Bluegills

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Assessment of the Immune Response to Posthodiplostomum minimum Infection in Bluegills

Olamide S. Olayinka Eastern Illinois University

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Recommended Citation Olayinka, Olamide S., "Assessment of the Immune Response to Posthodiplostomum minimum Infection in Bluegills" (2020). Masters Theses. 4823. https://thekeep.eiu.edu/theses/4823

This Dissertation/Thesis is brought to you for free and open access by the Student Theses & Publications at The Keep. It has been accepted for inclusion in Masters Theses by an authorized administrator of The Keep. For more information, please contact [email protected]. ASSESSMENT OF THE IMMUNE RESPONSE TO POSTHODIPLOSTOMUM

MINIMUM INFECTION IN BLUEGILLS

Olamide S. Olayinka

D.V.M., University of Ibadan, 2015

(Under the Direction of Dr Jeffrey R. Laursen)

A Thesis

Submitted for the Requirements for the Degree of

Master of Science

Department of Biological Sciences

Eastern Illinois University

August 2020

ABSTRACT

Bluegills (Lepomis macrochrius) are common intermediate hosts for white grub

(Posthodiplostomum minimum). They tolerate heavy infections with minimal effect on condition and continue to accumulate metacercariae as they age. This suggests that any immune response to this parasite might not be effective. This study was conducted to better understand the immune mechanisms underlying P. minimum infection in bluegills.

Infected organs (liver, kidney, and heart) were examined histologically, and serum from infected fish was tested for antibodies to white grub. Juvenile flukes were recovered from isolated metacercarial cysts. Polyclonal antibodies were produced in mice against white grub homogenate, bluegill serum, bluegill Immunoglobulins (IgM), green sunfish serum and bluegill mucus. These were used in serological assays (dot blots, Indirect ELISA, and

Western Blots) to examine the immunological response of infected bluegills to the white grub antigen prep, as well as potential cross reactivity with other trematode antigens.

Histological assessment of infected tissues revealed P. minimum in double walled cysts surrounded by a clear zone and mild leucocytic infiltration. Melano-macrophage centers, characterized by brown pigmentation of cells, were observed in the surrounding tissues. In the heart, P. minimum metacercariae were confined to the epicardium.

There were shared epitopes between the white grub antigen prep and fish proteins, when tested using mouse antibodies against fish tissues. Although, mouse anti-white grub did not recognize fish proteins, it bound to shared epitopes in the white grub and yellow grub

(Clinostomum sp.) preps. The ELISA assay proved unreliable. Absorbance values showed a general decrease to background levels across a serial dilution series. However, there was

high variability between fish, and there was no difference in absorbance values between heavily versus lightly infected fish. I attempted to identify specific proteins that were recognized by serum from infected bluegills using western blot analysis. Rather than seeing additional bands when bluegill serum was added prior to mouse anti-bluegill serum, results revealed no bands. This implied that the ELISA absorbance was not due to fish antibodies binding to white grub antigens.

Thus, we could not detect specific systemic antibodies to P. minimum in infected fish, and this effective absence may have contributed to the persistent heavy infections commonly observed. Conversely, melanization an innate immune mechanism may be the prominent immunological reaction to white grub infection in bluegills. An examination of skin mucus is recommended, as this may provide insights into mucosal antibodies and their potential roles in P. minimum infection.

ACKNOWLEDGEMENTS

First and foremost, I am deeply grateful to my major advisor, Dr Jeffery Laursen, and other members of my thesis committee (Dr Barbara Carlsward and Dr Eloy Martinez) for their unflinching support throughout my research and academic pursuits at Eastern Illinois

University. This work would have been impossible without their constant guidance and suggestions.

I would also like to extend gratitude to members of staff of the Department of Biological

Sciences for their assistance during my fish sampling, mice husbandry and procurement of materials for the study. Derrick Douros, Cassi Carpenter and Sandy Baumgartner were instrumental in the success of the work.

I would like to appreciate the Graduate School for providing funds for this study through the Research and Creativity Grant award. This supported the purchase of materials that were not readily available in the department.

Lastly, my sincere appreciation goes to my family and friends for their immense support all through my years of study. I am thankful to God for life, friends and family, and a memorable sojourn during my graduate studies at Eastern Illinois University.

TABLE OF CONTENTS

ABSTRACT 2

ACKNOWLEDGEMENTS 4

LIST OF TABLES 7

LIST OF FIGURES 8

INTRODUCTION 12

Sunfishes 13

Common Digenean Trematodes of Sunfishes 14

White grub 14

Yellow Grub 16

Fish Immune system 17

The Innate Immune System 18

The Adaptive (Acquired) Immune system 19

T lymphocytes 20

B lymphocytes 21

Immune Response to Parasitic Infections in Fish 21

Immune Evasion Mechanisms by Fish Parasites 24

METHODS 27

Fish Sampling 27

Tissue Processing and Staining 28

White grub Antigen Preparation 28

Mice Immunization 29

Dot Blot 30

SDS Polyacrylamide Gel Electrophoresis and Western Blotting 30

Indirect ELISA 30

RESULTS 32

Posthodiplostomum minimum Infection in Bluegills and Green sunfish 32

Pathology 32

Gross 32

Histology 33

Protein Analysis 33

Dot blots 34

Indirect ELISA 34

Western blot analysis 35

DISCUSSION 58

LITERATURE CITED

LIST OF TABLES

Table 1: Common Examples of Parasite Evasion Mechanisms in Fish

Table 2: Site and Age distribution of Posthodiplostomum minimum intensity in bluegills.

Table 3: ELISA table showing changes in absorbance values with dilutions of bluegill sera.

Red rows and black rows denote bluegills with low and high P. minimum

infections, respectively. Cut off point is the highest absorbance value recorded

from the control wells.

LIST OF FIGURES

Figure 1: Heavy burden of Posthodiplostomum minimum infection in an adult Bluegill.

Metacercarial cysts appear as numerous multifocal white pinpoint areas on the liver

surface as well as the kidney capsule. Parasite cysts can be seen adhering to the

epicardial surface of the heart (H).

Figure 2: Posthodiplostomum minimum metacercariae isolated from the kidney. The double

layered cysts are intact with viable young flukes waiting to be excysted. An empty

cyst (E) can be observed with a crack in its shell through which the fluke excysted.

Figure 3: Liver infected with Posthodiplostomum minimum metacercariae.

(a) P. minimum within the detached double layered cyst wall (blue arrowheads) in the

liver parenchyma. Normal hepatic structure (N) can be observed. Scale bar = 100

µm

(b) Melanization (black arrowheads). Empty spaces surrounded by cyst walls can be

seen (E); Scale bar = 100 µm

tissue (arrowheads); Scale bar = 50 µm

(d) Mild cellular infiltration (red arrow) around the cyst wall; Scale bar = 50 µm

(e) P. minimum structure with a thick body walls and a prominent posterior sucker

(PS), Scale bar = 50 µm

(f) Shards of proteinaceous cyst wall (blue arrows), Scale bar = 50 µm

(g) Melanization within the haptic parenchyma (arrowheads), Scale bar = 50 µm

Figure 4: Posthodiplostomum minimum infection in the kidney 8

(a) P. minimum metacercariae in the renal tissue parenchyma(i) and the renal periphery

(ii); Scale bar = 100 µm

(b) Mild compression (arrowheads) and cellular infiltration (arrows) in the adjoining

renal tissue. Scale bar = 50 µm

bar = 50 µm

(d) The separated double layered cyst wall (black arrows) bordering the parasite.

Melanization is not as pronounced (green arrow); Scale bar = 50 µm

Figure 5: P. minimum in the heart

(a) Accummulation of metacercariae in the epicardium (E). Melanization can be

observed (arrows); Scale bar = 100 µm

(b) Parasites are confined to the epicardium (E). The myocardium (M) appears normal;

Scale bar = 100 µm

Figure 6: Gel electrophoresis of proteins with Coomassie staining. (L) Ladder (a) white grub

(b) bluegill fish sera (c) Bluegill fish IgM (d) bluegill mucus (e) green sunfish sera.

Figure 7: Dot blot showing serum antibodies were sensitive to their respective antigens but

showed problems with specificity. Antigen dots are numbered (1) white grub (2)

bluegill serum (3) mucus (4) green sunfish serum (5) bluegill IgM; The primary

antibodies (from mice) in which the strips were incubated are lettered (a) mouse

anti-white grub (b) mouse anti-bluegill serum (c) mouse anti-green sunfish serum

(d) mouse anti- mucus (e) mouse anti- bluegill IgM. Only the mouse anti-white

grub antibody (a) was specific.

Figure 8: Representative ELISA results showing decrease in absorbance with increasing

bluegill sera dilution. The x-axis denotes a 1:2 dilution series of bluegill serum

starting at 1:20. The y-axis is the absorbance at 405nm. Serum was obtained from

bluegills infected with P. minimum. BGS7 is serum from a bluegill showing light

infection, while CC2, CC14 and LC6, are sera from severely infected bluegills. The

cutoff point is defined as the highest absorbance recorded from the control wells.

Figure 9: Western Blot of mice antibodies with their corresponding antigens. (L) ladder (a)

yellow grub (b) white grub, (c, d) fish sera were incubated in mouse antibodies –

mouse anti – white grub (a, b), mouse anti-bluegill serum (c) and mouse anti-

bluegill IgM (d). Multiple individual bands indicating antigen- antibody interaction

can be observed in most of the strips.

Figure 10: Western Blot testing sensitivity and specificity of mouse anti – white grub

antibodies. (L) molecular weight ladder (a) Total protein stain for white grub

antigen (Indian Ink) showing an approximately 9 protein bands. Other blots –

White grub (b), Yellow grub (c) and fish sera (d) blots were incubated in mouse

anti-white grub.

Figure 11: Western Blot showing reaction between white grub antigen prep and mice

antibodies. All strips were blotted with white grub. The mouse antibodies include

(a) mouse anti-white grub, (b) mouse anti- bluegill serum, (c) mouse anti – GSF (d)

mouse anti – IgM.

Figure 12: Western Blot showing binding of mouse anti – bluegill IgM antibodies to fish

serum (a), and white grub antigen (b). Blots were incubated in mouse anti- bluegill

IgM. (L) ladder.

Figure 13: Western Blot assessing binding of fish antibodies. Strips (a, b & c) were blotted

with white grub while (d) was blotted with yellow grub. The strips were incubated

in pooled sera of highly infected bluegills (a, b, d) and green sunfish (c) prior to

incubation in mouse anti- IgM (a, b, d) and mouse anti- GSF (c).

INTRODUCTION

Posthodiplostomum minimum (white grub) is a digenetic trematode that commonly infects freshwater fishes. It was historically thought to be a host species generalist (Paperna &

Dzikowski, 2006), but recent studies (Boone et al., 2018; Lane et al., 2015; Locke et al.,

2010) have suggested that, in fact, several strains do show host specificity and may represent separate species. The bluegill (Lepomis macrochirus), a member of the sunfish family, is one of the most commonly infected species, serving as second intermediate host in the parasite life cycle. They support high parasite burdens and show persistent infections, which intensify with fish age (Ferguson, 1943; 1938, 1956). This implies that there is little protective immunity against P. minimum infections in bluegills.

Fishes have a complex immune system including both innate and adaptive mechanisms.

They are able to fight off infections by harnessing cellular defense mechanisms and protective antibodies (Wilson et al. 1997; Zhao et al. 2002). Thus, it remains unclear why the immune response in bluegills is not effective against P. minimum infection. Conversely, green sunfish (Lepomis cyanellus) that often cohabit with infected bluegills are less often infected with P. minimum. It is not clear if this low susceptibility, is due to host specificity by the parasite or an effective immune response in green sunfish.

Other digenetic trematodes, such as the yellow grub (Clinostomum marginatum) and black spot (Uvulifer ambloplitis) have also been reported to infect these sunfishes. However, it is not clear whether or how immune responses to these other trematodes overlap with P. minimum.

This study was aimed at understanding the immune mechanism underlying

Posthodiplostomum minimum infection in bluegills. Specific goals were to:

• Evaluate the immunologic response of bluegills to P. minimum infections, using

histologic assessment and serum antibody detection.

• Assess any correlation between specific antibody titers and parasite intensity.

• Compare grub antigens to assess the extent of immune cross-reactivity.

Sunfishes

Sunfishes belong to the family Centrarchidae, which also includes bass and crappies. All are widely distributed, occurring naturally in freshwater bodies across the United States.

They are largely found in fertile lakes with lush vegetation. Common sunfishes in Illinois include the bluegill (Lepomis macrochirus), the green sunfish (Lepomis cyanellus), the pumpkin seed (Lepomis gibbosus), the orange spotted sunfish (Lepomis humilis), the longear sunfish (Lepomis megalotis) and the redear sunfish (Lepomis microlophus) (Illinois

Department of Natural Resources, 2020). Of interest in this study are the bluegill and the green sunfish, both of which are abundant in ponds and lakes across Illinois. They are important game fish which are native to the United States and Canada (Pfieger, 1997), and are widely utilized as source of food and for recreation. They are well adapted to pond life and provide a sustained forage base for larger predators such as the large-mouth bass.

Bluegills and green sunfish are widely stocked, and their abundance and high fecundity allows for a liberal recreational harvest. They have also frequently served as aquatic models in physiological studies and ecology research.

Like most organisms, the bluegill and green sunfish suffer from infectious and non- infectious diseases. Common infectious agents include viruses, bacteria, fungi, and parasites including protozoans and helminths (Hoffman, 1999) such as trematodes. This study involves a group of digenetic trematodes whose larval "grubs" are found in the skin, 13

muscle, and viscera of various freshwater fishes including sunfishes. They include white grub (Posthodiplostomum minimum), yellow grub (Clinostomum marginatum) and black spot (Uvulifer ambloplitis). Their presence makes fish undesirable for consumption, resulting in diminished angling effort. Grub infections are usually asymptomatic and do not pose serious health concerns except in very heavy burdens where they may disrupt physiological functions, and cause death (Wisenden et., al, 2012; Pracheil and Muzzail,

2010). These parasites may also alter the behavior of fish, making them more susceptible to predation. For instance, Diplostomum spathaecum metacercarial infections in the eye causes cataract and vision loss, which may result in reduced activity and diminished escape response (Seppälä et al., 2006).

Common Digenean Trematodes of Bluegills

White Grub

White grub (Posthodiplostomum minimum) is commonly found in freshwater fishes across

North America. It is small, about 1mm in length, and may require microscopic examination to be identified within tissues. The definitive host for white grub is the great blue heron

(Ardea herodias). Adult worm complete sexual development and reproduce in the heron’s intestine. Eggs are released and passed in feces, hatch in water to form the miracidia, whose success depend on locating and infecting a Physa snail intermediate host. Successive sporocyst and cercaria larval generations develop in the snail via asexual reproduction.

Snails release free-swimming tailed cercaria, and fish are infected when cercariae penetrate the skin and are carried by the circulatory system to the visceral organs. The cercariae encyst in these organs, and the cycle is completed when a bird feeds on the fish (Spall and

Summerfelt, 1969; Hoffman, 1999). In P. minimum infection, the double-walled cyst is

comprised of an inner layer of parasite origin and a fish host reaction that forms the outer fibrous capsule (Bogitsh, 1962; Mitchell et al., 1982).

P. minimum was historically thought to be a host species generalist and two subspecies were recognized based on their host preferences – P. minimum ssp. minimum that usually infects cyprinid fishes e.g minnows and carps, and P. minimum ssp. centrarchi that commonly parasitize centrarchids (sunfishes and basses). However, recent evidence has noted there are cryptic species that do show host specificity (Boone et al., 2018; Lane et al., 2015; Locke et al., 2010)

White grubs predominantly affect the liver, kidneys, heart, and in rare cases, the spleen, intestine, ovary and air bladder of freshwater fishes (Kvach et al., 2017; Zhang et al., 2018,

Calhoun et.al, 2018; Hamouda & Bazh, 2019). Infections are usually asymptomatic and infected fishes usually show persistent infections that accumulate with age. However, heavy white grub infections may cause deleterious effects such as stunted growth, significant weight loss, and increased mortality (Hoffman, 1956; Smitherman, 1968;

Wisenden et., al, 2012; Pracheil & Muzzail, 2010). Encystation of metacercariae in the liver parenchyma may result in hepatocellular compression, cellular degeneration, and hepatic necrosis, ultimately resulting in death (Lane & Morris, 2010). Hoffman and

Hutcheson (1970) reported a rare case of white grub that caused exophthalmia and mortality in striped bass (Morone saxatilis). In fathead minnows, white grub infections resulted in the accumulation of body fluids, displaced organs, and ruptured tissues

(Mitchell et al., 1982). There are also indications that breaks in the skin as a result of cercarial penetration may predispose fish to secondary bacterial and fungal infections.

Rossiter and Davidson (2018) have recently indicated that habitat, host size and species specificity are the strongest predictors of P. minimum infection intensities. In a host specificity study conducted by Lane et al. (2015), the prevalence of P. mimimum was 100% in bluegills with highest infections in the kidney, while the prevalence was 57% in white crappie with most infections in the liver. Other studies have reported varying intensity and prevalence of P. minimum infections in bluegills and green sunfish based on age, size, weight etc. Boone et al. (2018) recorded an 86 % prevalence of white grub infection in bluegills collected from the Illinois portion of the Ohio River drainage. Both Lane et al.

(2015) and Boone et al. (2018) reported there was a strong positive correlation between infection intensity and age. Ferguson (1943) and Hoffman (1956) thought that white grub infections in bluegills did not elicit an immune response as larval trematodes accumulate over the lifespan of its host.

Green sunfish (L. cyanellus) from the same habitat as bluegills are rarely infected by P. minimum. Evidence can be drawn from the study by Lane et al., (2015), which showed a

15% prevalence in green sunfish, with infections limited to the liver and heart. A similar prevalence pattern was observed in the Sangamon River with over 90% prevalence in bluegills while green sunfish were seldom infected. (White et al. 2019). These studies highlight the difference in infection rate between two closely related fish species. More importantly, it remains unclear whether this difference is due to variation in host specificity by the parasite or differences in the parasite immune response.

Yellow Grub

Yellow grubs (Clinostomum marginatum) are larger parasites (3-8 mm) commonly found in sunfish and bass (Olsen 1962) and have been shown to display seasonal variations. 16

(Hazen & Esch, 1978). They are usually found burrowed under the skin (subcutaneous) or within the muscle tissues (intramuscular). They are visible to the naked eye after filleting due to their large size and color. The definitive host for yellow grub is predominantly the great blue heron (Ardea herodias). Adult worms live and sexually mature in the throat of the bird. During feeding, when the heron plunges its beak into water, eggs are released and these hatch into miracidia under suitable conditions. On locating a Planorbella snail, a miracidium infects the snail to form germinal sacs, followed by successive asexual reproduction to release tailed cercaria. Free-swimming cercaria locate and penetrate the skin of the fish hosts and then embed in the fish musculature to form metacercaria (yellow grub). Fish-eating birds complete the lifecycle when they consume yellow, grub-infected fishes (Edney, 1950).

Yellow grub is asymptomatic in nature and does not cause any noticeable effects in fish except in heavy infections. They can live in fish for up to 4years (Elliot & Russert, 1949).

The unsightly appearance may, however, be unpleasant to consumers resulting in waste and economic losses.

Fish Immune System

Generally, there are two fundamental divisions of the immune system – innate and adaptive immunity. These function in unison, particularly in higher vertebrates. Fishes share a similar organization of immune system with most vertebrates and are the first vertebrate species to evolve an adaptive immune system. Thus, they are important to evolutionary biologists as a comparative reference group in the understanding of the evolution and general principles of the immune system (van Muiswinkel, 1993; Lieschke & Trede, 2009).

It is important to note that although jawless fish (e.g. the lamprey) are believed to not have 17

an adaptive immune system, they possess lymphocyte-like cells that have variable receptors and immunological functions, similar to the lymphocytes of jawed vertebrates.

The cells of the immune system in fish are mainly lymphoid in origin. The main lymphoid organs – cephalic kidney, spleen, thymus and mucosa associated lymphoid tissues, are arranged by reticular networks to provide a structure for the cellular components of the immune system (Miller et al., 1985; Salinas et al., 2011). Although the liver is not considered a typical immune organ in fish, it secretes proteins that are important to the innate immune system (Salinas et al., 2011). The absence of bone marrow and lymph nodes in teleost fishes has confined hematopoietic functions to the cephalic kidney (Meseguer et al., 1994).

The Innate Immune System

The innate immune system is the most primitive immune mechanism and broadly serves as the first line of defense. Although non-specific and short-lived, it is crucial to the immune defense in multicellular animals. It includes immunologically active mucosal barriers that span the external body surface, gills, and intestines, protecting the body against invasions from foreign agents (Watts, Munday, & Burke, 2001). Fish skin presents physical barriers, which include the epidermis, scales and mucus; and chemical barriers, such as antibacterial peptides, epithelial lysozymes, defensins and lectins; as well as microbiological barriers, formed by the normal microbial flora in the mucous membranes

(Gómez & Balcázar, 2008).

The fish innate immune system is comprised of humoral factors and cellular defense mechanisms. Humoral factors include interferons (antiviral activity), complement factors,

lysozymes, clotting transferrin, anti-proteases, digestive enzymes, natural antibodies (lack pathogen specificity), cytokines (TNF-β, Interleukins) and chemokines in plasma, serum, mucus and mucosal membranes (Magnadóttir, 2006). Cellular components include polymorphonuclear leucocytes such as granulocytes (neutrophils, eosinophils), macrophages, and natural killer cells. These cells are particularly important in mediating inflammatory processes as they coordinate with soluble humoral products to mobilize cells and carry out immune functions (C. Bayne & Gerwick, 2001; Lieschke & Trede, 2009).

Inflammation is an important component of the fish innate immune system, and the process is similar to that in mammals. Granulocytes arrive first at a site to destroy infectious agents, and this is mediated by toll-like receptors that recognize conserved molecular patterns common to pathogen groups (PAMPS) (Secombes & Wang, 2012; Uribe, Folch, Enríquez,

& Moran, 2011). The interaction stimulates the release of chemokines that results in vasodilation and mobilization of other inflammatory cells to the site of infection. Cellular remains and debris are phagocytosed and digested by the macrophages (Magnadóttir, 2006)

The Adaptive (Acquired) Immune System

The adaptive immune system also includes humoral and cell-mediated components. This system produces a specific, delayed but long-lived immune response, which is more rapid and exuberant on subsequent infections with similar pathogens (immunological memory).

The specific nature of the adaptive immunity is a function of lymphocytes – agranular cells with variable receptors that identify specific pathogens.

The two major classes of lymphocytes in fish are the T and B lymphocytes. Like in mammals, fish T cells express variable receptors (TCR) on their surface while B cells have

immunoglobulins (Ig) of varying isotypes (Scapigliati, 2013; Scapigliati, Fausto, &

Picchietti, 2018). T cell receptors are always membrane-bound, while immunoglobulins may be membrane-bound or free in serum. The variability of the cellular receptors depends on the recombination activating genes (RAG1 and RAG2), which direct the functional rearrangement of the Immunoglobulin VDJ gene segments, resulting in receptor diversity

(M. D. Cooper & Alder, 2006).

T lymphocytes

There are two major classes of T cells in fish, based on their surface receptor subtypes:

CD4 (helper) T cells and CD8 (cytotoxic) T cells, which play important roles in the fish adaptive immune response (Nakanishi, Shibasaki, & Matsuura, 2015). A subpopulation of fish helper T cells, the regulatory T cell, has also been described based on the gene expression patterns in these cells (Kasheta et al., 2017; Nakanishi, Toda, Shibasaki, &

Somamoto, 2011; Nuñez Ortiz et al., 2014; Takizawa et al., 2016).

Mature T lymphocytes are chiefly found in the mucosa-associated lymphoid tissues of the intestine and gills, where they carry out diverse functions (Tafalla, Leal, Yamaguchi, &

Fischer, 2016). Recent evidence has suggested that the intestine may be the primary lymphoid organ for T cells in fish. Cells are produced in the intestine and are activated in the gills (Boschi et al., 2011; Picchietti, Terribili, Mastrolia, Scapigliati, & Abelli, 1997;

Rombout, Huttenhuis, Picchietti, & Scapigliati, 2005; Scapigliati et al., 2018). Memory T cells have also been reported in fish, evidenced by cytokine modulations of T cell proliferation and responses in vaccinated carp (Hohn & Petrie-Hanson, 2012; Piazzon et al., 2015).

B lymphocytes

B cells produce immunoglobulins (Ig). There are three known functional Ig classes in fish.

IgM is a tetrameric protein (450 kDa) which is the most prevalent and has been reported in high titers in the serum/plasma. IgT/IgZ (Danilova et al., 2005; Hansel et al., 2005) is a monomeric protein (170 kDa) associated with intestinal mucosal immunity (Zhang et al.,

2010). A third class, IgD, is also a monomer (150 kDa) (Wilson et al., 1997), but its physiological functions are unknown.

Serum antibody titers in fish vary with species, size, sexual maturity, physiological events

(e.g. stress, spawning), and habitat conditions. Higher antibody titers have frequently been observed with vaccinations and chronic infections (Wilson et al. 1997; Zhao et al. 2002).

Antibody producing B cells have also been found in the skin, gills, and intestines. The immune response at mucosal surfaces is especially valuable due to the close contact with the external environment (Uribe et al., 2011). Antibodies may also function as anti- adhesins to prevent bacterial adherence at the mucosa interface (Davidson et.al, 1997; Ellis,

2001).

Immune Response to Parasitic Infections in Fish

The success of a parasite infection depends on the interaction between parasite factors (host specificity) and host factors (immune response) (Poulin, Krasnov, & Mouillot, 2011).

Roggers-Lowery et al. (2007) reported that mucosal and systemic antibody responses against infections in fish may vary based on the nature of the antigen, route of exposure or inherent nature of the host’s immune system. This may account for the variations in the intensity of specific infections across fish species/individuals in a habitat.

The fish immune system has evolved different strategies of innate immune responses to combat parasite infections, depending on the nature and route of infection. A common approach against intestinal parasites is hyperplasia of mucus producing cells and increased smooth muscle contraction to expel parasites through the anus (Dezfuli, Bosi, DePasquale,

Manera, & Giari, 2016). When a parasite is attached, as seen in helminths, mucus production is accompanied by secretion of chemokines and mobilization of inflammatory cells at the site of attachment (Harris & Loke, 2017). The release of parasite antigens may result in an extensive inflammatory response that can lead to destruction of host tissues. In this case, the immune system can “wall-off” the infection, preventing parasite migration and tissue destruction. An example is the encysted metacercaria of Tinca tinca in the heart, gills, and kidney, which triggers a granuloma formation that is characterized by melano- macrophagic centers and granulocytes (Dezfuli et al., 2013). Once encapsulated, the parasite elicits a minimal immune response (Olson & Pierce, 1991). A similar immune- protective strategy has been found in black spot disease, which is the cause of the peppered appearance on infected fish skin (Lemly & Esch, 1984).

Some studies have reported a positive correlation between metazoan parasite infection and splenic size, based on the premise that a larger spleen reflects an ongoing immunological response (Skarstein, Folstad, & Liljedal, 2001; Rohlenová et al., 2011). Other studies found no correlation between the two variables (Rohlenová & Šimková, 2010; Vainikka,

Taskinen, Löytynoja, Jokinen, & Kortet, 2009).

Acquired immunity plays a role as well, and often works in harmony with the innate response. For example, resistance to Ichthyophthirius multifilis (parasitic ciliate on skin) re-infections was associated with the presence of humoral antibodies in the mucus and sera 22

of the fish (Maki & Dickerson, 2003; X. Wang, Clark, Noe, & Dickerson, 2002). The exact immune mechanism involved cutaneous IgT and a local inflammatory response. (Kurt

Buchmann, 2019; Dickerson & Clark, 1998; Q. Wang, Yu, Zhang, & Xu, 2019). Other evidence of immunological competence against parasitic trematodes has been reported.

Roberts et al. (2005) concluded that the blood fluke, Sanguinicola inermis, infection elicits specific antibody production and complement activity in common carp. In a recent study, an experimental infection of bluegills with the trematode larva, Ribeiroia ondatrae, showed acute hemorrhage, leukocytosis, neutrophilia and a chronic granulomatous inflammation that destroyed 85% of encysted metacercaria within 4 weeks (Calhoun, Schaffer, Gregory,

Hardy, & Johnson, 2015). Hoglund and Thuvander (1990) also reported a gradual decrease in Diplostomum spathaecum metacercaria recovery following a 12-week experimental cercarial infection in rainbow trout. The results showed neutrophilia, monocytosis, and an increase in total antibody titers. However, a specific antibody response was not detected when the serum was tested against parasite antigens in an ELISA, suggesting that immune responses may be largely cell mediated. In a different experiment in immunized rainbow trout, activated macrophages destroyed infective larval stages of Diplostomum spathaecum in the presence of specific antibodies, leading to significant reduction in the infection intensity (Whyte, Chappell & Secombes, 2006). All of these studies emphasized the success of immune-protective strategies of fish species against parasites.

In other cases, immune reactions in fish may not be protective such that there is a continuous colonization of fish organs by new infections despite an immunological response. Ferguson (1943) and Hoffman (1956) have reported that white grub infections in bluegills do not elicit an effective immune response as larval trematodes accumulate over

the lifespan of its host. A study on Discocotyle sagittata trematode infection in rainbow trout (O. mykiss) reported the detection of specific antibodies in sera, with titers that showed no correlation with infection intensity (Rubio-Godoy, Sigh, & Buchmann, 2003).

Another study in common carp evaluated the difference in immune response to

Bothriocephalus acheilognathi between natural and experimental infections

(intraperitoneal injection of parasite extract) (Nie & Hoole, 1999). Results showed a higher specific antibody titer and plasma cell count in natural infections than in the experimental infections. However, these values were not significantly different from the uninfected control group. There are speculations that these non-protective responses may represent the complex interplay between host immunomodulation and parasite evasion mechanisms

(Buchmann, 2012).

Immune Evasion Mechanisms by Fish Parasites

Fish parasites have developed several evasion mechanisms to thrive in the presence of an active immune response. These mechanisms have helped the parasites to avoid the overarching activities of the immune cells and their defense products.

Some examples of parasite evasion mechanisms and corresponding fish parasites utilizing each mechanism, are listed in Table 1.

Evasion Mechanism Example of Fish Literature Parasites

Predilection for immune- Diplostomum Whyte et al., 1991 privileged sites such as the CNS spathaceum in the eye. and sensory organs.

Anatomical sequestration in host Posthodiplostomum (Sitjà-Bobadilla, 2008) tissues minimum metacercariae in the liver and kidney

Intracellular disguise in cells Kudoa ovivora in the Swearer & Robertson, oocytes 1999

Antigenic variation Trypanosoma spp Pays 2006 & Lischke 2000

Parasite migration and behavioral Icthyophthirius Clark 1996 changes multifilis on the skin

Resistance to humoral and cellular Diplostomum Mikes & Man, 2003 factors pseudospathaceum

Immunosuppression Ligula intestinalis Hoole & Arme, 1983

Fast development Trypanosoma borelli Bowden et al., 2007

Exploitation of the host immune Icthyophthirius Matthews, 2005 reaction multifilis

Table 1: Common examples of parasite evasion mechanisms in fish.

From the current understanding of trematode life cycles and the fish immune system, I should be able to detect an immune response to Posthodiplostomum minimum infection in bluegills. A cutaneous immune response should be elicited during cercarial penetration of the skin. Also, cercarial migration through the circulatory and lymphatic system should trigger a systemic response that could prevent or minimize parasite encystation. However, the prevalence of P. minimum and its accumulation with age suggests that the immune response may not be protective. This may be possible if the parasite had evolved a strategy to evade the immune system. Because there are not enough studies that have investigated this hypothesis, we are unsure of the role of the immune response to this parasite.

Thus, my study was aimed at better understanding the nature of immunologic response to

P. minimum infection in bluegill. Histologic sections of affected organs will describe the pathology and cellular response to the parasite infection, including any active inflammatory

response. In addition, I designed immunoassays to quantify P. minimum antibody titers in serum. The detection of parasite-specific antibodies will indicate an antibody response produced against the parasite. Moreover, antibody titers should correlate with intensity of infection as determined by necropsy. I also compared white grub to yellow grub antigens, as any cross reactions between these common worms would provide information on the reliability of serologic tests as a means of evaluating infection status in white grub infections.

METHODS

Fish sampling

Fish samples (bluegill (Lepomis macrochirus) and green sunfish (Lepomis cyanellus)) were collected from rivers and lakes around Charleston, IL. Fish were collected by hook and line from Carmen pond on the campus of Eastern Illinois University and via electrofishing using standard, boat-mounted electrofishing gear with pulsed DC and maintaining 3.5-4 A

(Reynolds 1996), in the Lake Charleston side channel reservoir. They were transported on ice to the laboratory, where each fish was weighed, and its length measured from the snout to the longer lobe of the caudal fins. Fish with length less than 15 cm and/or weight less than 60 g were recorded as young, while others that measured ≥ 15 cm in length and/or 60 g in size were noted as adults. Mucus was scraped off the skin surface and centrifuged, and the resulting clear supernatant was collected. Blood was collected from the caudal vein using 26-gauge needles, allowed to clot, and centrifuged to obtain serum. Some serum was subjected to ammonium sulfate precipitation to concentrate immunoglobulins (IgM).

Mucus, serum, and immunoglobulins were stored in sterile tubes at -20°C.

The fish carcass was dissected by making a ventral, mid-line incision that extended proximally from the anus to the gill openings. With the aid of a dissecting microscope, the prevalence (percentage of infected fish in the sample) and infection abundance (number of metacercariae per individual fish) of Posthodiplostomum minimum infection was determined. An infection scoring criterion was established – fish with no visible metacercariae in commonly affected organs were scored as “no” infection. Those with less than 50 total metacercarial cysts distributed across affected organs, were scored as “low”

infection, while infection with more than 50 metacercarial were scored as “high” infection.

Affected organs were harvested to isolate metacercarial cysts and for tissue processing.

Tissue Processing and Staining

Harvested fish tissues (liver, kidney, and heart) were carefully dissected, cut into blocks, and fixed in FAA (5% formaldehyde, 50% ethanol and 5% glacial acetic acid) for 8 weeks.

These were then washed in several changes of 70% ethanol followed by dehydration via a graded t-butyl/ethyl alcohol series and embedded in paraffin (Paul & Mukti, 2017).

Paraffin sections were made at 10 µm on an AO rotary microtome, and the resulting ribbons were mounted on slides in a warm water bath containing adhesive glue (Sta-On). Slides were dried overnight at 30°C.

Mounted ribbons were cleared in two changes of limonene (5 mins each) and then rehydrated in a graded ethanol series (100%, 95%, 70%; 5 mins each). Ferric ammonium sulfate (4%) was used as a mordant for 30 min, and sections were stained in 2.5% hematoxylin for 5mins. A bluing step of 2 drops household ammonia in water was carried out before staining in 1% eosin in 70% ethanol for 3min. Stained slides were dehydrated in a graded ethanol series (70%, 95%, 100%) and cleared in limonene. A cover slip was mounted using Permount, and slides were air-dried for 48 hr.

White Grub Antigen Preparation

P. minimum metacercarial cysts isolated from fish tissues were thoroughly washed in phosphate buffered saline (PBS) to remove fish components. The cysts were then treated in an acidic enzymatic medium (10% acid pepsin in Ringer’s solution) at 37°C for 1 hr.

Using a sterile pipette, treated metacercarial cysts were transferred to 1% sodium

bicarbonate (emergence and activation medium) at 37°C for 1 hr. Excysted flukes were transferred into fresh saline solution, while any intact metacercarial cysts were subjected to gentle manipulations with a dissecting pin to release the flukes. Isolated flukes were washed thoroughly in saline to exclude cyst particles.

Excysted flukes (approx. 250 μl) were homogenized in 500 μl normal saline using plastic pestles and centrifuged at 3000 g. The supernatant (white grub antigen) was collected and stored at -20°C. Protein concentration was determined using the Bradford protein assay.

Following antigen preparation, the resulting protein concentration of the white grub antigen was 3.67 mg/mL. Yellow grub (Clinostomum marginatum) metacercariae were isolated from perch (Perca flava) collected from Eagle Lake, Ontario, and subjected to similar treatment to prepare the yellow grub antigen.

Mice Immunization

In-bred mice (C57Bl6) were raised in the EIU Biology Animal House at optimum conditions of temperature and humidity and provided with feed and water ad libitum.

Groups of 12 mice were immunized with one of the following antigens: white grub prep, bluegill serum, bluegill mucus, bluegill IgM, and green sunfish serum. White grub antigen prep was reconstituted to 0.2 µg/µL while sera were subjected to 1:2 dilution.

Immunization protocol involved subcutaneous injections of 0.1 ml volumes containing antigen in Freund’s complete adjuvant (1:1) at week 1. Booster antigen doses were suspended in Freund’s incomplete adjuvant (1:1), and 0.1 ml volume of each antigen prep was administered at weeks 4 and 7. Blood was collected by cardiac puncture at the end of week 8, allowed to clot, and was centrifuged at 3000 g. Serum obtained was stored at -

20°C. 29

Dot blot

2 µl of each antigen was placed on nitrocellulose membrane and allowed to dry for 5 min.

The membrane was then blocked with Blotto (5% powdered milk in PBS) for 1 hr, followed by incubation in primary antibody then HRP-conjugated secondary antibody for 1 hr each.

This was then incubated in a substrate solution (4% chloronaphthol containing 3% hydrogen peroxide) until a purple color was observed. Each incubation step was followed by a thorough washing step in PBS.

SDS Polyacrylamide Gel Electrophoresis (SDS PAGE) and Western Blotting

Antigens were mixed with equal volume of 2X sample buffer (Sigma-Aldrich S3401) and heated in boiling water for 5 min. The treated proteins were loaded onto a 10% polyacrylamide gel (BIO-RAD) in a mini-PROTEAN Tetra electrophoretic cell containing

BIO-RAD Premixed Tris running buffer (1X Tris/Glycine/SDS) and run at 200 V for 35 min. The gel was retrieved and then equilibrated in BIO-RAD Premixed transfer buffer

(1X Tris/Glycine) containing methanol, for 15 min. Proteins were transferred to a nitrocellulose membrane using a semi-dry transfer apparatus (BIO-RAD Trans-Blot SD® semi-dry transfer cell). The blotting membrane was subsequently treated in a similar fashion as the dot blots. Separated proteins on nitrocellulose were probed using mouse antibodies to evaluate antigen-antibody specificity and identify cross-reacting proteins.

Indirect ELISA

White grub antigen was diluted in carbonate buffer (1.59 g Na2CO3 and 2.93 g NaHCO3 per liter, pH 9.6). 100 µL of diluted antigen (10 μg/mL) was pipetted into wells in a 96- well microtitre plate, and incubated overnight at 4°C. The coated plate was washed with 30

PBS and blocked with Blotto for 1 hr. Fish test serum (100 µL; 1:20 dilution) was pipetted into the first well in a row, then serially diluted (1:2) across the row to a final dilution of

1:20,840 ??? in well 11, and allowed to incubate on a rocker at room temperature for 1 hr.

This was followed by 1-hr incubations in mouse anti-bluegill serum antibodies (1:100 dilution) and HRP-conjugated goat anti-mouse antibody (1:2000 dilution). The addition of substrate (ABTS) resulted in a green color and the reaction was stopped by adding 1% SDS solution. Each plate included controls, each missing one component (white grub antigen, bluegill serum or mouse anti-bluegill serum) to establish background reactions. The color absorbance was measured using an ELISA plate reader at 405 nm. Each fish was run in duplicate, and each plate contained both heavily infected and uninfected or lightly infected fish.

RESULTS

Posthodiplostomum minimum Infection in Bluegills and Green Sunfish

A total of 87 bluegills (30 g - 136 g) were collected from Lake Charleston (n=63) and EIU

Carmen pond (n=24). Necropsy revealed P. minimum metacercaria in 80 bluegills while the remaining 7 showed no detectable parasites, for a total prevalence of 92%. Using the described scoring criteria, 60 of the infected bluegills showed a high infection intensity while 20 showed low infection intensity (Table 2). The Lake Charleston side channel had an approximate P. minimum prevalence of 98%, 81% of which were heavy infections, while Carmen pond had a prevalence of 75% with 37.5% heavy infection. Across infected bluegills, the kidney showed the highest infection intensity among examined organs, while the liver showed the least infection intensity.

8 green sunfish were collected from the EIU campus pond and these showed no infection with Posthodiplostomum minimum.

Gross Pathology

The kidney, liver, and heart were the most commonly affected organs. Liver and kidney tissues were friable on gross dissection. Metacercarial cysts appeared as numerous multifocal white pinpoint areas on the liver surface as well as the kidney capsule. The consistency of the heart was unaffected, but parasite cysts were seen adhering to the epicardial surface (Fig 1). At higher magnification of dissociated cysts, viable juvenile flukes are visible in clear walled cysts that were not adherent to the parasite. (Fig 2).

Histology

Liver tissue showed P. minimum in double-walled cysts surrounded by a clear zone and a thin peripheral layer of inflammatory cells. Melano-macrophage centers, characterized by brown pigmentation of cells (melanization), were observed in the surrounding tissue (Figs

3(a) – (g)) together with apparently normal hepatocytes.

Kidney tissue showed parasites in cyst walls similar to that observed in the liver. There was mild compression of kidney cells adjoining the cyst walls and mild leucocytic infiltrates. Melanization was observed but not as pronounced as in the liver (Figs 4(a) –

(d)). Normal kidney structures including the glomerular capillaries and nephron tubules were observed.

Examination of the heart tissue showed intense parasite aggregation within, and confined to, the connective tissue as well as melano-macrophage centers at the periphery of the heart

(epicardium). Striated cardiac muscle (myocardium) was normal and numerous nucleated red cells were observed (Figs 5(a) – (c)).

Protein analysis

I separated the white grub antigen alongside with other antigens in an SDS PAGE to compare proteins from white grub and fish proteins (Fig 6). The white grub showed bands ranging between 30 and 250 kD in size. The bluegill fish sera and green sunfish sera each showed multiple bands between 25 and 250 kD. The bluegill fish IgM showed bands between 25 and 150 kD. The bluegill mucus showed approximately 12 bands between 25 and 150 kD.

Dot blot

Hyperimmune serum antibodies were sensitive to their respective antigens, but some showed problems with specificity (Fig 7). As expected, serum obtained from mice immunized with bluegill sera (ms anti-bgs), green sunfish sera (ms anti-gsf), bluegill mucus (ms anti-mucus) and bluegill IgM (ms anti-IgM) each recognized their respective antigens. However, there were shared proteins or epitopes between white grub antigen prep and the fish antigens, as shown by the moderate reaction when mouse anti-bluegill serum was exposed to white grub antigen dots, and a more pronounced reaction when white grub dot was incubated in mouse anti-bluegill mucus. Conversely, serum obtained from mice immunized with white grub contained antibodies that recognized white grub only, indicating that fish tissue antigens had been removed from the cysts before grinding.

Indirect ELISA

I developed an indirect ELISA to test for P. minimum specific antibody in infected bluegills. Absorbance values (A405) declined with 1:2 serial dilutions with high variability between fish (Fig. 8). To determine a cutoff value, each plate included controls missing one component (either white grub antigen, bluegill serum or mouse anti-bluegill serum) to establish background reactions. The highest absorbance value recorded from these controls was selected as the minimum absorbance value, above which a test was considered positive. The cut off point for each microtiter plate as well as the highest dilution of bluegill sera with an absorbance value above the cutoff point, is shown in Table 3. Due to high variability in absorbance across serial dilutions, antibody titers (highest dilution with absorbance value that is above cutoff value and falls in the linear portion of the graph where absorbance varies with serum dilution) were indeterminate.

The high absorbance values above controls in initial 1:20 dilution suggest that certain components in bluegill serum (perhaps antibodies) was bound to white grub antigen, and this interaction decreased across the 1:2 dilution series down to background levels.

However, there was no difference in absorbance between heavily infected and lightly infected bluegills.

Western Blot Analysis

I used Western blots to identify specific protein bands recognized by the mouse antibodies and fish sera. Each mouse antibody was sensitive to its antigen. (Fig 9). The mouse anti – white grub recognized several shared bands in the white grub and yellow grub antigens

(sensitive) but not the fish sera (specific), confirming dot blot results (Fig 10). However, as with dot blots, the mouse anti bluegill and, to a lesser extent, mouse- anti IgM recognized white grub antigens as well as their respective antigens (Figs. 11, 12). This raised concerns of low specificity. Both mouse anti-fish antibodies recognized a protein (approximately

150 kD in size) in the white grub blots (Fig 10), which was also present in the fish sera (Fig

12).

Because ELISA results indicated that bluegill sera bound to white grub antigen, I attempted to identify the specific grub antigens that were recognized by bluegill antibodies. This was done by incubating the white grub blots first in fish sera (1:20 dilution), and subsequently in mouse anti-IgM or mouse anti-bluegill serum to look for additional bands not seen by mouse anti-fish serum alone (Fig 12). Results showed no protein bands on the blots using individual fish or pooled serum from different bluegills (Fig 13). This suggested that grub- specific antibodies may, in fact, not be present in the bluegill serum. In addition, there was

increased background on the blot membranes (Fig 13) suggesting that the bluegill sera may be bound to Blotto.

In this light, it may be possible that the absorbance recorded from the ELISA assay was a result of non-specific binding of bluegill sera to the microtiter plate and/or Blotto, rather than antibody binding to white grub.

Figures

P. minimum Medium – High Low Infection No infection infection in Infection Bluegills Juvenile Adult Juvenile Adult Juvenile Adult

Lake 2 49 1 10 0 1 63 Charleston

Carmen Pond 1 8 1 8 3 3 24

3 57 2 18 3 4 87

Table 2: Site and age distribution of Posthodiplostomum minimum intensity in bluegills.

Gross Pathology

H L

Figure 1: Heavy burden of Posthodiplostomum minimum infection in an adult Bluegill. The liver (L) and kidney (K) are friable. Metacercarial cysts appear as numerous multifocal white pinpoint areas on the liver surface as well as the kidney capsule. The consistency of the heart (H) is unaffected and parasite cysts can be seen adhering to the epicardial surface.

Figure 2: Posthodiplostomum minimum metacercaria isolated from the kidney. The double layered cysts are intact with viable young flukes waiting to be excysted. An empty cyst (E) can be observed with a crack in its shell through which the fluke excysted.

Histology

An examination of the commonly affected organ was carried to identify cellular changes in response to P. minimum infection.

Figure 3: Liver infected with Posthodiplostomum minimum metacercaria.

(a) Postodiplostomum sp. within the detached double-layered cyst wall

(arrowheads) in the liver parenchyma. Normal hepatic structure (N) can be

observed. Scale bar = 100 µm

(b) Melanization (black arrowheads). Empty spaces bounded by cyst walls can be seen

(E). Scale bar = 100 µm

(c) Melanization in the adjoining clear zone between the cyst wall and the parencymal tissue (arrowheads). Scale bars = 50 µm

(d) Mild cellular infiltration (red arrow) around the cyst wall. Scale bar = 50 µm

(e) P. minimum structure with a thick body walls and a prominent posterior sucker (PS).

Scale bar = 50 µm

(f) Shards of proteinaceous cyst wall (blue arrows). Scale bar = 50 µm

(g) Melanization within the haptic parenchyma (arrowheads). Scale bar = 50 µm

Figure 4: Posthodiplostomum minimum infection in the kidney

(i) (ii)

(a) P. minimum metacercariae in the renal tissue parenchyma (i) and the renal

periphery (ii)..Scale bars = 100 µm

(b) Mild compression (blue arrowheads) and cellular infiltration (arrows) in the adjoining renal tissue. Scale bar = 50 µm

N C

bar = 50 µm

(d) The separated double layered cyst wall (black arrows) bordering the parasite.

Melanization is not as pronounced (green arrow). Scale bar = 50 µm

Figure 5: P. minimum in the heart

(a) Accummulation of metacercariae in the epicardium (E). Melanization can be observed

(arrows); Scale bar = 100 µm

M M

(b) Parasites are confined to the epicardium (E). The myocardium (M) appears normal

Scale bar = 100 µm

Protein analysis

L a b c d e a

mw (kD)

250 150

100 75

Figure 6: Gel electrophoresis of proteins with Coomassie staining. (L) Ladder (a) white grub (b) bluegill fish sera (c) Bluegill fish IgM (d) bluegill mucus (e) green sunfish sera.

Dot blot

a b c d e

Figure 7: Dot blot showing serum antibodies were sensitive to their respective antigens but showed problems with specificity. Antigen dots are numbered (1) white grub (2) bluegill serum (3) mucus (4) green sunfish serum (5) bluegill IgM; The primary antibodies (from mice) in which the strips were incubated are lettered (a) mouse anti-white grub (b) mouse anti-bluegill serum (c) mouse anti-green sunfish serum (d) mouse anti- mucus (e) mouse anti- bluegill IgM. Only the mouse anti-white grub antibody was specific (a)

Indirect ELISA

PLATE ID FISH ID. ABSORBANCE Dilution at (1:20) (A405) Background

PLATE 1 SBG4 0.861 1:5120

(Cut off point = 0.226) LC4 0.577 1:1280

LC11 0.771 1:1280

LC2 0.597 1:1280

PLATE 2 BGS3 0.842 1:20480

(Cut off point = 0.294) LC5 0.666 1:1280

LC18 0.695 1:2560

CC3 0.792 1:20480

PLATE 3 SBG2 0.559 1:20480

(Cut off point = 0.156) CC6 0.611 1:20480

CC5 0.604 1:10240

CC18 0.632 1:5120

PLATE 4 BGS7 0.686 1:2560

(Cut off point = 0.246) CC2 0.465 1:5120

CC14 0.516 1:5120

LC6 0.833 1:2560

PLATE 5 CC11 0.289 1:160

(Cut off point = 0.14) CC9 0.326 1:320

BGS6 0.303 1:160

LC14 0.3 1:320

PLATE 6 CC7 0.536 1:1280

(Cut off point = 0.152) BGS4 0.342 1:320

CC8 0.335 1:160

LC8 0.394 1:1280

PLATE 7 LC3 0.4 1:20480

(Cutoff point = 0.137) SBG13 0.469 1:20480

LC7 0.427 1:20480

CC13 0.5095 1:20480

Table 3: ELISA table showing changes in absorbance values with dilutions of bluegill sera.

Red rows and black rows denote bluegills with low and high P. minimum infections, respectively. Cut off point is the highest absorbance value recorded from the control wells.

ELISA showing changes in absorbance in a bluegill sera dilution series (Plate 3)

0.7

0.6

0.5

) 405 0.4

0.3 AAbsorbance (A

0.2 Cut off = 0.156

0.1

0 0 1 2 3 4 5 6 7 8 9 10 11 12 1:2 Dilution series starting at 1:20

SBG2 CC6 CC5 CC18 cut off

Figure 8: Representative ELISA results showing decrease in absorbance with increasing bluegill sera dilution. The x-axis denotes a 1:2 dilution series of bluegill serum starting at

1:20. The y-axis is the absorbance recorded at each dilution. Serum was obtained from bluegills infected with P. minimum. SBG2 is serum from a bluegill showing light infection, while CC6, CC5 and CC18, are sera from severely infected bluegills. The cutoff point is defined as the highest absorbance recorded from the control wells.

Western analysis

L a b c d L Mw (kD) Mw (kD)

250 150 100 75 75 50

5037

25 25 20 15

Figure 9: Activity of mice antibodies with the corresponding antigens. (L) ladder (a) yellow grub (b) white grub, (c, d) fish sera were incubated in mouse antibodies –mouse anti – white grub (a, b), mouse anti-bluegill serum (c) and mouse anti- bluegill IgM (d). Clear individual bands indicating antigen- antibody interaction can be observed in most of the strips.

L a b c d

Mw(kD)

Figure 10: Sensitivity and specificity of mouse anti – white grub. (L) molecular weight ladder (a) Total protein stain for white grub antigen (Indian Ink) showing an approximate

(9) protein bands. Other blots – White grub (b), Yellow grub (c) and fish sera (d) blots were incubated in mouse anti-white grub.

L a b c d

Mw (kD)

250 150

100 75

Figure 11: Immune reaction between white grub antigen prep and mice antibodies. All strips were blotted with white grub. The mouse antibodies include (a) mouse anti-white grub, (b) mouse anti- bluegill serum, (c) mouse anti – GSF (d) mouse anti – IgM.

L a b

Mw (kD)

250 150

100 75

20 15

Figure 12: Comparison of mouse anti – bluegill IgM activity against fish serum (a), and white grub antigen (b). Blots were incubated in mouse anti- bluegill IgM. (L) ladder.

L a b c d

Mw (kD)

Figure 13: Strips (a, b & c) were blotted with white grub while (d) was blotted with yellow grub. The strips were incubated in pooled sera of highly infected bluegills (a, b, d) and green sunfish (c) prior to incubation in mouse anti- IgM (a, b, d) and mouse anti- GSF (c).

DISCUSSION

The total prevalence of Posthodiplostomum minimum infection in bluegills in this study was very high (92%, Table 2). Among infected fish, 93.5% were adults, most (75%) of which were heavily infected. Because more adult bluegills were collected (91%), it was expected that most of the recorded P. minimum infections would be from this age group.

Posthodiplostomum minimum was highly prevalent in both sampling sites but the intensity of infection varied considerably between the sites. Since sampling was not uniform between sites, it was impractical to compare P. minimum infection between fish age groups, and sampling sites. The high prevalence recorded at both sites suggests that there were intermediate snail and definitive bird hosts capable of consistently propagating the P. minimum lifecycle in both habitats.

Bluegills infected with P. minimum have been reported to show reduced activity and a diminished escape response (Seppälä et al., 2006), predisposing them to increased predation by birds. This induced behavioral change in the fish host may be an evolutionary advantage that increases the likelihood of successful transmission of the parasite to the definitive host (Auld & Tinsley, 2015). The Posthodiplostomum sp. lifecycle requires conditions suitable for trematode eggs to hatch, miracidiae to locate Physa snails, and released cercariae to find and penetrate susceptible fish that are eventually consumed by a definitive host bird (Spall and Summerfelt, 1969; Hoffman, 1999). The lower infection rate recorded in EIU’s Carmen pond may be attributed to the sampling technique. If heavily infected fishes show reduced activity and are more likely to be outcompeted in the search for food, the hook and line fishing technique might have artificially selected for the more active and exploratory bluegills (Biro & Post, 2008) that are likely to have lower burdens 58

of P. minimum. In my study, the limited number of green sunfish collected were uninfected with white grub. This agrees with previous reports that have suggested that the green sunfish possess some resistance against P. minimum infection (Boone et al., 2018;

Lane, Spier, Wiederholt, & Meagher, 2015; White etal., 2019). Avault and Smitherman

(1965) described the differences in susceptibility to P. minimum among sunfish species.

An experimental infection of fish with metacercariae produced a disproportionate burden of infection in bluegills and bluegill hybrids whereas, no infection was recorded in green sunfish. However, it remains unclear if this resistance was due to parasite preferences or certain host immunological factors.

The indirect ELISA proved unreliable. Absorbance values showed a general decrease to background levels across the dilution series, with a high variability between bluegills. This suggested that some component in bluegill sera, presumably immunoglobulins, interacted with proteins in the white grub prep. The highly variable absorbance values and lack of confirmed negative controls, hindered my ability to accurately determine a titer to quantify the antigen antibody reaction. All fish in this study were collected from sites where the parasitic fluke was present. Therefore, even the negative control fish that did not contain cysts at necropsy may have, in fact, been exposed to the parasite. For this same reason, the closely related green sunfish could not be considered as true negative since they were collected from the same habitat as infected bluegills.

Nevertheless, there was no correlation between infection intensity and absorbance values since there was no difference in absorbance values between heavily burdened and lightly burdened bluegills. This implied that antibody detection may not be a suitable diagnostic test to determine the intensity of Posthodiplostomum minimum infection. A study on 59

Discocotyle sagittata infection in the rainbow trout (O. mykiss) reported the detection of specific antibodies in sera, which showed a similar pattern of titers that showed no correlation with infection intensity (Rubio-Godoy et al., 2003). Because antibodies persist long after infections, antibody testing is perhaps best suited for testing prior exposure and cannot accurately discern active infections nor detect if, and to what extent, specific antibodies are protective. Also, the time lag between antigenic exposure and the development of an immune response may contribute to false negatives, and affects the sensitivity of antibody detection tests (Jacofsky, Jacofsky, & Jacofsky, 2020). In the case of P. minimum, the persistent infection, re-infection, and its accumulation with age (Fellis and Esch, 2006; Pracheil and Muzzal, 2009), suggest that any immunoglobulins bound to parasite components, may not be protective.

Because they are generally easier to develop, antibody testing assays are frequently employed in the sero-surveillance of epidemics such as HIV (Curtis et al., 2012; Metacalf et al, 2016) and SARS CoV-2 (CDC, May 2020). However, an antigen detection assay such as a capture ELISA, which aims to test for and quantify antigens, rather than antibodies, might be more suitable to determine active infections and/or intensity. This was demonstrated in the test for Dirofilaria immitis, the causative agent of heartworm in dogs.

The first ELISA tests that were designed to detect circulating antibodies to Dirofilaria immitis had extremely low accuracy (30% false negative), and significant cross reactivity

(40% false positives) with Dipetalonema reconditum. These unreliable tests were later withdrawn from the market and replaced with ELISA tests that detect circulating adult

Dirofilaria antigens in the blood. The newer tests were highly sensitive (4% false negatives) and specific (2% false positives)(Calvert & Ridge, 2007; Ely & Courtney, 1987)

and are still in use. For my study, a capture ELISA would require coating the microtiter wells with a white grub-specific antibody (coating/capture antibody) to bind and immobilize the white grub antigen in the sample. Then, a second white-grub specific antibody would be used to detect the antigen from the test specimen. However, unlike heartworm, where the antigens can be recovered from blood, this would likely require euthanizing the fish, harvesting its tissues, and artificial digestion of infected tissues to obtain the antigen sample. This would make it useful as a post-mortem diagnostic tool only and has no benefit over gross dissection. An additional limitation of the capture ELISA is its requirement for two specific antibodies that do not compete for the same antigenic sites.

Because there are no commercially available monoclonal anti-white grub antibodies to serve as capture antibody, an antigen detection capture ELISA was not feasible for this study. The RT-PCR to test for the parasite’s nucleic acids may be another option (Won et al., 2016). However, unless it could detect nucleic acids from circulating cercarial stages in blood samples, obtaining samples may be a drawback because P. minimum parasites are typically encysted in host tissues.

Given the unreliable ELISA results, I used dot blots and western blots to assess the sensitivity and specificity of mouse antibodies generated. These assays showed that there were shared epitopes between the white grub antigen prep and the fish proteins.

Specifically, mouse-anti bluegill serum recognized both fish serum and white grub antigens. This could be a result of the migratory phase of the cercaria during which the parasite acquired host antigens or made them themselves. Acquisition of host proteins may serve as an evasion mechanism directed at circumventing the host immune defenses (Sitjà-

Bobadilla, 2008). It has been reported that Schistosoma sp. masked itself from the host

immune responses by incorporating host proteins (Yazdanbakhsh & Sacks, 2010). Another study by Bayne, Boswell, & Yui, (1987) recorded widespread antigenic cross-reactivity between Biomphalaria glabrata plasma proteins and its trematode parasite, Schistosoma mansoni. Alternatively, since the fibrous outer wall of the P. minimum metacercaria was formed from fish tissue reaction (Bogitsch, 1962), it is also plausible that remnants of fish proteins may have persisted in the white grub antigen prep. However, the high specificity of the mouse anti-white grub to the white grub antigen prep implied that fish antigens were not acquired or had been removed during antigen preparation. Given this, it appears the mouse anti-bluegill hyperimmune serum was highly sensitive in its ability to detect low amounts of fish proteins that may have persisted in the antigen prep. However, any persistent fish remnants in the white grub prep may not have been sufficient to elicit an immune response in the mice. The immunogenicity of an antigen can be influenced by the dose (antigen load), molecular weight, and complexity of the introduced antigen

(Doevendans & Schellekens, 2019).

Mouse anti-white grub antibodies did recognize shared epitopes in the white grub and yellow grub antigen preps. Because these are both complex mixtures from trematode parasites that share similar morphology and complex lifecycles, it was not surprising that there was cross-reactivity. Cross reactive antigens are common among trematodes and cross-protective advantages have been demonstrated between Fasciola sp., Paragonimus sp., and Schistosoma sp. (Hillyer & Serrano, 1983). Although this undermines the specificity of the anti-white grub antibodies and its use in antibody evaluation, it would provide a broader spectrum of cross-protection if antibodies were protective. In addition,

it can be inferred that anti-yellow grub antibodies in fish sera would also detect white grub proteins, contributing to false positives in my study.

Because ELISA results indicated that bluegill sera bound the white grub antigen, I used western blot analysis in an attempt to identify specific proteins that were recognized by serum from infected bluegills. However, when bluegill serum was added prior to mouse anti-bluegill, rather than seeing additional bands over mouse anti bluegill alone, no bands were seen. Rather, there was high background staining of the nitrocellulose. These negative results suggested that grub-specific antibodies were not binding to white grub antigens, implying that the ELISA results may be indeterminate. False positives are a common occurrence with antibody-testing ELISA assays using polyclonal antibodies (Rao,

Gupta, Sayal, Ohri, & Kher, 1997; Rakel et al., 2015). Ineffective blocking or washing of the microtiter plates, and antibodies cross-reacting with the blocking solution can give rise to false positives. The characteristic resistance of polyclonal antibodies to rigorous washing suggests that non-specific binding of the mouse anti-bluegill sera antibodies to other components in the immune complex assembly can be maintained (Ivell, Teerds, &

Hoffman, 2014). The absorbance values recorded from the ELISA might be a result of non- specific binding of bluegill serum to Blotto or the microtiter plate. Thus, in spite of controls, it is possible that the polyclonal mouse antibodies interacted with white grub and/or blocking solution resulting in high backgrounds in the ELISA that were not as readily apparent in the less sensitive dot blot. Non-specific antibodies produced to other infections could also cross-react with similar antigens in the sample, thus contributing to false positives.

Given this, I suspect that the high burdens of infection might not be a result of the non- protective nature of antibodies, but an effective absence of specific antibodies. A study by

Hoglund and Thuvander (1990) reported the absence of a specific antibody response when infected fish serum was tested against Diplostomum spathaecum antigens in an ELISA. It may be possible that the cercaria stage is not present in the blood or lymphatic circulation for a duration long enough, to elicit an elaborate humoral immune response. Furthermore, cercarial encystation may limit antigen expression resulting in immunoregulation. This evasion strategy has been reported among fish parasites (Sitjà-Bobadilla, 2008). Hoglund and Thuvander (1990) suggested that the immune responses in Diplostomum spathaecum might be cell-mediated instead.

Because the adaptive immune response in fish is poorly developed, the apparent lack of specific antibodies is not unexplainable. An examination of pathological changes provided some insights into the roles of the well-developed fish innate immune system in

Posthodiplostomum minimum infection. Despite heavy infections and space-occupying lesions produced by the metacercariae, bluegills have been reported to maintain excellent body condition scores (White et al., 2019). Fish in this study were also in good condition, despite high parasite burdens. This may be a result of the regenerative abilities of the liver and kidney in teleosts to accommodate increasing infections (Hoppe et al., 2015;

Michalopoulos, 2007). In addition, the observed aggregation of P. minimum in the epicardium showed that the integrity of the heart muscle was little affected. Histological findings showed normal cellular structures in affected tissues and a thin layer of cellular infiltration, suggesting that the inflammatory response to P. minimum was mild. This could be due to the immunomodulatory activity of the parasites, which may be advantageous to

the fish host (Sitjà-Bobadilla, 2008), since an extensive inflammatory response typically results in tissue damage. Furthermore, metabolic energy can be traded off to fish growth and spawning activities, rather than eliciting an immune response (Rohlenová & Šimková,

2010). These findings might explain why the bluegill can tolerate very high burdens of infection while maintaining normal physiological functions. This may be evolutionarily important for the success of trematode transmission since infected fish must live long enough to be preyed upon by the bird host.

The most notable change observed in affected tissues was melanization. This is an innate immune mechanism that plays an immunological role in invertebrates and ectothermic vertebrates like fish. During infections, melano-macrophages recognize microbial pathogen associated molecular patterns (PAMPs such as LPS, peptidoglycans, structural carbohydrates, flagellin, dsRNA, etc.) and trigger a cascade of reactions that activates the phenoloxidase system. Activated phenoxidase catalyzes the hydroxylation of tyrosine, and oxidation of diphenol with the subsequent formation of melanin. Melanin is then secreted by the macrophages to encapsulate invaders and limit damage to the host (Kurt Buchmann,

2014; E. L. Cooper, 2010). A number of melano-macrophage centers were observed in the surrounding tissues, similar to previous reports of P. minimum-induced histological changes in fathead minnows (Mitchell, Smith, & Hoffman, 1982). This is similar to the characteristic peppered appearance of the skin in blackspot (Uvulifer ambloplitis) infections. Another fish innate mechanism, characterized by an expression of induced nitric oxide synthase (iNOs) in fish skin has been reported in Ichthyophthirius multifilis as well

(Sigh, Lindenstrom, & Buchmann, 2004).

In this study, I did not detect a specific antibody response to Posthodiplostomum minimum 65

infection in bluegills. The lack of a true negative control, in the form of fish known never to have been exposed, was a potential problem. However, because there was no difference in absorbance between heavy burdened and light burdened bluegills, a true negative control would have no impact on the ability to differentiate parasite burdens.

I chose to use an indirect ELISA because it is most appropriate for rapid screening of habitats to test for endemicity of white grubs. It may be useful to prepare a monoclonal capture antibody to specific structural proteins on the parasite and combine this with anti- white grub polyclonal detection antibody in a capture ELISA assay. Although labor- intensive and expensive, this could provide a better picture of the infection intensity and the immune response. This would likely be a purely academic exercise, however. It is possible that some non-specific, antimicrobial peptides may be present in green sunfish, but not bluegills, potentially explaining why the green sunfish is resistant to P. minimum

Finally, skin mucus has been shown to be an important component in protection from pathogens in fish. It contains immunoglobulins and bio-reactive molecules (Reverter,

Tapissier-Bontemps, Lecchini, Banaigs, & Sasal, 2018). Since infection by white grub is via cercarial penetration of the skin, mucus from both sunfish species should be tested for any potential in protection.

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