CILIATE XENOMAS IN VIRGINICA

FROM GREAT BAY, NEW HAMPSHIRE

By

EMILY SCARPA McGURK

A thesis submitted to the

Graduate School – Camden

Rutgers, The State University of New Jersey

in partial fulfillment of the requirements

for the degree of Master of Science

Graduate Program in Biology

and approved by

______

Dr. William Saidel

______

Dr. David Bushek

______

Dr. Susan Ford

Camden, New Jersey; May 2013

ABSTRACT OF THE THESIS

Ciliate xenomas in Crassostrea virginica from Great Bay, New Hampshire

By EMILY SCARPA McGURK

Thesis Director:

William Saidel

During routine histological examination of (Crassostrea virginica) from

Great Bay, New Hampshire, a high prevalence and intensity of ciliate xenomas has been noted since 1997. Xenomas are hypertrophic lesions on the gills of bivalve molluscs caused by . Although not known to cause mortality in oysters, xenomas have not previously been reported at this high level of abundance. The objectives of this study were to characterize the xenomas, classify the ciliates, and gather baseline epizootiological data with correlations to environmental and biological parameters.

Upon gross examination, xenomas appeared as white nodules located in the gill tissue, up to 3 mm in diameter, occasionally fusing into large masses along the gill filaments. Light microscopy of histological sections revealed xenomas located in the gill water tubes that often occupied the entire cross sectional area. Higher magnification revealed dual nuclei, eight kineties, and conjugation. Transmission electron microscopy

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revealed dual nuclei that vary in density, a maximum of twenty cilia in each kinety radiating from the oral apparatus to the posterior, and a 9+2 axoneme structure within the cilia. Sequencing of the 18S rRNA gene produced a unique sequence not present in

GenBank. These gill ciliates are generally listed as Sphenophrya dosiniae (Order

Rhynchodida) and although there are no representatives of Rhynchodida in GenBank’s database, all similar matches were within the class . Since 1997, xenoma prevalence has fluctuated with peaks in 2000, 2004, and 2011. Infected oysters generally contained <30 xenomas, but 2.1% contained >100, sharply contrasting the rare prevalence and low intensity reported elsewhere. Prevalence increased with oyster size, leveling off near 50% in oysters >60mm. Infection intensity peaked in 70-90mm oysters.

Individually, oyster condition was not associated with xenoma intensity, but sites with oysters in higher condition generally had a greater prevalence and intensity of xenoma infections. Seasonal data indicated an infection cycle increasing from summer to fall, peaking at 55-65% in November and dropping to <10% by spring. The oyster population at Great Bay, NH warrants further examination to understand the mechanisms and conditions controlling xenoma formation, as well as the possible effects of a changing climate.

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ACKNOWLEDGEMENTS

I would like to express my deep appreciation to Dr. David Bushek, for his guidance and assistance in my research and writing. I would not have begun this journey without his encouragement. I would also like to express my gratitude to Dr. Susan Ford and Dr. William Saidel, for their mentorship, thought provoking questions, and of course, for their patience!

I would also like to recognize Dr. Raymond Grizzle, of the Jackson Estuarine

Laboratory, who donated his time and efforts to collect samples for me throughout the year, and Bruce Smith, of the New Hampshire and Game department, who made sure that their annual samples arrived with plenty of extra oysters for me to examine.

Both were very accommodating in the accrual of any background information and data that could be of use.

I am indebted to my colleagues at the Haskin Shellfish Research Lab, those still around as well as those who have come and gone, who were instrumental in keeping me focused, providing either a second opinion or empathy as needed.

Finally, I dedicate this thesis to my family for their endless support and encouragement, my best friend for distracting me when I most needed a break, and my husband for his patience and understanding throughout.

iv TABLE OF CONTENTS

Abstract ….…………………………………………………..…… ii

Acknowledgements ...... ….....…………………………………………… iv

Introduction: Oyster Parasitology and Xenomas ...... ……..….……..…… 1

Background ...... ……………………………………………..……. 3

Methods ...... …………………………………………….…… 8

Results & Discussion ...... ….…………………………………..……….. 13

Conclusion ...... ………………………………………….………. 20

Figures ...... ……….……………………………………..…… 24

Tables ...... ……….……………………………………..…… 42

References ...... …………………………………………………... 50 1

INTRODUCTION: OYSTER PARASITOLOGY AND XENOMAS

Parasites have historically presented problems for shellfisheries and .

For example, (causative agent of MSX) and marinus

(causative agent of dermo disease) have devastated oyster populations on the east coast of the United States (Ford and Tripp, 1996) while such as Marteilia spp.,

Bonamia spp. and Perkinsus olseni have caused massive mortality of shellfish worldwide

(Bower et al., 1999; Sparks, 2005). When first detected in 1957, MSX had caused 90-

95% mortality in eastern oysters, Crassostrea virginica, which had been planted on leased grounds in lower Delaware Bay. Subsequently, H. nelsoni has been found from

Nova Scotia, Canada to Biscayne Bay, Florida (Burreson and Ford, 2004). Dermo disease has caused 50% mortality in C. virginica populations (Carnegie, 2009). The European flat oyster, Ostrea edulis, in Brittany, France experienced 50-90% mortality by Marteilia refringens infection in the 1960s, and then again by Bonamia ostreae in the early 1980s

(Sparks, 2005). The 1970’s mortality of the Sydney rock oyster, C. commercialis, in

Queensland, Australia was caused by M. sydneyi (Sparks, 2005).

Recent publications have attributed marine diseases over the past few decades to both compromised immunity caused by physiological stress of climate change or pollution, as well as direct disease transmission facilitated by human activity (Harvell et al., 1999; Lafferty et al., 2004; Mydlarz et al., 2006). It is possible that increased vigilance has led to increased reporting of disease events. During routine histological

2 examination of oysters (Crassostrea virginica) from Great Bay, New Hampshire, by the

Haskin Shellfish Research Laboratory, a high prevalence and intensity of ciliate xenomas, as described below, has been noted since 1997. The xenoma infections were at a greater prevalence and intensity than observed in other regions, possibly a signal of an emerging disease problem. Despite the abundance of information about several parasites of eastern oysters (C. virginica), relatively little is known about xenomas. Xenomas are not known to cause mortality, but they previously have not been reported at prevalence above 2.5%, and given the devastating effects that parasites can exert on shellfish populations, proactive investigation of changes in parasite prevalence is imperative.

Oysters in Great Bay, NH have suffered substantial reductions in both distribution and abundance due to pollution, overharvest, and siltation (Bolster, 2002). During the mid-1990s, an outbreak of MSX and Dermo disease caused further decline of the population. In response to these recent mortalities, and as part of restoration efforts, a program of annual population survey and disease monitoring by the New Hampshire Fish and Game Department was initiated in 1995. As part of this monitoring, oyster samples collected from one to six sites were sent each fall to Rutgers University’s Haskin

Shellfish Research Laboratory (HSRL) for histopathology analyses. In the early 2000s, it became apparent that an unusually high prevalence of xenomas was present in the Great

Bay oyster population. In 2005, the current study began to more thoroughly document the phenomenon of xenomas in Great Bay oysters.

The purpose of this study was to characterize both the ciliates and the xenomas that they form and to gather baseline epizootiological data that would reveal correlations with various environmental and biological parameters to begin to elucidate the cause and

3 effect of the unusual abundance of xenomas in oysters from Great Bay, New Hampshire.

The first objective was to characterize the xenomas and the pathological effect on the host through gross examination of the gills as well as light microscopy (LM) and transmission electron microscopy (TEM). The second objective was to classify the of ciliate involved via LM and TEM, as well as by sequencing a portion of the

DNA amplified from the xenomas. The third objective was to gather baseline epizootiological data by identifying temporal and spatial patterns of infection in Great

Bay, NH. Patterns were analyzed and correlated with several parameters, including oyster size, condition index, and prevalence of other parasites and disease.

BACKGROUND

Xenomas are intracellular hypertrophic lesions that can be found on the gills of fish and bivalve molluscs. They occur when a parasite enters a host and proliferates, causing the host cell to grow abnormally large. In fish, xenomas are commonly a symptom of microsporidial gill disease and contain microsporidial spores (Morrison and

Sprague, 1981). Xenomas in bivalve molluscs, however, are caused by ciliates and have been observed in both mussels and oysters. For example, the zebra mussel Dreissena polymorpha commonly harbors the ciliates Conchophthirus acuminatus, Conchophthirus klimentinus, Hypocomagalma dreissenae, Sphenophrya dreissenae, and Sphenophrya naumiana (Laruelle et al., 1999). In contrast, ciliates that parasitize the gills of oysters are typically reported to be Sphenophrya dosiniae (Lauckner, 1983; Bower et al., 1994).

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The phylum Ciliophora is distinguished from other by four main characteristics (Corliss, 1979 & 1985; Small and Lynn, 1985; Lynn and Corliss, 1991).

The first and most readily apparent characteristic is a heterokaryotic nuclear condition, or nuclear dualism (i.e., each individual contains a small micronucleus and a large macronucleus, Figure 1). In this condition the micronucleus contains condensed, physiologically inactive chromatin and the macronucleus contains dispersed and condensed chromatin that is active in gene expression throughout the cell cycle. The second characteristic is the presence of cilia or ciliary organelles in at least one life stage

(Figure 1). The third characteristic is a complex kinetidal, or infraciliature, system, which is made up of single, paired, or multiple kinetosomes, also known as basal bodies, with microtubules and microfibrils and external ciliature arranged in various patterns (Figure

1). The fourth characteristic of the phylum Ciliophora is sexual conjugation in which two individuals temporarily fuse and exchange nuclear material from the micronucleus to replace the original macronucleus in order to increase genetic diversity in the subsequent generation.

Ciliates typically have some type of mouth, as well as membrane-bound flattened sacs underlying the plasma membrane called pellicular alveoli and tubular mitochondrial cristae. Generally, motile cilia contain the 9+2 axoneme structure of microtubules, while non-motile cilia contain the 9+0 axoneme structure (Figure 1d, Dawe et al., 2007). Body size may range from 10 to 4,500 µm in length and 103 to 1010 µm3 in cell volume. Body shape can vary from spherical to highly elongate. Ciliates may be free-living or symbiotic, free-swimming to sessile, and occur in a wide range of diverse habitats (Lynn and Corliss, 1991). Although ciliates may be autotrophic or mixotrophic, most ciliate

5 species are heterotrophs using bactivory, herbivory or carnivory to obtain food by a variety of mechanisms from filter feeding to raptorial predation. In fact, after water quality and habitat characteristics, the most commonly cited ecological factor controlling ciliate distribution and abundance is the availability of a specific food source (Lynn and

Corliss, 1991).

Symbiotic ciliate species are less common than free-living species, typically inhabiting marine and freshwater invertebrates as fairly innocuous endocommensals, ectocommensals, or epibionts (Lynn and Corliss, 1991). Although many species of ciliates live in some association with a host species, few species of ciliates are described as definitively harmful to their host and recognized as pathogenic parasites.

In the life cycle of most ciliates, a trophont stage divides by binary , but rather than dividing symmetrogenically, or longitudinally, ciliates usually exhibit division either along the kineties, otherwise called perkinetal division, or transversely across the kineties, called homothetogenic division (Lynn and Corliss, 1991). The two individuals created then develop to resemble the parental form and repeat the cycle. Some symbiotic species do, however, have polymorphic life cycles that contain stages that may or may not be symbiotic and sometimes involve reproductive cysts.

The ciliate fossil record has not provided much phylogenetic information.

Although ciliates are likely to have originated in the Precambrian era, there are no known fossils from this time, and relatively few in more recent times (Lynn and Corliss, 1991).

The morphological diversity of the phylum Ciliophora complicates identification attempts and has led to a number of suggested classification methods varying in the choice of characters and methods of analysis. Today, the most commonly accepted

6 classification places emphasis on patterns of infraciliature and kinetidal diversity, and applies the structural conservatism hypothesis, which proposes that the conservation of a structure over time is inversely related to the level of biological organization in an . The classification scheme used below is based on somatic kinetidal features as well as oral structures, life cycle information, and molecular data, when available, and is summarized in Table 1.

There are 11 monophyletic classes: Heterotrichea (Stein, 1859), Spirotrichea

(Bütschli, 1889), (Schewiakoff, 1896), (Jankowski, 1964),

Karyorelictea (Corliss, 1974), (de Puytorac et al., 1974),

Phyllopharyngea (de Puytorac et al., 1974), (Small and Lynn, 1981),

Litostomatea (Small and Lynn, 1981), (Small and Lynn, 1981), and

Plagiopylea (Small and Lynn, 1985). The classification of ciliates has been debated over the years, and while both morphological and molecular classification schemes agree on the classes Heterotrichea, , Phyllopharyngea, Colpodea, , and

Nassophorea, there is some disagreement between morphological and molecular methods of classification when describing the classes Spirotrichea, Prostomatea, Armophorea,

Oligohymenophorea, and Plagiopylea (Lynn, 2003; Table 1).

The heterotrichs have well-developed postciliary microtubular ribbons and utilize extramacronuclear microtubules to aid in division of macronuclei (Diener, Burchill and

Burton, 1983). The karyorelicteans are generally long, flat, vermiform ciliates that live interstitially in marine sand (Lynn and Corliss, 1991). The colpodeans have a curving body profile and inhabit freshwater and edaphic, or soil, habitats. The litostomates are made up of non-toxic endosymbionts of , as well as carnivorous and

7 protistivorous free-living ciliates that use toxicysts to immobilize prey. The nassophoreans are rare and inhabit benthic marine and freshwater areas, feeding on cyanobacteria and photosynthetic protists. Dense oral polykinetids, areas defined by the presence of numerous cilia, characterize the spirotrichs, a highly diverse group (Lynn,

1991). The prostomates have simple oral ciliature at or near the anterior end and are planktonic and benthic in marine and freshwater habitats. However, this class is not supported by small subunit rRNA gene sequence analysis (Lynn, 1991; Lynn and Corliss,

1991). The oligohymenophoreans have few oral polykinetids on one side of the cytostome, but vary widely in morphology otherwise (Lynn and Corliss, 1991). The class

Oligohymenophorea is not typically supported by small subunit rRNA gene sequences

(Lynn, 1991). Armophorates contain methanogenic endosymbionts within hydrogenosomes and can be tightly classified by small subunit rRNA gene sequences, but include groups with a somewhat diverse kinetid arrangement (van Hoek et al., 2000;

Lynn, 1991). Plagiopylates also contain hydrogenosomes and are well classified by small subunit rRNA (Lynn and Struder-Kypke, 2002), but cannot be identified based on kinetid morphology.

The ciliates contained within the xenomas investigated in this study are likely to be of the Phyllopharyngea class. Phyllopharyngeans have radially arranged, “leaf-like” microtubular ribbons lining the cytopharynx, a non-ciliated tube leading from the cell mouth, or cytostome, into the inner of the organism (Lynn and Corliss, 1991).

The ciliated stage contains somatic monokinetids that lack or have a reduced transverse microtubule ribbon and have a distinctively shaped, laterally directed kinetodesmal fibril.

Although classification schemes diverge here, there seem to be five subclasses within the

8 class Phyllopharyngea: Chonotrichia, Suctoria, Cyrtophoria, Phyllopharyngia, and

Rhynchodia. The Chonotrichia consist of sessile ectocommensals that live on the appendages of various . The Suctoria are a tentacled, carnivorous group that feeds on other ciliates in marine and freshwater habitats as well as within the digestive tract of various vertebrates. The Phyllopharyngia, which sometimes includes the

Cyrtophoria, are free-living or ectoparasitic in sewage treatment plants, aquaculture facilities, and aquatic mammals. The Rhychodia are obligate parasites of tunicates and bivalve molluscs, and reportedly consume the epithelial and gill tissues of their hosts.

The rhynchodids Sphenophrya spp. parasitize bivalve molluscs.

These characteristics will be used to describe and help classify the ciliates contained within the xenomas of oysters from Great Bay, New Hampshire.

METHODS

Objective 1: Characterize xenomas and pathology

Samples (n = 20 to 30 oysters per site) were collected each fall between 1997 and

2012 by staff at the University of New Hampshire’s Jackson Estuarine Laboratory and

New Hampshire’s Fish and Game Department with oyster tongs or by hand from among the following sites: Adam’s Point, Nannie Island, Oyster River, Piscataqua River,

Squamscott River, and Woodman Point (Figure 2, Tables 2 and 3). Samples were shipped overnight to HSRL. Shell height of each oyster, defined as the greatest distance from hinge to bill, was measured to the nearest 0.01 mm and recorded. Oysters were then

9 shucked and the entire gill surface of each oyster examined to enumerate macroscopically detectable xenomas. This gross quantification was not performed on samples collected prior to 2005. The rectum and a piece of mantle from above the labial palps were carefully dissected and incubated in Ray’s Fluid Thioglycollate Medium (RFTM) for seven days to detect Perkinsus sp. infections (Bushek et al., 1994).

A cross section of each animal containing digestive gland, gonad, mantle, and gill was preserved in Davidson’s fixative (Table 4) for 48 hours, and processed by routine histological methods (Howard et al., 2004) for light microscopy (LM). Briefly, the section was dehydrated and cleared in an ascending series of ethanol, xylene, and molten paraffin baths (Table 5), before being embedded in paraffin blocks. Five-micrometer sections of each oyster were mounted on slides and stained with hematoxylin and eosin

(Tables 6 and 7) and then examined under light microscopy for xenomas and any other parasites or pathological conditions. Because Great Bay oysters were processed and examined by another technician prior to 2005, archived slides were re-examined for consistency of diagnosis and quantification. Number of xenomas per oyster was recorded and compared spatially across sites and longitudinally by time.

Ten xenomas from the fall 2008 collections were prepared for transmission electron microscopy (TEM) following a modification of previously described protocols

(Smolarz, Renault, and Wolowicz, 2006; McGourty et al., 2007). Whole xenomas were excised and preserved in 2.5% glutaraldehyde buffered in 0.1 M cacodylate buffer (pH

7.39) for 2 hours at 4°C. Samples were then fixed in 1% osmium tetroxide in buffer for one hour, then rinsed three times in buffer and stained in 0.5% uranyl acetate in distilled water for 1 hour. Samples were dehydrated through an ascending ethanol series (50%,

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70%, 95%, and 100%), then infiltrated and embedded in a plastic mixture with propylene oxide as an intermediate fluid. Blocks were sectioned at 70-80 nm thickness and mounted on grids for viewing and photography via a Zeiss 902A TEM using an electron energy loss spectrometer (EELS) system to enhance contrast. Negatives were scanned at 800-

1200 dpi and contrast-enhanced using Photoshop©.

Objective 2: Classify causative agent

The collection and microscopy methods used to classify the species of ciliate involved in formation of xenomas in oysters were the same as those described for light microscopy and transmission electron microscopy in the previous section.

For molecular identification of ciliate species, samples of xenomas collected during the monthly sampling were excised from gill tissue and preserved in 95% ethanol.

DNA was extracted using a Qiagen DNEasy and Tissue kit. The 18S rRNA gene was amplified using the 384F (5’ YTB GAT GGT AGT GTA TTG GA) and 1147R (5’

GAC GGT ATC TRA TCG TCT TT) primers developed for ciliate identification

(Dopheide et al. 2008). These primers have successfully amplified ciliates in seven of the eleven classes of Ciliophora, with the exception of heterotrichs, armophoreans, karyorelicteans, and plagiopleans, none of which are likely to be the class of the target organism due to major differences in habitat and morphology. The 25µl polymerase chain reaction (PCR) mixture contained 1x PCR buffer, 2mM MgCl2, 200 µM BSA, 1 U

AmpliTaq DNA polymerase (all Applied Biosystems), 100 µM deoxynucleoside triphosphates (Invitrogen), 0.2 µM each of forward and reverse primers, and 1 µl template DNA. The schedule for the thermal cycler was as follows: initial denaturation at

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94°C for 5 minutes; 30 cycles at 94°C for 45 seconds, 55°C for 60 seconds, and 72°C for

90 seconds; with final extension at 72°C for 7 minutes before being held at 4°C

(Dopheide et al., 2008). The expected 750 bp product was detected by electrophoresis and stained with SYBRgreen (Invitrogen). Amplified samples were sent to the High

Throughput Sequencing Center at University of Washington (www.htseq.org) for purification and sequencing. Sequence quality was assessed by visual inspection of sequence chromatograms, and alignment of sequences was performed using the Vector

NTI program from Invitrogen. A BLAST search was used to find matching or similar nucleotide sequences in the GenBank sequence database to the resulting 554 bp consensus sequence.

Objective 3: Gather baseline epizootiological data

Prior to 2005, xenomas were exclusively identified from histological sections.

From 2005 to 2012, macroscopically visible xenomas were also counted on the entire gill surface prior to dissection and fixation. These numbers were compared to the results from histological sections to evaluate the reliability and efficiency of each method in detection and quantification of xenomas. Based on the results described below, it was clear that macroscopic examination was an efficient method of detection and quantification of xenomas and was utilized, where appropriate, for the purposes of this study.

To examine spatial and temporal patterns, fall samples were compared across years and across regions (i.e., river sites versus open bay sites, Figure 2). In addition, monthly samples were collected from June to November 2008 and from May to

December 2009 from Nannie Island, a site in Great Bay previously determined to have a

12 high prevalence of xenomas in annual fall samples. These oysters (n = 40-67 each month) were measured to determine size frequencies and xenomas were quantified by macroscopic examination. Whole gills of infected oysters were dissected, preserved in

Davidson’s fixative, and stored in 70% ethanol for future histological analysis.

Condition index was calculated for samples collected in fall 2009 and 2010 from all six sites in Great Bay (Adams Point, Nannie Island, Woodman Point, Oyster River,

Squamscott River, and Piscataqua River) to determine if either the presence of the parasites was affecting oyster health or if poor condition allows oysters to more easily become infected. Condition index methods were adapted from Lawrence and Scott

(1982). Oysters were weighed whole (W) and shucked. The number of xenomas was then recorded and the oyster was tagged and fixed whole for 48 hours. Valves were air dried and weighed (V). Fixed tissues were weighed before (w) and after (h) a 5mm section was removed for histological examination. The remaining tissue was then dried in an oven at

60°C for 2 weeks before obtaining a final dry weight (d).

Condition index was calculated by the following equations:

calculated dry weight = (d / h) * w

condition index = [calculated dry weight / (W - V)] * 100

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RESULTS AND DISCUSSION

Characterization of xenomas and classification of the ciliate species

In the oysters collected from Great Bay, NH, xenomas were commonly visible to the naked eye and appeared as white nodules of varying size located in the gill tissue

(Figure 3a), measuring up to 3 mm in diameter. In many cases, scalloped or otherwise damaged gills were observed in oysters with either no xenomas or xenomas located along the outermost edge of the gill (Figure 3b).

Light microscopy of histological sections revealed that the xenomas observed in

Great Bay oysters were located in the gill water tubes, and were often large enough to occupy the entire cross sectional area (Figure 4a,b). The number of parasites within the xenoma cross-section varied from just a few to thousands and the diameter of each xenoma ranged from 30 to 3,000 µm (Figure 4c). Nuclear dualism was apparent in most ciliates examined and five to eight rows of cilia, or kineties, were visible on several individuals under the light microscope (Figure 5), but were not visible in all individuals.

No cilia were observed on morphologically similar gill ciliates found unassociated with a xenoma. Fused ciliates, suggesting conjugation, were also observed under light microscopy (Figure 5b).

TEM further revealed the division of the ciliate nucleoplasm into the macronucleus and micronucleus, and that these two compartments vary in density (Figure

6). While cilia were not observed in all individuals, a maximum of twenty cilia could be

14 counted in each kinety when present (Figure 7a). The kineties radiate from the oral apparatus to the posterior end of the organism (Figure 7a) and emerge from a nematodesmal fibril (Figure 7a,b), both of which indicate that these cilia are used primarily for feeding as nematodesma are bundles of microtubules that support organelles associated with the cytostome or cytopharynx. Additionally, cilia appear to be motile, with a 9+2 microtubule axoneme structure (Figure 7b), rather than the non-motile 9+0 structure containing no central microtubule pair (Figure 1d, top).

The 18S rRNA gene sequence obtained from this study (GenBank accession number KC798064) had no sequence matches in GenBank and none close enough for identification. Only nine of the eleven classes of Ciliophora were represented in the database (Table 1) and the order Rhynchodida, to which the genus Sphenophrya belongs, currently has no representatives. However, the highest alignment scores were found with eight clones of uncultured ciliates recovered from the pallial fluid of the deep-sea bivalves Bathymodiolus thermophilus and Calyptogena magnifica. These ciliates are also in the class Phyllopharyngea, but from a different subclass (Suctoria) than the genus

Sphenophrya. The query coverage ranged from 66-68%, meaning that only 66-68% of the target sequence was covered by the one found. E-values, which describe the number of random matches that could be expected in a BLAST search, ranged from 1x10-39 to

2x10-30, indicating a very small likelihood of the sequence matches resulting by chance.

Ciliates are, however, a relatively large phylogenetic group. The closest 100 matches included ciliates in the class Phyllopharyngea and ranged across subclasses Chonotrichia,

Cyrtophoria, Phyllopharyngia, and Suctoria (Figure 8).

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Baseline epizootiological data

During the course of this study, it became apparent that macroscopic examination of gills may detect heavy infections more accurately than microscopic examination via histology. The presence or absence of xenomas was equally detected by both macroscopic and microscopic methods in 74% of the samples. However, in those samples in which the methods each produced different results (n = 210), 87% (n = 182) were missed by the microscopic method, but detected by the macroscopic method whereas only 13% (n = 28) were missed by the macroscopic method, but detected by microscopy.

Chi-square analysis shows that these results are significantly different from a random distribution (Table 8). Furthermore, counts of xenomas in each individual were typically an order of magnitude greater from the macroscopic method than from the regular histological method (Figure 9). This most likely results from the small amount of tissue included in histological sections and uneven distribution of xenomas. As xenomas were not uniformly scattered across the gills, the likelihood of a 5µm-thick histological section landing directly on a representative section of infected gill is small. These results show that in locations such as Great Bay, NH where xenomas tend to be large enough to detect macroscopically, the more efficient method of gross examination that scans the entire gill area is the best option. However, because gross examination does not enable detection of the much more common small xenomas found in most oyster populations at low prevalence and intensity, it would not be advisable as an initial detection method because it would miss most, if not all, instances without large, highly prevalent xenomas.

Over the past 13 years, xenoma prevalence in Great Bay, NH has fluctuated, with peaks of 40-50% average prevalence in 2000, 2004, and 2011 (Figure 10). Fall

16 prevalence varied according to site as well as by year. The high degree of variability can be partially attributed to extreme peaks at a single site in a given year (e.g., Nannie Island had prevalence of 82% in 2004, while prevalence at other sites was between 29% and

49%). Prevalence and intensity tracked each other during the study and there was a tendency for river sites to be lower than mid-bay sites in both metrics, but this was not consistent with exceptions occurring in 2008, 2011, and 2012 (Figure 11). River sites did not experience the extreme peaks in intensity in 2000 and 2004 that were observed at mid-bay sites. When examined macroscopically, oysters (n = 2,062) generally contained fewer than 30 xenomas, but those with infections often contained between 2 and 10 xenomas, and 2.1% of oysters collected in the annual fall surveys contained greater than

100 xenomas (Figure 12).

To explore multi-parasite dynamics, xenoma infections were compared separately to MSX and dermo infections in individual oysters using the Pearson correlation coefficient. Only individuals in which at least one of the diseases in the analysis was present were used. No correlation was detected between xenoma abundance and dermo infection intensity (Figure 13, r-square = 0.03, p-value = 0.4). There was a weak, but significant negative correlation between xenomas and MSX infection intensity (Figure

13, r-square = -0.2, p-value = 0.0003). This negative association may result from parasite competition or inhibition, but reason for the relationship is unclear and warrants further investigation.

The greatest number of xenomas was observed in oysters between 70-90 mm in shell height (Figure 14a). Although Piscataqua River and Squamscott River contained the lowest and highest (respectively) mean shell heights, these sites had the lowest

17 prevalence and intensity of infections (Figure 14b,c). The relationships between shell height and mean prevalence, and shell height and mean intensity, were fitted using a plant disease Weibull model with an offset and a membrane transport model, respectively, via the ZunZun.com Curve Fitting Program (www.zunzun.com). Prevalence increased with size but leveled off between 42% and 53% once oysters reached a shell height of 60 mm

(Figure 15; y=1–exp(-1.0*((x(-1.19*105))/(1.19*105)4.75*10^3)+(-5.12*10-1), r2 = 0.94, p- value = 9.2*10-5). Infection intensity was greatest for oysters that were 70-90 mm in shell height, with a peak of 16.46 ± 2.96 (SEM) xenomas per oyster at 80 mm (Figure 15; y=((1.08*102)(x-1.64))/(x2+(-1.54*102)x+(6.35*103), r2 = 0.93, p-value = 0.0002). The low prevalence and intensity of infections in small oysters is likely a result of less exposure due to a lower filtration capacity and a shorter period of exposure compared to larger, older individuals. Likewise, the peak in xenoma infection intensity in oysters in the 70-90 mm size range may be indicative of a higher filtration capacity and longer period of exposure. Therefore, it is not necessary to invoke differences in immune capacity to explain differences between small and moderately sized oysters. It is unclear, however, why the largest oysters retained high prevalence while having a decreased intensity of infections. This decline in intensity without a decline in prevalence could indicate some level of defense response or mortality caused by heavy infections. A study following groups of infected cohorts would help determine how oyster size/age affects infection status and determine whether or not ciliate xenomas are associated with mortality. Additional studies, such as a common garden experiment using native and non- native oysters, are needed to determine if the patterns of prevalence and intensity are phenotypic or genotypic responses (Gaffney and Bushek, 1996).

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In general, oyster condition index can vary seasonally and is strongly influenced by a number of environmental factors, spawning stage, and disease burden (Sakuda,

1966; Haven, 1962; Newell, 1985; Barber et al., 1988; Davies et al., 1988; Crosby and

Roberts, 1990; Roper et al., 1991; Roesijadi, 1996). Wargo and Ford (1993) observed a mean condition index of 11.3 in disease resistant stocks on aquaculture racks in Delaware

Bay, Austin et al. (1993) observed a condition index range of 6.0-11.0 in the

Rappahannock River in , and McLean and Abbe (2008) observed a mean condition index of 8.03 in oysters held in the discharge of a nuclear power plant in

Chesapeake Bay. Condition index of individual oysters, collected from Great Bay, NH in the fall of 2009 and 2010, ranged from 2.1 to 29.5 with a mean of 8.0. The Pearson correlation coefficient for all data combined from all sites for fall of 2008 and 2009 showed no significant relationship between condition index and xenomas (r = -0.02, p- value = 0.7). These data include a large number of oysters without any detectable xenomas that vary widely in condition. As noted above, site characteristics (salinity regime, food quality and quantity, etc.) can dramatically affect condition indices, therefore, when examined by site with the uninfected oysters excluded, the three mid-bay sites where infections were generally highest and most prevalent (Adams Point, Nannie

Island, and Woodman Point) all display a negative relationship between condition index and xenoma intensity (Figure 16a-c). Although the relationships are not particularly strong, they indicate that oysters on mid-bay reefs with more macroscopically visible xenomas tend to have a lower condition index. The other three sites that exist around the fringes of the Great Bay system (Oyster River, Piscataqua River and Squamscott River) did not display this relationship, largely because there were many fewer infected oysters

19 and those that were infected often had relatively light infections. For example, the Oyster

River sample had mostly light infections (less than 10 xenomas, Figure 16d) while there were few infected oysters at either the Piscataqua or Squamscott River sites (n = 6 and 3, respectively; Figure 16e,f). Most high intensity infections (>75 xenomas per oyster) coincided with condition index scores of less than 10 and oysters with a condition index of 15 or higher never had more than 14 xenomas. Overall, these observations indicate that extensive xenoma infections may be detrimental to oyster health.

Curiously, the sites that have a higher mean condition index also tend to have a higher xenoma prevalence (mid-bay sites and Oyster River) while the sites that have a lower mean condition index also had a lower prevalence (Piscataqua River and

Squamscott River, Figure 16g). High condition index and xenoma prevalence at the mid- bay sites and Oyster River may indicate that environmental parameters at these sites are such that the oysters are not negatively impacted by infection, and that the ciliates are able to fare better in a healthier host. The low condition index and xenoma prevalence at the Piscataqua & Squamscott River sites could indicate that either the environmental parameters favor neither the oysters nor the ciliates at these sites, or possibly that these oysters are more susceptible to mortality from xenoma infection. This may also be an effect of greater lag time in the tidal cycle, resulting in a better chance for transmission of parasites, although water flow data have yet to be collected in Great Bay.

Seasonal data from 2008 and 2009 indicated an infection cycle increasing from summer to fall (Figure 17). Prevalence of xenomas peaked at 55-65% in November during both years and dropped to less than 10% during the spring and early summer.

Intensity of infections followed the same pattern. However, the infection intensity

20 observed in 2009 was greater than intensity in 2008 (Figure 17). The late fall peaks are not directly related to mean temperature or salinity lagged by one month prior to collection (Figure 18a,b), although a relationship may emerge with a greater lag and more consistently collected data. This seasonality may be indicative of behavioral changes in the oyster host through progression of annual feeding and spawning cycles, or it may indicate an optimum time of year for ciliates to take advantage of oyster susceptibility. It may also be related to an as yet undocumented seasonality of the ciliates’ life cycle and population dynamics. The diminished peaks in intensity of infections from oysters located in the tributary rivers indicate that salinity fluctuations may influence xenoma infections. Further examination of meteorological data may help explain spatial patterns of xenoma infection as they relate to salinity regimes as well as the peaks in annual prevalence and intensity in 2000, 2004, and 2011.

CONCLUSION

Oysters are filter-feeding , which exposes them to invasion by a wide variety and large number of potential pathogens. This study investigated the occurrence of highly prevalent and unusually large intracellular parasite colonies, or xenomas, in oysters from Great Bay, New Hampshire. Microscopic analyses confirmed the inclusion of the causative organisms in the phylum Ciliophora because they possess a heterokaryotic nuclear condition, with both a macronucleus and a micronucleus.

Furthermore, early life stages possess cilia that are arranged in a complex infraciliature

21 system, with kineties radiating away from the oral apparatus in a patch on one surface of the organism. Light microscopy also revealed fused cells that suggest the occurrence of conjugation for . The lack of cilia on individuals located on gill tissue outside of the xenoma may indicate that reproduction is occurring more rapidly within the xenoma than outside of it, consequently exhibiting more early life stages inside the xenoma.

Oyster gill ciliates are virtually always listed as Sphenophrya dosiniae (Lauckner,

1983; Bower et al., 1994). Morphological characteristics such as dual nuclei, cilia, and observation of conjugation confirm that the parasites found within oyster xenomas are of the phylum Ciliophora. Several morphological characteristics support classification of the xenoma ciliates in the class Phyllopharyngea. The 100 closest GenBank matches to sequences obtained from xenomas in this study were all within the class

Phyllopharyngea, although the database does not contain the sequences of more closely related species. This classification is also supported by morphological traits. The oral apparatus is a single ingestatory tube. The bud stage is ciliated, but adults are not. The symbiotic lifestyle on the gills of oysters indicate that these ciliates are members of the order Rhynchodida, and this is also apparent from morphological observations of cilia that are located only in a ventral patch, used for feeding and absent in adults. This is consistent with the literature, which describes all ciliates living symbiotically or parasitically with bivalve molluscs as rhynchodids (Lauckner, 1983; Bower et al., 1994).

As no genetic sequences had been previously submitted to GenBank for the order

Rhynchodida, the sequence obtained in this study, accession number KC798064, will stand as a representative for future molecular identification of bivalve ciliates.

22

In contrast to the high prevalence and intensity of xenoma infection found in

Great Bay oysters, xenomas are relatively rare and found at very low intensity at other locations. While approximately 50% prevalence was routinely observed in New

Hampshire, with 2% of the oysters containing up to 500 xenomas, Lauckner (1983) reported a total xenoma prevalence of less than 1% in upper Chesapeake Bay oysters and recent fall surveys in Delaware Bay, NJ found only 0.008% prevalence in 2010 and 2011

(HSRL, 2011 and 2012). Moreover, no oyster from either the Chesapeake study or the

Delaware Bay survey had more than two xenomas present in a histological tissue section.

In a study that sampled Crassostrea rhizophorae from Todos os Santos Bay,

Bahia, Brazil over the period of 1975 to 1979, ciliates were only detected in 1978 and only at 2% prevalence (Nascimento et al., 1986), with no mention of xenoma presence. In contrast, Boehs et al. (2009) provide the first report of xenomas in C. rhizophorae from

Camamu Bay, Bahia, Brazil at 2.5% prevalence with one to two xenomas visible per histological section. Although this is a much lower prevalence and intensity than that found in New Hampshire, it may represent an increasing trend in ciliate infections corresponding to the apparent global trend of increasing marine disease suggested by

Harvell et al. (1999).

Parasites are subject to environmental constraints just as any other organism.

Those with short life cycles and generation times likely have seasonal patterns of fluctuating abundance. For example, infections of Perkinsus marinus in oysters from

Chesapeake Bay increase from spring to peak in late summer or fall due largely to water temperature (Ragone and Burreson, 1993), but are still present, sometimes at almost undetectable levels, throughout the year. Similarly, ciliate xenomas can exist at low

23 levels of abundance and intensity at many locations. Indeed, ciliates exist in the gills of oysters throughout the world without causing xenomas (Bower et al., 1994). Until we understand what controls the proliferation of xenomas in oysters from Great Bay, NH, there is an unknown risk of the spread of this phenomenon.

Because we currently have no data on the effects that heavy xenoma infection can have on an oyster population, more information about their particular optimal and confining environmental parameters is essential, and more comprehensive research must be carried out to understand the mechanisms and conditions controlling xenoma formation, as well as the possible effects of a changing climate. Further research could be efficiently carried out with the use of macroscopic gross examination of gill tissue, as well as transplant experiments to explore whether it is the environment, the virulence of the distinct ciliate populations, the susceptibility of a local oyster population, or a combination thereof, that is allowing this phenomenon to occur.

24

a b c d d a a

Figure 1: a-c) Examples of hypothetical ciliates showing both the consistent characteristics and variable morphology. Ci: cilia, K: kinety, Ma: macronucleus, Mi: micronucleus, Mo: mouth. d) Microtubule configurations within cilia. Top to bottom: non-motile 9+0 axoneme structure, motile 9+2 axoneme structure, basal body.

25

Figure 2: Oyster sampling sites located at various points in and around Great Bay, NH. Nannie Island, Woodman Point, and Adam’s Point are designated as the “mid-bay sites.”

26

a

b

Figure 3. Gross examination of oyster gill reveals multiple xenomas of varying sizes (a, arrows) as well as scalloped gills (b, arrows).

27

a b

3mm

Figure 2. Histological sections of oysters with (a) a b

2 xenomas and (b) multiple xenomas filling entire

gill area.

1.5mm

Figure 2. Histological sect ions of oysters with (a)

2 xenomas and (b) multiple xenomac s filling entire

gill area.

Figure 4. Histological sections of oysters infected with ciliate xenomas (arrows). a) Two xenomas. b) Multiple xenomas filling the entire gill area. c) Large xenoma.

28

a

8µm

b

10µm

Figure 5: Up to eight rows of cilia (thin solid arrows), dual nuclei (dashed arrows), and fused ciliates (thick arrow) were observed in histological sections of xenomas under the light microscope.

29

a

b

Figure 6: The macronucleus (Ma) and micronucleus (Mi) can be seen in detail with TEM. a) Kineties (between black arrows) and basal bodies (white arrow) were observed in some individuals while b) no cilia were observed in others, although this may be due to the angle or level of the section. 30

a

Nd

b

Nd

Figure 7: a) Approximately twenty cilia (C) per kinety (K), radiating from the oral apparatus (Or) to the posterior end of the organism and emerging from a nematodesmal fibril (Nd), were observed under TEM. b) Axoneme structure arranged in 9 peripheral microtubule pairs surrounding a central pair. Figure 8: Neighbor-joining tree showing phylogenetic relationships between the target sequence derived from xenomas (unknown) and a condensed list of the closest 100 sequence matches Subclasses listed on the right. 31 32

Macro. Xenoma count

500 450 400 350 300 250 R² = 0.33188 200 Macro count 150 100 50 0 0 10 20 30 40 50 Micro count

Figure 9: Comparison of macroscopic (gross examination) and microscopic (histological examination) methods of detection. Note that when none were observed macroscopically, the maximum counted by the microscopy method was 6, however when none were observed microscopically, up to 130 were counted macroscopically. Removal of outliers increases R2 from 0.3 to 0.4. Samples included are fall collections from 2005 through 2012. 33

0.9 12 Prevalence 0.8 Intensity 10 0.7 0.6 8 0.5 6 0.4 0.3 4

0.2 Mean xenoma count 2 Percent prevalence 0.1 0.0 0

Figure 10: Long-term mean xenoma prevalence and intensity (=/- SEM). Intensity is defined as the xenoma count. Fall samples collected Sept-Nov 1997-2012 and examined by histology to display the entire time series. 34

1.0 16 Midbay Prevalence River Prevalence 0.9 Midbay Intensity River Intensity 14 0.8 12 0.7 10 0.6

0.5 8

0.4 6 0.3 4 Mean xenoma count Percent prevalence 0.2 2 0.1

0.0 0

Figure 11: Percent prevalence (blue) and infection intensity (red), both with SEM, averaged across mid-bay sites (Adams Point, Nannie Island, and Woodman Point; solid lines) and river sites (Oyster River, Piscataqua River, and Squamscott River; dashed lines). Data are fall samples collected Sept-Nov 1997-2012; microscopic counts used for the purposes of giving a longer time series.

35

0.9 0.8 2005 2006 2007 0.7 0.6 2008 2009 2010 0.5 2011 2012 0.4 0.3 0.2

Percent of annual samples 0.1 0.0 0 1 2 - 10 11 - 30 31 - 60 60 - 100 Abundance per oyster

Figure 12: Infection intensity distribution from macroscopic examination as a percentage of all oysters examined. Grey area indicates the average of all years, 2005-2012. Samples collected Sept-Dec 2005-2012 (n=2,062).

36

6 5 DERMO MSX 5 4 4 3 3 2 R² = 0.0004 MSX score Dermo score 2

1 1 R² = 0.02781 0 0 0 100 200 300 400 500 Macro xenoma abundance

Figure 13: Correlations between xenoma infection intensity and either dermo or MSX infection.

37

500 a Adams Point 400 Nannie Island Woodman Point 300 Oyster River 200 Piscataqua River

Xenoma count Squamscott River 100

0 20 40 60 80 100 120 140 160 Shell height (mm) 0.6 b 0.5

0.4

0.3

0.2

Percent prevalence 0.1

0.0 60 65 70 75 80 85 90 95 Mean shell height (mm) 30 25 c 20 15 10 5 Mean xenomas/oyster 0 60 65 70 75 80 85 90 95 Mean shell height (mm)

Figure 14: Relationship between xenoma prevalence/intensity and size by collection site. a) Individual data. b and c) site averages with SEM for prevalence and intensity, respectively. Data are from Fall 2005-2012 macroscopic counts.

38

0.6 25 a prevalence 0.5 intensity 20

0.4 15 0.3 10 0.2 Percent prevalence 5 Mean xenoma count 0.1

0.0 0 20 40 60 80 100 120 Shell height (mm)

b c

Figure 15: Macroscopic percent prevalence and intensity with SEM (a) of xenoma infections across the size range of oysters with 95% confidence intervals (b,c). Fall samples, 2005-2012; macroscopic counts.

39

a d 20 20 Adams Point Oyster River 15 15

10 10

5 5 R² = 0.02882

Condition index R² = 0.04947 0 0 1 10 100 1 10 100 b 20 20 e Nannie Island Piscataqua River 15 15

10 10

5 5

Condition index R² = 0.03727 R² = 0.10044 0 0 1 10 100 1 10 100 c 20 20 f Woodman Point Squamscott River 15 15 10 10 5 5 R² = 0.07657 R² = 0.38425 Condition index 0 0 1 10 100 1 10 100 # xenomas # xenomas 0.8 g 0.6

0.4

0.2

Xenoma prevalence 0.0 6 7 8 9 10 11 Mean condition index

Figure 16: Relationship between xenoma prevalence/intensity and oyster condition by collection site. a) Mean condition index (SEM) versus xenoma prevalence and b-g) individual xenoma counts and condition index. Fall 2009 and 2010, gross counts.

40

80% 30 Prevalence 70% Intensity 25

60% 20 50% 15 40% 10 30% 5

20% Mean xenomas per oyster Percent prevalence 10% 0

0% -5 Jul-09 Jul-08 Jan-09 Jun-09 Jun-08 Oct-09 Oct-08 Sep-09 Feb-09 Sep-08 Dec-09 Apr-09 Dec-08 Aug-09 Nov-09 Mar-09 Aug-08 Nov-08 May-09

Figure 17: Seasonal trends observed in samples collected throughout the year. Dashed line indicates a lapse in data more than one month. Macroscopic counts only from Nannie Island/Woodman Point, June 2008 – Dec 2009.

41

0.7 30 a Prevalence 0.6 25 Intensity 0.5 20

0.4 15 0.3

Percent prevalence 10

0.2 Mean xenomas/oyster

5 0.1

0 0 8 10 12 14 16 18 20 22 24 Temperature (C)

0.7 30 b 0.6 Prevalence 25

0.5 Intensity 20

0.4 15 0.3

Percent prevalence 10

0.2 Mean xenomas/oyster

5 0.1

0 0 15 17 19 21 23 25 27 29 Salinity

Figure 18: Percent prevalence and infection intensity across a) 30 day mean temperature and b) 30 day mean salinity.

42

Table 1: Summary of the classes of Ciliophora based on somatic kinetidal features as well as oral structures, life cycle information, and molecular data, when available. Agreement and discrepancies between morphological and molecular distinctions are noted. Asterisks indicate those classes not represented in the GenBank database.

Morph/mol Classes agreement Habitat Morphology

Heterotrichea yes* Diverse Well developed postciliary microtubular ribbons; extramacronuclear microtubules aid in division of macronuclei Spirotrichea no Diverse Dense oral polykinetids; highly diverse Prostomatea no Planktonic & Simple oral ciliature at or near the anterior benthic; marine end & freshwater Armophorea no Diverse Contain methanogenic endosymbionts within hydrogenosomes; somewhat diverse kinetid arrangements Karyorelictea yes* Interstitial in Generally long, flat, vermiform marine sand Oligohymenophorea no Diverse Few oral polykinetids on one side of the cytostome; morphologically diverse Phyllopharyngea yes Diverse Radial microtubular ribbons; reduced transverse microtubule ribbon; laterally directed kinetodesmal fibril Colpodea yes Freshwater & Curving body profile soil Litostomatea yes Endosymbionts Endosymbionts are non-toxic, free-living or free-living ciliates are carnivorous and protistivorous and use toxicysts Nassophorea yes Benthic marine Feed on cyanobacteria and photosynthetic & freshwater protists Plagiopylea no Diverse Contain hydrogenosomes; kinetid morphologically highly diverse

43

Table 2: Latitude and longitude of collection sites within Great Bay, New Hampshire.

Adam’s Point 43°05’31.91”N 70°51’52.84”W Nannie Island 43°04’09.39”N 70°51’46.24”W Woodman Point 43°04’17.77”N 70°51’42.40”W Oyster River 43°07’58.88”N 70°54’06.56”W Piscataqua River 43°06’50.03”N 70°48’09.15”W Squamscott River 43°03’08.40”N 70°54’43.74”W

44

Table 3: Sites collected each year and the methods employed for detection and quantification. Site codes: AP = Adams Point, NI = Nannie Island, WP = Woodman Point, OR = Oyster River, PR = Piscataqua River, SR = Squamscott River.

Histological Gross exam AP NI WP OR PR SR counts counts 1997 X X X X X 1998 X X X X X 1999 X X 2000 X X X X 2001 X X X X 2002 X X X X 2003 X X 2004 X X X X 2005 X X X X X X 2006 X X X X X X X X 2007 X X X X X X X 2008 X X X X X X X X 2009 X X X X X X X X 2010 X X X X X X X X 2011 X X X X X X X 2012 X X X X X X X X

45

Table 4: Davidson’s fixative recipe.

% by volume Seawater 30 Ethanol 30 Formaldehyde 20 Acetic acid 10 Glycerol 10

46

Table 5: Tissue processing schedule for dehydration and clearing of tissues in preparation for embedding in paraffin blocks.

Step Reagent Time (hours) 1 95% ethanol 2 2 100% ethanol 1.5 3 100% ethanol 1 4 100% ethanol 1 5 100% ethanol 1 6 100% ethanol 1 7 100% ethanol 1 8 Xylene 2 9 Xylene 1.75 10 Xylene 1.5 11 Melted paraffin 2 12 Melted paraffin 2

47

Table 6: Procedure for the standard hematoxylin and eosin stain.

Deparaffinize Xylene 5 min Xylene 2 min Hydrate to water 100% ethanol 15 dips 100% ethanol 15 dips 95% ethanol 15 dips 95% ethanol 15 dips Running tap water 5 min Stain Ferric alum mordant 10 min Running tap water Quick dip Hematoxylin 30-45 min Running tap water 3 min Blue NaHCO2 2 min Running tap water 3 min Counterstain Eosin Y 2.5 min Differentiate eosin & dehydrate 95% ethanol 6 quick dips 95% ethanol 6 quick dips 100% ethanol 10 dips 100% ethanol 10 dips Xylene 3 min Xylene 5 min Xylene 5 min Xylene 5 min

48

Table 7: Recipes for solutions used in the hematoxylin and eosin staining procedure.

Ferric alum mordant Eosin Y stock 25g ferric ammonium sulfate crystals 800ml 95% ethanol 500ml distilled water 200ml distilled water 10g Eosin Y powder Groat/Weigert hematoxylin 245ml distilled water Eosin Y working solution 5ml sulfuric acid 100ml Eosin Y stock 5g ferric ammonium sulfate crystals 590ml 95% ethanol 245ml 95% ethanol 110ml distilled water 2.5g hematoxylin powder 4ml acetic acid

NaHCO2 1 scoop (~1teaspoon) 500ml distilled water

49

Table 8: Two-way contingency table for validation of the use of macroscopic detection methods versus microscopic examination; expected values are in italics, individual χ2 values are in parentheses. χ2=199.993, df=1, P(χ2>199.993)=0.0000

Macro + - 182 28 210 + 94.25 115.75 (81.69) (66.52) Micro 182 419 601 - 269.75 331.25 (28.54) (23.24) 364 447 811

50

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