Eukaryotic communities in epizootic shell disease lesions of the American lobster (Homarus americanus, H. Milne Edwards).
KEY WORDS: denaturing gradient gel electrophoresis, 18S rRNA gene, epibionts, American lobster, Homarus americanus
Various prokaryotic and eukaryotic organisms have been shown to form epibiotic communities on the surfaces of aquatic arthropods (e.g., Fernandez-Leborans & Von Rintelen 2007, Normant et al. 2007, Di Camillo et al. 2008, Fernandez-Leborans & Gabilondo 2008). Commensal or parasitic association of epibionts with their host constitutes biofouling, which may contribute to significant losses to the fishery industry. This can occur directly, by biofouling of the animals themselves (Rodriguez & Ibarra-Obando 2008), or indirectly, by fouling of cages, nets, filters, and other equipment (Braithwaite & McEvoy 2005). Most commonly, biofouling epibionts are studied by direct observation or aided by light or electron microscopy. This limits such surveys to metazoans or large protozoans (mostly ciliates). Indeed, ciliates are by far the most frequently described group found among epibionts of arthropods (e.g., Fernandez-Leborans 2003, Fernandez-Leborans & Von Rintelen 2007, Fernandez-Leborans & Gabilondo 2008, Lopez-Tellez et al. 2009). Metazoans are very diverse and include hydrozoans (Fernandez-Leborans & Gabilondo 2006, Di Camillo et al. 2008, Fernandez-Leborans & Gabilondo 2008), rotifers (Fernandez-Leborans & Von Rintelen 2007), bryozoans (Di Camillo et al. 2008), entoprocts (Fernandez-Leborans & Gabilondo 2006, Fernandez-Leborans & Gabilondo 2008), nematodes (Normant et al. 2007), annelids (Schejter & Spivak 2005, Fernandez-Leborans & Gabilondo 2006, Normant et al. 2007, Fernandez-Leborans & Gabilondo 2008), barnacles (Key et al. 1996, Girl & Wicksten 2001, Fernandez-Leborans 2003, Schejter & Spivak 2005, Fernandez-Leborans & Gabilondo 2006, Fernandez-Leborans & Gabilondo 2008), and other crustaceans (Normant et al. 2007) and molluscs (Normant et al. 2007).
Few studies exist on epibionts of commercially important lobsters--specifically, the American lobster, Homarus americanus. The only group of healthy lobster epibionts, which recently received attention for the research community, is symbiotic Cycliophora associated with lobster mouthparts (Obst & Funch 2006, Obst et al. 2006). Apart from lobsters held in captivity (one can easily observe barnacles on lobsters kept in public aquaria), wild lobsters usually do not develop an extensive epibiotic community. An exception would be lobsters afflicted by disease. For example Dove et al. (2004) have observed extensive epibiosis in lobsters with excretory calcinosis.
Currently, the most prevalent diseases of wild lobsters are various form of shell disease (syndrome). Shell disease was first identified in the American lobster (H. americanus H. Milne Edwards, 1837) as early as the 1930s (Hess 1937). Beginning in the late 1990s, a new form of shell disease of the American lobster termed epizootic shell disease (ESD) was described (Smolowitz et al. 2002). This disease has caused serious losses to the lobster fishery and natural populations on the northeast coast of the United States (Smolowitz et al. 2005). From 2000 to 2004, the Massachusetts Division of Marine Fisheries underwent a survey of wild lobster populations at 6 different locations in Massachusetts and found an average prevalence of 8% of females and 3% of males, but prevalence was as high as 28% at one southern location in 2003 (Glenn & Pugh 2006). Lobsters with ESD will often survive by mounting an effective immune response, primarily consisting of melanization of edges of the lesions, proliferation of an inflammatory membrane underlying the affected carapace, or by "molting out" of the disease (Smolowitz et al. 2005). However, affected lobsters are unmarketable because of the grotesque appearance of the shell lesions. ESD is characterized by severe, diffuse, deep erosions of the cephalothorax and the abdomen that, unlike other known forms of shell disease, spread irregularly over the dorsal carapace of the animal (Smolowitz et al. 2005). Another distinction from the more classic presentations of shell disease is that chitin degradation by chitinoclastic bacteria appears to occur minimally in the lesions (Chistoserdov et al. 2005, Smolowitz et al. 2005).
Although the etiology of ESD is not conclusive, it has been well established that high numbers of bacteria are present in the shell lesions and likely contribute to the biofouling of the carapace (Chistoserdov et al. 2005, Smolowitz et al. 2005). Thus, the bacterial communities of these lesions have been described by culture and observed directly using electron microscopy and histopathology (Hsu & Smolowitz 2003, Chistoserdov et al. 2005, Smolowitz et al. 2005). It is believed that the lipolytic, proteolytic, and, to some extent, chitinivorous activities of these bacteria degrade the carapace, producing the lesions. Interestingly, bacteria were not the only organisms observed in shell lesions. Smolowitz et al. (2005) described the presence of secondary micro-organisms, including nematodes, protistans, large and small protozoa, and barnacles, of which their role in ESD is unknown.
Although most investigations of microbial communities in ESD lesions have involved bacterial culture, little is known of the eukaryotic organisms present in the lesions. This study attempts to analyze the presence of eukaryotic micro-organisms in the lesions of lobsters with ESD in an attempt to understand better the complexity of the community and possibly to identify any eukaryotic microbes associated with initial stages of lesion development. Primers designed to target highly conserved regions of the eukaryotic 18S rRNA gene were used to amplify DNA from the lesion community. These amplicons were then separated based on sequence using denaturing gradient gel electrophoresis (DGGE), and sequenced for identification. Initially used to characterize bacterial communities by Muyzer et al. (1993), this technique has been an effective means of analyzing eukaryotic communities as well (Diez et al. 2001, Gast et al. 2004, Brad et al. 2008, Hatamoto et al. 2008).
MATERIALS AND METHODS
Lobster Collection, Lesion Sampling, and DNA Extraction
A total of 32 lobsters were collected from 2001 to 2008 at 5 different locations: Eastern and Central Long Island Sound (ELIS and CLIS, respectively), NY; Buzzard's Bay, MA (BB); Kittery, ME (KMA); and Rhode Island (RI), and were sent to our laboratory by courier as live animals (Table 1). Two Rhode Island lobsters had both lesion and healthy carapace samples taken. Status as diseased or healthy was determined by visual observation of ESD lesions according to the guidelines of the Maine Department of Fisheries. Any healthy lobsters were not visually showing signs of ESD, but may have been unhealthy in any other respect. The presence/absence of a visible epibiotic community on healthy surfaces and lesions was also noted.
The animals were first rinsed with sterile seawater over the entire carapace. Only lesions on the cephalothorax were sampled. Shell sampling for both affected and healthy carapace consisted of scraping with a sterile razor blade of the carapace or lesion and collecting the scraped material in sterile 50 mM Tris/ethylenediamine tetra-acetic acid (EDTA) buffer (pH, 8.8) (Sigma-Aldrich, Inc., St. Louis, MO). Egg white lysozyme (Amresco, Solon, OH) was added to the shell samples to a final concentration of 1 mg/mL and the mixture was then incubated at 37[degrees]C for 30 min. A sodium dodecyl sulfate solution was added to a 2% final concentration followed by addition of proteinase K (Fisher Bioreagents, Fair Lawn, NJ) at 1.25 mg/ mL and incubated at 50[degrees]C for 15 min. The samples were then subjected to 3 freeze-thaw cycles at 50[degrees]C then -80[degrees]C, and bead beating in a Mini-Beadbeater-8 with 0.1 mm Zirconia/Silica Beads (BioSpec Products, Bartlesville, OK). DNA was extracted using phenol/chloroform. Phenol (Sigma-Aldrich Inc., St. Louis, MO) was added at a 1:1 ratio, then centrifuged for 5 min at 5,000 g, and the aqueous phase was removed. A 1:1 volume of chloroform (FisherBiotech, Fair Lawn, NJ) was added and spun at 5,000 g for 15 min. DNA was precipitated from this aqueous phase by adding 1/10 volume of 3 M sodium acetate (Sigma-Aldrich, Inc., St. Louis, MO) and 2.2 x volume of 100% cold ethanol. The sample was then frozen overnight at -80[degrees]C and spun at 5,000 g for 35 min in an Eppendorf microcentrifuge. The DNA pellet was then washed with 70% ethanol, dried, and resuspended in 200 [micro]L sterile [ddH.sub.2]0. This raw DNA was then purified using the MoBio PowerClean DNA Clean Up Kit (MoBio Laboratories, Carlsbad, CA) according to the manufacturer's instructions.
Polymerase Chain Reaction of Eukaryotie Community DNA
For DGGE analysis, DNA samples were amplified directly using primers 960F (5'-GGCTTAATTTGACTCAACRCG-3') and 1200R (5'-GGGCATCACAGACCTG-3') (Gast et al. 2004). The forward primer 960F contained a 40-bp 5' GC-clamp attached (5'-CGCCCGCCGCGCCCCGCGCCCGTCCCGC CGCCCCCGCCCC-3') to produce amplicons suitable for DGGE analysis. This polymerase chain reaction (PCR) contained 25 [micro]L GoTaq Green Master Mix (Promega, Madison, WI), 1.5 [micro]M of the forward and 0.5 [micro]M of the reverse primers, an additional 1.5 mM Mg[Cl.sub.2], and 1.5 [micro]L template (approximately 200 ng) in a 50-[micro]L reaction volume. The thermocycling consisted of 2 initial cycles of denaturation at 95[degrees]C for 1 min, annealing at 65[degrees]C for 45 sec, and extension at 72[degrees]C for 45 sec. This was then followed by 10 cycles of denaturing at 95[degrees]C for 1 min, annealing at 65[degrees]C for 1 min, and extension at 72[degrees]C for 1 min. Finally, the cycling concluded with 25 cycles of the same temperatures and times except for annealing at 55[degrees]C. PCR products were verified for amplification by electrophoresis and ethidium bromide staining prior to DGGE (Sambrook et al. 1989).
Denaturing Gradient Gel Electrophoresis and Band Processing
DGGE was carried out using a CBS Scientific DGGE system (CBS Scientific Co., Del Mar, CA) in 1 x Tris-acetate EDTA buffer (pH, 7.5) (Muyzer et al. 1993) at 60[degrees]C. The DGGE gel was 6% polyacrylamide (dimensions, 20 cm x 17.6 cm and 1.5 mm thick) and contained an increasing denaturant concentration (7 M urea and 40% formamide) of 20-80%. Electrophoresis was carried out at 80 V for 14 h. After electrophoresis, the gel was stained with ethidium bromide (0.5 [micro]g/mL) for 20 min and visualized using a transilluminator. Bands of interest were excised from the gel using a sterile razor blade and placed into a sterile microcentrifuge tube with 0.2 g 1-mm sterile Zicronia/ Silicia beads (Biospec Products) and 500 [micro]L sterile [ddH.sub.2]O. The excised acrylamide/bead mixture was incubated at -80[degrees]C, homogenized, and then bead beaten in a Mini Beadbeater (Biospec Products) at high speed for 3 min. The sample was then kept at 4[degrees]C overnight to allow diffusion of DNA.
A 1.5-[micro]L aliquot of the aqueous portion of the homogenized band sample was then used for reamplification with the primers 960F (no GC-clamp) and 1,200R. This reaction contained 25 [micro]L GoTaq Green Master Mix, 0.5 [micro]M of each forward and reverse primer, and 1.5 [micro]L of the DNA sample. The PCR conditions were the same as outlined for primers 960F GC-clamp and 1,200R above. The PCR product was then purified with a Wizard SV Gel and PCR Clean Up System (Promega). This purified product was then sequenced with the forward primer 960F using a BigDye terminator cycle sequencing kit (v3.1; Applied Biosystems, Foster City, CA). Sequences were then searched against the GenBank database using BLAST to determine their phylogenetic affiliation.
Lesions of all diseased lobsters contained amplifiable eukaryotic communities (Figs. 1 and 2). Two of the 4 healthy lobsters (KMA13 and RI4) and both of the healthy carapace scrapings (lobsters RI2 and RI3) from diseased animals also contained eukaryotic communities (Fig. 2). However, the samples collected from healthy surfaces of the RI lobsters (RI2, RI3, and RI4) produced only a single band, which belonged to H. americanus itself (Fig. 2, band H). The KME samples showed a high level similarity within the group, with at least 4 common bands seen in all samples (Fig. 2). Eukaryotic communities in the ELIS and CLIS lesion samples showed a lesser but still relatively high degree of similarity within their group with at least 3 common bands observed in all. The BB lesion samples also contained very similar eukaryote communities. However, DGGE patterns generated for lobster lesion material collected from different locations are very dissimilar. Overall, the estimated eukaryotic diversity seen as the number of bands in the DGGE gels is highest in the LIS samples, with up to 18 bands observed in ELIS6 (Fig. 1). The diversity is also high in the KME samples, with approximately 14 bands observed in KME6 and KME 1 (Fig. 2). The estimated diversity of the RI samples is far lower, with as few as 2 bands observed in the diseased individual RI3 (Fig. 2).
[FIGURE 1 OMITTED]
[FIGURE 2 OMITTED]
The eukaryotic communities of ESD lesions were highly diverse, with the presence of both micro- and macro-eukaryotes. Of particular interest to this study were bands that were present in a number of lesion samples from a variety of sampling sources, because these may represent organisms with a central role in ESD development and biofouling (Table 2). The marine aquatic nematode Geomonhystera disjuncta was observed in 26 of 28 lesion samples and from all locations sampled. The only 2 lobsters that did not have this nematode in their lesions were RI2 and RI3. Bands at up to 4 different positions in the DGGE gel produced sequences related to this organism (e.g., ELIS 2, band 1 in Fig. 1). Sequences belonging to the barnacle Wanella millepore were found in 10 different lesion samples, but only from 2 locations. One commonly encountered group of microscopic eukaryotes was the stramenopiles. Labyrinthuloides haliotidis and Aplanochytrium sp. were detected in 6 lesion samples from 3 different locations, whereas Atkinsiella dubia was detected in 4 samples from 2 locations, and Paraphysomonas foraminifera was detected in 2 samples from 2 locations. The marine copepod Ecbathyrion prolixicauda was found in 5 separate lesion samples from 2 locations. Sequences related to the Ascomycete fungus Colletotrichum sp., the colonial bryozoan Callopora sp., and an uncultured marine eukaryote were detected in 4 different samples from various locations. Other nematodes such as Enoploides brunettii and Calomicrolaimus parahonestus were detected in a few samples, but E. brunettii appeared to be restricted to the KMA sample group.
There was also a highly diverse group of eukaryotic organisms detected in only a single lesion sample (Table 3). Representatives of the stramenopiles were again encountered, as well as members of Rhizaria and Alveolata. Metazoans were a highly diverse group, including bryozoans, nematodes, copepod crustaceans, Platyhelminthes, barnacles, arthropods, and molluscs. Of the 4 healthy lobsters and 2 healthy carapace samples, only 1 individual produced an observable community (excluding the band for H. americanus), which consisted of predominantly stramenopiles and the known lobster commensal copepod Histriobdella homari (Fig. 2).
The only healthy lobster that harbored an epibiotic eukaryotic community was KMA13. This community consisted of 3 unique members (2 brown algae and a polychaete) and a copepod closely related to E. prolixicauda.
Colonization of aquatic arthropod surfaces by epibionts depends on a number of factors. The first is the frequency of molt. It is observed that frequently molting animals tend to have smaller epibiotic communities or no community at all (Schejter & Spivak 2005). The ability of an animal to groom itself will also be an important factor in whether epibionts are allowed to colonize animal surfaces. This, of course, will depend on the individual animal's behavior. Some crustaceans tend to groom themselves (Bauer 1981, Bauer 2002), whereas others decorate themselves further, promoting the development of epibiotic communities (Parapar et al. 1997, Schejter & Spivak 2005). Finally, physical properties of the carapace, such as hydrophobicity (sometimes measured as "wettability") may also be important. Although Becker et al. (2000) claimed that there is little correlation between the carapace wettability and epibiotic community formation, they found that the crustacean species with the least wettable carapace were colonized by the fewest bacteria and no other colonizers were observed.
A healthy lobster carapace shows little microbial growth on its surface (Hsu & Smolowitz 2003, Chistoserdov et al. 2005). Using DGGE analyses, an epibiotic eukaryotic community was found on the healthy surface of only 1 lobster (KMA13). Notably, all lobsters used in this work were just above legal size or smaller, meaning that they molt approximately once per year for males and nonovigerous females, and once per 2 y for ovigerous females. It likely takes less time than a full molt cycle in these lobsters for a bacterial biofilm accompanied by eukaryotic epibionts to establish a biofouling community. Furthermore, lobsters are known to groom themselves, but not particularly intensively (Bauer 1981). Therefore, it is possible that the lobster shell surface is hydrophobic enough and/or produces antimicrobial agents to keep it clean.
An unexplored aspect of ESD is the existence of eukaryotic organisms in the shell lesions and the role they may play in the severe biofouling of the carapace and ESD lesion development. To date, only histological identification of unidentified protists and nematodes in ESD lesions of the American lobster exists (Smolowitz et al. 2005). This study showed that eukaryotic micro- and macro-organisms make up a large component of the community of ESD lesions and are represented across broad taxa of Metazoa, Fungi, and Protista. Although the diversity of eukaryotic communities cannot be directly determined by counting bands in DGGE, because of the potential for multiple bands produced for a single species (Diez et al. 2001, Zijnge et al. 2006), certain ESD lesion samples had as high as 7 different eukaryotes identified (samples ELIS2, CLIS1, BB9, KMA1, and KMA10). In contrast, samples RI2 and RI3 generated only 3 and 2 visible bands on DGGE gels, respectively (including that of H. americanus; Fig. 2). Grossly, the lesions of these 2 lobsters were highly calcified and melanized, indicating they may not have been undergoing an active case of ESD. It is possible that these lobsters had either recently overcome the disease and were in the process of healing, were in initial stages of lesion development, or had an altogether different form of shell disease.
The high epibiont community variability observed supports the notion proposed by Smolowitz et al. (2005) that most eukaryotic organisms present in the lesions are secondary opportunistic invaders. It is likely that the eukaryotes identified are either attracted to the components of the already degraded cuticular matrix or are possibly grazing on bacteria present in early lesions. Also possible is that the degradation of the carapace by bacteria creates a rough surface with microtopographies that allow for colonization of biofouling organisms and epibionts. The surface microtopography of Cancer pagarus has been shown to prevent effectively the colonization of certain biofouling organisms (Bers & Wahl 2004). The degradation of the carapace surface that occurs with the progression of shell disease would also decrease the wettability of the carapace, allowing for epibionts to colonize crustaceans with shell disease effectively.
There were a total of 13 eukaryotic organisms present in more than 1 lesion, 12 of which were present in more than 1 location. These organisms are of particular interest in the context of the etiology of ESD, because their presence in lesions from various locations may indicate they have evolved a specific niche for ESD lesions. One particular organism of interest is the nematode G. disjuncta. This nematode was found in 26 of 28 lesion communities and was absent only in samples from the lobsters RI2 and RI3, which had minimal eukaryotic communities and, as mentioned earlier, may not have had a typical case of ESD. G. disjuncta is a known bacterivorous nematode (Zolda & Hanel 2007); therefore, it is probable that G. disjuncta is not contributing to the etiology of the disease, but has adapted to colonize the lesions readily and feed on the available bacteria already present. Similarly, members of the Enoploides nematodes are also known predators of other nematodes (Moens et al. 2000). Thus, E. brunettii detected in ESD lesions may be opportunistically feeding on the presence of other nematodes in the lesions, including G. disjuncta. Larval stages of the barnacles W. millipore and Tetraclitella divisa were also commonly encountered in ESD lesions. Only 1 lobster, CLIS1, exhibited visible barnacles in ESD lesions. Barnacles do contribute to biofouling on the carapace of even healthy lobsters, which are limited in their mobility. The marine copepod Ecbathyrion prolixicauda detected in 5 samples is related to the parasitic marine copepod of salmon Lepeophtheirus salmonis and may exist in a similar parasitic existence in ESD lesions (Huys et al. 2006). The Ascomycete fungus Colletotrichium sp., detected in 4 samples, is likely involved in the carapace biofouling as fungi have been observed in a variety of shell diseases of Crustacea (Burns et al. 1979). Of specific interest to lobster shell disease, Fusarium spp. have been implicated as a causative agent of burn spot shell disease (Stewart 1984, Sindermann 1988). Both Fusarium sp. and Colletotrichum are members of the class Sordariomycetes and may be specifically involved in the biofouling observed in various forms of lobster shell disease.
Seven different stramenopile protists were detected in the lesions. The stramenopiles L. haliotidis and Aplanochytrium sp. were observed in 6 different ESD lesion samples from a variety of locations. Smolowitz et al. (2005) were able to observe directly the presence of labyrinthomorphidlike protistans in some ESD lesions and proposed that these organisms colonized the lesion after the degradation by bacteria. In this study, stramenopiles were commonly detected in ESD lesions, but only sporadically, supporting the notion that they are likely opportunistic. However, some representatives of the Aplanochytrium stramenopiles are known parasites of algae and sea grasses, and L. haliotidis is a known pathogen of abalone (Bower 1987, Moto et al. 2003). Although not necessarily involved in the etiology of ESD, it is likely that these stramenopiles are collectively contributing significantly to the biofouling of the lobster carapace.
This study demonstrated that eukaryotic micro- and macroorganisms make up a large component of the lesion community of lobsters with ESD. It is important to understand the role that these eukaryotic organisms play in the development of ESD lesions, specifically those organisms that are found in a high proportion of samples. Most of these eukaryotes were not universally detected and are, therefore, likely secondary opportunistic invaders. However, they may be specifically important in the pathology of ESD in its later stages. The severity of the lesions increases with the duration of the molt cycle and it is possible that some of these opportunistic eukaryotes may contribute to the severe lesions seen later in the molt cycle (Smolowitz et al. 2005). These severe ESD lesions render lobsters virtually unmarketable. Thus, further work should be focused on determining when these eukaryotes invade ESD lesions and how the lobsters acquire them.
We thank Penelope Howell (Connecticut Department of Environmental Protection), Carl LoBue (New York Department of Environmental Conservation), Bruce Estrella (Massachusetts Division of Marine Fisheries), David Kaselauskas (Maine fisherman), Carl Wilson (Maine Department of Marine Resources), and Kathy Castro (University of Rhode Island) for the gift of lobsters for this research. This research is conducted under auspices of the New England Research Initiative: Lobster Shell Disease and it was supported by grant NA06NMF4720100 from the National Marine Fisheries Service of the U.S. Department of Commerce.
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ROBERT A. QUINN, (1) ROXANNA SMOLOWITZ (2) AND ANDREI CHISTOSERDOV (1) *
(1) University of Louisiana-Lafayette, Biology Department, P.O. Box 42451, Lafayette, LA; (2) Aquatic Diagnostic Laboratory, Marine and Natural Science Bldg., Rm 246, Roger Williams University, One Old Ferry Road, Bristol, RI 02809
* Corresponding author. E-mail: ayc6160@Louisiana.edu
TABLE 1. Location and year of collection, and disease severity of lobsters sampled for this study. Year of No. of Site (Abbr.) Sampling Lobsters Coding Fishers Island, Eastern 2001 7 ELIS2-4, ELIS6, Long Island Sound, NY ELIS11, ELIS13-14 (ELIS) Smith Point, Central 2001 1 CLIS1 Long Island Sound, NY (CLIS) Buzzards Bay, MA (BB) 2002 6 BBS-10 Rhode Island, 2008 2008 4 RI1 trawl survey (RI) RI2, RI3 RI4 Kittery, ME (KME) 2004 13 KMA1, KMA2, KMA5, KMA7-8 KMA3-4, KMA6, KMA9-10 KMA11-13 Site (Abbr.) Disease State Fishers Island, Eastern Severe shell lesions Long Island Sound, NY (ELIS) Smith Point, Central Severe shell lesions Long Island Sound, NY (CLIS) Buzzards Bay, MA (BB) Severe shell lesions Rhode Island, 2008 Severe shell lesions on trawl survey (RI) tail, melanized lesions on carapace Minor, melanized lesions around ducts, most likely not ESD; healthy carapace samples Healthy animal Kittery, ME (KME) Severe shell lesions Moderate shell lesions Healthy animals TABLE 2. List of eukaryotes commonly identified in DGGE gels from more than one individual lobster. No. of Common Accession Lobsters Band No. No. Location (n = 28) 1 FJ842398 ELIS, CLIS, BB, 26 KMA, RI 2 FJ842399 BB, KMA 10 3 FJ842400 ELIS, CLIS, BB, 6 4 FJ842401 ELIS, CLIS, RI 6 5 FJ842402 BB, KMA 5 6 FJ842403 ELIS, CLIS 4 7 FJ842404 ELIS, CLIS 4 8 FJ842405 ELIS, BB 4 9 FJ842406 KMA 4 10 FJ842407 BB, RI 3 11 FJ842408 ELIS, BB 3 12 FJ842409 ELIS, CLIS 2 Gel not BB, KMA 5 shown Common Nearest Relative Band No. (Accession No.) 1 Geomonhystera disjuncta (AJ966485.1) 2 Wanella milleporae (AM497906.1) 3 Labyrinthuloides haliotidis (U21338.1) 4 Aplanochytrium sp. S2128 (EU851173.1) 5 Ecbathyrion prolixicauda (EU851173.1) 6 Colletotrichum sp. CPCC 480,565 (EU827606.1) 7 Uncultured marine eukaryote clone SA2_2D2 (EF527606.1) 8 Callopora sp. (FJ009104.1) 9 Enoploides brunettii (AY854193.1) 10 Atkinsiella dubia (AB284575.1) 11 Calomicrolaimus parahonestus (AY854218.1) 12 Paraphysomonas foraminifera (AF174376.1) Gel not Tetraclitella divisa shown (AY520637.1) Identity to Common Closest Band No. Inferred Phylogeny Relative (%) 1 Nematoda, Chromadorea, 99 Monhysterida 2 Athropoda, Maxillipoda, 99 Sessilia 3 Prbotista, stramenopiles, 97 Labyrinthulomycetes 4 Protista, stramenopiles, 98 Thraustochytriaceae 5 Arthropoda, Maxillipoda, 95 Siphonostomatoida 6 Fungi, Ascomycota, 100 Sordariomycetes 7 Rhizaria, Cercozoa, 95 Cercomonadida 8 Bryozoa, Cheilostomatida, 100 Calloporidae 9 Nematoda, Enoplea, Enoplida 92 10 Protista, stramenopiles, 94 Oomycetes, Lagenidiales 11 Nematoda, Chromadorea, 90 Desmodorida 12 Protista, Stramenopiles, 100 Chrysophyceae Gel not Arthropoda, Maxillopoda, shown Sessilia TABLE 3. List of eukaryotes identified in ESD lesions. Figure Band No. Nearest Relative Lobster No. (Accession No.) (Accession No.) ELIS2 1 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (F7842410) Protaspis grandis (DQ303944.1) 3 (FJ842401) Aplanochytrium sp. S2128 (EU851173.1) 4 (FJ842400) Labyrinthuloides haliotidis (U21338.1) 5 (FJ842409) Paraphysomonas foraminifera (AF174376.1) 6 (FJ842403) Colletotrichum sp. CPCC 480,565 (EU827606.1) 7 (FJ842405) Callopora sp. (FJ009104.1) ELIS4 1 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842400) Labyrinthuloides haliotidis (U21338.1) 3 (FJ842402) Colletotrichum sp. CPCC 480,565 (EU827606.1) ELIS6 1 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842408) Calomicrolaimus parahonestus (AY854218.1) 3 (FJ842404) Uncultured eukaryote clone SA2_2D2 (EF527606.1) 4 (FJ842401) Aplanochytrium sp. 52128 (EU851173.1) 5 (FJ842413) Nitzschia closterium (EF553459.1) 6 (FJ842409) Paraphysomonas foraminifera (AF174376.1) CLIS1 1 1 (FJ842417) Parauronema virginianum (AY392128.1) 2 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 3 (FJ842401) Aplanochytrium sp. 52128 (EU851173.1) 4 (FJ842400) Labyrinthuloides haliotidis (U21338.1) 5 (FJ842409) Paraphysomonas foraminifera (AF174376.1) 6 (FJ842402) Colletotrichum sp. CPCC 480,565 (EU827606.1) 7 (FJ842405) Callopora sp. (FJ009104.1) BB8 1 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842407) Atkinsiella dubia (AB284575.1) 3 (FJ842408) Calomicrolaimus parahonestus (AY854218.1) BB9 1 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842407) Atkin.siella dubia (AB284575.1) 3 (FJ842402) Ecbathyrion proliricauda (EU851173.1) 4 (FJ919377) Paratrichodorus minor (AM269897.1) 5 (FJ842415) Vermiliopsis striaticeps (DQ317133.1) 6 (FJ842399) Wanella milleporae (AM497906.1) BB10 1 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842399) Wanella milleporae (AM497906.1) 3 (FJ842416) Bryocamptus pygmaeus (AY627015.1) RIl 2 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842418) Maehrenthalia agilis (AJ312273.1) 3 (FJ919378) Homarus americanus (AY743945.1) RI2 2 1 (FJ842407) Atkinsiella dubia (AB284575.1) 2 (FJ842401) Aplanochytrium sp. S2128 (EU851173.1) KMA1 2 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842418) Maehrenthalia agilis (AJ312273.1) 3 (FJ842419) Balanus balanus (AY520628.1) 4 (FJ919378) Homarus americanus (AY743945.1) 5 (FJ842399) Wanella milleporae (AM497906.1) 6 (FJ919379) Megabalanus stultus (AM497924.2) 7 (FJ919380) Achelia sawayai (DQ389916.1) KMA2 2 1 (FJ919381) Enoplus meridionalis (Y16914.1) 2 (FJ842420) Paramphiascella fulvofasciata (EU380293.1) KMA4 2 1 (FJ842402) Ecbathyrion proli.cicauda (EU85l173.1) 2 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 3 (FJ842399) Wanella milleporae (AM497906.1) 4 (FJ842421) Trichonaya hirsuta (EU289830.1) KMAS 2 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ842406) Enoploides brunettii (AY854193.1) 3 (FJ842422) Lingula anatina (U08331.1) 4 (FJ842414) Navicula tripunctata (AM502028.1) KMA6 2 1 (FJ842399) Wanella milleporae (AM497906.1) KMA8 2 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ919382) Tripyla sp. (EF197733.1) KMA9 2 1 (FJ842398) Geomonhystera disjuncta (AJ966485.1) 2 (FJ919378) Homarus americanus (AY743945.1) KMA10 2 1 (FJ919383) Uncultured eukaryote (DQ104595.1) 2 (FJ842424) Ptychopera westbladi (AY775770.1) 3 (FJ842423) Didymorchis sp. (AY157182.1) 4 (FJ842399) Wanella milleporae (AM497906.1) 5 (FJ842402) Echathyrion prolixicauda (EU851173.1) KMA13 2 1 (FJ842425) Histriobdella homari (AY527053.1) 2 (FJ919378) Homarus americanus (AY743945.1) 3 (FJ919384) Scambicornus sp. (AY627011.1) 4 (FJ842426) Phaeosiphoniella cryophila (AB117926.1) 5 (FJ842427) Pleurocladia lacustris (AY307397.1) Band No. Identity to (Accession No.) Relative (%) Inferred Phylogeny 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (F7842410) 92 Rhizaria, Cercozoa, Thaumatomonadida 3 (FJ842401) 99 Protista, stramenopiles, Thraustochytriaceae 4 (FJ842400) 97 Protista, stramenopiles, Labyrinthulomycetes 5 (FJ842409) 100 Arthropoda, Maxillopoda, Sessilia 6 (FJ842403) 99 Fungi, Ascomycota, Sordariomycetes 7 (FJ842405) 100 Bryozoa, Cheilostomatida, Calloporidae 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ842400) 97 Protista, stramenopiles, Labyrinthulomycetes 3 (FJ842402) 99 Fungi, Ascomycota, Sordariomycetes 1 (FJ842398) 100 Nematoda, Chromadorea, Monhysterida 2 (FJ842408) 90 Nematoda, Chromadorea, Desmodorida 3 (FJ842404) 95 Rhizaria, Cercozoa, Cercomonadida 4 (FJ842401) 99 Protista, stramenopiles, Thraustochytriaceae 5 (FJ842413) 93 Stramenopiles, Bacillariophyta, Bacillariophyceae 6 (FJ842409) 100 Arthropoda, Maxillopoda, Sessilia 1 (FJ842417) 95 Alveolata, Oligohymenophorea, Philasterida 2 (FJ842398) 100 Nematoda, Chromadorea, Monhysterida 3 (FJ842401) 99 Protista, stramenopiles, Thraustochytriaceae 4 (FJ842400) 97 Protista, stramenopiles, Labyrinthulomycetes 5 (FJ842409) 100 Arthropoda, Maxillopoda, Sessilia 6 (FJ842402) 99 Fungi, Ascomycota, Sordariomycetes 7 (FJ842405) 100 Bryozoa, Cheilostomatida, Calloporidae 1 (FJ842398) 92 Metazoa, Platyhelminthes, Turbellaria, Rhabdocoela 2 (FJ842407) 94 Protista, stramenopiles, Oomycetes, Lagenidiales 3 (FJ842408) 90 Nematoda, Chromadorea, Desmodorida 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ842407) 94 Protista, stramenopiles, Oomycetes, Lagenidiales 3 (FJ842402) 93 Arthropoda, Maxillipoda, Siphonostomatoida 4 (FJ919377) 85 Nematoda, Enoplea, Trichodoroidea 5 (FJ842415) 98 Metazoa, Annelida, Polychaeta, Sabellida 6 (FJ842399) 99 Athropoda, Maxillipoda, Sessilia 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ842399) 99 Athropoda, Maxillipoda, Sessilia 3 (FJ842416) 94 Arthropoda, Maxillopoda, Copepoda, Harpacticoida 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ842418) 96 Metazoa, Platyhelminthes, Turbellaria, Rhabdocoela 3 (FJ919378) 100 Arthropoda, Crustacea, Decapoda 1 (FJ842407) 94 Protista, stramenopiles, Oomycetes, Lagenidiales 2 (FJ842401) 99 Protista, stramenopiles, Thraustochytriaceae 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ842418) 96 Metazoa, Platyhelminthes, Turbellaria, Rhahdocoela 3 (FJ842419) 90 Metazoa, Arthropoda, Maxillopoda, Sessilia 4 (FJ919378) 100 Arthropoda, Crustacea, Decapoda 5 (FJ842399) 99 Athropoda, Maxillipoda, Sessilia 6 (FJ919379) 86 Metazoa, Arthropoda, Maxillopoda, Sessilia 7 (FJ919380) 84 Metazoa, Arthropoda, Pycnogonida, Sessilia 1 (FJ919381) 91 Metazoa, Nematoda, Enoploidea 2 (FJ842420) 91 Arthropoda, Maxillipoda, Harpacticoida 1 (FJ842402) 93 Arthropoda, Maxillipoda, Siphonostomatoida 2 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 3 (FJ842399) 99 Athropoda, Maxillipoda, Sessilia 4 (FJ842421) 95 Metazoa, Mollusca, Bivalvia, Mytiloida 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ842406) 92 Metazoa, Nematoda, Enoploidea 3 (FJ842422) 94 Metazoa, Brachiopoda, Lingulata, Lingulida 4 (FJ842414) 92 Stramenopiles, Bacillariophyta, Bacil/ariophyceae 1 (FJ842399) 99 Athropoda, Maxillipoda, Sessilia 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ919382) 91 Nematoda, Enoplea, Trypilidae 1 (FJ842398) 99 Nematoda, Chromadorea, Monhysterida 2 (FJ919378) 100 Arthropoda, Crustacea, Decapoda 1 (FJ919383) 89 Eukarya 2 (FJ842424) 93 Metazoa, Platyhelminthes, Turbellaria, Rhabdocoela 3 (FJ842423) 92 Metazoa, Platyhelminthes, Turbellaria, Didymorchiidae 4 (FJ842399) 99 Athropoda, Maxillipoda, Sessilia 5 (FJ842402) 93 Arthropoda, Maxillipoda, Siphonostomatoida 1 (FJ842425) 96 Metazoa, Annelida, Polychaeta, Histriobdellidae 2 (FJ919378) 100 Arthropoda, Crustacea, Decapoda 3 (FJ919384) 95 Metazoa, Arthropoda, Copepoda, Synapticolidae 4 (FJ842426) 98 Stramenopiles, Phaeophyceae Tilopteridales 5 (FJ842427) 99 Stramenopiles, Phaeophyceae, Ectocarpales
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|Author:||Quinn, Robert A.; Smolowitz, Roxanna; Chistoserdov, Andrei|
|Publication:||Journal of Shellfish Research|
|Date:||Dec 1, 2009|
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