BIOL 4900 Honours Thesis Towards an understanding of shallow water marine hydroids on Cape Breton coastlines
Matthew S. A. Penney Cape Breton University Sydney, Nova Scotia April, 2017
Supervisor: Dr. Timothy A. Rawlings
Towards an understanding of shallow water marine hydroids on Cape Breton coastlines
Matthew Penney
April 2017
A thesis submitted in partial fulfillment of the requirements for BIOL4900 in the Department of Biology, Cape Breton University, Sydney, Nova Scotia, Canada.
The undersigned grants permission for the Cape Breton University Library to store and provide access to this thesis and for the Chair of the Department of Biology, upon request, to allow extensive copying of the material contained in this thesis, in whole or in part, for the purposes of teaching and research by faculty and students of Cape Breton University.
Requests for permission to copy or make other use of the material contained in this thesis, in whole or in part, should be addressed to:
Chair, Department of Biology Cape Breton University P.O. Box 5300 Sydney, NS B1P 6L2 Canada
Student Signature: ______Date: ______Table Of Contents
Abstract ...... 1 Introduction ...... 3
Materials & Methods ...... 8 Sample Collection ...... 8 Isolation, Processing, & Fixing...... 9 Morphological Analysis ...... 9 Molecular Analysis ...... 10 Phylogenetic Analysis ...... 14
Results ...... 16 Sample Collection ...... 16 16S Amplification & Sequencing ...... 17 Identifications ...... 18 Species Profiles ...... 18
Discussion...... 26 Comparison To Regional Surveys ...... 26 Genetic Connectivity ...... 26 Clarification of Species-Level Identities ...... 28 Monophyly of Bougainvillia ...... 30 Study Limitations ...... 30
Future Studies ...... 32 Conclusion ...... 33 Figures ...... 34 Tables ...... 50 Acknowledgments ...... 55 References ...... 56
Abstract Distributions of North Atlantic marine animals can be expected to shift in response to climate change. Tracking distributions temporally requires an understanding of what species are currently present. Hydroids (Cnidaria, Hydrozoa, Hydroidolina) are conspicuous taxa in the fouling community of shallow water marine environments in Cape Breton, yet surveys of them specific to Cape Breton are sparse and outdated. Consequently, the identity and diversity of local species needs to be clarified. For this study, hydroids were sampled from shallow water environments around Cape Breton and preserved for morphological and molecular analysis. Key morphological features including tentacle anatomy and arrangement, gonophore anatomy, colony structure, and the presence of hydrothecae were examined for family and genus level designations. Sequences were generated for the 16S rRNA barcoding gene and used in BLAST searches to find strong genetic matches on GenBank. Molecular phylogenetic analyses were then performed to refine identifications and determine genetic relationships to other populations. The use of both molecular and morphological analyses has enabled the identification of thirteen taxa to at least genus level and eleven to species level. In total, 14 out of 18 sequences strongly matched GenBank sequences, with similarities ranging from 98%-100%. Two taxa,
Podocorynoides minima and Pachycordyle michaeli, do not appear to have been recorded previously in the region. Phylogenetic analysis of genetic connectivity indicated strong genetic relationships with conspecifics in the United States and more distant relationships to conspecifics in Europe. Additionally, it revealed potential cryptic species in Obelia dichotoma. Two genera,
Hydractinia and Sarsia, have yet to be identified to species level. Having reliable morphological and molecular databases to identify species, especially for Hydrozoa, is important for monitoring changing distributions over time. This study represents the first stage in the development of a useful dataset for identifying local hydroids using 16S sequence data and accompanying
1 | P a g e morphological descriptions.
2 | P a g e
Introduction
Hydrozoa is a class within the phylum Cnidaria (Collins, 2002; Daly et al., 2007).
Hydrozoans are characterized by septae-lacking polyps which have a chitinous perisarc. In species with a medusoid life stage, reproduction occurs via budding and medusae have a velum and dual nerve rings (Bouillon et al., 2006; Daly et al., 2007). Symmetry of hydrozoans is tetramerous, polymerous, or biradial (Bouillon and Boero, 2000). Members are typically carnivorous, though some feed on phytoplankton and bacteria, and can be heavy consumers of larvae and other marine organisms (Bouillon et al., 2006).
The Subclass Hydroidolina is one of two major clades within Hydrozoa, along with
Trachylina (Collins, 2002). It includes three major orders: Anthoathecata, Leptothecata, and
Siphonophora (Collins, 2000). According to Collins (2000; 2002), Hydroidolina traditionally includes five main groups: Capitata, e.g. Sarsia tubulosa (Schuchert 2010); Filifera, e.g.
Bougainvillia muscus (Schuchert, 2007); Hydridae, e.g. the genus Hydra (Schuchert, 2010);
Leptomedusa, e.g. the genus Obelia (Cornelius, 1995a); and Siphonophora, e.g. Physalia physalis (Ulster Museum, 1997). However, the monophyly of Filifera has been disputed
(Cartwright et al., 2008).
There are solitary and colonial species of hydroids. Colonial varieties can exhibit specialized polyps, e.g. gastrozooids for feeding, gonozooids for reproduction, and dactylozooids for defense, all connected via stolons with a continuous gastrovascular canal; the stolon is usually covered by a perisarc. Many hydroids are typically benthic, a notable exception being the clade Siphonophora (Bouillon et al., 2006). Reproduction is indirect, with an asexual polyp stage budding off sexual adult medusae. Hydroids are largely polymorphic, the only exception being
3 | P a g e the clade Hydridae (Collins, 2002), and also have a diverse cnidome, with 21 forms of cnidocysts in total (Bouillon and Boero, 2000).
Hydroid species can inhabit a wide range of salinities, as evidenced by the euryhaline
Cordylophora caspia (Folino-Rorem and Indelicato, 2005) and marine species such as
Plumularia setacea (Schuchert, 2014). They can also be found ranging from shallow to deep waters, with one record of hydroids fouling deep-sea instruments located at a 3690m depth
(Blanco et al., 2013). Benthic hydroids have a high capacity to foul many types of surfaces, which allows them to disperse over extensive distances on drift algae/artificial structures and ship hulls; they can also disperse via ballast water or their pelagic forms (Blanco et al., 2013;
Calder et al., 2014). In general, they hold a preference for hard surfaces over soft surfaces and tend to avoid sand or silt bottoms (Calder, 1991).
Hydromedusae are considered the most diverse and widespread group of gelatinous marine zooplankton (Boero et al., 2008; Miglietta et al., 2008, cited by Laakmann and Holst,
2013). Additionally, the aforementioned Hydra, which is a model organism, has been used in several research fields and even has a fully sequenced nuclear genome (Nawrocki et al., 2013).
Despite this, hydroids have remained largely understudied and many basic questions about hydroid systematics, diversity, distribution, and genetic connectivity remain unanswered (Collins
2002; Collins et al. 2006, cited by Daly et al., 2007). Hydroids may be overlooked in research efforts because some, e.g. benthic hydroids, are small in size and some species look the same to the untrained eye (Gili and Hughes, 1995). Some species may also be lacking important characteristics for identification, such as gonophores, at the time of collection. Collection is also made difficult by some specimens being firmly attached to the benthos.
4 | P a g e
A focus within hydroid research involves regional biodiversity studies around the world, including New Zealand (Schuchert, 1996), Bransfield Strait in Antarctica (Cantero and Ramil,
2006), the Arctic Ocean (Ronowicz et al., 2015), three benthic marine habitats in Taiwan (Tseng et al., 2014), and Hawaii (Calder, 2010). Within the Atlantic Ocean, surveys have included comparisons across biogeographic regions (Medel and Lopez-Gonzales, 1998) as well as more focused studies of the Mid-Atlantic Ridge (Calder and Vervoort, 1998), Cape Cod (Calder, 1975) and Europe (Schuchert, 2008, 2010). In the western Atlantic, hydroids have been studied in Cape
Cod and Georges Bank, and one survey has been completed recently in the Bay of Fundy
(Calder, 1975; Calder, in press; Bollens et al., 2001; Concelman et al. 2001). North of this region, however, there is very limited information on hydroid diversity beyond a list of marine invertebrate species compiled for the Gulf of St. Lawrence (Brunel et al., 1998). For Cape Breton in particular, specific work done on hydroids is very limited and outdated (Whiteaves, 1874;
Fraser, 1926), lacking genetic characterization of species collected and using outdated taxonomic groupings. Thus, we currently lack a modern understanding of hydroid species, diversity, and genetics in the marine habitats of Cape Breton.
For this study, I will examine hydroids of near-shore shallow-water marine habitats, both benthic and pelagic, from various environments around Cape Breton. My primary focus will be to assess the presence, diversity, and genetic characteristics of hydroid species in select local habitats. I will also attempt to use these data in distance-based phylogenetic analyses to discern relationships between local hydroids and other N. Atlantic populations, focusing primarily on relations to European varieties. The combined use of morphological and molecular techniques provides the potential to explore these phenomena (Dawson and Jacobs, 2001; Lindeque et al.,
2013; Rodriquez et al., 2014; Schuchert, 2014).
5 | P a g e
With an ever-increasing database of DNA barcodes and sequences for hydroids in
GenBank, the study will include the use of DNA sequences as molecular taxonomic units to compile molecular data, with a focus on the popular 16S mitochondrial rRNA gene region.
Morphological characteristics will also be used in collaboration with molecular techniques to refine or confirm assessments using 16S sequence data by focusing on several key morphological features of hydroids in the North Atantic and Europe. Sampling locations will be opportunistic and determined based on access to locations expected to have an abundance of hydroids, including but not limited to Lingan Bay Zostera marina beds and the Bras d’Or Lakes.
Molecular techniques are useful to overcome shortfalls faced in morphological taxonomy, examples being the limiting number of phylogenetically-informative characters in simple-bodied organisms, high phenotypic plasticity, difficulties matching morphologically disparate life stages, and challenges involving damaged specimens (Ortman et al., 2010; Hardy et al, 2011; Laakmann and Holst, 2013). Fast evolving molecular markers can also be useful for determining phylogenetic relatedness, linking the aforementioned life stages, and assigning species identities (Hardy et al., 2011). However, molecular tools can have their own limitations.
In the case of the DNA barcoding gene Cytochrome oxidase subunit I (COI), slow rates of mitochondrial evolution in cnidarians may result in low levels of inter-species genetic differentiation, making it ineffective for species-level identification (Zheng et al., 2014 and references therein). Combining the two methods provides two semi-independent data sets to compare for purposes of consensus.
Since hydroids are also significant fouling organisms in aquaculture and fisheries, particularly for commercially farmed molluscs, a good understanding of the species present in
6 | P a g e
Cape Breton should be beneficial for local fisheries, as well. My results also benefit biologists interested in hydroid systematics and evolution, since they can provide descriptions and sequences of local hydroid populations, help demonstrate patterns of relatedness among populations of species, and possibly provide hints of past population expansions and cryptic species. My specimens and collection records may also be useful for the Atlantic Reference
Centre and the Nova Scotia Museum of Natural History.
7 | P a g e
Materials & Methods
Sample Collection
Collection was performed in the Fall from September to November of 2016; sampling locations were determined by opportunity and accessibility (Fig. 1). Hydroids have been recorded in marine and estuarine habitats (Folino-Rorem and Indelicato, 2005; Schuchert, 2014), so sites ranging in salinities were targeted for collection. Regions with hard substrates, e.g. rocky intertidal, were chosen because hydroids are known to prefer hard substrates (Calder, 1991).
Shells, grasses, rocks, and other substrates were examined when present. Soft-sediment environments with plants (e.g. seagrass) or macroalgae were also examined because hydroids can grow on marine macroalgae and seagrass. (Cornelius, 1995b); floating wracks of algae and seagrass were also examined for hydroids. Sampling locations were mainly intertidal, although some subtidal habitats were also sampled (Table 1).
As specimens are often difficult to identify in the field (Gili and Hughes, 1995), volunteer collectors were instructed to examine sites for “white fuzzy growth” on hard surfaces. Polypoid collection in intertidal zones was performed by hand, forceps, or using dipnets to scrape substrates. Samples were obtained from subtidal habitats using SCUBA by Dr. Hatcher and colleagues. Zostera marina and Fucus blades, mollusc shells, and crab parts were also collected when polyps were observed on them. Collected samples, and substrates when possible, were placed in individual bags with site water and returned to the lab immediately.
To collect medusae, plankton tows were performed using a telescopic pole with a 254µm mesh net. The size was used to minimize collection of non-hydroid plankton, as suggested by
Cornelius (1995a). Plankton tows were done slowly for 10 minutes at 1m depth walking back
8 | P a g e and forth along piers or docks. Time of day for plankton tows varied. The net was brought slowly up out of the water and the collecting end was drained into plastic containers filled with site water. The net was then rinsed to retrieve any residual sample.
Isolation, Processing, & Fixing
Field collected samples were transported to the lab, preferably in a whirl pack filled with site water for polyps or a plastic container for plankton tows, and then examined under a dissecting microscope (Wild Heerbrugg 113079) for polyps or medusae. Specimens identified as hydroid polyps or medusae were then isolated into individual plastic Petri dishes. The Petri dishes were then transported to a dissecting scope with a camera attachment (Olympus SZH-
ILLD) and photographed using ImagePro Express software to generate a photographic record.
Once photographed, specimens were returned to the lab and fixed in preservative. Specimens were preserved in glass vials or jars of 95% ethanol for molecular analysis and 10% formalin for morphological analysis. Where possible, specimens were preserved in both fixatives. Medusoid specimens preserved in ethanol were pooled based on morphology. Specimens were not anaesthetized prior to formalin preservation.
Morphological Analysis
Morphological descriptions and identifications mainly followed diagnostics outlined in
The European Athecate Hydroids and their Medusae (Schuchert 2007, 2008, 2010) and with
Parts 1 and 2 of The European Thecate Hydroids and their Medusae (Cornelius, 1995a,
Cornelius, 1995b). Replicates were also shipped to Dr. Calder for species confirmation and deposition in the Cnidarian collections at the Royal Ontario Museum.
Major features were identified and used to categorize specimens down to the family and genus taxon levels. Features of polypoid forms included the presence or absence of hydranth
9 | P a g e covering (thecate/athecate), the presence of tentacle whorls, number of whorls, gonophore structure, morphology of the hydrotheca and gonotheca when present, presence and morphology of hydrothecal teeth, tentacle morphology, hydrocaulus branching pattern, and base morphology
(stolonal/encrusting). All hydroids encountered in this study were colonial. Examples of important features are presented in Figure 2.
Features of medusoid forms noted included medusa shape (saucer, bell-shaped, cubic), number of tentacles around bell margin, tentacle morphology, arrangement of ocelli on basal tentacle bulbs (if visible), presence of apical process, tentacle branching, morphology of manubrium, presence of manubrial bulbs or flaps, and location of gonads when present. All types of medusae encountered in this study had four radial canals (Figure 3).
Molecular Analysis
PCR Primer Selection
PCR primers for the 16S mitochondrial rRNA gene were chosen from selected publications in which this gene region was successfully amplified in Hydrozoans. The primers chosen were Cnidarian specific and amplified a 600bp fragment. CNID-16SF: 5'-TCG ACT GTT
TAC CAA AAA CAT AGC-3' and CNID-16SR: 5'-ACG GAA TGA ACT CAA ATC ATG
TAA G-3' (Cunningham and Buss, 1993).
Prior to DNA extraction, a collection record was completed for each specimen to indicate tentative ID, life stage, collection site and date, and the specimen code. Individual specimens were assigned DNA codes to track taxa through molecular processing. The DNA code was
10 | P a g e formatted as MMDDYY-#, with MMDDYY determined by the extraction date and # determined by order of processing. Up to six specimens were processed for DNA extraction at one time.
Tissue samples were isolated from EtOH-preserved specimens in clean plastic Petri dishes filled with 95% ethanol. Isolated tissue was then transferred to a separate labelled Petri dish filled with 95% ethanol using clean tools or micropipettors, depending on life stage.
Separate procedures were developed for polypoid and medusoid specimens from this point. For polypoid samples, 1.5ml blue eppendorf mortar tubes were filled with 180µl ATL buffer prior to tissue addition. Tissues were damp-dried with fresh sterile Kimwipes beforehand to remove residual ethanol anded and then ground using a blue sterile plastic pestle. For medusoid samples, medusae were pipetted directly into 1.5ml blue eppendorf tubes and left open upright in a clean tube rack covered by a fresh Kimwipe to allow evaporation of ethanol. 180µl of ATL buffer was pipetted into tubes once samples were sufficiently dry. Medusoid samples were not ground.
DNA extraction was performed using Qiagen DNeasy Blood and Tissue kits (QIAGEN
Inc., Mississauga, ON, #62804) following manufacturer-recommended protocols. The only modification was the elution volume, which was reduced to 50µl of elution buffer. This kit has successfully extracted DNA from Cnidarians in other studies (Schuchert, 2005; Cartwright et al.,
2008; Martinez et al., 2010).
Once extracted, 5µl of each 45µl DNA solution was run on a 0.8% TBE Agarose electrophoresis gel (40ml 1x TBE buffer, 0.32g agar powder, 4µl Sybersafe.) in 1x TBE buffer for evidence of extraction of total DNA. 1.5µl of 6xLB loading buffer was added to 5µl samples before loading. A wide-range molecular weight standard (GelPilot Wide Range Ladder #239125)
11 | P a g e was used for these gels. Gels were examined and photographed under UV light using
AlphaImager once electrophoresis was completed (Fig. 4A).
DNA solutions were then prepared for PCR using either the Promega GoTaq Hot Start or
Phusion High Affinity DNA Polymerase PCR Master Mixes.
Individual GoTaq reactions were performed in 25μl volumes and contained 12.5µl of
GoTaq with added buffer, 0.5µl of 10µM of both forward and reverse PCR primers, and 1-2µl of
DNA. Solution was then topped up to 25µl with sdH20. Reactions were performed in an MJ Mini
Personal ThermoCycler (Bio-Rad) using the following protocols from Cantero et al. (2010):
Initial denaturation at 94oC for 3 min, followed by 38 cycles of denaturation (92oC, 30s), annealing (52oC, 30s), and synthesis (72oC, 1 min) ending with a final elongation at 72oC for 10 min. Reactions were then held at 4oC until removed.
Phusion DNA Polymerase PCR Mix, with a high-affinity enzyme and a proofreading enzyme, was used for samples which failed to amplify or were otherwise problematic with
GoTaq PCR Mix. Individual reactions were performed in 25µl volumes and contained 12.5µl
Phusion DNA Polmerase mix, 1.25µl of 10µM of each PCR primer, 9µl dH20, and 1µl of DNA solution. Reactions were performed in a ThermoCycler. Protocols were as follows: Initial denaturation at 98oC for 30s, followed by 34 cycles of denaturation (98oC, 10s), annealing
(62.1oC, 30s), and synthesis (72oC, 30s) ending with a final elongation at 72oC for 10 min.
Reactions were then held at 4oC until removed.
12 | P a g e
PCR Gel, Cleanup, & Sequencing
After PCR amplification, a small amount of product (5μl of 25μl PCR rxn) was run on a
1% agarose electrophoresis gel (40ml 1x TBE Buffer, 0.40g agar, 4µl Sybersafe) in 1X TBE buffer to check for PCR products of the expected size. A 100bp ladder (GelPilot 100bp Plus,
#239045) was used as a molecular weight standard in these gels. Gels were examined and photographed under UV light using AlphaImager (Fig. 4B). When amplifications were successful, PCR solutions were purified using the Qiaquick PCR Purification Kit (#28104). Total
DNA concentration was measured using Nanodrop ND-1000 from Thermo Scientific, measured in ng/µl.
For sequencing, 7.7µl sequencing reactions were prepared in individual 200µl PCR tubes arranged in tube strips. Reaction mixtures consisted of 0.7µl of 7.14µM sequencing primer solution and 7ul of a DNA solution diluted to contain between 40-50ng of DNA. Reaction mixtures were then shipped to the Center for Applied Genomics and the Toronto Sick Kids hospital in Ontario to undergo Sanger sequencing.
Since PCR primers did not produce usable sequence, a suite of sequencing primers for the 16S rRNA gene were developed in Geneious 9.1.5 by looking for highly conserved regions of sequence in Hydrozoan taxa relative to outgroup taxa from other Cnidarians (Scyphozoa,
Cubozoa, Staurozoa, Anthozoa) and non-Cnidarian taxa. The complete list of sequencing primers is provided in Table 2.
13 | P a g e
Sequence Assembly & Tentative Identification
DNA sequence data produced by the Center for Applied Genomics were edited, assembled, and analyzed in Geneious 9.1.5. For sequence assembly, unreadable regions where it was impossible to determine individual nucleotides were removed from both ends of a sequence of chromatograms. Any uncertain base reads were either estimated based on the strongest signal in the position or left undefined if chromatogram peaks were ambiguous. Complementary forward and reverse sequences of individual specimens were then used to generate consensus sequences using De Novo Assembly. Additional uncertain bases were clarified at this stage based on consensus among aligned sequences. Primers were then removed from the sequence by searching for and deleting the sequences corresponding to the forward and reverse PCR primers.
The final sequence with primers removed was determined to be the final consensus sequence.
Final consensus sequences were run through a BLAST (Basic Linear Alignment Search
Tool) search on GenBank to determine tentative specimen identifications. Matches with a 98% or higher % sequence similarity were assumed to be conspecific.
Phylogenetic Analysis
Clarification of Species Identifications
Tentative identifications from BLAST searches were used to search for additional sequences of conspecifics in GenBank. Sequences of relatively similar nucleotide lengths to my consensus sequences were preferred where possible. Sequences of taxa closely related to the tentative identification (e.g. taxa in the same genus) were also included to test robustness of the
14 | P a g e tentative identification. Outgroup taxa, closely related to the tentative identification, were also included where known.
Consensus sequences were assembled into a multiple alignment with GenBank sequences of conspecifics and heterospecifics along with an outgroup (IUB Cost Matrix, Gap open cost=15,
Gap extend cost=6.66). Phylogenetic trees were constructed using a Jukes-Cantor Neighbor-
Joining distance analysis in Geneious.
Resulting phylogenetic trees were examined for relationships between my consensus sequence(s) and other taxa included in the tree. A sister grouping between my sequence and another sequence matching the tentative identification was interpreted as additional confirmation of the tentative identification.
Analysis of Genetic Connectivity
Similar to the previous analysis, a search was performed for each taxa on GenBank to determine the number of available conspecific sequences for comparison. Sequences of relatively similar nucleotide lengths were preferred where possible. Taxa selected for analysis were required to have at least their country of origin recorded. Where possible, sequences of specimens sampled from the North American and European Atlantic coastlines were preferred.
For taxa with a sufficient number of conspecific sequences, a multiple alignment and phylogenetic tree were generated using the same parameters as described previously.
Resulting phylogenetic trees were analyzed by examining relative positions of samples and geographic locations. Interpretations of these results were based on sister-groupings, branch distances, and clade formations by continent. Specifically, though not exclusively, the interest
15 | P a g e was the relationship between hydroids from Cape Breton to other geological locations in North
America, such as Europe. If, for example, there was minimal branch distance between geologically disparate samples, it was interpreted as potential indication of recent human- mediated dispersal.
Results
Sample Collection
From September to November of 2016, hydroids were successfully sampled from 17 different locations around Cape Breton Island (Table 2). Hydroids were found in rocky intertidal regions, along several docks, in soft-sediment intertidal regions, and in subtidal habitats. Habitat salinity ranged from marine to estuarine. Subtidal sampling was done to a maximum depth of approximately 7.6m. On 22 Feb 2017, one additional sample was collected by Dr. Rawlings on the exposed shore of Dominion Beach on a blade of kelp.
Colonies were also found inhabiting floating Zostera marina blades in East Bay.
Instructing volunteer collectors to examine for “white fuzzy growth” directed them mainly to members of the Campanulariidae (Penney, pers. obs.). A specimen of Hydractinia was found growing on the claws of a Rock Crab, Cancer irroratus, in the sheltered region of Port Morien.
Members of the family Campanulariidae were collected on shells of Mytilus mussels in Lingan
Bay, near the eelgrass beds, in addition to Zostera marina blades.
Plankton tows were performed at the Old Sydney Yacht Club, the North Sydney Yacht
Club, Main á Dieu, the Lingan Bay eelgrass beds and Fisherman's Wharf, the South Bar
Government Dock, and Fourchu. Collectively, medusoid forms of genera Obelia, Bougainvillia,
Lizzia, and Podocorynoides were sampled in the plankton. An actinula larva was also collected
16 | P a g e in the plankton tow from Fourchu, but it is unknown if this larva was from a hydroid (Penney, pers. obs.).
16S Amplification & Sequencing
Partial sequences of the 16S rRNA gene were successfully amplified for 21 specimens using either GoTaq Hot Start or Phusion High Fidelity DNA Polymerase PCR kits. GoTaq was generally very successful in amplifying partial fragments of the 16S rRNA gene from DNA samples. For five samples, however, the use of GoTaq PCR Mix failed to generate a significant
DNA yield even when the amount of DNA in the PCR reaction was increased to 2µl. The average DNA concentration for all five samples was 5.45ng/µl after GoTaq amplification. The use of Phusion High Fidelity DNA Polymerase Mix significantly increased the DNA yield for all five samples (Fig. 5), increasing the average DNA concentration per sample to 11.36ng/µl.
The two PCR primers, CNID 16SF and 16SR, generated unusable sequences when used for sequencing. Individual nucleotide signals were difficult to distinguish and multiple signals were present for the same nucleotide position. Consequently, new sequencing primers (HYD-
16SF, HYD-16SR-LONG, HYD-16SR-SHORT-ALL, and HYD-16SR-SHORT-TG) were developed using known hydrozoan sequences to obtain usable 16S rRNA sequence (Table 2).
The results of the sequencing reactions using these primers are displayed in Table 3. The forward primer successfully generated good quality sequence data from all species tested, but the results of the reverse primers varied with species.
17 | P a g e
Identifications
Initial Identifications
Many of the BLAST searches successfully identified species identity of hydroid specimens, with matches ranging in similarity from 98%-100% (Table 4). However, for two taxa the highest scoring GenBank sequences were only 95% similar, suggesting there were no comparable species-level matches in GenBank. These results were further analyzed using morphological examination of preserved materials and photographic records and the results of molecular phylogenetic analyses. Additional confirmation for many specimens was provided by
Dr. Calder. Some identifications could not be determined to species level.
Final Identifications
The final list of identifications for this study included 13 taxa with 11 identified to the species level. This list represented a collection of 11 species, 12 genera, and 7 families supported by morphological and/or molecular evidence (Table 5). Specimens of Podocorynoides minima also fell below the 98% threshold, but these identifications were further supported by morphological analysis and confirmed by Dr. Calder (Calder, pers. comm.). A photographic representation of the life stages identified for each individual species is presented in Figure 6.
Species Profiles
The following species-level profiles represent summaries of information collected on five of the 13 species identified using morphological and molecular analyses. The five species were
18 | P a g e chosen based on availability of sequences for comparison and appropriateness for comparisons to other taxa.
Gonothyraea loveni Order: Leptothecata Family: Campanulariidae
Morphological Description: Thecate hydroid. Erect colonies. Colony length varies, generally around approx. 18mm long. Vestigial medusa buds hanging from gonophores. Hydrotheca with toothed rim; indents in teeth. Internodal segments not curved. Gonotheca flute-shaped as opposed to vase-shaped (see Obelia dichotoma). Stolonal. Hydrothecal pedicels annulated.
Habitat and Distribution: Polypoid colonies collected from estuarine habitat of East Bay colonizing detatched blades of Zostera marina on 07 Sept 2016.
Nucleotide Sequence Data: 16S sequence, 587 nucleotides long, from sample collected on 07
Sept 2016 in East Bay.
Phylogenetic Analysis
Taxa Clarification
G. loveni is currently the only accepted species in the genus Gonothyraea (Cornelius,
1992b). In accordance with this, it is the only species of the genus Gonothyraea with representative 16S sequences on GenBank. My sequence showed a 100% match to one sequence
(KX665257) sequence over 587 base pairs.
Genetic Connectivity
19 | P a g e
In a distance-based phylogenetic analysis using Obelia bidentata (AY789815) and
Campanularia volubilis (AY789804), as outgroups, sequence from this study formed a clade with conspecific 16S sequences from Plymouth, USA (KX665257) and Dennis, USA
(AY789826), placing it with other North American sequences (Fig. 7).
Clava multicornis Order: Anthoathecata Family: Hydractiniidae
Morphological Description: Athecate hydroid, stolonal. Gonophores not present. Hydranths in two samples, one collected from tidepool and another collected from a wave-exposed rock.
Significant difference in sizes. Shorter polyps approx. 2-3mm in length from base to hypostome, larger polyps approx. 4-5mm. Hydranths approx. 0.5mm wide at widest point (slightly contracted in this measurement). Specimens greatly contractile. Hypostome rounded, clearer than body.
Hydranth coloured orange. Tentacles do not appear to be arranged in whorls. Taper off at ends.
Restricted to mid-upper region of hydranth, not extending into hypostome.
Nucleotide Sequence Data: 16S sequence, 590 nucleotides in length, from specimen collected on
11 Nov 2016 from Fourchu.
Habitat and Distribution: Polypoid colonies collected from Fourchu rocky intertidal. Scraped off bedrock. Tide pool: temperature 11.2 oC, salinity 29.4 ppt.
Phylogenetic Analysis
Taxon Clarification
20 | P a g e
C. multicornis is the only species of the genus Clava with representative 16S sequences on GenBank. My sample showed very strong genetic similarity to other conspecifics on
Genbank, matching 100% to one sequence (FJ214440) over 99% of query coverage.
Genetic Connectivity
The resulting distance analysis (Fig. 8) indicated that my specimen was more closely related to conspecifics sampled on the American coastline than those in European waters.
Interestingly, there appears to be a potential translocation event of an American C. multicornis to the European shoreline. Subsequent analyses using different outgroups, including Podocoryna carnea (FJ14469), Hydractinia echinata (FJ214556), Clavactinia gallensis (FJ214377), and
Janaria mirabilis (FJ214555) did not yield identical trees. Sequence FJ214444 appeared as a sister group to the clade containing the local C. multicornis sequence with J. mirabilis as the outgroup. However, the clade containing FJ214440. FJ214441, and FJ214442 with the sequence from this study was highly conserved in all trees tested including, yet not visible in, Figure 8.
The three sequences forming a clade with the study sequence all come from Woods Hole, USA.
Obelia dichotoma Order: Leptothecata Family: Campanulariidae
Morphological Description: Thecate hydroid with a cup-shaped hydrotheca and vase-shaped gonotheca with end narrowed in (Fig. 2D). Colonies from Lingan vary greatly in length. One sample had colonies ranged between 2mm to approx. 5mm in size. Gonophore present with medusal buds inside. Slight curvature to hydrocaulus. Hydrothecal rim smooth. Hydranth with
21 | P a g e rounded hypostome. Filiform tentacles in one whorl. Colonies stolonal and demonstrate slight alternating curving in some hydrocauli, while others appear more straight.
Habitat & Distribution: Polypoid colonies collected in the Lingan Bay eelgrass beds on 11 Sept
2016 and 21 Sept 2016 from the Barra Strait, placing them in marine and estuarine environments.
Nucleotide Sequence Data: 16S sequences generated from the Barra Strait specimen collected on
21 Sept 2016 (1 sequence) and the Lingan Bay eelgrass bed specimens collected on 11 Sept 2016
(2 sequences). All 589 nucleotides in length.
Phylogenetic Analysis
Taxon Clarification
Although the three sequences did not match within the same clade, all three grouped in clades with other sequences of O. dichotoma (Fig. 9).
Genetic Connectivity
The three 16S sequences extracted in this study did not form a single clade when analyzed with other sequences of O. dichotoma (Fig. 10). This result was consistent using both
Campanularia hincksii (KP776784) and Clytia gracilis (KX665352) as outgroups. Two of the O. dichotoma sequences from this study paired most closely with a sequence from Argentina
(KX665331) while the other paired most closely with a sequence from Bourne, USA
(KX665316). There was, oddly enough, an additional clade of O. dichotoma separated from the
22 | P a g e clades containing study sequences by two monophyletic clades, one of O. genticulata and one of
O. bidentate (Fig. 9).
Dynamena pumila Order: Leptothecata Family: Sertulariidae
Morphological Description: Thecate hydroid with erect colonies, specimen without gonophores,
Tubular hydrotheca with single point on rim. Colonies growing to about 10mm in length, but usually shorter. Hydrothecae branch out in opposite pairings along hydrocaulus. Perisarc brown, hydranth off-white when present (Fig. 2E). Opposite pairs of hydrotheca forming an upside- down triangular shape. Width of pairing about 1mm from rim to rim. Hydrotheca with two points on rim. Colony stolonal with individual non-branching stems. Stiff stalk, contractile hydranth.
Habitat & Distribution: Polypoid colonies collected from Main á Dieu low rocky intertidal zone on 18 Sept 2016 growing on blades of Fucus serratus. Growing in heavily wave-exposed site.
Nucleotide Sequence Data: 16S sequence, 575 nulceotides in length, from specimen collected on
18 Sept 2016 from Main á Dieu.
Phylogenetic Analysis
Taxon Clarification
Only one sequence of D. pumila (AY787902) was available on GenBank. When placed into a phylogenetic tree with the sequence from this study and seven other sequences including six other Dynamena species and an outgroup, sequence AY787902 formed a sister group with
23 | P a g e the sequence from this study (Fig. 11). Sequences were 99% similar over 99% of their sequences. Outgroups Sertularia moluccana (FJ550494) and Amphisbetia operculata (FJ550489) were used. Resulting sister group pairing was consistent between both trees.
Genetic Connectivity
Due to insufficient sequence data from GenBank, genetic connectivity analysis could not be performed for this specimen.
Pachycordyle michaeli Order: Anthoathecata Family: Bougainvillidae
Morphological Description: Athecate hydroid with long filiform tentacles. Unsure of number of tentacle whorls, but tentacles seem to be restricted to region below hypostome. Hydranth on single stalk rising from stolon (Fig. 2A). Approx. 4mm long from stolon base to tip of hypostome. Male has sporosacs present along hydrocaulus. No medusa buds. Bend in stalk below hydranth at least once. Hydranth colored red with white hypostome. Perisarc present, but does not form pseudohydrotheca. Specimens contractile.
Habitat & Distribution: Polypoid colonies collected on 11 Sept 2016 from the Lingan Bay eelgrass beds, an estuarine habitat. Polyp collected from North Sydney Yacht Club on 28 Oct
2016 falls into this species, as well.
Nucleotide Sequence Data: 16S sequence from specimen collected on 11 Sept 2016 from Lingan
Bay eelgrass beds. Second sequence collected from North Sydney Yacht Club 28 Oct 2016. Both sequences 590 nucleotides in length. Both sequences are 100% identical to one another.
24 | P a g e
Sequences differ from sequence of P. pusilla provided by Dr. Schuchert by 7.4% over 593 base pairs.
Phylogenetic Analysis
Taxon Clarification
Not surprisingly, 16S sequences obtained from my specimens in Lingan Bay eelgrass beds and the North Sydney Yacht Club formed a grouping (Fig. 12). This grouping formed a sister group with sequences of Bougainvillia muscus, and this clade was sister to the sequence from Pachycordyle pusilla, making the pair more closely related to B. muscus based on 16S data
(see Discussion). Interestingly, the sequence of Rhizorhagium arenosum formed a sister grouping with several uncertain species of Bougainvillia. Sequences of Pachycordyle pusilla and
Rhizorhagium arenosum were provided by Dr. Peter Schuchert.
Additionally, the sequences labelled as “Bougain Study Sequence” in Figure 12 formed a clade with the B. muscus sequences apart from the ones in the sister group to Dr. Schuchert’s P. pusilla.
Genetic Connectivity
Due to insufficient sequence data from GenBank, genetic connectivity analysis could not be performed for this specimen.
25 | P a g e
Discussion
Comparison To Regional Surveys
Compared to recent surveys in and around Nova Scotia (Brunel, 1989; Calder, unpublished), 11 of the 13 taxa identified in this study have been recorded in the region previously with the exceptions of Podocorynoides minima and Pachycordyle michaeli, the latter of which was identified in Maine several decades ago by Berrill (1948). The species diversity recorded here is low relative to other regional surveys. This is almost certainly a reflection of both the non-exhaustive nature of this study and its limited temporal and spatial scopes. For instance, there are species of hydroids which do not inhabit the intertidal zone and only occur in deeper water (Blanco et al., 2013), meaning additional species may exist outside of our sampling range. Also, the survey period of this study only spanned from September to November, limiting the project seasonally. Certain hydroids are known to have seasonality in certain regions, e.g.
Gonothyraea loveni (Cornelius, 1992b; Calder, personal correspondence), which leaves much potential for greater species diversity if sampling was expanded across seasons.
Genetic Connectivity
For the species where genetic connectivity analyses were performed, specimens collected generally appeared to be more closely related to conspecifics on the North American coast as opposed to Europe or other parts of the world. The clearest representation of this phenomenon was seen in the genetic connectivity results from Clava multicornis, which formed a distinctive
NA clade and a European Clade, excluding one sequence from New York positioned within the
European clade (Fig. 8). This result is consistent with the phylogenetic analysis performed in a
26 | P a g e more comprehensive analysis of C. multicornis sequences from GenBank (Miglietta et al., 2008).
Although the exact placement of individual sequences was not fully conserved in additional phylogenetic trees with different outgroups, the relationship between the study sequence and sequences from North America was consistent, leading to the conclusion that our population of
C. multicornis is not the result of a recent human-mediated introduction in this region. There is certainly the possibility that the sequence from New York is a human-mediated introduction, considering the capacity of hydroids to foul hard surfaces such as ship hulls (Blanco et al., 2013;
Calder et al., 2014); however, this could represent a case of natural range expansion on free- floating debris, such as algal rafts (Miglietta et al., 2008). A similar result was seen with
Gonothyraea loveni (Fig. 7). It is worth noting that because G. loveni was found on a detached blade of eelgrass floating in East Bay (Penney, pers. obs.), it is possible the colony originally came from another region in the Bras d’Or Lakes.
The connectivity results of Obelia dichotoma were also very interesting given the considerable genetic difference between the specimens collected from the Lingan Bay eelgrass beds and the specimen from the Barra Strait. These specimens failed to group together and instead formed sister relationships with other O. dichotoma sequences extracted from GenBank
(Fig. 10). My phylogenetic analysis also failed to return a monophyletic clade of O. dichotoma relative to O. genticulata and O. bidentata. The monophyly of O. dichotoma, and even the genus
Obelia itself, was disputed in the source paper for the O. dichotoma GenBank sequences (Cunha et al., 2017), which is consistent with the results presented in Figures 9 and 10. In Figures 1-3 of
Cunha et al. (2017), which are parsimonious consensus trees based on molecular markers 16S,
18S, 28S, and COI, individual clades of O. dicotoma are found scattered throughout the tree in a
27 | P a g e polyphyletic manner, which is consistent with our results based only on 16S data. This result suggests that local populations of O. dichotoma may include a cryptic species.
Clarification of Species-Level Identities
The best GenBank match for the specimen of Hydractinia found in Port Morien was to a sequence of H. echinata from the United Kingdom (KX355405). This result is intriguing, as the distribution of H. echinata is believed to be restricted to Europe (Calder, in press). This result was not due to insufficient 16S sequence data on GenBank, as this particular genus is well- studied and well represented in this database (Cunningham and Buss, 1993; Miglietta et al.,
2008). This, instead, may reflect human-mediated dispersal or algal rafting across the Atlantic.
The discovery of this specimen of Hydractinia on the claw of a Rock Crab, Cancer irroratus, is also unusual, especially if it truly is H. echinata. This species is known to use hermit crabs as hosts by colonizing their shells (Miglietta et al., 2008). However, given the uncertainty in species-level taxonomy in the genus Hydractinia (Miglietta et al., 2008), this specimen was not assigned a species identification in this study.
Most of the hydroids encountered in this study have been recorded previously in the region (Brunel, 1998). One of the exceptions to this was the Pachycordyle michaeli collected from the eelgrass beds of Lingan Bay, and North Sydney. The identity Pachycordyle michaeli was proposed for the specimen from the Lingan Bay eelgrass beds by both Dr. Schuchert and Dr.
Calder (Schuchert, pers. comm.; Calder, pers. comm.). This was after BLAST results and phylogenetic analyses (Fig. 12) indicated the closest genetic match to be Bougainvillia muscus, which was inconsistent with morphological data. Morphological data also strongly support the
28 | P a g e identification of P. michaeli based on Schuchert (2007). Molecular analysis was confounded because the only 16S sequence available for comparison was a P. pusilla sequence provided by
Dr. Schuchert, as no other Pachycordyle sequences were available on GenBank. Also, over a stretch of 593 base pairs, the sequence from the Lingan Bay eelgrass specimen differed from
Pachycordyle pusilla by 7.4%, which falls out of the range for intraspecific variation but within that of intrageneric differences. This suggests that Pachycordyle pusilla is not the correct identification, either. At this time, the specimen is identified as P. michaeli based on strong morphological evidence; to confirm this, ethanol-preserved tissue of P. michaeli from Maine is required for genetic comparison to the Lingan Bay sequence.
The identification of Sarsia was challenging in both morphological and molecular analyses. Morphological analysis of Sarsia to species level was confounded by the lack of gonophores on collected polyps, which are often necessary for species identification (Schuchert,
2001). DNA sequencing, while partly successful, was impeded by a 7-9 base-pair weak homopolymer region in the 16S gene fragment (Fig. 13) which disrupted sequence reads downstream from this region. It was initially hypothesized that this was due to sequence errors made during GoTaq PCR amplification through the homopolymer region. Phusion High Affinity
DNA Polymerase PCR Mix, which includes a proofreading enzyme, was used to rectify this, but did not prove to be successful.
To explore sequencing issues in Sarsia further, the sequences in this study were aligned with 16S sequences from several species of Sarsia to compare homopolymer regions (Fig. 14).
Some homopolymer regions were broken up by point mutations to either adenine or cytosine, which may explain why they were fully sequenced without issue. However, other sequences with
29 | P a g e the same number and arrangement of nucleotides in the region as the study sequence appeared to have been successfully amplified to over 600 base pairs, an example being GQ395328. This indicates that there are possibly effective ways to amplify and sequence this region. It may be useful to contact the authors of these studies to better understand how they resolved this problem.
Due to these complications, Sarsia was unable to be assigned a species identification.
Monophyly of Bougainvillia
The genus Bougainvillia is strongly supported by morphological characters (Vannucci &
Rees, 1960). Nevertheless, results of molecular phylogenetic analysis appear to challenge the monophyly of this genus. As seen in Figure 12, two clades, one containing Pachycordyle pusilla and another containing Bougainvillia species from an unpublished study (He et al., unpubl.) paired with Rhizorhagium arenosum, are positioned between clades of Bougainvillia sequences in the phylogenetic tree, not as sister to a monophyletic Bougainvillia clade. This is suggestive that the genus Bougainvillia, when examined using 16S sequence data, is not monophyletic. A close relationship between P. pusilla and Bougainvillia has been noted by Schuchert (2007), who even suggested that placing P. pusilla in the genus Pachycordyle may be incorrect. The sister grouping of R. arenosum with a clade of Bougainvillia also suggests that there are potential misidentifications of specimens as Bougainvillia when they may be another species. Additional examination with other molecular markers should be performed to further analyze this phenomenon and determine if it is restricted to the 16S rRNA gene region.
Study Limitations
The accuracy of morphological analysis in this study was potentially compromised by a number of limitations, including not anesthetizing samples prior to fixing in 10% formalin
30 | P a g e despite the recommendations of Cornelius (1995a). In formalin preserved samples, hydranths of thecate polyps were often contracted into hydrothecae and athecate hydroids were occasionally contracted, which obstructed important details of the polyps. Also, specimens without gonophores present were limiting for this analysis. This was the case for the morphological analysis of Sarsia (Schuchert, 2001). Keeping live specimens of hydroids and rearing them to generate specimens with gonophores is an option to overcome this, but it is also possible to re- sample sites where a particular species was encountered at different time intervals to see if specimens with gonophores can be obtained.
Molecular analyses, especially for genetic connectivity, were often limited by insufficient sequence data on GenBank. This also made species-level determinations of some taxa difficult.
More well-studied hydroids such as species of Obelia and Hydractinia had many sequences submitted. Pachycordyle pusilla only had one 16S sequence submitted to GenBank and was the only representative of the genus. As seen with Pachycordyle michaeli, this limited the reliability of BLAST searches for identification. Additionally, Dynamena pumila had only one corresponding sequence available in GenBank. It may be necessary to contact individual researchers for future studies, as they may have unpublished sequence data available.
31 | P a g e
Future Studies
Future studies that expand on this research could use additional mitochondrial genes and nuclear gene markers in phylogenetic analyses. Studies which focus on the genetic relationships among hydroids usually employ more than one gene region, and typically incorporate a mitochondrial and nuclear gene region (Martinez et al., 2010; Cunha et al., 2017). Although there has been some contention regarding the usefulness of the COI barcoding gene in certain cnidarians (Zheng et al., 2014 and references therein), sequences of COI for hydroids are still available on GenBank and BOLD (Barcode Of Life Database) and might provide an additional fast-evolving molecular marker. Additional molecular data will allow for testing of the robustness of the 16S molecular analyses presented here.
The focus of this study has been to provide a temporal benchmark of species diversity for members of Hydroidolina in Cape Breton given the expectation of changes in species distributions in response to climate change. However, it is also important to have a thorough understanding of the sensitivities of different species to environmental conditions, particularly given the potential use of hydroids as bioindicators (Jones, 2002). Environmental conditions have been documented to lead to cases of morphological plasticity of hydroids, e.g.
Hydractiniidae colonies based on substratum, Dynamena pumila hydrotheca shape based on wave exposure, and Clava multicornis hydranth size based on wave exposure (Cornelius, 1995b;
Miglietta et al., 2008). Hence, it is important to understand the sensitivities of species to different environments, both to avoid misidentification and to begin measuring responses of species to changing environmental conditions.
32 | P a g e
Conclusion
The combined use of molecular and morphological analyses with molecular phylogenetics in this study has allowed for the identification of eleven taxa to species level and thirteen to at least the genus level. This assemblage of taxa reflects many of the common species that can be found in shallow estuarine and marine habitats in Cape Breton; additionally, it represents the first set of 16S rRNA gene sequences for hydroids in Cape Breton. The hydroids identified here also provide a temporal benchmark of species present, which will be useful in future studies assessing changing species distributions in association with climate change.
Finally, this study represents the first stage in the development of a useful dataset for identifying local hydroids using 16S sequence data and accompanying morphological descriptions.
33 | P a g e
Figures
Source: https://www.google.ca/maps/@46.2971511,-60.5937506,279378m/data=!3m1!1e3
Figure 1. Google Maps image of Cape Breton Island showing relative positions of sampling sites. Sites were chosen based on opportunity and accessibility. Red dots indicate sampling sites.
34 | P a g e
w A B E
hyp r h t sta
g h
1mm 1mm C gn D ht
t ht sto h
r 1mm 1mm 0.5mm Figure 2. Photographs highlighting important characteristics for morphological analysis of polypoid life stages. h=hydranth, hyp=hypostome, t=tentacle, w=whorl of tentacles, ht=hydrotheca, r=hydrothecal rim, g=gonophore, gn=gonotheca, sto=stolon, sta=stalk. Taxa include Pachycordyle michaeli (A), Hydractinia spp. (B), Obelia genticulata (C), Obelia dichotoma (D), and Dynamena pumila (E).
35 | P a g e
A B
m mb g
n
n t 1mm C 1mm
D E
o n g t m n
0.5mm 0.5mm 1mm t
Figure 3. Photographs highlighting important characteristics for morphological analysis of medusoid stages. g=gonad, m=manubrium, mb=manubrial bulb, n=tentacle node, o=ocelli, t=tentacle. Taxa include Podocorynoides minima (A), Lizzia blondina (B), Bougainvillia muscus (C), and Obelia genticulata (D&E). Features not shown in (A) but present in other samples include manubrial bulbs, ovular gonads on manubrium, and filiform tentacles. Structure not shown in (B) but present in other samples is a shallow round apical process. Radial canals not visible in photos, but observed in all specimens except B. muscus (radial canals not visible); four canals in each specimen (see Results).
36 | P a g e
1 2 3 4 5 6 - L 1 2 3 4 L 5 6 + -
A B
Figure 4. DNA (A) and PCR (B) gels for a set of samples extracted on 19 Jan 2017 and amplified using GoTaq Master Mix conditions on 20 Jan 2017. Size standards used were GelPilot Wide Range Ladder and GelPilot 100bp Plus Ladder, respectively. In the DNA extraction gel (A) the DNA is shown as a faint smear (highlighted by box in lane 3). PCR products (B) were estimated to be 650bp in size. Positive (+) and negative (-) controls are also included in the PCR gel.
1 2 3 4 L X 5 - + - + 1 2 3 4 5 L
A B
Figure 5. Comparison of yields from PCR reactions on the same samples using two different kits, GoTaq (A) and Phusion (B). PCR amplifications were performed on 23 Feb 2017 and 23 Mar 2017, respectively. The GenePilot 100bp Plus Ladder is shown in both gels. Strength of individual bands, i.e. brightness, is positively correlated with the DNA yield. Faint bands are highlighted. Numbers indicate the five samples amplified in both PCR conditions. Also shown in (A) are the negative (-) and positive (+) controls and one sample (X) that was discontinued in further analysis. All bands estimated to be 650bp in size.
37 | P a g e
Figure 6. Photographic representations of specimens corresponding to study identifications where possible. Clytia gracilis lacked a physical specimen and does not have a photographic
record. Species are listed in alphabetical order by genus. Taxon Identification Image of Life Stage(s) Bougainvillia muscus
Clava multicornis
Clytia gracilis N/A Dynamena pumila
38 | P a g e
Ectopleura larynx
Gonothyraea loveni
Hydractina spp.
39 | P a g e
Lizzia blondina
Obelia dichotoma
Obelia genticulata
Pachycordyle michaeli
40 | P a g e
Podocorynoides minima
Sarsia spp.
41 | P a g e
OG
Canada
(KX665257)
USA: Dennis (AY789826)
(KX665351)
(KT266616)
(AY789827) France: Roscoff
(FJ550480)
Figure 7. Jukes-Cantor Neighbor-Joining (NJ) distance analysis of 16S rRNA sequence data from 7 specimens of Gonothyraea loveni. Obelia bidentata (AY789815) was used as an outgroup (OG). Sequences of G. loveni were extracted from GenBank, accession numbers included. Scale bar included. The sample labelled “Canada” is the sequence from this study.
42 | P a g e
OG
(FJ214439)
(FJ214443)
(FJ214438)
(EU305471)
Europe (FJ214444)
Canada
(FJ214442)
(FJ214440)
(FJ214441)
Figure 8. Jukes-Cantor Neighbor-Joining (NJ) distance analysis of 16S rRNA sequence data from 9 specimens of Clava multicornis. Clavactinia gallensis (FJ214377) was used as an outgroup (OG). Sequences of C. multicornis were extracted from GenBank, accession numbers included. Scale bar included. The sample labelled “Canada” is the sequence from this study.
43 | P a g e
OG
(KM603472)
(KX665347)
(KX665309) 0 (AY789815)
(AY789816)
(KX665266) 0 (AY530362)
(AY530364)
(AY530363)
(FJ550481)
Study Sequence*
(KX665330)
Study Sequence*
Study Sequence*
(KX665325)
(KX665359)
Figure 9. Jukes-Cantor Neighbor-Joining (NJ) distance analysis for 16S rRNA sequence data from 16 specimens of the genus Obelia. Gonothyraea loveni (FJ550480) was used as an outgroup (OG). Sequences were extracted from GenBank, accession numbers included. Scale bar included. Sequences from this study are labelled “Study Sequence.”
44 | P a g e
OG
(KX665309)
(KX665322)
(KX665324)
(KX665347)
(KX665360)
(KM603472)
Canada
(KX665316)
(KX665330)
(KX665288)
(KX665359)
(KX665297)
(KX665325)
(KX665326)
Canada
Canada
(KX665331)
Figure 10. Jukes-Cantor Neighbor-Joining (NJ) distance analysis of genetic connectivity for 16S rRNA sequence data from 17 specimens of Obelia dichotoma. Clytia gracilia (KX665352) was used as an outgroup. Sequences of O. dichotoma were extracted from GenBank, accession numbers included. Scale bar included. Samples labelled as “Canada” are sequences from this study.
45 | P a g e
OG
(JF898011)
(JF898013)
(AY787909)
(JF898003)
(KT286609)
Study Sequence
(AY787902)
(KT266610)
Figure 11. Jukes-Cantor Neighbor-Joining (NJ) distance analysis of 16S rRNA sequence data from 8 specimens of the genus Dynamena. Amphisbetia operculata (FJ550489) was used as an outgroup (OG). Sequences were extracted from GenBank, accession numbers included. Scale bar included. Sequence from this study is labelled as “Study Sequence.”
46 | P a g e
OG (AM183128) (KP776760) (KP776799) (AM411412) (AM411413) (AM183127) (KJ660344) (KJ660345) (EU305470) (JQ715892) (JQ715893) (JQ715894) (JQ715890) (JQ715891) (KF962406) (KF962411) (KF962412) (KF962414) (KF962415) (KF962413)
Pachy Study Sequence Pachy Study Sequence (KT266606) (AM411409) (AM411410) (AM411411) (AM411408) Bougain Study Sequence (EU999220) Bougain Study Sequence (AM41 1408) (KX355427) (AY787880)) (AM183126) ) ) ) Figure 12. Jukes-Cantor Neighbor-Joining (NJ) distance analysis based on 16S rRNA sequence data from 14 Bougainvillidae specimens. Bimeria vestita (KT266604) was used as an outgroup (OG). All sequences of Bougainvillia were extracted from GenBank, accession numbers included. Sequences from this study are labelled one of two ways: “Pachy Study Sequence” if they group closely with Pachycordyle, or “Bougain Study Sequence” if they group closely with Bougainvillia. Sequences of Pachycordyle pusilla and Rhizorhagium arenosum provided by Dr. Peter Schuchert. Scale bar included.
47 | P a g e
A B C
Figure 13. Segment of forward (A) and reverse (B) sequence from Sarsia sample presented in Geneious showing the problematic homopolymer region and the sequence around it. Sequences amplified using GoTaq PCR amplification. Regions were extracted using HYD-16SF and HYD-16SR- LONG primers, respectively. Sequencing reactions move from left to right in this figure. Sequence is readable (A) until homopolymer region is encountered (B), after which mixed signalling results in unusable sequence (C).
48 | P a g e
Consensus
Study Sequence-G Study Sequence-P S. striata (GQ395328) S. apicula (GQ395330) S. tubulosa (EU876548) S. tubulosa (GQ395327) S. tubulosa (AJ878720) S. tubulosa (AY512545) S. marii (AY512544) S. lovenii (AJ608796) S. lovenii (AY787876) S. princeps (EU876549) S. mirabilis (AY512548) S. tubulosa (AB720902)
Figure 14. De Novo assembly of 16S rRNA sequence regions containing the long weak homopolymer (highlighted with box) from several different species of Sarsia. The consensus sequence is along the top with the support level for individual nucleotide positions below it. The sequences from this study are labelled “Study Sequence-G” and “Study Sequence-P,” with G and P referring to the type of PCR amplification used. G=GoTaq, P=Phusion. Other sequences extracted from GenBank.
49 | P a g e
Tables
Table 1. Summary of sampling dates and habitat types for all regions sampled over the course of this study.
Locations Date(s) Sampled Habitat Type Lingan Bay Eelgrass Beds 07/30/16, 09/11/16 Estuarine Dominion Beach 02/22/17 Marine Lingan Bay Fisherman's Wharf 09/13/16 Estuarine East Bay 09/07/16 Estuarine Old Sydney Yacht Club 09/16/16, 09/23/16, Marine 10/07/16 Little Bras d'Or Channel 09/13/16 Estuarine St. Patrick's Channel 09/27/16 Estuarine Great Bras d'Or Channel (inner) 09/15/16 Estuarine Great Bras d'Or Channel (outer) 09/16/16 Estuarine Main a Dieu 09/18/16 Marine Little Narrows 09/20/16 Estuarine Barra Strait 09/21/16 Estuarine Port Morien 09/25/16 Marine St. Peter's Bay 09/27/16 Estuarine South Bar Government Dock 10/02/16 Marine North Sydney Yacht Club 10/28/16 Marine Fourchu 11/11/16 Marine
50 | P a g e
Table 2. List of sequencing primers developed for this study. Primers were designed in Geneious 9.1.5.
Primer Name Sequence (5’ – 3’) NB Designer HYDROZ-16SF GGTGTRACCTGCCCARTG 18 Tim Rawlings HYDROZ-16SR- TTAAAGGTCGAACAGACCT 19 Tim Rawlings LONG HYDROZ-16SR- AACATAGAGGTGACAAACTT 20 Tim Rawlings SHORT-ALL HYDROZ-16SR- CATAGAGGTGACAAACTTTG 20 Tim Rawlings SHORT-TG
51 | P a g e
Table 3. A record of successful and unsuccessful attempts to sequence PCR amplifications of the 16S rRNA gene in various specimens using custom sequencing primers. Species are listed in code form, with the capital letter representing genus and the small letter representing species. Unknown species have an “x” to code for species name. Podocorynoides minima was an additional letter to distinguish it from Pachycordyle michaeli: Bougainvillia muscus (Bm), Clava multicornis (Cm), Clytia gracilis (Cg), Dynamena pumila (Dp), Ectopleura larynx (El), Gonothyraea loveni (Gl), Hydractinia spp (Hx), Lizzia blondina (Lb), Obelia dichotoma (Ob), O. genticulata (Og), Pachycordyle michaeli (Pm), Podocorynoides minima (Pom), Sarsia spp. (Sx).
Primer Successful Sequencing Unsuccessful Sequencing
HYD-16SF Bm, Cm, El, Dp, El, Gl, Hx, Lb, Od, Og, Pm, Pom, Sx HYD-16SR-LONG Bm, Cm, El, Gl, Hx, Lb, Od, Dp, Pm Og, Pom, Sx HYD-16SR-SHORT-ALL Bm, Cg, Cm, Dp, El, Gl, Hx, Lb, Pom Od, Og, Pm, Sx HYD-16SR-SHORT-TG Bm, El Lb, Og, Pom, Sx
52 | P a g e
Table 4. Tentative identifications from BLAST searches of my 16S sequence data based on the highest percentage match found in GenBank. Best matches presented here may not
represent final identifications. Best Match In Location Collected Sequence %Match %Query Accession Genbank Length (bp) Cover # of Match Bougainvillia muscus Lingan Bay Eelgrass 590 95 100 AM183126 Bougainvillia muscus St. Patrick's Channel 590 99 100 AM411408 Bougainvillia muscus St. Patrick's Channel 494* 100 97 EU999220 Bougainvillia muscus North Sydney Yacht 590 95 100 AM183126 Club Clava multicornis Fourchu 590 100 99 FJ214440 Clytia gracilis Lingan Bay Wharf 496* 98 98 KX665334 Dynamena pumila Main a Dieu 575 99 99 AY787902 Ectopleura larynx Great Bras d'Or 567 99 100 EU876545 Channel Gonothyraea loveni East Bay 587 100 100 KX665257 Hydractina echinata Port Morien 587 99 100 KX355405 Lizzia blondina Old Sydney Yacht 580 99 99 AM411423 Club Obelia dichotoma Lingan Bay Eelgrass 589 99 100 KX665359 Obelia dichotoma Lingan Bay Eelgrass 589 99 100 KX665359 Obelia dichotoma Barra Strait 589 99 100 KX665330 Obelia genticulata Lingan Bay Wharf 589 99 100 KX665358 Podocorynoides Old Sydney Yacht 581 96 100 AM411420 minima Club Podocorynoides Lingan Bay Wharf 581 97 97 AM183125 minima Sarsia lovenii, Sarsia Fourchu 474*** 98, 98 99, 99 KT809333, tubulosa** GQ395329 *Asterisk indicates non-consensus sequence based only on reverse primer sequences. **BLAST search turned up to equally likely matches, i.e. had same match, query cover, and E-values. ***Consensus sequence generated with truncated sequence data.
53 | P a g e
Table 5. Final list of taxon identifications in this study made using morphological and/or molecular datasets.
Species Identification Family Location(s) Found Life Stages Found Bougainvillia muscus Bougainvillidae St. Patrick's Polyp, Medusa Channel Clava multicornis Hydractiniidae Fourchu Polyp Clytia gracilis Campanulariidae Lingan Bay Docks Unknown* Dynamena pumila Sertulariidae Main a Dieu Polyp Ectopleura larynx Tubulariidae Great Bras d'Or Polyp Channel Gonothyraea loveni Campanulariidae East Bay Polyp Hydractina spp. Hydractiniidae Port Morien Polyp Lizzia blondina Rathkeidae Old Sydney Yacht Medusa Club Obelia dichotoma Campanulariidae Lingan Bay Polyp Eelgrass Beds, Barra Strait Obelia genticulata Campanulariidae Dominion Beach Polyp and Medusa Pachycordyle michaeli Bougainvillidae Lingan Bay Polyp Eelgrass Beds, North Sydney Yacht Club Podocorynoides minima Rathkeidae Old Sydney Yacht Medusa Club, Lingan Bay Wharf Sarsia spp. Corynidae Fourchu Polyp *Identification is based on match in GenBank of partial sequence extracted from DNA sample of Podocorynoides minima. Life stage is thus unknown.
54 | P a g e
Acknowledgments
I would like to extend my sincerest thanks first and foremost to my supervisor, Dr.
Timothy A. Rawlings, for taking me on as an Honours student and for his continued encouragement and support throughout the process. I would also like to thank committee member Dr. Dale R. Calder for his infinite wisdom on hydroids, assistance with morphological analysis and identification, and for providing me with resources I would not have otherwise had.
I also want to thank committee member Dr. Bruce G. Hatcher for his efforts in collecting hydroids in the Bras d’Or and allowing use of his facilities.
I would also like to thank Dr. Vielka Salazar and Dr. Michael Tanchak for the use of their laboratories for molecular analysis, Dr. Peter Schuchert for his contributions, my labmates and fellow Honours students Robyn Novorolsky, Stephen Williams, and Frank Sinclair for their support, and anyone else kind enough to bring me hydroids for this project.
55 | P a g e
References
Berrill, N. J. 1948. The life cycle of Aselomaris michaeli, a new gymnoblastic hydroid. The Biological Bulletin. 95(3): 289-295. Web. Available from http://www.biodiversitylibrary.org/part/3765#/summary Blanco, R., Shields, M.A., Jamieson, A.J. 2013. Macrofouling of deep-sea instrumentation after three years at 3690 m depth in the Charlie Gibbs fracture zone, mid-Atlantic ridge, with emphasis on hydroids (Cnidaria: Hydrozoa). Deep-Sea Research II. 98: 370-373. Web. DOI: 10.1016/j.dsr2.2013.01.019 Bollens, S.M., Horgan, E., Concelman, S., Madin, L.P., Gallager, S.M., Butler, M. 2001. Planktonic hydroids on Georges Bank: effects of mixing and food supply on feeding and growth. Deep-Sea Research II. 48: 659-672. Web. Available from https://research.vancouver.wsu.edu/sites/research.vancouver.wsu.edu/files/Bollens-Deep- Sea%20Research%20II%2048%20659-672%20pdf.pdf Bouillon, J., Gravili, C., Pages, F., Gili, J., Boero, F. 2006. An introduction to Hydrozoa. Tome 194. Paris: Publications Scientifiques du Muséum. Web. Available from ResearchGate https://www.researchgate.net/publication/231575419_An_Introduction_to_Hydrozoa Bouillon, J., Boero, F. 2000. The Hydrozoa: a new classification in the light of old knowledge. Thalassia Salentina. 24:1-43. Web. Available from http://siba- ese.unile.it/index.php/thalassiasal/article/viewFile/i15910725v24p3/1885 Brunel, P., Bosse, L., Lamarche, G. 1998. Catalogue of the marine invertebrates of the estuary and Gulf of Saint Lawrence. Canadian Special Publication of Fisheries and Aquatic Sciences. 126. P. 64-75. Print. Calder, D.R. In press. Hydroids of the Bay of Fundy, northeastern North America. Zootaxa.
Calder, D.R. Unpublished. Hydroids of Atlantic Canada. Royal Ontario Museum.
Calder, D.R. 1975. Biotic census of Cape Cod Bay: hydroids. The Biological Bulletin. 149: 287- 315. Web. Available from http://www.journals.uchicago.edu/doi/pdfplus/10.2307/1540528 Calder, D.R. 1991. Associations between hydroid species assemblages and substrate types in the mangal at Twin Cays, Belize. Canadian Journal of Zoology. 69:2067-2074. Web. DOI: 10.1139/z91-288 Calder, D.R., Vervoort, W. 1998. Some hydroids (Cnidaria: Hydrozoa) from the Mid-Atlantic Ridge in the North Atlantic Ocean. Zoologische Verhandelingen. Leiden. 319: 1-65. Web. Available from ResearchGate https://www.researchgate.net/publication/245536295_Calder_DR_and_W_Vervoort_199
56 | P a g e
8_Some_hydroids_Cnidaria_Hydrozoa_from_the_Mid- Atlantic_Ridge_in_the_North_Atlantic_Ocean_Zoologische_Verhandelingen_Leiden_31 9_1-65 Calder, D.R. 2010. Some anthoathecate hydroids and limnopolyps (Cnidaria, Hydrozoa) from the Hawaiian archipelago. Zootaxa. 2590: 1-91. Available from ResearchGate https://www.researchgate.net/publication/266017097_Some_anthoathecate_hydroids_and _limnopolyps_Cnidaria_Hydrozoa_from_the_Hawaiian_archipelago Calder, D.R., Choong, H.H.C., Carlton, J.T., Chapman, J.W., Miller, J.A., Geller, J. 2014. Hydroids (Cnidaria: Hydrozoa) from Japanese tsunami marine debris washing ashore in the northwestern United States). Aquatic Invasions. 9(4): 425-440. Web. Available from http://dx.doi.org/10.3391/ai.2014.9.4.02 Cantero, A.L.P., Sentandreu, V., Latorre, A. 2010. Phylogenetic relationships of the endemic Antarctic benthic hydroids (Cnidaria, Hydrozoa): what does the mitochondrial 16S rRNA tell us about it? Polar Biology. 33: 41-57. Web. DOI: 10.1007/s00300-009-0683-5 Cantero, A.L.P., Ramil, F. 2006. Benthic hydroids associated with volcanic structures from Bransfield Strait (Antarctica) collected by the Spanish Antarctic expedition GEBRAP96. Deep-Sea Research II. 53: 949-958. Web. DOI: 10.1016/j.dsr2.2006.02.007 Cartwright, P., Evans, N.M., Dunn, C.W., Marques, A.C., Miglietta, M.P., Schuchert, P., Collins, A.G. 2008. Phylogenetics of Hydroidolina (Hydrozoa: Cnidaria). Journal of the Marine Biological Association of the United Kingdom. Pgs. 1-10. Web. DOI: 10.1017/S0025315408002257 Collins, A.G. 2002. Phylogeny of Medusozoa and the evolution of cnidarian life cycles. Journal of Evolutionary Biology. 15: 418-432. Web. Available from http://onlinelibrary.wiley.com/doi/10.1046/j.1420-9101.2002.00403.x/full Collins, A.G., Schuchert, P., Marques, A.C., Jankowski, T., Medina, M., Schierwater, B. 2006. Medusozoan phylogeny and character evolution clarified by new large and small subunit rDNA data and an assessment of the utility of phylogenetic mixture models. Systems Biology. 55(1): 97-115. Web. DOI: 10.1080/10635150500433615 Concelman, S., Bollens, S.M., Sullivan, B.K., Madin, L.P., Horgan, E., Bulter, M., Keuren, D.V. 2001. Distribution, abundance and benthic-pelagic coupling of suspended hydroids on Georges Bank. Deep-Sea Research II. 48: 645-658. Web. Available from http://dx.doi.org/10.1016/S0967-0645(00)00091-6 Cornelius, P.F. 1995a. North-West European thecate hydroids and their medusae, Part 1: Introduction, Laodiceidae to Haleciidae. Shrewsbury: Field Studies Council. ISBN: 1 85153 254 4. Print. 1-347. Cornelius, P.F. 1995b. North-West European thecate hydroids and their medusae, Part 2: Sertulariidae to Campanulariidae. Shrewsbury: Field Studies Council. ISBN: 1 85153 255 2. Print. 1-386.
57 | P a g e
Cunha, A.F., Collins, A.G., and Marques, A.C. 2017. Phylogenetic relationships of Proboscoida Broch, 1910 (Cnidaria, Hydrozoa): Are traditional morphological diagnostic characters relevant for the delimitation of lineages at the species, genus, and family levels? Molecular Phylogenetics and Evolution. 106: 118–135. Web. Available from http://www.sciencedirect.com/science/article/pii/S1055790316302378 Cunningham, C.W., Buss, L.W. 1993. Molecular evidence for multiple episodes of paedomorphosis in the Family Hydractiniidae. Biochemical Systematics and Ecology. 21: 57-67. Web. Available from http://sites.biology.duke.edu/cunningham/pdfs/CunninghamBuss1993.pdf Daly, M., Brugler, M.R., Cartwright, P., Collins, A.G., Dawson, M.N., Fautin, D.G., France, S.C., McFadden, C.S., Opresko, D.M., Rodriguez, E., Romano, S.L., Stake, J.L. 2007. The phylum Cnidaria: a review of phylogenetic patterns and diversity 300 years after Linnaeus. Zootaxa 1668: 127-182. Web. Available from http://www.mapress.com/zootaxa/2007f/zt01668p182.pdf Dawson, M.N, Jacobs, D.K. 2001. Molecular evidence for cryptic species of Aurelia aurita (Cnidaria, Scyphozoa). The Biological Bulletin. 200: 92-96. Web. Available from http://www.biolbull.org/content/200/1/92.full Folino-Rorem, N.C., Indelicato, J. 2005. Controlling biofouling caused by the colonial hydroid Cordylophora Caspia. Water Research. 39: 2731-2737. Web. DOI: 10.1016/j.watres.2005.04.053 Fraser, C.M. 1926. No. 12: The hydroids of the Cheticamp expedition of 1917. Contributions to Canadian Biology and Fisheries. 3(1): 323-329. Web. doi: 10.1139/f26-012 Gili, J.M., Hughes, R.G. 1995. The ecology of marine benthic hydroids. Oceanography and Marine Biology: an Annual Review. 33: 351-426. Web. Available from ResearchGate https://www.researchgate.net/publication/236123829_The_Ecology_of_marine_benthic_ hydroids Hardy, S.M., Carr, C.M., Hardman, M., Steinke, D., Corstorphine, E., Mah, C. 2011. Biodiversity and phylogeography of Arctic marine fauna: insights from molecular tools. Marine Biodiversity. 41: 195-210. Web. DOI: 10.1007/s12526-010-0056-x He, J.R., Zheng, L.M., Lin, Y.S., Zhang, W.J., Cao, W.Q. Unpublished. DNA bardocing
Medusozoa of China Seas. Dept. of Marine Biological Science and Technology, Xiamen
University.
Jones, T. 2002. Biology and ecology of athecate hydroids in Biscayne Bay, Florida: potential use as bioindicators. Dissertation (Ph.D): University of Miami. Ann Arbor: ProQuest Information and Learning Company.
58 | P a g e
Laakmann, S., Holst, S. 2013. Emphasizing the diversity of North Sea hydromedusae by combining morphological and molecular methods. Journal of Plankton Research. 36(1): 64-76. Web. DOI: 10.1093/plankt/fbt078 Lindeque, P.K., Parry, H.E., Harmer, R.A., Somerfield, P.J., Atkinson, A. 2013. Next generation sequencing reveals the hidden diversity of zooplankton assemblages. PLoS ONE. 8(11): 1-14. Web. DOI: 10.1371/journal.pone.0081327 Martinez, D.E., Iniguez, A.R., Percell, K.M., Willner, J.B., Signorovitch, J., Campbell, R.D. 2010. Phylogeny and biogeography of Hydra (Cnidaria: Hydridae) using mitochondrial and nuclear DNA sequences. Molecular Phylogenetics and Evolution. 57: 403-410. Web. DOI: 10.1016/j.ympev.2010.06.016 Medel, M.D., Lopez-Gonzales, P.J. 1998. Distribution patterns in Atlantic hydroids. Zoologische Verhandelingen. Leiden. 323: 155-168. Web. Available from http://repository.naturalis.nl/document/149179 Miglietta, M.P., Schuchert, P., Cunningham, C.W. 2008. Reconciling genealogical and morphological species in a worldwide study of the Family Hydractiniidae (Cnidaria, Hydrozoa). Zoologica Scripta. 38(4): 403-430. Web. DOI: 10.1111/j.1463- 6409.2008.00376.x Nawrocki, A.M., Collins, A.G., Hirano, Y.M., Schuchert, P., Cartwright, P. 2013. Phylogenetic placement of Hydra and relationships within Aplanulata (Cnidaria: Hydrozoa). Molecular Phylogenetics and Evolution. 67: 60-71. Web. Available from doi:10.1016/j.ympev.2012.12.016 Ortman, B.D., Bucklin, A., Pages, F., Youngbluth, M. 2010. DNA barcoding the Medusozoa using mtCOI. Deep-Sea Research II. 57: 2148-2156. Web. DOI: 10.1016/j.dsr2.2010.09.017 Rodriquez, E., Barbeitos, M.S., Brugler, M.R., Crowley, L.M., Grajales, A., Gusmao, L., Haussermann, V., Reft, A., Daly, M. 2014. Hidden among sea anemones: the first comprehensive phylogenetic reconstruction of the Order Actiniaria (Cnidaria, Anthozoa, Hexacorallia) reveals a novel group of hexacorals. PLoS ONE. 9(5): 1-17. Web. DOI: 10.1371/journal.pone.0096998 Ronowicz, M., Kuklinski, P., Mapstone, G.M. 2015. Trends in the diversity, distribution, and life history strategy of Arctic Hydrozoa (Cnidaria). PLoS ONE. 10(3): 1-19. Web. Available from http://dx.doi.org/10.1371/journal.pone.0120204 Schuchert, P. 1996. The marine fauna of New Zealand: athecate hydroids and their medusa (Cnidaria: Hydrozoa). Wellington: New Zealand Oceanographic Institute. ISSN: 0083- 7903, 106. Web. Available from ResearchGate https://www.researchgate.net/publication/229439242_The_marine_fauna_of_New_Zeala nd_athecate_hydroids_and_their_medusae_Cnidaria_Hydrozoa
59 | P a g e
Schuchet, P. 2001. Survey of the family Corynidae (Cnidaria, Hydrozoa). Revue Suisse de Zoologie. 108(4): 739-878. Web. Available from ResearchGate https://www.researchgate.net/publication/229439255_Survey_of_the_family_Corynidae_ Cnidaria_Hydrozoa Schuchert, P. 2005. Species boundaries in the hydrozoan genus Coryne. Molecular Phylogenetics and Evolution. 36: 194-199. Web. Available from ResearchGate https://www.researchgate.net/publication/7836448_Species_boundaries_in_the_hydrozoa n_genus_Coryne?ev=prf_pub Schuchert, P. 2007. The European athecate hydroids and their medusae (Hydrozoa, Cnidaria): Filifera part 2. Revue Suisse de Zoologie. 114(2): 195-396. Web. Available from ResearchGate https://www.researchgate.net/publication/229439193_The_European_athecate_hydroids_ and_their_medusae_Hydrozoa_Cnidaria_Filifera_Part_2 Schuchert, P. 2008. The European athecate hydroids and their medusae (Hydrozoa, Cnidaria): Filifera part 3. Revue Suisse de Zoologie. 115(2): 221-302. Web. Available from http://www.biodiversitylibrary.org/page/41183893#page/5/mode/1up Schuchert, P. 2010. The European athecate hydroids and their medusa (Hydrozoa, Cnidaria: Capitata part 2. Revue Suisse De Zoologie. 117(3): 337-555. Web. Available from ResearchGate https://www.researchgate.net/publication/229439705_The_European_athecate_hydroids_ and_their_medusae_Hydrozoa_Cnidaria_Capitata_Part_2?ev=prf_pub Schuchert, P. 2014. High genetic diversity in the hydroid Plumularia setacea: a multitude of cryptic species or extensive population subdivision? Molecular Phylogenetics and Evolution. 76: 1-9. Web. Available from doi:10.1016/j.ympev.2014.02.020 Tseng, L., Wu, C., Twan, W., Tang, Z., Hwang, J. 2014. Hydroids (Cnidaria, Hydrozoa) from marine environments in Taiwan. Zoological Studies. 53: 1-7. Web. Available from ResearchGate https://www.researchgate.net/publication/275487597_Hydroids_Cnidaria_Hydrozoa_fro m_marine_environments_in_Taiwan Ulster Museum and the Marine Conservation Society. 1997. The species directory of the marine fauna of the British Isles and surrounding seas. 1st ed. Edited by Howson, C.M., and Picton, B.E. Belfast (UK): Ulster Musem & The Marine Conservation Society. Web. Available from www.vliz.be/imisdocs/publications/68694.pdf Vanucci, M., Rees, W. J. 1961. A revision of the genus Bougainvillia (Anthomedusae). Boletim Do Instituto Oceanografico. Sao Paulo. 9: 57-100. Web. Available from ResearchGate https://www.researchgate.net/publication/262440310_A_revision_of_the_genus_Bougain villia_Anthomedusae
60 | P a g e
Whiteaves, J.F. 1874. On recent deep-sea dredging operations in the Gulf of St. Lawrence. The American Journal of Science and Arts. 7: 1-12. Web. Available from https://play.google.com/books/reader?id=_CkrAAAAYAAJ&printsec=frontcover&outpu t=reader&hl=en&pg=GBS.PA1 Zheng, L., He, J., Lin, Y., Cao, W., Zhang, W. 2014. 16S rRNA is a better choice than COI for DNA barcoding hydrozoans in the coastal waters of China. Acta Oceanologica Sinica. 33(4): 55-76. Web. DOI: 10.1007/s13131-014-0415-8
61 | P a g e