Thompson Rivers University

Developmental and environmental variation in the microbial
communities of the newly hatched juvenile marine snail, Nucella ostrina.
UREAP Project

Tallis Dixon
Primary supervisor: Dr. Louis Gosselin
Co-supervisor: Dr. Eric Bottos
September 20, 2024

INTRODUCTION
The microbiome plays an important role in maintaining the health and fitness of
marine invertebrates by facilitating immune defence, aiding in digestion, and enabling
adaptation to environmental conditions (Khan et al. 2018). Marine invertebrates
harbour a diverse variety of bacterial communities, predominantly within their
digestive systems as well as on the external surfaces of the body. Some bacteria found
on the body surface of marine invertebrates have beneficial properties, others can be
harmful pathogens, and some may have no functional benefit to the animal at all. The
bacteria associated with marine invertebrates have been found to alter the animal’s
vulnerability to pathogens and predators (Menabit et al. 2024). In addition, studies
using PCR (polymerase chain reaction) sequencing and synthesis techniques have
identified bioactive metabolites with antimicrobial properties in marine invertebrates (
El Samak et al. 2018), an important consideration given the current increase in
antibiotic resistance among medically important pathogens. There is also a growing
concern that climate change, which is causing a rise in seawater temperature
(Iwabuchi & Gosselin 2019), could disrupt the natural microbial communities
associated with marine invertebrates. For instance, Khan et al. (2018) discovered that
bacteria in the genera Clostridiales and Verrucomicrobiales were present in high
densities on the gills and in the gut of heat-resistant oysters, and those bacteria
improved the oyster’s ability to respond to heat; however, Acidobacteriales and

Betaproteobacteriales were potential pathogens and were found more frequently on
heat-susceptible oysters. Additionally, PCR analysis of sponge microbiomes before and
after heat stress events revealed the presence of transient bacteria on vulnerable

sponges, resulting in fatalities of some of those sponges (De Castro-Fernández et al.
2024). Warming oceans can also affect the gut microbiome of marine invertebrates,
particularly the abundance of disease-causing bacteria (Liu et al. 2023); Liu et al.
(2023) found microbial diversity and abundance increased under warming
temperatures in the laboratory, and to a lesser extent in the field. Evidence to date
therefore indicates that the microbial community associated with marine invertebrates
plays an important role in the survival and growth of these animals.
Though several studies have examined microbial diversity and activity on marine
invertebrates, most have focussed on the gut microbiome, and all studies have
focussed on adult invertebrates. No study to date has investigated the outer
microbiome of early juveniles, which is known to be the most vulnerable phase of the
life cycle of marine invertebrates (Gosselin & Qian 1997). In addition, a majority of
microbial studies have focussed on invertebrates reared in aquaculture facilities; far
fewer studies have examined the microbial community associated with wild
invertebrates in natural habitats. This project, carried out in part at the Bamfield
Marine Sciences Center, seeks to characterize the microbiome of early juvenile marine
invertebrates living in natural habitats. The specific goal of the project was to
determine an effective methodology for the extraction and amplification of bacterial
DNA from both the juvenile and adult stages of the snail, Nucella ostrina.

METHODS
Field collection at BMSC

The first part of the study aimed to collect juvenile and adult N. ostrina from
their natural habitats and preserve them for subsequent analysis of their microbiome.
Field collection was carried out from May 15th to June 30th 2024 in Barkley Sound, near
the Bamfield Marine Sciences Center (BMSC) on the west coast of Vancouver Island.
Adult snails and egg capsules of N. ostrina were collected by hand, using sterilized
gloves and tweezers, from the intertidal zone of an island within the Ross Islets (48°
52’ 18.8” N, 125° 09’ 43.7” W) and brought back to the BMSC laboratory, where they
were placed immediately into holding tanks containing water collected from their field
site.

Hatching and preservation of snails for DNA extraction
The next step was to obtain newly hatched snails from the egg capsules
collected from the field and to preserve snails for eventual microbiome analysis. The
egg capsules and adults were placed together in cleaned tanks; these tanks contained
seawater collected from their field site to ensure the snails remained exposed to the
same microbes and conditions they were exposed to in the field. Hatchlings emerged
from the egg capsules over the following 36 h into the same water as the adults,
ensuring consistency across the microbes the snails were exposed to. After a sufficient
number of juveniles hatched, both adults and juveniles were then placed for 48h into a
different clean tank containing seawater from the field site to allow a microbiome to
develop on the surfaces of the juvenile snails. Snails were then placed in sterilized
centrifuge tubes containing either 5 adults or 20 juveniles, and frozen at -20˚C for 24 h
to euthanize. Seawater from the tanks was also collected and frozen for subsequent

DNA analysis to determine whether the seawater microbiome is significantly different
from the snail microbiome.

DNA extraction
The second part of this study aimed to develop a method to extract DNA from
the outside of both the juvenile and adult snails. DNA extraction and laboratory work
was conducted at the Microbial Ecology and TRUGen sequencing laboratory at
Thompson Rivers University from July to September 2024. Juveniles and adult snails
were thawed and washed three times in sterile seawater by vortexing, and then the
seawater was removed via pipet and placed in separate collection tubes. This was
performed to remove transient bacteria in the surroundings that are not part of the
snail’s microbiome. ATL buffer was used to remove the outer microbiome; snails were
placed in the buffer for 30-40 minutes at 60 °C, then the surrounding buffer containing
the outer microbiome was removed into a separate 2mL microcentrifuge tube. Sterile
seawater was used to wash the remaining ATL buffer off the snails, then using
sterilized tweezers the body of each adult snail was removed from its shell to allow the
following buffers to reach body tissues normally located under the shell. The samples
of ATL buffer containing DNA from wash water, seawater, or the outer body surface
were all centrifuged at 14,000 rpm for 10 min into a pellet, and the supernatant ATL
buffer was then poured off. The pellet samples were also rinsed with sterile seawater
to remove all ATL buffer for future steps.

DNA purification
Amplification of DNA from the pellet samples was carried out using the E.Z.N.A
Mollusc & Insect DNA Kit® (Omega Bio-Tek), which is designed for the efficient

recovery of genomic DNA from molluscs and was chosen for its ability to break down
complex mucopolysaccharides that are present within the mucus of snails (Belouhova
et al., 2022). Mucopolysaccharides are negatively charged polysaccharide compounds
that are believed to function in cell signalling and other biochemical processes (Casale
& Crane, 2019). Their negative charge allows them to interact with a variety of proteins
and ions as well as interacting with DNA polymerases (Casale & Crane, 2019, Sidstedt
et al. 2020). The binding of mucopolysaccharides could impede the activity of the
polymerase stopping effective transcription (Sidstedt et al. 2020). The following steps
were applied as described by the manufacturer of the E.Z.N.A Mollusc & Insect DNA Kit.
The procedure relies on the properties of the cationic detergent, cetyltrimethyl
ammonium bromide (CTAB), present in the ML1 buffer to break down and dissolve
membrane lipids. Proteinase K digests proteins, histones and nucleases. Together,
these compounds lyse the cell and release the DNA. Samples were incubated at 60˚C
for 30 min or until soluble, as proteinase K is the most effective at this temperature,
maximizing its ability to degrade proteins surrounding the DNA. Once soluble, a
solution of chloroform:isoamyl alcohol (24:1) was added to separate the DNA and RNA
from the mucopolysaccharides that inhibit transcription. This resulted in two distinct
layers: a top layer containing RNA and DNA (which was removed into separate tubes),
and a lower layer (which was discarded) containing proteins, lipids, other organic
molecules; in between these layers was a central milky interface containing denatured
proteins and debris. BL buffer and RNase A were then added to the tube containing
the RNA and DNA. BL buffer helps promote the binding of the DNA to the silica
column, as it contains chaotropic salts that disrupt hydrogen bonds allowing the DNA

to bind more easily. RNase A is an enzyme that cleaves the phosphodiester bonds
between ribonucleotides, selectively destroying all the RNA in the sample. Following
this, ethanol was then added to precipitate DNA and help it bind to the silica; DNA is
soluble in water because of its negatively charged backbone, but in ethanol the
hydration shell is disrupted allowing the DNA to precipitate out of the solution. This
allows the DNA to bind onto the silica spin column, to isolate and discard other liquids.
HBC buffer is then added to disrupt hydrogen bonds after the DNA has bound, aiding
to denature any remaining proteins and increasing the affinity of DNA for the column.
A DNA wash buffer was then used to remove residual proteins, polysaccharides, lipids,
salts and organic solvents, and an elution buffer heated to 70˚C rehydrated the DNA,
releasing it from the silica into the collection tube below.

Analysis of extracted DNA
This step served to isolate the 16S rRNA bacterial gene sequence by thermal
cycle amplification to visualize the presence of bacterial DNA in the samples. This gene
was targeted in sequencing due to being highly conserved across bacterial phyla
allowing for comparThons. The samples containing the outer microbiome and the
samples containing the entire microbiome from the adult snails were diluted to reduce
the amount of DNA in the sample, as high concentrations inhibit binding of primers
during PCR. The outer samples were diluted in sterile PCR water to a 1:5 ratio, and
samples of entire homogenized snails were diluted to a 1:10 ratio.
Each purified DNA sample was processed using PCR to amplify the 16S rRNA
gene. The DNA along with Taq polymerase (a type of polymerase that can withstand
high temperatures), the 341F primer, and the 806R primer were run through 30 cycles

in the thermocycler. These samples were then run on a 1% agarose gel to confirm the
amplification of this gene region and presence of bacterial DNA.

RESULTS

The initial PCR trials using the DNeasy PowerBiofilm Kit resulted in the majority
of samples not producing visible DNA bands (Figures 1, 2). This was likely due to the kit
not containing chloroform:isoamyl alcohol (24:1), a critical step in the removal of the

Figure 1. Agarose gel of juvenile extractions at BMSC. From left to right, ladder,
positive control, negative control, sterile seawater (2nd control), Seawater, wash
water, outer of juvenile (2 reps), entire homogenized juveniles (2 reps).

mucopolysaccharides that can inhibit DNA polymerase. Additional trials in which the
positive control (which is known to work) was added into the juvenile samples also
produced no visible DNA bands (Figure 2), confirming that inhibition was the issue

rather than an absence of bacterial DNA. The analyses of the 5 different species of
marine invertebrate helped to determine if the lack of DNA from N. ostrina samples
was specific to this species or was indicative of a broader problem with the extraction
methodology. The results showed no band for echinoderms and ascidians (the species
with increased mucus production), two bands in cnidarians, and a band for both
crustaceans and mollusc (Figure 2).

Figure 2. Agarose gel of juvenile extractions at BMSC. From left to right, ladder,
positive control, negative control, homogenized juvenile spiked with positive control,
juvenile unspiked, adult (N.ostrina), Echinoderm, Cnidarian, Ascidian, Crustacean,
Mollusc (mussel).

The next DNA extraction trials, using the E.Z.N.A Mollusc & Insect DNA Kit,
resulted in visible DNA bands for all samples of juvenile and adult N. ostrina (Figures 2,

3, 4). Bands of ~450 base pairs were visible, as expected for the amplification of the
16S rRNA gene sequence. The 16S rRNA gene has a known sequence length and,
based on the primers we selected (341F and 806R), it should result in a band at ~450
base pairs long if bacterial DNA is present and has been amplified.

Figure 3. Agarose gel of diluted extractions from adult snails. The top row of the gel
left to right, ladder, positive control, negative control and 12 replications of the outer
microbiome. The lower row shows 12 replicates of the entire homogenized adults both
outer and inner.

Figure 4. Agarose gel of extraction from juvenile snails. the top row in the gel left to
right, ladder, positive control, negative control and 12 replications of the outer
microbiome. The lower row shows the replications of the entire homogenized juveniles

Figure 5. Agarose gel of extraction on seawater. the gel left to right, ladder, positive
control, negative control and 12 replications of the outer microbiome. Well 9, is blank and
did not get a good extraction possibly due to errors in the extraction process or insufficient
DNA in that sample.
DISCUSSION
This study revealed that the E.Z.N.A Mollusc & Insect DNA Kit is more effective for
extracting and amplifying microbial DNA from the outer surface of Nucella ostrina than the
DNeasy PowerBiofilm Kit. Initial trials using the DNeasy PowerBiofilm Kit failed to produce
consistently visible DNA bands on electrophoresis gels following PCR, suggesting that this
kit is not able to remove inhibitors such as mucopolysaccharides during DNA extraction,
adding challenges for the transcription of DNA during PCR amplification. As seen with
other mucus-rich marine invertebrates such as echinoderms, DNA amplification was not
successful even when spiked with positive controls, revealing the presence of inhibitors in
the mucus of these organisms. The E.Z.N.A Mollusc & Insect DNA Kit includes

chloroform:isoamyl alcohol (24:1) step, which separates DNA and RNA from the other
proteins, polysaccharides and organic compounds, allowing for cleaner isolation of
DNA and RNA. This was essential in obtaining clear bands of~450 bp during gel
electrophoresis that would confirm the successful amplification of the 16S rRNA gene
sequence. The inclusion of the chloroform:isoamyl alcohol (24:1) step was responsible
for the difference between these two kits. This suggests that the removal of
mucopolysaccharides is critical for successfully extracting and amplifying microbial
DNA from mucus-rich invertebrates.
Following the extraction and amplification of microbial DNA, the next step in
this study would be to sequence the 16S rRNA gene in order to identify the bacterial
phyla found on the outside of snails and compare the microbiome associated with the
two life stages (juvenile & adult). In addition, by comparing the outer microbiome of
the snails, the inner microbiome of the snails, the wash water microbes from the snails
and the seawater microbes in their environment, we could determine if the outer
microbiome is significantly different from the surrounding water and the inner
microbiome. A comparative analysis here would also confirm whether the method of
extracting the outer microbiome is working and what communities of microbes it
contains.
This study demonstrates the importance of selecting appropriate DNA
extraction methods when working to extract DNA from different conditions, such as
the mucus-rich surface of N. ostrina. The E.Z.N.A Mollusc & Insect DNA Kit was shown
to be the most effective in extracting amplifiable bacterial DNA from both juveniles
and adult snails, largely due to its ability to remove mucopolysaccharides and other

possible inhibitors of PCR. The presence of distinct DNA bands at the expected size
range confirms the successful isolation and amplification of the 16S rRNA gene,
leading the way for future DNA sequencing of the microbial communities.
These findings contribute to the understanding of the microbial ecology
associated with N. ostrina, but also provide a foundation for future research, including
phyla characterization through sequencing to better understand interactions between
marine invertebrates and their associated microbiomes. Understanding these
relationships is crucial for assessing the potential impacts of environmental changes
on the health and survival of marine invertebrates such as N. ostrina.
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