Available via license: CC BY 4.0
Content may be subject to copyright.
Page 1/23
Impacts of near‐future ocean warming on microbial
community composition of the stomach of the
soft‐bottom sea star Luidia clathrata (Say)
(Echinodermata: Asteroidea)
Michelle D Curtis ( curtismi@uab.edu )
The University of Alabama at Birmingham https://orcid.org/0000-0001-8360-8991
Casey D Morrow
The University of Alabama at Birmingham
James B McClintock
The University of Alabama at Birmingham
Research Article
Keywords: Asteroidea, climate change, ocean warming, microbiome, 16s, cardiac stomach
DOI: https://doi.org/10.21203/rs.3.rs-306104/v1
License: This work is licensed under a Creative Commons Attribution 4.0 International License.
Read Full License
Page 2/23
Abstract
There is growing evidence that environmental changes caused by climate change can impact the
microbiome of marine invertebrates. Such changes can have important implications for the overall health
of the host. In the present study we investigated the impact of chronic exposure to an ambient (28°C) and
a predicted mid- (30°C) and end-of-century (32°C) seawater temperature on microbiome modication in
tissues of the cardiac stomach of the abundant predatory sea star Luidia clathrata collected in
September 2018 from Apalachee Bay, Florida (29°58’N, 84°19’W) in the northern Gulf of Mexico (GOM).
Diversity (Shannon index) was lowest among the microbial community of stomach tissue when
compared to the microbiome of the articial sea star feed, and aquarium sand and seawater across all
three experimental temperature treatments. Moreover, the stomach microbial community composition
was distinct between each of the four sample types. Exposure to the highest experimental temperature
treatment (32°C) resulted in a signicant modication of the composition of the microbial community in
stomach and sand samples, but not in seawater samples when compared to those from the current mean
ambient GOM temperature (28°C). Importantly, at the most elevated temperature the stomach
microbiome shifted from a Vibrio sp. dominated community to a more diverse community with higher
proportions of additional taxa including Delftia sp. and Pseudomonas sp. This microbiome shift could
impact the digestive functionality and ultimately the health of L. clathrata, a key soft-bottom predator in
the northern GOM.
Introduction
Mean ocean surface temperatures have risen approximately 0.6°C over the last century due to global
climate change and are projected to warm up to 4°C by the year 2100 (IPCC, 2019). This warming has
already impacted the structure of benthic marine invertebrate communities by forcing shifts in
geographic distributions of various species (Schiel et al. 2004; Parmesan 2006), introducing non-native
species (Przeslawski et al. 2008), and decreasing tness of select species subject to thermal stress
(Pörtner 2008). Ectothermic marine invertebrates living in shallow waters at or near their upper thermal
tolerance are most at risk for deleterious impacts with continued ocean warming (Pörtner 2002; Peck et
al. 2008).
Ocean warming has also been linked to increased prevalence of marine pathogens (Harvell et al. 2002)
and as a result some organisms are increasingly susceptible to disease. This is particularly the case in
marine ectotherms including sea urchins and sea stars (Tracy et al. 2019). For example, the large, multi-
armed predatory sea star
Pycnopodia helianthoides
has experienced widescale population declines along
the Pacic coast of North America resulting from an outbreak of sea star wasting disease thought to be
linked to ocean warming (Harvell et al. 2019). Ocean warming has also been associated with increased
morbidity and mortality from disease in the keystone sea star
Pisaster ochraceus
(Bates et al. 2009;
Eisenlord et al. 2016; Kohl et al. 2016). Exposure to increased seawater temperature can induce more
benign impacts in stars including accelerated growth and elevated feeding rates (Watts and Lawrence
1986, 1990; Eckert et al. 2000; Gooding et al. 2009; Fly et al. 2012; Monaco et al. 2014). However,
Page 3/23
sublethal stress impacts from elevated temperature may also occur including higher metabolic demand,
decreased immune response (Watts and Lawrence 1986, 1990), and arm autotomy in sea stars (Barker et
al. 1992; Haramoto et al. 2007; Barker and Scheibling 2008).
Microbial communities impact a variety of aspects of host physiological function through their inuence
on host behavior, contribution to metabolic functioning, and maintenance or enhancement of immune
response (reviewed in Holmes et al. 2011; Kinross et al. 2011; Vuong et al. 2017). Conversely, microbial
community dynamics are affected by the genetics, behavior, and physiological state of the host (Spor et
al. 2011; Foster et al. 2015; Gomez et al. 2017). This delicate balance between host-microbiome
interactions can be disrupted by changes in host physiology or by changes in the external environment
(Lynch and Hsiao 2019). Disruption of this balance can have deleterious consequences for the host,
which may in turn lead to signicant ecological impacts if the host is an important player in the
community. A recent example of host-microbiome discordance with ecological implications is illustrated
by the negative impacts of sea star wasting disease on the epidermal microbiome of the keystone sea
star
Pisaster ochraceus
. Lloyd and Pespeni (2018) reported a decrease in overall epidermal microbial
species richness in
P. ochraceus
, suggesting a shift away from bacterial taxa with functional attributes
that supported an immune response to pathogenic microbial typical of sea star wasting disease. In a
separate comparative study, disease progression similarly led to a shift in the microbial community
structure of the gonad, body wall, tube feet, and pyloric ceca of four species of coralivorous crown-of-
thorns sea stars in the genus
Acanthaster
(Høj et al. 2018).
The microbial community of the vertebrate gut also plays a crucial role in host-microbiome dynamics and
host health (Kinross et al. 2011; D’Argenio and Salvatore 2015; Shreiner et al. 2015). The same key roles
of microbes are likely to occur in marine invertebrate digestive systems. For example, Brothers et al.
(2018) found that chronic exposure to an elevated seawater temperature aligned with predicted near-
future ocean warming signicantly altered the presumptive functionality of the gut microbiome of the
common subtropical sea urchin
Lytechinus variegatus
. Importantly, Brothers et al. (2018) found that the
predicted microbial metagenomics impacted were those related to host physiological processes of
membrane transport and metabolism of amino acids and carbohydrates, suggesting that individuals are
likely to face physiological challenges as seawater temperatures continue to rise. Moreover,
L. variegatus
evaluated in this study were collected from the northern Gulf of Mexico (GOM) where population
migration northward to escape currently rising seawater temperature is not feasible. If the ultimate
outcome of changes in host microbiome structure and functionality is an increase in sub-lethal or lethal
impacts on ecologically important species such as sea urchins and sea stars, then there will be important
downstream ecological consequences on benthic community structure. Despite the potential signicance
of rapid anthropogenic climate warming impacting the gut microbiome of a model representative sea
urchin that plays a key role in seagrass ecology (Brothers et al. 2018), to date, no studies have similarly
examined temperature‐sensitivity of the gut (stomach) microbiome of a model sea star.
Luidia clathrata
is a key predatory, soft-bottom sea star in the family Luidiidae that has the appropriate
attributes to make it a model species for the study of the impacts of ocean warming on the stomach
Page 4/23
microbiome. The species is widely distributed and can be found in abundance in near‐shore habitats
throughout the GOM, as well as nearshore regions of the southeastern Atlantic coast of the U.S. north to
North Carolina, coastal environments of northeastern South America, and throughout the Caribbean
(Hendler 1995; Lawrence 2013; Lawrence et al. 2013). Its high abundance in shallow (< 40 m depth),
protected, bays along the northern GOM, a region projected to experience disproportionately high rates of
ocean warming and lacking northward refugia (Przeslawski et al. 2008). Accordingly,
L. clathrata
sampled from this region provide an excellent model for the examination of prospective climate‐induced
impacts of ocean warming.
In contrast to the complex intestinal tract of the sea urchin
L. variegatus
(Holland 2013), the digestive
morphology of
L. clathrata
is comparatively simple. Essentially the digestive system consists of the
mouth, cardiac stomach, and ve pyloric ceca, the latter used largely in nal digestive processes and
nutrient storage (Jangoux 1982). Unlike many sea stars that extrude the cardiac stomach from the
mouth, members of Luidiidae are generally considered to feed intraorally by depressing the oral disc into
the sediment and guiding prey into the mouth with the tube feet (McClintock and Lawrence 1981).
Interestingly, McClintock et al. (1983) observed extraoral feeding in
L. clathrata
via a modest extrusion of
the cardiac stomach both in the eld and under experimental laboratory conditions. In the latter case,
powdered food added to sand induced extraoral feeding. A capacity for extraoral feeding distinguishes
L.
clathrata
from other luidiid sea stars in that it allows for the exploitation of detrital food resources in
addition to infaunal and epifaunal prey.
Despite the established importance of the microbiome in host health and the presumption that the
impacts of climate change-induced temperature stress are likely to have negative impacts on aspects of
whole animal physiology and behavior, current knowledge of how near-future ocean warming may impact
host microbial community structure in marine invertebrates is largely lacking. The purpose of the present
study was to experimentally evaluate the potential effects of chronic exposure to predicted near-future
seawater temperatures on the stomach microbiome of the common sea star
L. clathrata
sampled from a
population in the northern GOM where there no refugia from rising seawater temperatures.
Material And Methods
Acquisition and Husbandry
Adult, non arm-regenerating
Luidia clathrata
(mean arm radius (R) ± SE = 65.2 ± 1.8 mm, n = 30) were
collected by standard otter trawl (1 cm mesh size) from Apalachee Bay, Florida (29°58’N, 84°19’W) in
early September, 2018. Immediately following collection, each individual was placed individually into a
plastic bag containing ambient seawater. The bags containing sea stars were placed into 80-L LoBoy©
Styrofoam coolers (10 bagged sea stars/cooler) and shipped overnight to the University of Alabama at
Birmingham (UAB). Each cooler was supplied with two 1-lb frozen gel ice packs (ChillPak©, Greiner Bio-
One International). Upon arrival at UAB, sea stars were transported to a climate-controlled room. Bags
containing individual sea stars and the seawater they were collected from were immediately oated in
Page 5/23
seawater in experimental tanks at a temperature of 25°C. The room was placed on a 12:12 h light‐dark
cycle for the duration of an adjustment period (see below) and throughout the subsequent experiment.
To facilitate gradual acclimatization to experimental tank conditions, the seawater in each plastic bag
containing a sea star was replaced with articial seawater (25°C, salinity 27; previously prepared with
Instant Ocean® mixed with reverse‐osmosis water ltered through a 4‐stage Aquafx Barracuda RO/DI
ltration system). Seawater replacements were carried out over a period of 3 h via a series of repeated
seawater exchanges. Following this adjustment period, each of the 30 individuals was randomly
assigned (using a random number generator in Microsoft Excel) to fteen 38‐L glass aquaria each
divided equally (each half = 25.4X12.7X30.5 cm) by a solid plexiglass plate that was sealed in place with
standard aquarium silicone sealant. For purposes of clarity, ‘half aquaria’ will hereafter be referred to as
‘separate aquaria’. Accordingly, the nal assignment of sea stars was one individual per aquarium.
Temperature was maintained in aquaria for each of three temperature treatments by submerging aquaria
in large, oval water baths (99X147X61 cm) consisting of 568-L Rubbermaid® structural foam stock pools
perched on cement blocks. Water temperature in each of the three water baths was controlled using a
submersible aquarium heater (EHEIM Jager 250 W TruTemp). Each aquarium was supplied with a 3-cm
deep layer of ne‐grain sand (Nature’s Ocean® Substrate, Marine White), a re‐circulating water pump
(Aqueon® Power Filter 10), and air line tubing with an air stone driven by a Coralife® Super Luft SL–65
high pressure aquarium pump.
Sea stars were held in their respective aquaria at 25°C for a two‐week holding period without food prior to
initiating the experiment. This allowed individuals to adjust to the aquaria and to standardize their
nutritional state. Throughout this two-week period and over the subsequent holding period and
experiments, water temperature, salinity, and dissolved oxygen were monitored daily using a YSI™ ProPlus
meter. Moreover, to evaluate and maintain water quality, nitrate, nitrite, ammonia, and pH levels were
measured weekly with an API™ Aquarium Pharmaceuticals liquid test kit. Large (1/3 aquarium volume),
water exchanges were conducted weekly, and smaller water exchanges were conducted as necessary.
Following the initial two-week holding period, the water temperature in all three water baths was slowly
elevated from 25 to 28°C over a period of 34 days. During this period, and during the subsequent three
temperature exposure treatments, each sea star was fed an
ad libitum
diet of formulated sea star feed
(formulation of feed provided by Addison Lawrence, Texas A&M University). Feed was proffered to each
individual every three days and food not consumed within a 14-h period was removed from each
aquarium by gloved hand. During this 34-day temperature adjustment period there were a small number
of sea stars in the aquaria that suffered mortality: one in one water bath, and two in each of the other two
water baths.
Prior to initiating experimental temperature treatments, three Time-Zero groups of two individuals from
each of the three water baths (n = 2X3 water baths = 6 sea stars) were randomly selected using the
random number generator in Microsoft Excel to establish a baseline microbial community. The cardiac
stomach of each sea star was excised by lifting the stomach tissue with sterile forceps and then cutting it
Page 6/23
free with sterile, dissecting scissors. Samples of sand and seawater were simultaneously collected from
each aquarium to evaluate their associated microbial communities as controls. The sand samples were
each collected by scraping sand into a sterile 50 mL falcon tube. Each sand sample was processed by
using a sterilized scoopula to collect approximately 1 g wet sand from the center of each falcon tube and
transferring it directly into a sterile Eppendorf tube for DNA extraction. Seawater samples were prepared
by vacuum ltering 60 mL of seawater through a sterile, 0.22-µm, Millipore membrane (EMD Millipore
Corporation). Stomach, sand, and seawater samples were all immediately frozen and stored at ‐20°C until
DNA extraction.
Over the following 14 days, water temperatures in two of the three water baths were slowly increased to a
projected mid-century (30°C) and end-of-century (32°C) ocean temperature based on predictions for the
northern GOM (IPCC, 2019). Individuals in the control (ambient) temperature treatment were maintained
at 28°C, the present-time mean July-September
in situ
temperature for the northern GOM (National
Centers for Environmental Information, NOAA). After this 14-day period, all surviving individuals (n = 7 of
9, 6 of 8, and 4 of 8 for the 28°C, 30°C, and 32°C temperature treatments, respectively) were dissected and
cardiac stomach tissue collected and stored as described above. Importantly, all surviving individuals
examined in each temperature treatment appeared healthy and were behaviorally vigorous. Sand and
seawater samples were re-collected at the end of the experiment as above for later microbiome DNA
extractions, along with three samples of the formulated sea star feed.
Microbial DNA Purication and Sequence Analysis
Microbial metacommunity DNA was extracted using a Zymo Research Fecal DNA isolation kit (Catalog #
D6010). An amplicon library was constructed with PCR using unique barcoded primers (Kozich et al.
2013; Kumar et al. 2014) to target the hyper variable 4 (V4) region of the 16S rRNA gene (forward primer
V4: 5′AATGATACGGCGACCACCGAGATCTACACTATGGTAATTGTGTGCCAGCMGCCGCGGTAA-3′; reverse
primer V4:5′-CAAGAGAAGACGGCATACGAGATNNNNNNAGTCAGTCAGCCGGACTACHVGGGTWTCTAAT-3)
(Eurons Genomics, Inc., Huntsville, AL). PCR was conducted with the New England Biolabs LongAmp
Taq PCR kit (Catalog # E5200S) under the conditions outlined in Kumar et al. (2014). PCR products were
then electrophoresed on a 1.0% agarose/TRIS-borate-EDTA gel, visualized by UV illumination, excised,
and puried with a Qiagen QIAquick Gel Extraction Kit (Catalog # 2870) (Kumar et al. 2014).
Puried PCR products were quantied with Pico Green dye, adjusted to a 4 nM concentration, and
sequenced on the NextGen sequencing Illumina MiSeq™ platform using an Illumina paired 250-bp kit
(Caporaso et al. 2012; Kozich et al. 2013; Kumar et al. 2014). Two samples were excluded from
subsequent analysis due to insucient PCR product volume, bringing nal sample numbers for the 28°C,
30°C, and 32°C treatments to (n = 6, 5, and 5, respectively).
The Quantitative Insights into Microbial Ecology package (QIIME2 v2019.10.0; (Bolyen et al. 2019) was
used for all downstream data analysis. Demultiplexed sequence data was checked for quality using the
DADA2 plugin in QIIME2 (Callahan et al. 2016). Due to a decrease in quality score after 250 nucleotides,
the rst 13 nucleotides for each forward and reverse read were trimmed and the total read length was
Page 7/23
truncated to 250 nucleotides. Chimeric variants were removed and amplicon sequence variants (ASVs)
were identied at a 97% similarity threshold according to the DADA2 pipeline (Callahan et al. 2016).
Taxonomic assignments were made using reference sequences from the SILVA 132 database (Quast et
al. 2012) trimmed to the V4 region with the 515f-806r primers. Representative sequences were classied
using the “classify-sklearn” function of the QIIME2 feature-classier plugin (Bokulich et al. 2018) and non-
target sequences (mitochondria, chloroplast, archaea, eukaryota, unknown) were removed prior to
analysis. The remaining sequences were subsampled to a maximum depth of 24,573 to account for
variation in read depth using the “alpha-rarefacation” analysis command in the diversity plugin for
QIIME2. Rarefaction curves for α‐diversity in each sample were generated using the “alpha-rarefaction”
command in the Qiime2 diversity plugin. The normalized, raried ASVs were used for all subsequent
analysis (Hakim et al. 2015; Brothers et al. 2018).
Statistical Analyses
The “core metrics” command in the diversity plugin for QIIME2 was used to calculate microbial diversity
of ASVs within each community (α‐diversity) and between communities (β‐diversity). Shannon’s index
values for α‐diversity were compared between time zero and 28°C treatments and across all temperature
treatments (28°C, 30°C, and 32°C) within sample types (cardiac stomach, sand, seawater, and feed) using
Kruskal-Wallis test with Benjamini & Hochberg correction via the “alpha-group-signicance” command in
the diversity plugin in for QIIME2. β‐diversity was calculated using Bray-Curtis dissimilarity values
command for cluster analysis and visualized with principal components analysis (PCoA) using the
“diversity core-metrics” command in the diversity plugin for QIIME2. Bray-Curtis dissimilarity values for
β‐diversity were compared for each sample type between Time Zero and 28°C treatments and across all
temperature treatments using permutational multivariate analysis of variance (PERMANOVA) via the
“beta-group-signicance” command in the diversity plugin for QIIME2. A p-value of <0.05 was considered
signicant for all analyses. A script of all QIIME2 commands used for this study is included as an online
resource (see Electronic Supplementary Material 1).
Results
A total of 6,280,565 16S V4 rRNA gene sequences were initially obtained from sequencing. After
denoising, quality ltering, and removing non-target sequences the remaining 3,663,121 sequences were
assigned to 3,578 ASVs. Rarefaction curves for α‐diversity in each sample leveled off and indicated that
all sampling was performed to a sucient depth to represent microbial community diversity. α‐diversity
(Shannon Index) was highest in feed, and lowest in stomach tissue overall. Generally, α‐diversity
increased with increasing temperature in sand, decreased in seawater, and decreased in stomach tissue
from 28°C to 30°C, with a sharp increase at 32°C (Table 1).
Page 8/23
Table 1 Shannon diversity values for all sample types across all temperature treatments. Note: feed
samples were taken once at the beginning of the experimental period and all feed was held under
constant conditions throughout the experiment.
Time
Zero
28°C 30°C 32°C
Feed 13.54 ± 0.45
Seawater 4.49 ± 0.95 4.18 ± 0.92 3.53 ± 0.30 3.48 ± 0.43
Sand 5.66 ± 1.43 5.90 ± 0.42 5.96 ± 0.46 6.59 ± 0.43
Stomach 1.70 ± 0.57 1.15 ± 0.70 0.88 ± 0.52 2.14 ± 1.23
Kruskal-Wallis pairwise comparisons of α‐diversity revealed no statistically signicant change in
microbial diversity of any sample type between time zero and 28°C or between any temperature treatment
(Table 2).
Table 2 Kruskal-Wallis pairwise signicance for seawater, sand, and stomach samples between all
temperature treatments. Kruskal-Wallis p-values are shown with Benjamini and Hochberg correction.
Time
Zero – 28°C
28°C – 30°C 28°C – 32°C 30°C – 32°C
Seawater 0.674 0.231 0.130 0.774
Sand 0.573 0.633 0.070 0.102
Stomach 0.140 0.518 0.231 0.072
Vibrio
sp., the most abundant ASV overall, represented 40% of the microbial community across all sample
types and treatments. In stomach tissue,
Vibrio
sp. made up 94% of the microbial community at 28°C and
30°C and decreased to 66% at 32°C (Fig. 1). Approximately 98% of the bacterial community in stomach
tissue at 28°C was made up of four bacterial taxa:
Vibrio
sp. (94.25%),
Carboxylicivirga
sp. (3.01%),
Delftia
sp. (0.36%), and
Halodesulfovibrio
sp. (0.31%) (Fig. 1). Similarly, over 98% of the bacterial
community of stomach tissue at 30°C was made up of three bacterial taxa:
Vibrio
sp. (94.40%),
Delftia
sp. (2.62%), and
Pesudomonas
sp. (1.39%) (Fig. 1). The top three bacterial taxa were the same between
30°C and 32°C, but
Vibrio
sp. abundance decreased,
Delftia
sp. and
Pseudomonas
sp. abundance
increased, and the top ten most abundant taxa only made up approximately 94% of the total stomach
tissue bacterial community (Fig. 1). The most abundant taxa in the bacterial community of stomach
tissue at 32°C included:
Vibrio
sp. (66.34%),
Delftia
sp. (11.46%),
Pseudomonas
sp. (4.67%),
Halodesulfovibrio
sp. (3.99%),
Bacteroides thetaiotaomicron
(2.69%),
Bacteroides
sp. (1.73%),
Lachnospiraceae
NK4A136 group (1.07%),
Clostridium butyricum
(0.62%),
Phyllobacterium
sp. (0.55%),
and an unidentied member of the kingdom Bacteria (0.37%) (Fig. 1).
Page 9/23
The seawater bacterial community was primarily dominated by
Delftia
sp. and
Pseudomonas
sp., both of
which increased in abundance from 28°C (29.86% and 18.33%, respectively), to 30°C (35.54% and
22.86%, respectively), to 32°C (39.25% and 25.17%, respectively) (Fig. 1).
Phyllobacterium
sp. and
Methylobacterium
sp. were also present with increasing abundance with warming (3.25% and 1.58%,
respectively at 28°C, 4.35% and 2.14%, respectively at 30°C, and 4.77% and 2.60%, respectively at 32°C)
(Fig. 1). The most abundant bacterial taxa in sand at 28°C were
Vibrio
sp. (15.42%),
Arcobacter
sp.
(6.42%),
Halodesulfovibrio
sp. (5.92%),
Delftia
sp. (5.28%),
Photobacterium
sp. (4.37%), and
Motiliproteus
sp. (3.18%) (Fig. 1).
Vibrio
sp. was also the most abundant taxa in sand at 30°C (10.91%), followed by
Delftia
sp. (6.67%),
Photobacterium
sp. (6.33%),
Propioigenium
sp. (4.85%),
Pseudomonas
sp. (4.27%),
and
Shewanella
sp. (3.16%) (Fig. 1).
Photobacterium
sp. and
Vibrio
sp. were also among the top taxa in
the sand bacterial community at 32°C (5.95% and 3.63%, respectively), but the most abundant taxa at
this temperature was
Motiliproteus
sp. (9.07%) (Fig. 1).
Principal components analysis of Bray-Curtis dissimilarity values showed distinct clustering of microbial
community composition by sample type (Fig. 2). Permutational multivariate analysis of variance
(PERMANOVA) of Bray-Curtis dissimilarity values revealed signicantly different microbial community
composition between all sample types (p = 0.001, stomach–sand, stomach–seawater, stomach–feed,
sand–seawater, and sand–feed) except for seawater–feed (p = 0.152). Statistically signicant changes
were observed in microbial community composition between time zero and 28°C for sand and seawater,
but not for stomach tissue (PERMANOVA; sand [F = 1.855, q = 0.036, p = 0.030], seawater [F = 3.305, q =
0.009, p = 0.003], and stomach [F = 0.710, q = 0.731, p = 0.731]).
Importantly, comparison of Bray-Curtis dissimilarity values revealed signicant changes in stomach
tissue microbial community composition associated with warming between 28°C and 32°C
(PERMANOVA, F = 2.703, q = 0.045, p = 0.039), but not between 28°C and 30°C (PERMANOVA, F = 1.066,
q = 0.322, p = 0.318) or between 30°C and 32°C (PERMANOVA, F = 2.217, q = 0.123, p = 0.115).
Signicant changes in sand microbial community composition were also detected with warming between
28°C and 32°C and between 30°C and 32°C, but not between 28°C and 30°C (PERMANOVA; 28°C–32°C [F
= 2.621, q = 0.020, p = 0.013], 30–32°C [F = 1.874, q = 0.017, p = 0.010], and 28°C–30°C [F = 1.170, q =
0.216, p = 0.208]). No signicant difference in microbial community composition of seawater was
observed between any of the three temperature treatments (PERMANOVA; 28°C–30°C [F = 1.143, q =
0.252, p = 0.246], 28°C–32°C [F = 1.571, q = 0.115, p = 106], and 30–32°C [F = 1.235, q = 0.097, p =
0.087]).
Discussion
Microbial diversity detected in seawater, sand, and articial food in the present study were all higher than
that detected in the tissues of the cardiac stomach of the star
Luidia clathrata.
This was particularly
evident at experimental seawater temperatures of 28°C and 30°C. The highest levels of microbial diversity
occurred in seawater and sand (Table 1). This low level of microbial diversity in the stomach tissues of
L
clathrata
aligns well with other studies of echinoderms which have reported a generally low microbial
Page 10/23
diversity in stomach and digestive tissues of sea urchins (Hakim et al. 2016; Brothers et al. 2018), sea
cucumbers (Gao et al. 2014) and sea stars (Jackson et al. 2018). It is also important to note that, similar
to the present study, microbial communities characterized from various tissues of 12 species of sea stars
have been reported to be distinct from microbial communities of surrounding seawater (Jackson et al.
2018). In the present study, given that extraoral feeding is a common mode of feeding of
L. clathrata
under laboratory conditions (McClintock et al. 1983) and was also observed throughout the present study,
it is not unexpected that the composition of the microbial community in stomach tissue was more closely
aligned with that of the sand rather than seawater.
In the present study, the dominant class of bacteria making up the bulk of the microbial communities
across all sample types in all three experimental temperature treatments was Gammaproteobacteria. The
order Vibrionales dominated the stomach microbial community across all temperatures, while
Betaproteobacteriales and Pseudomonadales had the highest abundance in seawater across all
temperatures. Samples of aquarium sand had the highest diversity with Vibrionales and
Betaproteobacteriales being the two most abundant taxa.
The presence of
Vibrio
sp. as the primary component of the stomach microbiome in
L. clathrata
across
all three temperature treatments is unique for a sea star. Although high proportion of
Vibrio
spp. have
been reported in the intestine of the sea cucumber
Holothuria glaberrima
(Pagán-Jiménez et al. 2019), the
ophiuroid
Ophionema
sp. (Dilmore and Hood 1986), and the sea urchins
Strongylocentrotus
droebachiensis
and
Tripneustes ventricuosus
(Guerinot and Patriquin 1981), we believe ours is the rst
study to report this pattern in a sea star. Høj et al. (2018) reported small amounts
Vibrio
sp. in the
microbial community of the pyloric cecum of the tropical corallivorous sea star
Acanthaster plancii.
They
also reported a signicantly higher incidence of Vibrionaceae-related taxa in diseased individuals than in
healthy individuals. Jackson et al. (2018) did not detect
Vibrio
sp., but reported
Tenericutes
as the most
abundant microbial taxa in the pyloric caeca of the sea stars
Pycnopoida helanthoides, Crossaster
papposus, Orthasterias koehleri, Pteraster tesselatus, Linckia guildingii, Pentaceraster
spp., and
Linckia
laevigata
from the coasts of Washington, USA and Queensland, Australia. A 10- to 20-fold increase in
Vibrio
sp. and
Alteromonas
sp. was observed in the microbial communities of food pellets ingested by
the sea urchin
Lytechinus variegatus
held chronically under conditions of near-future warming (30°C)
(Brothers et al. 2018). An opposite trend was observed in the present study, as abundance of
Vibrio
sp.
declined in stomach tissues of
L. clathrata
chronically exposed to an end-of-century warming
temperature of 32°C.
High proportion of bacteria from the Vibrionaceae family is common in gut and stomach tissues of
mussels and marine arthropods (Preheim et al. 2011), particularly in crabs (Huq et al. 1986). Additionally,
marine copepods reared in an aquarium setting have been shown to have a higher abundance of
Vibrio
in
their gut than copepods from the natural environment (Sochard et al. 1979). Food availability has been
reported to affect the gut microbiome of the shrimp
Penaeus monodon
with higher abundance of
Vibrio
in the gut of individuals fed to those fed a restricted diet (Simon et al. 2020). Furthermore, Lokmer and
Wegner (2015) found thermal stress, but not exposure to
Vibrio
infection, signicantly altered the
Page 11/23
microbiome of the hemolymph of the temperate pacic oyster
Crassostrea gigas
. As some
Vibrio
spp. are
typically heat-resistant, rapidly growing opportunists (Bourne et al. 2016), it will be interesting to further
examine the role of this taxa in the stomach microbiome of luidiid and non-luidiid sea stars to determine
if their presence is indicative of disease or stress, or if
Vibrio
spp. are common members of gut bacterial
reservoir communities as seen in other echinoderms and invertebrates more generally (Webster and
Thomas 2016; Jackson et al. 2018).
The shift in the stomach microbial community composition of
L. clathrata
from ~94%
Vibrio
sp.
dominated (as observed in the 28°C and 30°C treatments) to higher proportions of
Delftia
sp. and
Pseudomonas
sp. in the highest temperature treatment (32°C) is noteworthy. This signicant shift in the
composition of the stomach’s microbial community at the predicted end-of-century temperature for the
northern GOM is consistent with that observed in the sea urchin
Lytechinus variegatus
(Brothers et al.
2018). However, the temperature induced trends for the major microbial taxa present are reversed in these
two echinoderms. Brothers et al. (2018) reported that under conditions of near-future warming, the
proportion of
Vibrio
sp. in the gut of
L. variegatus
increased relative to the other microbial taxa. In the
present study, the opposite trend in
Vibrio
sp. occurred with relative abundance trending down with
increasing temperature.
Pseudomonas
is known to possess antibacterial activity (Jayatilake et al. 1996)
with one strain reportedly having inhibitory properties against marine pathogenic
Vibrio
(Chythanya et al.
2002). It is unknown whether the shifts in the microbial communities we observed in the stomach of
L.
clathrata
held at elevated temperature are reversible with a reduction in temperature. However, there is
evidence this may not be the case. For example, Ramsby et al. (2018) reported a disruption in the
microbial community of the tropical sponge
Cliona orientalis
at 29°C and bleaching of
Symbiodinium
algae at 32°C, were both irreversible when the sponge was returned to its natural temperature of 23°C.
Microbial community composition of the tissues of marine invertebrates are critically important both in
the context of resistance to disease (Egan and Gardiner 2016; Lloyd and Pespeni 2018). Gudenkauf and
Hewson (2015) reported high proportions of both
Vibrio
sp. and
Pseudomonas
sp. in body-wall tissues of
the Pacic coast sea star
Pycnopodia helianthoides
infected with sea star wasting disease. In contrast,
these same microbial taxa were reported in lower proportions in the body-wall tissues of the Pacic
keystone sea star
Pisaster ochraceus
(Paine 1968) over the documented progression of sea star wasting
disease (Lloyd and Pespeni 2018). Gudenkauf and Hewson (2015) suggested that
Vibrio
sp.
and
Pseudomonas
sp. might represent an active portion of the holobiont that increases rapidly in proportion
when host tissues are compromised.
Delftia,
a genus of gram-negative facultative opportunistic pathogen
in the family Comamonadaceae, is known to inhabit marine environments. To date,
Delftia
has not been
reported in sea stars, although this group of gram+ bacteria has been detected in a non-pathogenic form
in adults of the hermatypic coral
Acropora gemmifera
(Zhou et al. 2017). Additionally,
Delftia acidovorans
has been shown to act as an opportunistic pathogen in European eels held in captivity, co-infecting the
eels along with
Pseudomonas anguilliseptica
(Andree et al. 2013).
In addition to considering the effect of stress-induced gut microbiome modication on overall health, it is
also important to consider the potential effects on nutrient processing and metabolism. Brothers et al.
Page 12/23
(2018) detected a signicant shift in predicted metabolic and membrane transport functions in the gut
microbiome under elevated temperature stress. This observation underlies the importance of
understanding how climate change-induced stressors such as ocean warming will affect the gut
microbiome and its associated functions. Although little is yet known about the role of the digestive
system microbial community in the processes of digestion and metabolism in sea stars, symbiotic gut
bacteria are known to play a role in amino acid provision for some terrestrial invertebrates including the
potato psyllids (Thao et al. 2000; Arp et al. 2014) and enchytraeid worms (Larsen et al. 2016).
While the present study focused on adult sea stars, it is noteworthy that the microbial community of sea
star larvae is distinct from that of the external environment and can display shifts with dietary changes
associated with larval growth and development (Carrier et al. 2018). Bipinnaria larvae of
Luidia
sp.
collected from plankton samples in the Northern Atlantic displayed a well-established microbial
community with the majority of bacteria isolated to the developing gastric region (Bosch 1992). Bosch
(1992) surmised that these symbiotic bacteria were likely associated with digestive processes in the
developing larvae. To date, the microbiome of bipinnaria and brachiolaria larvae of
L. clathrata
is
unknown. George et al. (1990) suggested that bacteria, in addition to dissolved nutrients and vitellogenic
reserves, might have played a role in nutrition of early development of
L. clathrata
bipinnaria larvae
during their laboratory-based experiment. The ndings of Bosch (1992) and George et al. (1990) warrant
further exploration of the community structure and functional role of the microbiome across larval,
juvenile, and adult life history phases in sea star development.
The simple anatomical structure and basal phyletic nature of the genus
Luidia
provides a unique
opportunity for developing an understanding of gut microbiome dynamics in sea stars. Although
members of
Luidia
lack the pyloric stomach found in higher sea star taxa, in addition to predatory
foraging
L. clathrata
exhibits extraoral feeding (McClintock et al. 1983). This behavior is unique among
the Paxillosid sea stars, affording
L. clathrata
the potential advantage of exploiting detrital food
resources in addition to infauna and epifauna. Moreover, it suggests a possible evolutionary link between
luidiid sea stars and higher taxa with more complex digestive anatomy (McClintock et al. 1983). Whether
the microbial community associated with the gut of
L. clatharata
is also representative of an evolutionary
branch between the guts of intraoral and extraoral sea stars remains to be determined.
As has been generally reported in the literature to date, gut microbial communities in healthy individuals
under non-stressful conditions have been reported to be relatively stable. Lloyd and Pespeni (2018)
reported disruption of microbial community stability and decrease in microbial richness in the epidermal
body wall of the sea star
Pisaster ochraceus
with progression of sea star wasting disease. This stability
was similarly observed in the gut microbial community of the sea urchin
Lytechinus variegatus
exposed
to near-future ocean warming (Brothers et al. 2018). Stomach bacterial community structure in
L.
clathrata
in the present study was altered by exposure to projected near-future warming. This is
consistent with the presumption stress allows for disruption of microbial community structure as
opportunistic pathogenic bacteria ourish at the expense of previously stable communities comprised of
a few bacterial taxa. Stressors such as increased seawater temperature are known to disrupt microbial
Page 13/23
community structure in marine environments and organisms by either increasing or decreasing overall
diversity (e.g., Brothers et al. 2018; Li et al. 2018; Ramsby et al. 2018; Alma et al. 2020; Botté et al. 2020;
Greenspan et al. 2020).
Conclusions
In the present study we report the rst account of a sea star gut microbiome community reacting to
projected ocean warming in a common sea star. Moreover, this is the rst description of the gut
microbiome in a sea star from a primitive Asteroid (family Luidiidae). The microbial community of the
tissues of the cardiac stomach of the common soft bottom sea star
Luidia clathrata
was determined to
be distinct from that of formulated feed, sand, and seawater that to which sea stars in the experimental
study were exposed. Of these four sample types, the stomach microbial community had the lowest
overall diversity, with the order Vibrionales,
Vibrio
in particular, dominating the microbiome across all
three temperature treatments (28, 30, 32°C). The shift in stomach microbiome to include higher
proportions of
Delftia
and
Pseudomonas
at 32°C is of note as
Vibrio, Delftia,
and
Pseudomonas
can all
be opportunistic pathogens with some
Pseudomonas
having activity against some marine pathogenic
Vibrio.
Further studies should be conducted to examine the microbial communities of
Luidia
and other
sea stars from the natural environment to see if
Vibrio
-domination is common in natural microbial
communities to determine how much the base community observed in this study might have been
inuenced by acclimatization to aquarium conditions.
Declarations
Compliance with Ethical Standards
Ethics approval Not applicable.
Consent to participate Not applicable.
Consent for publication Not applicable.
Acknowledgments
We thank Sabrina Heiser for her valuable assistance with sample processing. We also thank Dr. CJ
Brothers, Liam Van Der Pol, Dr. Guido Bonthond, and Dr. Stacy Kruger-Hadeld for helpful discussion of
experimental preparation and data analysis methods. We are grateful to Dr. Charles Amsler and Maggie
Amsler for use of laboratory facilities and equipment. We thank the reviewers for their suggestions.This
research was funded by an Endowed University Professorship in Polar and Marine Biology to Dr. James
McClintock.
Funding
Page 14/23
This research was funded by an Endowed University Professorship in Polar and Marine Biology to Dr.
James McClintock.
Conicts of interest/Competing interests
The authors have no conicts of interest or competing interests to declare.
Data/Code availability
Data will be provided upon reasonable request to the corresponding author [MDC]. A document outlining
the code used in analysis of the data for this project has been included as an electronic supplementary
le.
Authors' contributions
MDC and JBM conceived the study. MDC conducted the experiments, analyzed the data, and provided the
early draft of the manuscript. CDM oversaw and facilitated the microbiome sample processing and data
generation. MDC, JBM, and CDM worked together on later drafts to develop this manuscript.
References
Alma L, Kram KE, Holtgrieve GW, Barbarino A, Fiamengo CJ, Padilla-Gamiño JL (2020) Ocean
acidication and warming effects on the physiology, skeletal properties, and microbiome of the purple-
hinge rock scallop. Comp Biochem Physiol A Mol Integr Physiol 240:110579.
https://doi.org/10.1016/j.cbpa.2019.110579
Andree KB, Rodgers CJ, Furones D, Gisbert E (2013) Co-infection with
Pseudomonas anguilliseptica
and
Delftia acidovorans
in the European eel,
Anguilla anguilla
(L.): A case history of an illegally tracked
protected species. J Fish Dis 36(7):647–656. https://doi.org/10.1111/jfd.12066
Arp A, Munyaneza JE, Crosslin JM, Trumble J, Bextine B (2014) A global comparison of
Bactericera
cockerelli
(Hemiptera: Triozidae) microbial communities. Environ Entomol 43(2):344–352.
https://doi.org/10.1603/EN13256
Barker MF, Scheibling RE (2008) Rates of ssion, somatic growth and gonadal development of a
ssiparous sea star,
Allostichaster insignis
, in New Zealand. Mar Biol 153:815–824.
https://doi.org/10.1007/s00227-007-0854-2
Barker MF, Scheibling R, Mladenov P (1992) Seasonal changes in population structure of the ssiparous
asteroids
Allostichaster insignis
(Farquhar) and
Coscinasterias calamaria
(Gray). In: Scalera-Liaci L,
Canicatti C (eds) Echinoderm Research 1991. AA Balkema, Rotterdam, Netherlands, pp 191–195
Bates AE, Hilton BJ, Harley CDG (2009) Effects of temperature, season and locality on wasting disease in
the keystone predatory sea star
Pisaster ochraceus
. Dis Aquat Organ 86:245–251.
Page 15/23
https://doi.org/10.3354/dao02125
Bokulich NA, Kaehler BD, Rideout JR, Dillon M, Bolyen E, Knight R, Huttley GA, Gregory Caporaso J (2018)
Optimizing taxonomic classication of marker-gene amplicon sequences with QIIME 2’s q2-feature-
classier plugin. Microbiome 6:90. https://doi.org/10.1186/s40168-018-0470-z
Bolyen E, Rideout JR, Dillon MR, Bokulich NA, Abnet CC, Al-Ghalith GA, Alexander H, Alm EJ, Arumugam
M, Asnicar F, Bai Y, Bisanz JE, Bittinger K, Brejnrod A, Brislawn CJ, Brown CT, Callahan BJ, Caraballo-
Rodríguez AM, Chase J, Cope EK, Da Silva R, Diener C, Dorrestein PC, Douglas GM, Durall DM, Duvallet C,
Edwardson CF, Ernst M, Estaki M, Fouquier J, Gauglitz JM, Gibbons SM, Gibson DL, Gonzalez A, Gorlick K,
Guo J, Hillmann B, Holmes S, Holste H, Huttenhower C, Huttley GA, Janssen S, Jarmusch AK, Jiang L,
Kaehler BD, Kang KB, Keefe CR, Keim P, Kelley ST, Knights D, Koester I, Kosciolek T, Kreps J, Langille MGI,
Lee J, Ley R, Liu Y-X, Lofteld E, Lozupone C, Maher M, Marotz C, Martin BD, McDonald D, McIver LJ,
Melnik AV, Metcalf JL, Morgan SC, Morton JT, Naimey AT, Navas-Molina JA, Nothias LF, Orchanian SB,
Pearson T, Peoples SL, Petras D, Preuss ML, Pruesse E, Rasmussen LB, Rivers A, Robeson MS, Rosenthal
P, Segata N, Shaffer M, Shiffer A, Sinha R, Song SJ, Spear JR, Swafford AD, Thompson LR, Torres PJ,
Trinh P, Tripathi A, Turnbaugh PJ, Ul-Hasan S, van der Hooft JJJ, Vargas F, Vázquez-Baeza Y, Vogtmann E,
von Hippel M, Walters W, Wan Y, Wang M, Warren J, Weber KC, Williamson CHD, Willis AD, Xu ZZ,
Zaneveld JR, Zhang Y, Zhu Q, Knight R, Caporaso JG (2019) Reproducible, interactive, scalable and
extensible microbiome data science using QIIME 2. Nat Biotechnol 37:852–857.
https://doi.org/10.1038/s41587-019-0209-9
Bosch I (1992) Symbiosis between bacteria and oceanic clonal sea star larvae in the western North
Atlantic Ocean. Mar Biol 114:495–502. https://doi.org/10.1007/BF00350041
Botté ES, Luter HM, Marangon E, Patel F, Uthicke S, Webster NS (2020) Simulated future conditions of
ocean warming and acidication disrupt the microbiome of the calcifying foraminifera
Marginopora
vertebralis
across life stages. Environ Microbiol Rep 12(6):693–701. https://doi.org/10.1111/1758-
2229.12900
Bourne DG, Morrow KM, Webster NS (2016) Insights into the coral microbiome: Underpinning the health
and resilience of reef ecosystems. Annu Rev Microbiol 70:317–340. https://doi.org/10.1146/annurev-
micro-102215-095440
Brothers CJ, Van Der Pol WJ, Morrow CD, Hakim JA, Koo H, McClintock JB (2018) Ocean warming alters
predicted microbiome functionality in a common sea urchin. Proc R Soc B Biol Sci 285:20180340.
https://doi.org/10.1098/rspb.2018.0340
Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJA, Holmes SP (2016) DADA2: High-resolution
sample inference from Illumina amplicon data. Nat Methods 13:581–583.
https://doi.org/10.1038/nmeth.3869
Page 16/23
Caporaso JG, Lauber CL, Walters WA, Berg-Lyons D, Huntley J, Fierer N, Owens SM, Betley J, Fraser L,
Bauer M, Gormley N, Gilbert JA, Smith G, Knight R (2012) Ultra-high-throughput microbial community
analysis on the Illumina HiSeq and MiSeq platforms. ISME J 6:1621–1624.
https://doi.org/10.1038/ismej.2012.8
Carrier TJ, Wolfe K, Lopez K, Gall M, Janies DA, Byrne M, Reitzel AM (2018) Diet-induced shifts in the
crown-of-thorns (
Acanthaster
sp.) larval microbiome. Mar Biol 165:157. https://doi.org/10.1007/s00227-
018-3416-x
Chythanya R, Karunasagar I, Karunasagar I (2002) Inhibition of shrimp pathogenic vibrios by a marine
Pseudomonas
I-2 strain. Aquaculture 208(1–2):1–10. https://doi.org/10.1016/S0044-8486(01)00714-1
D’Argenio V, Salvatore F (2015) The role of the gut microbiome in the healthy adult status. Clin Chim Acta
451(A):97–102. https://doi.org/10.1016/j.cca.2015.01.003
Dilmore LA, Hood MA (1986) Vibrios of some deep-water invertebrates. FEMS Microbiol Lett 35(2–
3):221–224. https://doi.org/10.1111/j.1574-6968.1986.tb01531.x
Eckert GL, Engle JM, Kushner DJ (1999) Sea star disease and population declines at the Channel Islands.
Proc Fifth Channel Isl Symp 390–393
Egan S, Gardiner M (2016) Microbial dysbiosis: Rethinking disease in marine ecosystems. Front Microbiol
7:991. https://doi.org/10.3389/fmicb.2016.00991
Eisenlord ME, Groner ML, Yoshioka RM, Elliott J, Maynard J, Fradkin S, Turner M, Pyne K, Rivlin N, van
Hooidonk R, Harvell CD (2016) Ochre star mortality during the 2014 wasting disease epizootic: Role of
population size structure and temperature. Philos Trans R Soc B Biol Sci 371:20150212.
https://doi.org/10.1098/rstb.2015.0212
Fly EK, Monaco CJ, Pincebourde S, Tullis A (2012) The inuence of intertidal location and temperature on
the metabolic cost of emersion in
Pisaster ochraceus
. J Exp Mar Biol Ecol 422–423:20–28.
https://doi.org/10.1016/j.jembe.2012.04.007
Foster MC, Byrnes JEK, Reed DC (2015) Effects of ve southern California macroalgal diets on
consumption, growth, and gonad weight, in the purple sea urchin
Strongylocentrotus purpuratus
. PeerJ
3:e719. https://doi.org/10.7717/peerj.719
Gao F, Li F, Tan J, Yan J, Sun H (2014) Bacterial community composition in the gut content and ambient
sediment of sea cucumber
Apostichopus japonicus
revealed by 16S rRNA gene pyrosequencing. PLoS
ONE 9(6):e100092. https://doi.org/10.1371/journal.pone.0100092
George S, Lawrence J, Fenaux L (1990) Effect of starvation and the time to rst feeding on larvae of
Luidia clathrata
. In: De Rider, Dubois, Lahaye, Jangoux (eds) Echinoderm research. AA Balkema,
Rotterdam, Netherlands, pp 91–98
Page 17/23
Gomez A, Espinoza JL, Harkins DM, Leong P, Saffery R, Bockmann M, Torralba M, Kuelbs C, Kodukula R,
Inman J, Hughes T, Craig JM, Highlander SK, Jones MB, Dupont CL, Nelson KE (2017) Host genetic
control of the oral microbiome in health and disease. Cell Host Microbe 22(3):269–278.e3.
https://doi.org/10.1016/j.chom.2017.08.013
Gooding RA, Harley CD, Tang E (2009) Elevated water temperature and carbon dioxide concentration
increase the growth of a keystone echinoderm. Proc Natl Acad Sci 106(23):9316–9321.
https://doi.org/10.1073/pnas.0811143106
Greenspan SE, Migliorini GH, Lyra ML, Pontes MR, Carvalho T, Ribeiro LP, Moura-Campos D, Haddad CFB,
Toledo LF, Romero GQ, Becker CG (2020) Warming drives ecological community changes linked to host-
associated microbiome dysbiosis. Nat Clim Change 10:1057–1061. https://doi.org/10.1038/s41558-020-
0899-5
Gudenkauf BM, Hewson I (2015) Metatranscriptomic analysis of
Pycnopodia helianthoides
(Asteroidea)
affected by sea star wasting disease. PLoS ONE 10(5):e0128150.
https://doi.org/10.1371/journal.pone.0128150
Guerinot ML, Patriquin DG (1981) N2-xing vibrios isolated from the gastrointestinal tract of sea urchins.
Can J Microbiol 27(3):311–317. https://doi.org/10.1139/m81-048
Hakim JA, Koo H, Kumar R, Lefkowitz EJ, Morrow CD, Powell ML, Watts SA, Bej AK (2016) The gut
microbiome of the sea urchin,
Lytechinus variegatus
, from its natural habitat demonstrates selective
attributes of microbial taxa and predictive metabolic proles. FEMS Microbiol Ecol 92(9):w146.
https://doi.org/10.1093/femsec/w146
Haramoto S, Komatsu M, Yamazaki Y (2007) Patterns of asexual reproduction in the ssiparous seastar
Coscinasterias acutispina
(Asteroidea: Echinodermata) in Japan. Zoolog Sci 24(11):1075–1081.
https://doi.org/10.2108/zsj.24.1075
Harvell CD, Mitchell CE, Ward JR, Altizer S, Dobson AP, Ostfeld RS, Samuel MD (2002) Climate warming
and disease risks for terrestrial and marine biota. Science 296:2158–2162.
https://doi.org/10.1126/science.1063699
Harvell CD, Montecino-Latorre D, Caldwell JM, Burt JM, Bosley K, Keller A, Heron SF, Salomon AK, Lee L,
Pontier O, Pattengill-Semmens C, Gaydos JK (2019) Disease epidemic and a marine heat wave are
associated with the continental-scale collapse of a pivotal predator (
Pycnopodia helianthoides
). Sci Adv
5:eaau7042. https://doi.org/10.1126/sciadv.aau7042
Hendler G, Miller JE, Pawson DL, Kier PM (1995) Sea stars, sea urchins, and allies: Echinoderms of
Florida and the Caribbean. Smithsonian Institution Press, Washington
Page 18/23
Høj L, Levy N, Baillie BK, Clode PL, Strohmaier RC, Siboni N, Webster NS, Uthicke S, Bourne DG (2018)
Crown-of-thorns sea star
Acanthaster
cf.
solaris
has tissue-characteristic microbiomes with potential
roles in health and reproduction. Appl Environ Microbiol 84(13):e00181–18,
https://doi.org/10.1128/AEM.00181-18
Holland ND (2013) Digestive system in regular sea urchins. In: Lawrence JM (ed) Sea urchins: Biology
and ecology, 3rd edn. Elsevier, London, pp 119–133
Holmes E, Li JV, Athanasiou T, Ashraan H, Nicholson JK (2011) Understanding the role of gut
microbiome–host metabolic signal disruption in health and disease. Trends Microbiol 19(7):349–359.
https://doi.org/10.1016/j.tim.2011.05.006
Huq A, Huq SA, Grimes DJ, O’Brien M, Chu KH, Capuzzo JM, Colwell RR (1986) Colonization of the gut of
the blue crab (
Callinectes sapidus
) by
Vibrio cholerae
. Appl Environ Microbiol 52(3):586–588.
https://doi.org/10.1128/AEM.52.3.586-588.1986
Jackson EW, Pepe-Ranney C, Debenport SJ, Buckley DH, Hewson I (2018) The microbial landscape of sea
stars and the anatomical and interspecies variability of their microbiome. Front Microbiol 9:1829.
https://doi.org/10.3389/fmicb.2018.01829
Jangoux M (1982) Food and feeding mechanisms: Asteroidea. In: Jangoux M, Lawrence JM (eds)
Echinoderm nutrition. AA Balkema, Rotterdam, Netherlands, pp 117–159
Jayatilake GS, Thornton MP, Leonard AC, Grimwade JE, Baker BJ (1996) Metabolites from an Antarctic
Sponge-Associated Bacterium,
Pseudomonas aeruginosa
. J Nat Prod 59(3):293–296.
https://doi.org/10.1021/np960095b
Kinross JM, Darzi AW, Nicholson JK (2011) Gut microbiome-host interactions in health and disease.
Genome Med 3:14. https://doi.org/10.1186/gm228
Kohl WT, McClure TI, Miner BG (2016) Decreased temperature facilitates short-term sea star wasting
disease survival in the keystone intertidal sea star
Pisaster ochraceus
. PLoS ONE 11:e0153670.
https://doi.org/10.1371/journal.pone.0153670
Kozich JJ, Westcott SL, Baxter NT, Highlander SK, Schloss PD (2013) Development of a dual-index
sequencing strategy and curation pipeline for analyzing amplicon sequence data on the MiSeq Illumina
sequencing platform. Appl Environ Microbiol 79:5112–5120. https://doi.org/10.1128/AEM.01043-13
Kumar R, Eipers P, Little RB, Crowley M, Crossman DK, Lefkowitz EJ, Morrow CD (2014) Getting started
with microbiome analysis: Sample acquisition to bioinformatics. Curr Protoc Hum Genet 14(82):18.8.1–
29. https://doi.org/10.1002/0471142905.hg1808s82
Page 19/23
Larsen T, Ventura M, Maraldo K, Triadó-Margarit X, Casamayor EO, Wang YV, Andersen N, O’Brien DM
(2016) The dominant detritus-feeding invertebrate in Arctic peat soils derives its essential amino acids
from gut symbionts. J Anim Ecol 85(5):1275–1285. https://doi.org/10.1111/1365-2656.12563
Lawrence JM (2013) Starsh: Biology and ecology of the Asteroidea. JHU Press, Baltimore
Lawrence JM, Durán-González A, Solís-Marín FA, Herrera J, Ventura CRR (2013) Distribution of
Luidia
clathrata
and
Luidia lawrencei
(Echinodermata: Asteroidea) along the coast of the western Atlantic
Ocean, the Gulf of Mexico and the Caribbean Sea. Cah Biol Mar 54:525–529.
Li Y-F, Yang N, Liang X, Yoshida A, Osatomi K, Power D, Batista FM, Yang J-L (2018) Elevated seawater
temperatures decrease microbial diversity in the gut of
Mytilus coruscus
. Front Physiol 9:839.
https://doi.org/10.3389/fphys.2018.00839
Lloyd MM, Pespeni MH (2018) Microbiome shifts with onset and progression of sea star wasting disease
revealed through time course sampling. Sci Rep 8:16476. https://doi.org/10.1038/s41598-018-34697-w
Lokmer A, Mathias Wegner K (2015) Hemolymph microbiome of Pacic oysters in response to
temperature, temperature stress and infection. ISME J 9:670–682.
https://doi.org/10.1038/ismej.2014.160
Lynch JB, Hsiao EY (2019) Microbiomes as sources of emergent host phenotypes. Science
365(6460):1405–1409. https://doi.org/10.1126/science.aay0240
McClintock JB, Klinger TS, Lawrence JM (1983) Extraoral feeding in
Luidia clathrata
(Say)
(Echinodermata: Asteroidea). B Mar Sci 33(1):171–172
McClintock JB, Lawrence JM (1981) An optimization study on the feeding behavior of
Luldia clathrata
say (Echinodermata: Asteroidea). Mar Behav Physiol 7(4):263–275.
https://doi.org/10.1080/10236248109386987
Monaco CJ, Wethey DS, Helmuth B (2014) A dynamic energy budget (DEB) model for the keystone
predator
Pisaster ochraceus
. PLoS ONE 9:e104658. https://doi.org/10.1371/journal.pone.0104658
Pagán-Jiménez M, Ruiz-Calderón JF, Dominguez-Bello MG, García-Arrarás JE (2019) Characterization of
the intestinal microbiota of the sea cucumber
Holothuria glaberrima
. PLoS ONE 14:e0208011.
https://doi.org/10.1371/journal.pone.0208011
Parmesan C (2006) Ecological and evolutionary responses to recent climate change. Annu Rev Ecol Evol
Syst 37:637–669. https://doi.org/10.1146/annurev.ecolsys.37.091305.110100
Peck LS, Webb KE, Miller A, Clark MS, Hill T (2008) Temperature limits to activity, feeding and metabolism
in the Antarctic starsh
Odontaster validus
. Mar Ecol Prog Ser 358:181–189.
https://doi.org/10.3354/meps07336
Page 20/23
Pörtner HO (2002) Climate variations and the physiological basis of temperature dependent
biogeography: Systemic to molecular hierarchy of thermal tolerance in animals. Comp Biochem Physiol A
Mol Integr Physiol 132:739–761. https://doi.org/10.1016/S1095-6433(02)00045-4
Pörtner HO (2008) Ecosystem effects of ocean acidication in times of ocean warming: a physiologist’s
view. Mar Ecol Prog Ser 373:203–217. https://doi.org/10.3354/meps07768
Preheim SP, Boucher Y, Wildschutte H, David LA, Veneziano D, Alm EJ, Polz MF (2011) Metapopulation
structure of
Vibrionaceae
among coastal marine invertebrates. Environ Microbiol 13(1):265–275.
https://doi.org/10.1111/j.1462-2920.2010.02328.x
Przeslawski R, Ahyong S, Byrne M, Wörheide G, Hutchings P (2008) Beyond corals and sh: The effects of
climate change on noncoral benthic invertebrates of tropical reefs. Glob Change Biol 14(12):2773–2795.
https://doi.org/10.1111/j.1365-2486.2008.01693.x
Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, Peplies J, Glöckner FO (2012) The SILVA
ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res
41(D1):D590–D596. https://doi.org/10.1093/nar/gks1219
Ramsby BD, Hoogenboom MO, Whalan S, Webster NS (2018) Elevated seawater temperature disrupts the
microbiome of an ecologically important bioeroding sponge. Mol Ecol 27(8):2124–2137.
https://doi.org/10.1111/mec.14544
Schiel DR, Steinbeck JR, Foster MS (2004) Ten years of induced ocean warming causes comprehensive
changes in marine benthic communities. Ecology 85(7):1833–1839. https://doi.org/10.1890/03-3107
Shreiner AB, Kao JY, Young VB (2015) The gut microbiome in health and in disease. Curr Opin
Gastroenterol 31(1):69–75. https://doi.org/10.1097/MOG.0000000000000139
Simon CJ, Truong HH, Noble TH, Osborne SA, Wynne JW, Wade NM (2020) Microbial biomass, marine
invertebrate meals and feed restriction inuence the biological and gut microbiota response of shrimp
Penaeus monodon
. Aquaculture 520:734679. https://doi.org/10.1016/j.aquaculture.2019.734679
Sochard MR, Wilson DF, Austin B, Colwell RR (1979) Bacteria associated with the surface and gut of
marine copepods. Appl Environ Microbiol 37:750–759. https://doi.org/10.1128/AEM.37.4.750-759.1979
Spor A, Koren O, Ley R (2011) Unravelling the effects of the environment and host genotype on the gut
microbiome. Nat Rev Microbiol 9:279–290. https://doi.org/10.1038/nrmicro2540
Thao ML, Moran NA, Abbot P, Brennan EB, Burckhardt DH, Baumann P (2000) Cospeciation of Psyllids
and Their Primary Prokaryotic Endosymbionts. Appl Environ Microbiol 66:2898–2905.
https://doi.org/10.1128/AEM.66.7.2898-2905.2000
Page 21/23
Tracy AM, Pielmeier ML, Yoshioka RM, Heron SF, Harvell CD (2019) Increases and decreases in marine
disease reports in an era of global change. Proc R Soc B Biol Sci 286:20191718.
https://doi.org/10.1098/rspb.2019.1718
Vuong HE, Yano JM, Fung TC, Hsiao EY (2017) The Microbiome and host behavior. Annu Rev Neurosci
40:21–49. https://doi.org/10.1146/annurev-neuro-072116-031347
Watts SA, Lawrence JM (1986) Seasonal effects of temperature and salinity on the organismal activity of
the seastar
Luidia clathrata
(say) (Echinodermata: Asteroidea). Mar Behav Physiol 12(3):161–169.
https://doi.org/10.1080/10236248609378643
Watts SA, Lawrence JM (1990) The effect of temperature and salinity interactions on righting, feeding
and growth in the sea star
Luidia clathrata
(SAY). Mar Behav Physiol 17(3):159–165.
https://doi.org/10.1080/10236249009378765
Webster NS, Thomas T (2016) The sponge hologenome. mBio 7:e00135–16.
https://doi.org/10.1128/mBio.00135-16
Zhou G, Cai L, Yuan T, Tian R, Tong H, Zhang W, Jiang L, Guo M, Liu S, Qian P-Y, Huang H (2017)
Microbiome dynamics in early life stages of the scleractinian coral
Acropora gemmifera
in response to
elevated pCO2. Environ Microbiol 19(8):3342–3352. https://doi.org/10.1111/1462-2920.13840
Figures
Page 22/23
Figure 1
Relative abundance of the ten most abundant amplicon sequence variants (ASVs) from samples of feed
and Luidia clathrata stomach, seawater, sand, and feed at time zero and after two-week exposure to
warming. ASVs binned and taxonomy assigned using Qiime2, data sorted and stacked bar chart
constructed using Excel. Taxa represented by the most resolved level based on SILVA9 (v132) taxonomic
classication
Page 23/23
Figure 2
2-dimensional principal component analysis (PCoA) plot of Bray-Curtis dissimilarity for feed, and
stomach, sand, and seawater samples for Luidia clathrata at time zero and held under 28°C, 30°C, and
32°C for 2 weeks. Axis 1 explains 38.79% variance and axis 2 explains 22.47%. Graphed with Qiime2
Supplementary Files
This is a list of supplementary les associated with this preprint. Click to download.
ESM1.pdf
