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The Effects of Combined Ocean Acidification and Nanoplastic Exposures on the Embryonic Development of Antarctic Krill

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In aquatic environments, plastic pollution occurs concomitantly with anthropogenic climate stressors such as ocean acidification. Within the Southern Ocean, Antarctic krill (Euphausia Superba) support many marine predators and play a key role in the biogeochemical cycle. Ocean acidification and plastic pollution have been acknowledged to hinder Antarctic krill development and physiology in singularity, however potential multi-stressor effects of plastic particulates coupled with ocean acidification are unexplored. Furthermore, Antarctic krill may be especially vulnerable to plastic pollution due to their close association with sea-ice, a known plastic sink. Here, we investigate the behaviour of nanoplastic [spherical, aminated (NH2), and yellow-green fluorescent polystyrene nanoparticles] in Antarctic seawater and explore the single and combined effects of nanoplastic (160 nm radius, at a concentration of 2.5 μg ml–¹) and ocean acidification (pCO2 ∼900, pHT 7.7) on the embryonic development of Antarctic krill. Gravid female krill were collected in the Atlantic sector of the Southern Ocean (North Scotia Sea). Produced eggs were incubated at 0.5 °C in four treatments (control, nanoplastic, ocean acidification and the multi-stressor scenario of nanoplastic presence, and ocean acidification) and their embryonic development after 6 days, at the incubation endpoint, was determined. We observed that negatively charged nanoplastic particles suspended in seawater from the Scotia Sea aggregated to sizes exceeding the nanoscale after 24 h (1054.13 ± 53.49 nm). Further, we found that the proportion of embryos developing through the early stages to reach at least the limb bud stage was highest in the control treatment (21.84%) and lowest in the multi-stressor treatment (13.17%). Since the biological thresholds to any stressors can be altered by the presence of additional stressors, we propose that future nanoplastic ecotoxicology studies should consider the changing global ocean under future climate scenarios for assessments of their impact and highlight that determining the behaviour of nanoplastic particles used in incubation studies is critical to determining their toxicity.
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ORIGINAL RESEARCH
published: 03 August 2021
doi: 10.3389/fmars.2021.709763
Edited by:
Stefania Ancora,
University of Siena, Italy
Reviewed by:
Chiara Gambardella,
National Research Council (CNR), Italy
Arnaud Huvet,
Institut Français de Recherche pour
l’Exploitation de la Mer (IFREMER),
France
*Correspondence:
Emily Rowlands
emirow@bas.ac.uk
Clara Manno
clanno@bas.ac.uk
Specialty section:
This article was submitted to
Marine Pollution,
a section of the journal
Frontiers in Marine Science
Received: 14 May 2021
Accepted: 16 July 2021
Published: 03 August 2021
Citation:
Rowlands E, Galloway T, Cole M,
Lewis C, Peck V, Thorpe S and
Manno C (2021) The Effects
of Combined Ocean Acidification
and Nanoplastic Exposures on
the Embryonic Development
of Antarctic Krill.
Front. Mar. Sci. 8:709763.
doi: 10.3389/fmars.2021.709763
The Effects of Combined Ocean
Acidification and Nanoplastic
Exposures on the Embryonic
Development of Antarctic Krill
Emily Rowlands1,2*, Tamara Galloway2, Matthew Cole3, Ceri Lewis2, Victoria Peck1,
Sally Thorpe1and Clara Manno1*
1British Antarctic Survey, Cambridge, United Kingdom, 2College of Life and Environmental Sciences, University of Exeter,
Exeter, United Kingdom, 3Plymouth Marine Laboratory, Plymouth, United Kingdom
In aquatic environments, plastic pollution occurs concomitantly with anthropogenic
climate stressors such as ocean acidification. Within the Southern Ocean, Antarctic
krill (Euphausia Superba) support many marine predators and play a key role
in the biogeochemical cycle. Ocean acidification and plastic pollution have been
acknowledged to hinder Antarctic krill development and physiology in singularity,
however potential multi-stressor effects of plastic particulates coupled with ocean
acidification are unexplored. Furthermore, Antarctic krill may be especially vulnerable
to plastic pollution due to their close association with sea-ice, a known plastic sink.
Here, we investigate the behaviour of nanoplastic [spherical, aminated (NH2), and
yellow-green fluorescent polystyrene nanoparticles] in Antarctic seawater and explore
the single and combined effects of nanoplastic (160 nm radius, at a concentration
of 2.5 µg ml1) and ocean acidification (pCO2900, pHT7.7) on the embryonic
development of Antarctic krill. Gravid female krill were collected in the Atlantic sector
of the Southern Ocean (North Scotia Sea). Produced eggs were incubated at 0.5 C in
four treatments (control, nanoplastic, ocean acidification and the multi-stressor scenario
of nanoplastic presence, and ocean acidification) and their embryonic development
after 6 days, at the incubation endpoint, was determined. We observed that negatively
charged nanoplastic particles suspended in seawater from the Scotia Sea aggregated
to sizes exceeding the nanoscale after 24 h (1054.13 ±53.49 nm). Further, we found
that the proportion of embryos developing through the early stages to reach at least
the limb bud stage was highest in the control treatment (21.84%) and lowest in the
multi-stressor treatment (13.17%). Since the biological thresholds to any stressors can
be altered by the presence of additional stressors, we propose that future nanoplastic
ecotoxicology studies should consider the changing global ocean under future climate
scenarios for assessments of their impact and highlight that determining the behaviour
of nanoplastic particles used in incubation studies is critical to determining their toxicity.
Keywords: nanoparticle, plastic pollution, multi-stressor, Antarctic krill, Scotia Sea, embryonic development, egg
abnormality
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Rowlands et al. Antarctic Krill and Multi-Stressors
INTRODUCTION
In the Southern Ocean, plastic debris has been detected from
surface waters (e.g., Jones-Williams et al., 2020;Suaria et al.,
2020) to the sea-floor (e.g., Munari et al., 2017;Reed et al.,
2018), and across a range of Antarctic biota such as pelagic
amphipods (Jones-Williams et al., 2020), pelagic and demersal
fishes (Cannon et al., 2016), and benthic invertebrates (Sfriso
et al., 2020). Oceanic plastic exists in a continuum of sizes,
from the macro to nanoscale (Ter Halle et al., 2017), though
size classifications are still evolving. Here, we use the definition
provided by Hartmann et al. (2019), referring to a nanoplastic
size criterion of <1 micron (µm) to conform to existing
definitions for nanomaterials. Whilst the technical challenges
of determining exactly how much nanoscale plastic exists in
the natural environment have thus far limited the detection
of nanoplastic in the Southern Ocean, nanoplastic is predicted
to be as pervasive as microplastic (Alimi et al., 2017). Even
concerning the better studied microplastic, there is a paucity of
data in the Southern Ocean with studies specifically dedicated
to microplastic investigation emerging only from 2017 (Tirelli
et al., 2020). Consequently, determining the concentration of
plastic pollution in the region remains challenging (Cincinelli
et al., 2017;Isobe et al., 2017;Lacerda et al., 2019;Jones-
Williams et al., 2020;Suaria et al., 2020). Reports of microplastics
range from an absence (Kuklinskia et al., 2019) to dense
concentrations comparable to concentrations observed in the
Northern Hemisphere oceans. For example, Lacerda et al. (2019)
estimated a mean microplastic concentration in oceanic surface
waters of 1794 km2relative to the estimated density range for
over 70% of the world’s oceans of 1000–100,000 particles km2,
whilst Isobe et al. (2017) estimated microplastic particles in the
order of 100,000 km2through the entire water column at
sample sites closest to Antarctica. Nevertheless, across studies,
the abundance of pelagic microplastics in the Southern Ocean are
generally lower than global trends (Tirelli et al., 2020). However,
since Antarctica is further from major sources of plastic, transport
to the region may take longer and consequently the Southern
Ocean may have a larger proportion of plastics within the smaller
size classifications than other regions due to fragmentation en
route (Obbard, 2018).
As the smallest plastic fraction, nanoplastics are perhaps
more hazardous than their micro counterparts (Koelmans et al.,
2015). Their greater surface area to volume ratio compared to
larger plastics increases their reactivity in marine environments
(Tallec et al., 2019). Whilst plastics characteristically have
hydrophobic surfaces with no net electrostatic charge, this
changes rapidly in seawater as particles accumulate substances
from the water column and are subjected to natural processes
such as weathering (Galloway et al., 2018). Laboratory studies
have tried to assess nanoplastics with differing surface charges
(no-charge, carboxylated, and aminated) in simplified conditions
of natural seawater, and highlight that surface functionalisation
plays a critical role in the behaviours of particles, such as
agglomeration (Della Torre et al., 2014;Bergami et al., 2016;
Manfra et al., 2017;Tallec et al., 2019). Additionally, in-vivo
ecotoxicity studies show that the surface charge of nanoplastics
leads to differing exposure routes as well as differing biological
responses of marine biota at both cellular and organism level
(Corsi et al., 2020). Generally, negatively charged plastic particles
(such as carboxylated particles) with a high ionic strength
quickly agglomerate within seawater (Lee et al., 2019;Tallec
et al., 2019) forming micro-scale aggregates easily ingestible
by zooplankton, whilst positively charged particles (including
many aminated particles) tend to maintain their nano-specific
properties and size (Della Torre et al., 2014;Bergami et al., 2016;
Manfra et al., 2017;Pinsino et al., 2017). Aminated particles
retaining their smaller size facilitates translocation into cells
and triggers physiological disruption such as embryotoxicity
(Bergami et al., 2020). Consequently, aminated particles are
generally shown to be more toxic to cells and tissues in
comparison to carboxylated particles, or those with no surface
functionalisation (Corsi et al., 2020).
Nanoplastic may plausibly enter the Antarctic ecosystem
via both local and long-range sources (Rowlands et al.,
2021). However, research exploring the impacts of nanoplastic
on polar species is limited. Polystyrene spheres (40–50 nm,
at concentrations of 1 and 5 µg ml1) were observed to
impair immune responses of Antarctic sea urchin (Sterechinus
neumayeri) (Bergami et al., 2019), and with the same polymer
(50–60 nm, at a concentration of 2.5 µg ml1) sublethal effects
such as reduced swimming behaviour and increased exuviae
production were observed in juvenile Antarctic krill (Euphausia
superba) (Bergami et al., 2020).
Global trends of ocean acidification, which induce
fundamental changes in seawater chemistry including a reduction
of seawater pH and carbonate ion (CO2
3) concentrations, are
exacerbated in the Southern Ocean (Fabry et al., 2011;McBride
et al., 2014). Carbonate ions are naturally low in the region and
cold temperatures enhance the solubility of CO2, consequently
increasing the rate of ocean acidification. Additionally, upwelling
of hypercapnic (high CO2concentration) deep-sea water and
freshening of surface waters from glacial and sea-ice melt
contribute to further carbonate under-saturation and variations
in pH and pCO2(McNeil and Matear, 2008). Whilst earlier
investigations into the impact of ocean acidification focused
on calcifying biota (Orr et al., 2005), ocean acidification has
been shown to affect a number of physiological and behavioural
processes including acid-base regulation abilities and the energy
demands to maintain homeostasis (Lannig et al., 2010) and can
therefore be critical for non-calcifying organisms such as krill.
By the end of the century, models suggest that that in two key
Antarctic krill habitats of the Southern Ocean, the Scotia Sea
and Weddell Sea, surface water pCO2levels may reach 584 and
870 µatm, respectively, whilst pCO2may reach 1400 µatm at
depth (300–500 m) in the Weddell Sea (Kawaguchi et al., 2011).
Antarctic krill spawn negatively buoyant embryos at the ocean
surface which sink to depths of 700–1000 m before hatching,
and are therefore exposed to enhanced pCO2concentration at
depth (Quetin and Ross, 1984;Kawaguchi et al., 2011, 2013).
Investigating the impact of ocean acidification on Antarctic
krill embryos, Kawaguchi et al. (2011) observed no significant
effects on embryonic development at 1000 µatm pCO2, however
at 2000 µatm pCO2, embryonic development was inhibited
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Rowlands et al. Antarctic Krill and Multi-Stressors
before the gastrulation stage in 90% of cases and no hatching
was observed. Effects were suggested to be associated with
hypercapnia and the intracellular acid-base balance hindering
protein function, since early embryonic development does not
involve calcification.
In a multi-stressor scenario, harmful impacts can be additive,
antagonistic or synergistic where the magnitude of the combined
impact is equal to, less than or greater than the sum of the
singular effects combined, respectively (Norwood et al., 2003).
A meta-analysis exploring the effects of multiple abiotic stressors
on marine embryos and larvae determined that synergistic
interactions (65% of individual tests) were more common than
additive (17%) or antagonistic (17%) interactions (Przeslawski
et al., 2015). According to Alava et al. (2018) interactions between
climate change (e.g., temperature increases, pH decreases,
or pCO2increases) and pollutants may be either climate
change dominant, where the climate stressor leads to enhanced
susceptibility of biota to chemical toxicity, or contaminant
dominant, where chemical exposure leads to an increase in
climate stressor susceptibility. Ocean acidification has also been
shown to alter bioaccumulation rates of contaminants (Braune
et al., 2014;Shi et al., 2016;Alava et al., 2018) and lead
to antagonistic, additive or synergistic interactions with other
pollutants (Pascal et al., 2010;Benedetti et al., 2016;Lewis et al.,
2016;Wilson-McNeal et al., 2020). Another mechanism by which
plastic pollution and ocean acidification may interact in their
toxicity is via variations in seawater pH modifying the chemical
equilibrium of plastics. For example, Piccardo et al. (2020)
observed an ultrastructural change of microplastics exposed to
acidified conditions at the nanoscale-level, enabling release of
chemical additives from the plastic and enhancing ecotoxicity.
Despite interaction abilities, plastics are rarely considered in a
multi-stressor scenario. However, synergisms between plastic and
ocean acidification were observed to negatively impact marine
organisms in both the East China Sea (Wang et al., 2020) and in
temperate regions (Piccardo et al., 2020), impacting digestion and
growth rates, respectively. The potential cumulative impact of
these stressors on Antarctic organisms has not yet been explored.
Antarctic krill (E. superba), referred to hereafter as krill,
potentially have heightened vulnerabilities to multi-stressor
scenarios due to their physiology, behavioural traits and the
rapid environmental changes of their habitats (Rowlands et al.,
2021). The stenothermal species has weak genetic differentiation
(Zane and Patarnello, 2000) and must contend with a plethora
of stressors. As well as the developing embryos being exposed to
large fluctuations in carbonate chemistry, krill have potentially
elevated exposure risks to plastic debris due to a close
association with sea-ice, a known sink for plastic (Peeken et al.,
2018;Kelly et al., 2020). Furthermore, beyond a temperature
optimum, increasing temperatures are demonstrated to reduce
the growth rate of adults (Atkinson et al., 2006) and hatching
success of embryos (Perry et al., 2020). Combined stressors
are hypothesised to lead to increased energy expenditure for
krill embryos to maintain homeostasis (Perry et al., 2020)
and may impact a number of physiological, biochemical, and
molecular parameters (Kawaguchi et al., 2011). As the keystone
species of a highly productive system, the demand on krill
is high in this dominant yet simplistic diatom–zooplankton–
predator food chain (Murphy et al., 2007;Atkinson et al.,
2009) and their role in the biogeochemical cycle is significant
(Belcher et al., 2019;Manno et al., 2020). The potential impact
of multiple environmental stressors on krill development is
therefore critical to explore.
Here, we determine the behaviour of aminated polystyrene
nanoplastic in Antarctic seawater in terms of dispersion and
stability. We assess the ability of aminated nanoplastic to adhere
to and permeate the outermost membrane of the krill embryo
(the chorion) and explore the impact this may have on the
development of embryos from krill collected in the Scotia Sea,
within the Atlantic sector of the Southern Ocean. The sampling
location is a particularly high biomass region for krill, with
50% of all Southern Ocean post-larval biomass contained within
this sector (Atkinson et al., 2009). We focus our study on
krill embryonic development since early life stages are generally
considered the most vulnerable to anthropogenic stressors
(Kurihara, 2008). We extend existing studies on the sensitivity of
krill embryos to future ocean acidification conditions (Kawaguchi
et al., 2011, 2013) by exploring the potential cumulative effects of
nanoplastic coupled with ocean acidification.
MATERIALS AND METHODS
Behaviour of Nanoplastic in Antarctic
Seawater
To understand the behaviour (surface charge and aggregation
state) of polystyrene spheres during the incubation experiment,
we tested the stability of 170 nm diameter amino modified
polystyrene (PS-NH2) spheres (1% w/v, 2 ml, Spherotech) in
open ocean water collected from nearby to the krill collection site
(King Edward Point bay, South Georgia, 54.260S, 36.439W).
We compared the behaviour of nanoplastic in seawater with its
behaviour in Milli-Q water, having filtered both media through
a polycarbonate membrane (pore size of 100 nm). Nanoplastic
stock was vortexed for 30 s and bath sonicated for 60 s prior
to dilution. Dynamic Light Scatter (DLS) analyses (Zetasizer
Ultra; Malvern Instruments) determined the aggregation state
(polydispersity index, PDI), the hydrodynamic diameter of
particles/aggregates (Z-average; nm) and the mean surface
charge (ζ-potential; mV) of nanoplastic. Measurements were
performed in triplicate at 0.5C in 50 µg ml1nanoplastic
suspensions at T0 and T24h.
Krill Collection and Embryo Selection
Krill were collected aboard the RRS Discovery in January
2019 (Cruise DY098) using a targeted Rectangular Midwater
Trawl with a mouth area of 8 m2(RMT8) in the northern
Scotia Sea (outbound trawl 53.837S, 39.180W, inbound trawl
53.808S, 39.160W). Active and undamaged gravid females were
selected and transferred into individual 500 ml perforated plastic
containers within a 320 L holding tank within 30 min of the
net recovery. A constant flow of uncontaminated seawater from
the ship’s underway pump (6 m depth) ensured a turnover
of fresh oxygenated seawater and provided the females with a
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natural food supply (for full protocol, see Perry et al., 2020). As
broadcast-spawners, Antarctic krill shed their eggs freely into the
water (Gómez-Gutiérrez and Robinson, 2005). As such, mesh
with 1 mm apertures were attached 25 mm above the base
of each container to protect sinking eggs from cannibalism.
Krill were kept in darkened conditions and checked every
12 h for spawning.
To control for genetic variability within our incubation
experiment, eggs from the brood of a single female krill which
was selected at random were utilised. Embryos were selected
for incubation from the first spawned egg batch and displayed
a layer of fertilisation jelly surrounding the chorion (embryo
membrane). The spent female had a wet weight of 1.9 g and
length of 60 mm, as measured from the anterior edge of the eye
to tip of telson (Morris et al., 1998). Perry et al. (2020), carrying
out research during the same cruise, found that embryonic
development was not significantly related to wet weight or length
of the gravid female krill in the range of 1.25–2.1 g and length of
52–60 mm, respectively; hence we do not expect our results to be
affected by the size of the female krill.
Embryo Treatment Conditions
Four separate treatments (detailed below) were carried out
during the 6-day experiment on embryo development, with
each treatment performed in triplicate. Embryos (n= 673) were
split between four treatments, with three replicates (a random
subsample of 40–60 embryos) per treatment.
Control Treatment
Ambient sea water was collected from the ship’s underway
pump in the region that the krill were collected and filtered
with a 0.22 µm filter. Non-calibrated salinity measurements
were obtained from the ship-fitted system (Surfmet system)
output. The pHTwas measured directly within incubation wells
(Metrohm 826 pH meter, with a LL Primatrode, pH glass
electrode and a tris-buffer).
Nanoplastic Treatment
Initial dilutions of fluorescently labelled 160 nm amino (PS-NH2)
modified polystyrene nanoparticles (yellow-green, 1% w/v, 2 ml,
Spherotech) were centrifuged at 3000 revolutions per minute
(RPM) for 2 h at 4C and washed three times with deionized
water to remove the bacteriostatic preservatives. Nanoplastic
suspensions were prepared at a final concentration of 2.5 µg ml1
in filtered seawater. The final solutions were vortexed but not
sonicated prior to use. A concentration of 2.5 µg ml1was
chosen based on acute toxicity thresholds observed with other
marine zooplankton incubated with polystyrene nanoplastic
(Bergami et al., 2017;Manfra et al., 2017) and to align with
previous nanoplastic research on Antarctic krill (Bergami et al.,
2020). Aminated particles were chosen since they have been
shown to be more toxic to cells and tissues under other exposure
scenarios and are generally well dispersed under saline conditions
(Della Torre et al., 2014). We utilised fluorescent nanoparticles
with the intent of quantifying adhesion to embryo membranes
and internalisation of particles via fluorescent microscopy. We
later opted for alternative analyses due to difficulties with
autofluorescence coupled with new insight that fluorophore alone
can adhere to and accumulate within internal tissues of larvae
(Catarino et al., 2019).
Ocean Acidification Treatment
Ambient sea water was manipulated to represent a pHTof 7.7 and
pCO2level of 900 µatm that reflects future ocean acidification
states predicted within the depth range (700–1000 m) of krill
embryos (Quetin and Ross, 1984;Kawaguchi et al., 2011)
through adding a combination of acid (HCl) and base (Na2CO3).
The acid and base additions were calculated using seacarb
software (Lavigne and Gattuso, 2010). This calculation relies on
knowledge of the total alkalinity (TA), dissolved inorganic carbon
concentration (DIC) and pH of the seawater, although one of
these three parameters can be estimated if values for the other
two are known. In this instance, we based the calculation on
values determined for pHTand TA. The pHTwas measured as
per the control. TA was determined through applying a surface-
salinity (S) and temperature (TC) based algorithm, based on Lee
et al. (2006) and refined through spatially intensive carbonate
chemistry surveys in the region (M. P. Humphreys, personal
communication):
TA =683.41S9.139S21.37T0.896T210364.16
Sub-samples of seawater from the incubation set-up were fixed
with 2% mercuric chloride for shore-based carbonate chemistry
analysis (Table 1A). The TA and DIC of each sample was
measured simultaneously by a potentiometric titration system
using a technique based on the method of Edmond (1970), with
a closed cell described by Goyet et al. (1991). The accuracy
(3 mmol kg1for TA and 4 mmol kg1for DIC) was determined
by analysing Certified Reference Material (CRM) with known TA
and DIC concentrations. Carbonate saturation state with respect
to aragonite (a) was indirectly calculated from TA and DIC
data using the CO2SYS software (Lewis and Wallace, 1998)
with carbonate dissociation constants by Mehrbach et al. (1973)
refitted by Dickson and Millero (1987) and sulphate dissociation
constants by Dickson (1990).
Nanoplastic and Ocean Acidification Treatment
The multi-stress treatment followed the same method as that
of the nanoplastic treatment, with ambient seawater replaced
with water manipulated for the ocean acidification treatment, as
described above.
Experimental Set-up
Embryos were incubated at 0.5C in line with Jia et al. (2014)
who measured krill development from egg to early juvenile. Due
to its antibacterial properties, magnesium chloride (3332 µl L1)
was added to the seawater and ocean acidification solutions prior
to them being used to make up each of the four treatments.
Treatment solutions were added to 6-well culture plates (10 ml
well capacity) to a total volume of 8 ml and sealed with parafilm
to minimise CO2exchange with the atmosphere.
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Rowlands et al. Antarctic Krill and Multi-Stressors
TABLE 1 | Mean ±SD of carbonate system parameters of SW cultures: water temperature (T, C), total alkalinity (TA, mmol kg1), total CO2(TC, mmol kg1), partial
pressure of CO2(pCO2ppm), aragonite saturation state (a), salinity (S, %), and pHT.
A) Onshore carbonate chemistry T (C) S (%) TA (mmol kg1) TC (mmol kg1) pCO2(ppm) a pH
Control, NP 0.5 ±0.22 33.5 ±0.03 2311.28 ±11.50 2191.31 ±17 387 ±16 1.56 ±0.13 8.05 ±0.03
OA, OA & NP 0.5 ±0.24 33.5 ±0.04 2309.28 ±7.82 2297.12 ±17 902 ±18 0.68 ±0.09 7.70 ±0.02
B) Within incubation pHTmeasurements Day 1 Day 2 Day 3 Day 4 Day 5
Control 8.05 ±0.02 8.05 ±0.03 8.04 ±0.03 8.05 ±0.03 8.04 ±0.02
NP 8.06 ±0.03 8.04 ±0.04 8.04 ±0.04 8.06 ±0.02 8.03 ±0.03
OA 7.7 ±0.03 7.67 ±0.03 7.69 ±0.03 7.71 ±0.02 7.69 ±0.03
OA and NP 7.7 ±0.03 7.68 ±0.02 7.69 ±0.02 7.7 ±0.03 7.69 ±0.04
Experimental Monitoring
The pHTwas monitored throughout the incubation with direct
measurements every 24 h (Table 1B). At the midway point
(day 3) the incubation water for each treatment was replaced
to minimise the impact of embryo respiration on the dissolved
oxygen content of the water. Total oxygen consumption of 60
krill embryos for 6 days is predicted to be 15–30 µl (Ikeda, 1984;
Yoshida et al., 2004).
Embryos were imaged every 12 h throughout the 6-day
incubation (Olympus SZX16 stereomicroscope with a Canon
camera appended via a 0.5 magnification lens adaptor). Imaging
was carried out within a cold laboratory set at the lowest possible
temperature (2C) and imaging time was kept to a minimum
to limit temperature impact during observations. Hatched larvae
were removed from the experiments to prevent damage to the
remaining incubating embryos and preserved on the sixth day at
the experiment endpoint along with unhatched embryos.
Embryonic Analysis
Chorion Examination and Nanoplastic Internalisation
To test for adhesiveness of the nanoplastic spheres used in
the experiment, embryos from the incubation were preserved
in formalin (4%) fixative for onshore analyses. Post-cruise, we
investigated the chorions of a sub-sample of embryos from the
nanoplastic and multi-stress treatment (n= 10 per treatment)
using Scanning Electron Microscope (Zeiss GeminiSEM 500).
Embryos were first cleansed of the fixative with Milli-Q water
and subjected to an ethanol dehydration gradient. In addition,
to quantify any internalised nanoplastic spheres, a further
subsample of embryos from the nanoplastic and multi-stress
treatment (n= 10 per treatment) were digested (10% KOH, 60C,
48 h) and filter papers (polycarbonate, diameter 12 mm, pore size
100 nm) containing residual were SEM imaged.
TABLE 2 | Mean ±SD of behavioural parameters of polystyrene nanoplastics
(50 µg ml1) suspended in Milli-Q (mQW) and filtered seawater (FSW) over 24 h.
Medium Time (h) Z-average (nm) Polydispersity index (PDI)
mQW 0 173.47 ±2.8 0.028 ±0.011
24 173.17 ±2.24 0.028 ±0.028
SW 0 986.93 ±139.52 0.184 ±0.010
24 1054.13 ±53.49 0.193 ±0.016
Embryonic Development Scoring
Embryonic development at the end of the incubation period
was categorised into five stages comprising of single cell, two
cell, multi cell, limb bud, and nauplius using the photographs
of Jia et al. (2014) for identification, plus an additional category
for entrapped nauplii. Embryos classified as unidentifiable were
assumed to be damaged and removed from analyses (n= 12).
Data Analysis
Statistical analyses were conducted in R Studio version 1.4.1103.
A binomial regression was used to determine the probability of
embryos reaching the limb bud or later stage differing between
treatments with assumptions met, i.e., no multicollinearity,
linearity in the logit reported and independent error terms. The
optimal model was backwards selected from the full model,
which contained a three-way interaction between all measured
explanatory variables [i.e., treatment (control, nanoplastic, ocean
acidification, nanoplastic, and ocean acidification), individual
well (to account for variance between replicates), and number
of embryos per well (to account for the variance in the
number of embryos that were randomly pipetted into each well
plate)]. We also included two-way interaction terms for the
measured explanatory variables, primarily to determine whether
any relationship existed between treatment and the number of
embryos per well that may have influenced the probability output
of embryos reaching the limb bud or later stage. Additionally,
we ran the model with a random downsized equal sample
(n= 100 per treatment) which yielded the same results in terms
of the significant or non-significant effects, and consequently we
present the model utilising all available data.
RESULTS
Behaviour of Polystyrene Nanoplastic in
Antarctic Seawater
Results of the nanoplastic characterisation study are shown in
Table 2. DLS analyses revealed a consistent dispersion and
stability of nanoplastic in Milli-Q over 24 h at 0.5C as
indicated by the unaltered Z-average resembling manufacturer
specifications and low PDI values. In contrast, agglomerates
of nanoplastic particles were observed in seawater from the
outset, with agglomerates surpassing current upper thresholds
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for particles exhibiting nano-specific behaviours (1000 nm) after
24 h (Z-average, 1054.13 ±53.49 nm). Measurements of the
mean surface charge (ζ-potential) confirmed a negative charge
of the plastic spheres, with a value of 28.03 ±0.02 mV in
Milli-Q, and a smaller absolute value of 4.50 ±2.98 mV
observed in seawater.
Chorion Examination and Nanoplastic Internalisation
In the onshore analyses, we firstly confirmed that nanoplastics
were easily identifiable via SEM using nanoplastic stock filtered
onto a polycarbonate membrane (Figures 1A,B). Examining the
chorion of 20 embryos across the nanoplastic and multi-stress
treatment with the same SEM method revealed that nanoplastic
spheres did not adhere to the chorions (Figures 1C,D). It should
be noted that the corrugated surface of krill embryos left some
parts of the membrane inaccessible for examination. However,
since 20 embryos from nanoplastic treatments were assessed,
we would expect to observe some incidences of nanoplastic
attachment had the chorion exhibited any adhesive properties.
A few nanoplastic particles were observed from the residual of
digested embryos from both the nanoplastic and multi-stress
incubation (n= 10) rinsed only in Milli-Q (Figures 1E,F),
but no nanoplastic was identified in the residual of digested
embryos across treatments (n= 10) that underwent additional
cleansing steps with both Milli-Q and ethanol. Therefore, the
nanoplastic identified in the residual rinsed only with Milli-Q
is likely unadhered nanoplastic picked up from the incubation
water along with the embryos that withstood the Milli-Q bath
rather than nanoplastic that permeated the embryo membrane.
Embryonic Development
The proportion of embryos reaching at least the limb bud
development stage after 6 days of incubation was highest in
the control treatment (21.84%) and lowest in the multi-stress
treatment (13.17%) (Figure 2). The binomial regression model
reflects the data with the probability of embryos developing to
at least the limb bud stage significantly lower for the multi-
stress treatment compared with the control (B=1.09 ±0.39
SE, p= 0.006). No significant difference in embryo development
was observed for the singular nanoplastic (B=0.13 ±0.34 SE,
p= 0.689) or ocean acidification (B=0.04 ±0.37 SE, p= 0.920)
treatments compared to the control. The probability of reaching
the limb bud stage was positively correlated with number of
embryos per incubation well (B= 0.04 ±0.01 SE, p= 0.001)
but no interaction effect was observed between treatment and
number of eggs per well which was consequently removed from
the statistical model.
Total hatch success rate in the experiment was low at
<2% across all treatments (1.66% control, 1.30% nanoplastic,
1.44% ocean acidification, and 0.5% nanoplastic and ocean
acidification: Figure 2). These results reflect the low hatching
success (mean = 5.7 ±SD 2%) of embryos collected within the
same location of South Georgia during the same research cruise
also incubated at 0.5C (Perry et al., 2020). The limited number
of nauplii that did hatch did so from 4 days post-fertilisation in
line with Perry et al. (2020) who observed hatching from day
4 at 0.5C. Some developed nauplii became entrapped in the
membrane and failed to hatch (Figure 3).
DISCUSSION
We aimed to determine the vulnerability of Antarctic krill
embryos under singular and multi-stressor nanoplastic
and ocean acidification exposure scenarios. We found that
exposure of embryos to the multi-stressor treatment negatively
impacted development, significantly reducing the probability
of reaching the limb bud stage compared to the control
(B=1.09 ±0.39 SE, p= 0.006), with a reduction in reaching
the limb bud or later stage of 8.67%. These findings support
our hypothesis that a multi-stressor nanoplastic and ocean
acidification scenario is detrimental to krill embryogenesis, and
impacts are not necessarily addressed when approaching the
stressors in singularity.
We observed that whilst nanoplastic spheres remained stable
as individual unaggregated particles in Milli-Q water, within
seawater the behaviour of nanoplastic spheres changes quickly
with the formation of microscale aggregates. This aggregation
of nanospheres in seawater contradicts the typical properties
of amino modified particles which often have high stability
(Della Torre et al., 2014), strong positive charges and remain
dispersed due to a sufficiently high ζ-potential ensuring repulsive
mechanisms (Lin et al., 2010). Nonetheless, a negative charge
of aminated particles has been reported elsewhere when tested
in a seawater solution (González-Fernández et al., 2018).
A shift towards a smaller absolute value in the ζ-potential of
nanoplastic in seawater compared to Milli-Q as we observed
(4.50 ±2.98 mV in SW, 28.03 ±0.02 mV in mQW) has also
previously been documented (Della Torre et al., 2014;Lee et al.,
2019). This reduction in the negative charge of nanoplastics in
sea water may be attributed to cations such as Na+, K+, Ca2+,
and Mg2+being attracted to the surface of the nanoplastic as well
as due to proteins and other compounds (Lee et al., 2019). The
alteration in surface charge can consequently lead to instability in
the dispersal of nanoplastic and fast agglomeration since reduced
ζ-potential leads to attractive forces between colloids outweighing
the repulsive mechanisms, and the particles can then adhere
when they collide (Lin et al., 2010). This instability of nanoplastic
in seawater is an important consideration since particles lose their
nano-specific properties, such as their ability to permeate cells,
when the size of aggregates exceeds the nano scale, as observed
in our experiment.
When considering internalisation of nanoplastic within
our incubated embryos, we have inconclusive evidence
that permeation of the krill chorion occurred. Whilst some
nanoplastic particles were observed in the residual of the
digested embryos rinsed only in Milli-Q water, we believe this to
be unadhered nanoplastic picked up from the incubation water
along with the embryos that withstood the Milli-Q bath since
embryos cleansed more rigorously prior to digestion, i.e., through
additional cleansing steps with both Milli-Q water and ethanol,
showed no nanoplastic in digested residual. In opposition to
our findings, Lee et al. (2019) observed polystyrene nanoplastic
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Rowlands et al. Antarctic Krill and Multi-Stressors
FIGURE 1 | SEM images from nanoplastic investigations of the krill chorion and residual of digested krill embryos. (A) Nanoplastic stock diluted with Milli-Q on a
polycarbonate membrane. (B) Nanoplastic stock diluted with Milli-Q on polycarbonate membrane post treatment with10% KOH (60C, 48 h) to mimic the krill egg
digestion. (C) Chorion of a krill embryo from the control treatment. (D) Chorion of a krill embryo from the plastic treatment. (E) Digested (10% KOH, 60C, 48 h) krill
egg residual on a polycarbonate membrane of embryos Milli-Q cleansed. (F) Digested (10% KOH, 60C, 48 h) krill egg residual on polycarbonate membrane of
embryos Milli-Q and ethanol cleansed. SEM images are courtesy of Christian Hacker, University of Exeter.
internalisation in zebrafish embryos, with 50 nm particles
internalised to a much larger extent than 200 and 500 nm
particles. They also observed the hydrodynamic diameter of the
negatively charged particles to remain stable over 48 h, which
is likely attributable to the freshwater environment in which
they were tested since the behaviour of nanoplastic in freshwater
compared to seawater is known to differ (Koelmans et al.,
2015). In our incubation experiment, the lack of internalisation
of nanoplastic particles may be explained by using 160 nm
nanoplastics, which readily aggregated in Antarctic seawater to
the micro-scale.
We also found that the polystyrene nanoplastic spheres did
not attach to the krill chorion. Whilst nanoplastic adherence
to euphausiid chorions has not previously been explored, our
observation differs to that of Kashiwada (2006) who found
polystyrene nanoplastic (39.4 nm) adsorbed on the surface of
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Rowlands et al. Antarctic Krill and Multi-Stressors
FIGURE 2 | (A) Percentages of krill embryos at the early (single cell to multi cell) and late (limb bud to hatching) developmental stage within each triplicate after
6 days of incubation in treatment conditions. Treatment conditions include control, nanoplastic (NP), ocean acidification (OA), nanoplastic and ocean acidification (NP
and OA). Asterisk marks statistically significant difference from the control *<0.05 with results from a binomial GLM. (B) The proportion of krill embryos at the early
(single cell to multi cell) and late (limb bud to hatching) developmental stage after 6 days of incubation in treatment conditions. Treatment conditions include control,
nanoplastic (NP), ocean acidification (OA), nanoplastic and ocean acidification (NP and OA). Total embryos analysed across the three replicates per treatment were
119 (control), 103 (NP), 107 (OA), and 129 (NP and OA).
Japanese medaka (Oryzias latipes) embryos, and Lee et al. (2019)
who observed polystyrene nanoplastic (50–500 nm) attached
to the chorions of zebrafish embryos. Chorion physicality is
observed to vary across aquatic organisms. Eggs of some fish
species, for example, are observed to have an adhesive chorion
whilst others are harder and less adhesive (Goto et al., 2019).
Chorions are also known to differ between euphausiid species,
for example, Gómez-Gutiérrez et al. (2010) determined that
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Rowlands et al. Antarctic Krill and Multi-Stressors
FIGURE 3 | (A) Microscope image (Olympus SZX16 stereomicroscope with a Canon camera appended via a 0.5 magnification lens adaptor) of a newly spawned
krill egg from outer to inner embryo: (F) fertilisation jelly, (C) chorion (egg membrane), (PVS) perivitelline space, (VM) vitelline membrane, and (N) nucleus.
(B) Microscope images (Olympus SZX16 stereomicroscope with a Canon camera appended via a 0.5 magnification lens adaptor) of each development stage
categorised as: single cell, two cell, multiple cell, limb bud, entrapped nauplius, and nauplius.
the chorions of Euphausia pacifica eggs were firm, smooth
and elastic whilst Thysanoessa spinifera chorions were soft
and sticky with particles attached. We observed Antarctic
krill chorions to be firm, smooth and non-sticky which may
offer an explanation for the lack of nanoplastic adhered.
These differences in chorion characteristics, likely impacting
nanoplastic adherence, in turn may affect the bioavailability
and toxicity of plastic particulates, for example by membrane
breakages initiating damage (Nel et al., 2009). Therefore, we
suggest that chorion features of krill embryos might offer an
additional protection to the exposure of nanoplastic compared to
marine organisms (including other euphausiid species) showing
a chorion adhesive characteristic.
Regarding embryonic development, hatch success rates were
low (<2%) across treatments. A low hatch success rate is not
uncommon in Antarctic krill studies with hatch success varying
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Rowlands et al. Antarctic Krill and Multi-Stressors
from 0 to 100% (Yoshida et al., 2004;Perry et al., 2020) and
often reduced in field based observations, e.g., 0–54% (Perry
et al., 2020) and 0–48% (Yoshida et al., 2004) compared to
laboratory based observations. The causes of such variability
remain somewhat unclear. Yoshida et al. (2004) postulate the
high variation in krill hatching success observed in the field
observed can be explained by the condition of the female krill
when caught, further finding in a laboratory setting that hatch
success is influenced by fatty acid composition of embryos,
which is influenced by the diet of maternal krill. Yet neither
weight nor length of the gravid krill were observed to impact
hatch success in observations of Perry et al. (2020). Whilst
variability in hatch success of krill embryos is likely influenced
by an array of environmental factors, as well as individual krill
variability and requires further investigation, we focus on the
likelihood of handling effects driving the low hatch success
within our incubation. Our low hatch success rate resembled the
mean hatching success that Perry et al. (2020) observed in field
experiments with embryos collected on the same research cruise,
within the same location of South Georgia and incubated at the
same temperature of 0.5C (mean hatching success = 5.7 ±SD
2%). However, Perry et al. (2020) also observed high variability
in hatch success across multiple krill from different schools
(15% ±SD 17%) and therefore exclude handling effects and
experimental set-up as a driver of hatch success. We adopt
the same assumption since the gravid female that produced
the embryos used in our incubation experiment was kept in
the same conditions as other gravid females that produced
embryos with high hatch success (Perry et al., 2020). Moreover,
Antarctic krill embryos are generally observed to be robust
towards handling compared to embryos of other krill species
(Harrington and Ikeda, 1986). Consequently, it is our opinion
that the limited development to the later embryo stages which
occurred even in the control (<22%) and low hatch success rate
<2% is representative of Antarctic krill’s reproductive outputs.
Nonetheless, our experiment was limited by the availability
of gravid females during the research expedition and further
incubation experiments would be beneficial to understand
whether observed responses to treatments are reflective of the
wider krill population.
We observe no significant difference in development between
the singular nanoplastic treatment and control for Antarctic krill
embryos (B=0.13 ±0.34 SE, p= 0.689). Previous studies of
other species suggest the effect of nanoplastic on embryogenesis
is influenced dramatically by size and charge of the particles. As
examples, Tallec et al. (2018) assessed the impact of polystyrene
nanoplastic of varying sizes and functionality on Pacific oyster
(Crassostrea gigas) embryos, determining that positively charged
particles of the smallest size (50 nm) had the strongest toxicity,
though uptake of nanoplastic was not quantified. Further, Della
Torre et al. (2014) observed positively charged polystyrene
nanoplastic spheres (50 nm) caused severe developmental defects
in Mediterranean sea urchin (Paracentrotus lividus) embryos
while no embryotoxicity was observed for negatively charged
nanoplastic (40 nm). The reduced uptake and consequent
lower toxicity of negatively charged particles was attributed
to the formation of large aggregates in seawater, unlike
positive particles which remained dispersed. We expect the
same influences regarding surface charge and aggregation
(ζ-potential 4.50 ±2.98 mV with particle aggregation Z-average
1054.13 ±53.49 nm) for our findings.
We also observe no significant differences between the ocean
acidification (pCO2900 µatm) treatment and ambient control
(390 µatm) in agreement with Kawaguchi et al. (2011) who
observed no significant effect on krill embryonic development
at a similar pCO2(1000 µatm). Further, Kawaguchi et al.
(2013) determined that at pCO2levels above 1000 µatm a
sharp decline in hatch success is observed, reaching 20% of
control levels at 1500 µatm pCO2. Building on findings of
Kawaguchi et al. (2011, 2013), we show that when introducing
another environmental stressor (in this instance nanoplastic),
ocean acidification becomes an important consideration for
embryogenesis at a lower pCO2of 900 µatm. An understanding
as to how the pCO2thresholds alter in a multi-stressor
nanoplastic and ocean acidification scenario is therefore required.
Whilst multi-stressor plastic and ocean acidification
ecotoxicology studies are limited, Wang et al. (2020) determined
that ocean acidification conditions under pH 7.7 enhanced the
toxicity of 2 µm polystyrene microplastic to Korean mussels
(Mytilus coruscus) by significantly inhibiting digestive enzymes
and impacting oxidative responses. However, since the uptake
of plastic particulates was not quantified, the underlying cause
of the enhanced toxicity is unknown. In our experiment, since
nanoplastic was not observed to adhere to or penetrate the
chorion, we suggest an indirect effect on the developmental
impairment of krill embryos in a multi-stressor scenario, perhaps
associated with leachates from the polystyrene spheres. Martínez-
Gómez et al. (2017) concluded leachates from fluorescent
polystyrene microspheres (6 µm) caused a higher toxicity to sea
urchin (P. lividus) embryos than virgin and aged polystyrene
themselves, attributing toxicity to styrene/ethylene monomers,
as well as unknown toxic chemicals and additives (such as
antioxidant) from the polystyrene microsphere production. With
the same species, Oliviero et al. (2019) determined microplastic
flakes from PVC toys hindered larval development at the highest
dose used in their experiments (30 mg L1) with the main effect
being attributed to leached chemicals such as phthalates.
One possible explanation for embryo development being less
successful under the multi-stressor scenario (8.67% less than the
control) in our study is that the ocean acidification treatment
altered the speciation, bioavailability and therefore toxicity of
chemicals within the nanoplastic spheres under the reduced
pH condition, as previously observed for ionisable chemicals
(Millero et al., 2009). Among the different variables that may
influence leaching behaviour, Quina et al. (2009) determined
pH of the leachant solution to be the most important. Piccardo
et al. (2020) observed that sea urchin (P. lividus) larvae, exposed
to treatments of polyethylene terephthalate (PET) 0.1 mg ml1
at various sizes (small: 5–60 µm, medium: 61–499 µm, and
large: 500–3000 µm) and PET leachates, present a reduction of
limb length under more acidic conditions (pH 7.5) compared
to the control (pH 8.0). Adverse effects from the microplastic
treatments were also attributed to leaching of chemicals such as
nonyl-and octyl-phenols and bisphenol-A, common additives in
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Rowlands et al. Antarctic Krill and Multi-Stressors
plastic packaging, since plastic flake sizes were not compatible
with the feeding apparatus of the animal. The assumption was
further supported by spectral analysis which identified that under
a lower pH condition, intensity peaks of PET were reduced,
which were attributed to the release of chemical additives and
structural modifiers. In the case of our study, the low pH may
have altered the leaching of chemicals and/or fluorophores from
the fluorescent nanoparticles. Quantifying leachates was beyond
the scope of our study, however for broadening our knowledge
of nanoplastic toxicity, nanoplastic leachate treatments should
be considered in future ecotoxicology experiments and chemical
analyses conducted to validate leachate assumptions.
The reduction in embryo development we observed under the
multi-stressor condition of nanoplastic and ocean acidification
may also be associated with additional energy requirements to
maintain homeostasis. Acid/base regulations bear a substantial
energetic cost; for example, sea urchins (P. lividus) were observed
to have up to a 20-fold increase in gene upregulation, and
consequently an increased energy consumption to reach the same
developmental stage at a lower pH (7.5–7.0) compared to ambient
conditions (Martin et al., 2011). In the aforementioned Piccardo
et al. (2020) study, authors suggested that reduced growth of
sea urchin within in the ocean acidification/plastic treatments
could have been due to increased energy consumption induced
by pH stress which consequently enhanced the ecotoxicological
responses of the embryos to leachates, i.e., there was a
synergistic effect of an increased energy consumption and
gene transcription associated with chemical damages. To fully
understand the mechanisms driving developmental hindrance in
this multi-stressor Southern Ocean scenario, additional in-depth
physiological, biochemical, and molecular analyses are required
such as correlation with reactive oxygen species (ROS) (e.g., Lee
et al., 2019;Wang et al., 2020) and investigation of the impacts
on gene expression, cellular morphology and phagocytic activity
(e.g., Browne et al., 2008;Bergami et al., 2019).
CONCLUSION AND FUTURE DIRECTION
Ocean acidification is expected to increase in the future, with
Southern Ocean pCO2predicted to rise to 1400 µatm in the
depth range of Antarctic krill by the year 2100 (Kawaguchi
et al., 2011). Cumulative global plastic production is expected
to reach 34 billion metric tons by 2050 (Garside, 2019) with
an estimated 4.8–12.7 Mt entering the oceans as macroscopic
litter and microplastic particles each year (Geyer et al., 2017;
Worm et al., 2017). Regardless of increased plastic input into
oceanic systems, the degradation of existing oceanic plastic will
facilitate an increase in micro- and nano-scale plastic debris.
Here, our results show that the development of Antarctic krill
embryos is lowest under a multi-stress nanoplastic and ocean
acidification scenario, with a reduction of 9% in the probability
of embryos reaching later development stages in the observed
time period compared to the control. Consequently, results
are indicative of the need to consider plastic pollution within
a multi-stressor scenario when addressing early development
of Antarctic krill. However, results should be interpreted
with the understanding that more robust experiments are
required to determine whether findings are representative of
the wider Antarctic krill population. Additionally, further work
is required to quantify nanoplastic doses, sizes, and surface
charges and ocean acidification pCO2thresholds within a multi-
stress scenario. Moreover, nanoplastic multi-stress investigations
should not be confined to predicted future ocean acidification
conditions. Perry et al. (2020) found that water temperatures
above 3C markedly decreased krill hatching success. Ocean
warming is evidenced to lead to synergistic reactions with other
contaminants in Antarctic species such as Antarctic scallop
(Adamussium colbecki) (Benedetti et al., 2016) and stenothermal
Antarctic species, such as krill, may be more sensitive to
warming-chemical synergisms (Rowlands et al., 2021). Since the
biological thresholds to stressors can be lower in combination
compared to in singularity, future nanoplastic ecotoxicity studies
should begin to acknowledge the changing climate in terms of
both ocean acidification and ocean warming for a more realistic
representation of existing marine habitats.
DATA AVAILABILITY STATEMENT
The raw data supporting the conclusions of this article will be
made available by the authors, without undue reservation.
AUTHOR CONTRIBUTIONS
ER and CM undertook the conceptualisation, conducted the
experimental design and field experiments, and carried out the
original drafting of the manuscript. ER did the analysis of krill
eggs in the laboratory and data analysis. TG, MC, CL, VP, and ST
assisted with data interpretation and provided a general overview.
All authors contributed significantly to the redrafting, reviewing,
and editing of the manuscript.
FUNDING
ER was funded by a NERC GW4+scholarship NE/L002434/1.
TG was supported by grants NE/N006178 and NE/S003975/1.
CM, VP, and ST were supported by NERC funding to the British
Antarctic Survey “Ecosystems” and “Paleo Environments, Ice
Sheets and Climate Change” science programmes.
ACKNOWLEDGMENTS
We would like to acknowledge the British Antarctic Survey Polar
Ocean Ecosystem Time Series (POETS) project, and all involved
in krill collection. We thank Franki Perry, Kirstie-Jones Williams,
and Bjorg Apeland for assistance with fieldwork as well as RRS
Discovery officers and crew (DY098). For bio-imaging support
we thank Christian Hacker and for useful discussion on DLS
analyses, Diogo Fernandes. For support with onshore carbonate
chemistry analyses we also thank S. Sandrini.
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Frontiers in Marine Science | www.frontiersin.org 14 August 2021 | Volume 8 | Article 709763
... With most permanent research stations and tourist landings occurring on the coast (COMNAP, 2022;IAATO, 2021), local point sources of pollution are understudied in the remote inland regions of Antarctica (Aves et al., 2022). However plastic pollution, which is intertwined with the Antarctic food web, can have a wide range of ecological consequences from small nanoplastics hindering the development of Antarctic krill (Rowlands et al., 2021) to microplastic debris being found in the digestive tract of a range of Antarctic seabirds which may cause gastrointestinal tract blockages, toxicity and oxidative stress (Taurozzi and Scalici, 2024). ...
... Microplastic exposure may adversely impact the robustness of Antarctic species to environmental change, exacerbating existing stresses 41 . A study of the combined impacts of low pH conditions and nanoplastic exposure showed more suppressed development of krill embryos under the combined stresses than when considered separately 44 . Thus, to understand realistic microplastic toxicity thresholds and potential trophic cascade impacts through the pelagic food web, this pollutant should be considered within the context of other local environmental changes, not in isolation. ...
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... The publications that we found did not correspond to works related to nanometal of aquatic metazoans were subtracted of the analysis The search queries in Pubmed were "Transcriptomic" and "nanoparticle" and "aquatic" and "metazoa" for "any field" of the publication (at 05 June 2024). The publications that we found not corresponding to works related to transcriptomic after exposure to nanoparticle in aquatic metazoans were subtracted of the analysis Capolupo et al. 2021;Wang et al. 2021); (10) decrease in offspring quality, with reduction of oocyte quality and sperm mobility and reduced of larval growth (Lee et al. 2013;Besseling et al. 2014;Sussarellu et al. 2016); (11) neurotoxicity (Oliveira et al. 2013;Chen et al. 2017;Capolupo et al. 2021;Santos et al. 2021;Hoyo-Alvarez et al. 2022); (12) behavioral alterations in mobile organisms, such as swimming ability, speed, distance traveled and avoidance (Chen et al. 2020;Santos et al. 2021;Kim et al. 2022), and altered valve movement in bivalves (Bringer et al. 2021); (13) histopathological damages in organs as liver and digestive tract (Kim et al. 2022;Li et al. 2022a); (14) embryotoxicity also coexposed to acidification (Rowlands et al. 2021); and finally (15) mortality (Lee et al. 2013;Cole and Galloway 2015;Manno et al. 2022). ...
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