Impacts of microplastics on growth and health of hermatypic corals are species-specific

Environmental Pollution 254 (2019) 113074

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Impacts of microplastics on growth and health of hermatypic corals are species-specific* Jessica Reichert a, *, Angelina L. Arnold a, Mia O. Hoogenboom b, Patrick Schubert a, Thomas Wilke a a

Department of Animal Ecology and Systematics, Justus Liebig University Giessen, Heinrich-Buff-Ring 26-32 (IFZ), 35392 Giessen, Germany Australian Research Council Centre of Excellence for Coral Reef Studies and College of Science and Engineering, James Cook University, Townsville, Queensland 4811, Australia

b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 23 June 2019 Received in revised form 16 August 2019 Accepted 17 August 2019 Available online 19 August 2019

Coral reefs are increasingly affected by the consequences of global change such as increasing temperatures or pollution. Lately, microplastics (i.e., fragments < 5 mm) have been identified as another potential threat. While previous studies have assessed short-term effects caused by high concentrations of microplastics, nothing is known about the long-term effects of microplastics under realistic concentrations. Therefore, a microcosm study was conducted and corals of the genera Acropora, Pocillopora, Porites, and Heliopora were exposed to microplastics in a concentration of 200 particles L
1, relating to predicted pollution levels. Coral growth and health, as well as symbiont properties were studied over a period of six months. The exposure caused species-specific effects on coral growth and photosynthetic performance. Signs of compromised health were observed for Acropora and Pocillopora, those taxa that frequently interact with the particles. The results indicate elevated energy demands in the affected species, likely due to physical contact of the corals to the microplastics. The study shows that microplastic pollution can have negative impacts on hermatypic corals. These effects might amplify corals' susceptibility to other stressors, further contributing to community shifts in coral reef assemblages. © 2019 Elsevier Ltd. All rights reserved.

Keywords: Microcosm Microplastic Long-term exposure Reef-building corals Photosynthesis Polyethylene

1. Introduction The world's tropical coral reef ecosystems are increasingly affected by human impacts (Heron et al., 2016; Hoegh-Guldberg et al., 2007; Hughes et al., 2003). Their foundation species, hermatypic zooxanthellate corals, live in a mutualistic symbiosis with endocellular algae (Muscatine et al., 1981; Muscatine and Porter, 1977). Both, local (e.g., overfishing, coastal development, and pollution) and global stressors (e.g., elevated temperatures and ocean acidification) have caused severe damage to coral reef ecosystems during the last decades (Gardner et al., 2003; Hughes et al., 2018). Recently, plastic debris has been identified as a pollutant of rising concern for coral reefs as it is associated with increased susceptibility to diseases (Lamb et al., 2018).

* This paper has been recommended for acceptance by Eddy Y. Zeng. * Corresponding author. Department of Animal Ecology & Systematics, Justus Liebig University Giessen, Heinrich-Buff-Ring 26e32 (IFZ), 35392 Giessen, Germany. E-mail address: [email protected] (J. Reichert).

https://doi.org/10.1016/j.envpol.2019.113074 0269-7491/© 2019 Elsevier Ltd. All rights reserved.

Plastics make up 60e80% of marine litter (Derraik, 2002) and are ubiquitous in marine ecosystems (Browne et al., 2015; Eriksen et al., 2014; Van Cauwenberghe et al., 2013). Worldwide 309,000 tons of plastic debris are estimated to float at or just beneath the sea surface (Koelmans et al., 2017). Microplastics (defined as plastic fragments <5 mm; Andrady, 2011) are of special concern because they may contain chemical pollutants (Bakir et al., 2012; Lee et al., 2014; Teuten et al., 2009) or pathogens (Kirstein et al., 2016; Vir
sek et al., 2017), and are readily ingested by various marine organisms (Galloway et al., 2017; Wright et al., 2013b). Concentrations of microplastics in ocean waters are highly variable and are influenced by large and small scale circulation patterns and local pollution events (Critchell and Lambrechts, 2016; Hidalgo-Ruz et al., 2012). In highly polluted coastal areas, concentrations can reach up to 150 particles L
1 (Chae et al., 2015; Song et al., 2014). Moreover, current estimates likely underestimate true values due to logistical challenges in sampling small particles (Lusher et al., 2014; Miller et al., 2017). By the year 2100, worldwide concentrations are predicted to rise roughly by a factor of three (Koelmans et al., 2017). The chronic exposure of organisms to microplastics is associated

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with decreased uptake of natural prey (Murphy and Quinn, 2018), resulting in energy deficits (Wright et al., 2013a), reduced growth (Lo and Chan, 2018), and lower fecundity (Sussarellu et al., 2016). Microplastics are of particular concern for suspension feeders such as hermatypic corals. Corals can ingest and partly egest particles (Allen et al., 2017; Hall et al., 2015; Hankins et al., 2018; Reichert et al., 2018), but also respond to contact with particles using cleaning mechanisms, such as mucus release (Reichert et al., 2018). These mechanisms can be energetically costly and might also interfere with the normal heterotrophic feeding of the corals, thereby potentially affecting growth and fecundity. For cold-water corals it has been recently shown that macro- and microplastics affect growth rates, feeding, and behavior (Chapron et al., 2018; Mouchi et al., 2019). However, the effects of microplastic exposure on long-term growth and health performances of tropical hermatypic corals are poorly understood. Therefore, the general goal of this study was to assess the impact of microplastics on the physiological performance of reef-building corals. A six-months microcosm experiment was conducted with four species of coral families, commonly dominating coral assemblages in the Indo-Pacific region: Acropora muricata (Linnaeus, 1758), Pocillopora verrucosa (Ellis and Solander, 1786), Porites lutea Milne Edwards and Haime, 1851, and Heliopora coerulea (Pallas, 1766). In order to assess their responses under realistic microplastic conditions (Lenz et al., 2016), we chose a concentration of ~200 particles L
1, based on conservative estimations for polluted ocean surface waters by the year 2100 (Koelmans et al., 2017). Specifically, we assessed the effects of microplastics (I) on coral growth rates, (II) on photosynthetic performance, densities, and chlorophyll concentrations of the associated endocellular algae, and (III) on the corals' overall health. 2. Material and methods 2.1. Experimental settings and study species A six-month experiment was conducted to study the impact of microplastics on representatives of the major families of reefbuilding Indo-Pacific scleractinian corals, i.e., Acropora muricata (Acroporidae), Pocillopora verrucosa (Pocilloporidae), and Porites lutea (Poritidae). Additionally, Heliopora coerulea (Helioporidae) was included as a reef-building octocoral. The studied species have varying growth forms, ranging from massive (P. lutea) to columnar (H. coerulea), and branching (A. muricata, P. verrucosa). However, all species share comparable polyp sizes (1e2 mm diameter). Coral colonies were maintained at the aquarium facilities of Justus Liebig University Giessen under laboratory conditions (10:14 light:dark photoperiod, light intensity 200 mmol photons m
2 s
1, and temperature 26 ± 0.5  C) for at least six months prior to the experiment (for details on coral colonies see Table S1). Colonies were fragmented into experimental nubbins with a small angle grinder (Multitool 3000-15, Dremel, The Netherlands). Acropora, Pocillopora, and Heliopora nubbins consisted of 2e4 cm long terminal branches; Porites nubbins of small cubes with an edge length of approx. 2 cm. All nubbins were attached to self-made concrete bases (height: 2 cm, diameter: 4 cm) with a two-component glue (CoraFix SuperFast, Grotech, Germany). For Acropora, Pocillopora, and Porites, a total of 90 nubbins per species were cut equally from three original colonies. For Heliopora, 30 nubbins were cut from a single colony due to the lack of replicate colonies. After fragmentation, the nubbins were ID labeled, distributed equally among six tanks, and acclimated three weeks before the experiment commenced. As coral nubbins can be sensitive to handling, four additional nubbins per colony were generated as backups and not included a

priori in the analyses. These backup-nubbins were kept in one microplastic treatment and one control tank. All nubbins were distributed randomly within the tanks with a minimal distance of 5 cm to avoid direct interactions and minimize competition. Over the time of the experiment, a total of nine nubbins was replaced with backup nubbins due to handling-related damages. Additionally, one Acropora nubbin was excluded from the control treatment without replacement, due to the lack of further backup nubbins. 2.2. Tank setup and water system The experiment was conducted in six 80 L tanks (three microplastic treatment and three control tanks) as a closed laboratory microcosm experiment, connected to a 4000 L artificial seawater system. The system includes a large ‘buffer’ tank containing corals, fish, and a deep-sand bed, together with a protein skimmer and a calcium reactor (pH 6.2e6.4, coral rubble). All experimental tanks were connected to the system with an exchange rate of 120 L day
1 (equivalent to 150% of the tank volume) to assure similar water conditions in all tanks. The inflowing water was treated in a UV clarifier (RWUVC/78/4000, RuWal Aquatech, Italy; 33000 mW s
1 cm
2 at 4000 L h
1) to minimize pathogen loads entering the experimental tanks. The outflow of each tank was equipped with 65 mm filters to prevent the removal of plastic particles and a distribution within the flow-through system. Further, the outflowing water was filtered through a fleece membrane to remove microplastic particles smaller than 65 mm (i.e., originating from fragmentation processes over the course of the experiment). The temperature was maintained at 26 ± 0.2  C using 300 W heaters, controlled by an aquarium computer (Profilux 3, GHL Advanced Technology, Germany). The light was set with a 10:14 light:dark photoperiod, using two sets of four fluorescent tubes (two 80 W Aquablue Special, ATI, Germany; two 80 W Blue Plus, ATI, Germany), each extending over three tanks. Light intensity was adjusted to 135 mmol photons m
2 s
1 as measured at the base of each tank with a quantum sensor (measuring range: 410e655 nm, Apogee Quantum Flux MP-200, Apogee Instruments, USA). All tanks were equipped with a propeller pump (RW-8, Jebao, China; 700 L h
1) to generate horizontal currents. A second pump (S 400, Resun, China; 400 L h
1) generated a vertical current in order to submerge floating particles. Water flow in the tanks was established at 5e7 cm s
1 (OTT MF pro, HydroMet, Germany). To limit algae growth and detritus accumulation, ca. 50 small gastropods (Nassarius spp., Euplica spp., Turbo spp., and Stomatella auricula) were added to each experimental tank. 2.3. Microplastic treatments and concentrations High density polyethylene (PE) microplastic (density: 0.95 g cm
3, Novoplastic, Germany) was used for the experiment as it is one of the most common types of plastic in the oceans (Andrady, 2017). The particles were irregularly shaped and exhibited a rough surface structure, resembling natural secondary microplastics. Attenuated total reflection Fourier-transform infrared spectroscopy (ATR-FTIR) confirmed the polymer type. Particle properties (mean length, minimum length, maximum length, perimeter, planar surface area, and solidity; n ¼ 900, Fig. S1) were analyzed under a digital microscope (Keyence VHX-2000, Keyence, Japan) using the ‘particle analyzer’ plugin from the ‘biovoxxel’ toolbox in Fiji ImageJ (Brocher, 2015; Rasband, 1997). Particles ranged from 65 mm to 410 mm with a mean diameter of 175.5 ± 73.5 mm (mean ± SD), comparable to the size of particulate food commonly ingested by the studied corals. Prior to the addition, microplastics were incubated for 24 h in 70% (v/v) ethanol at room

J. Reichert et al. / Environmental Pollution 254 (2019) 113074

temperature for surface sterilization, followed by a washing step in micro-filtered (0.22 mm) autoclaved artificial seawater. Sterilized microplastics were added to the treatment tanks in a concentration of 2.5 mg L
1. Due to the positive buoyancy of a relatively high proportion of the added microplastic particles, the concentration of particles in the water column stabilised after 48 h at approx. 10% of the total amount added (~200 particles L
1 or 0.25 mg L
1). The concentration of bioavailable plastic particles (i.e., in the water surrounding the corals) was monitored at least every three weeks. To do so, 50 mL tank water was taken from the water column around the corals and filtered onto a 65 mm gauze filter (polyamide fabric, gravity filtration, 3e5 replicate measurements per tank). The particles on each filter were counted under a stereo microscope and the numbers were extrapolated to assess the concentration per L. Concentrations were maintained at 203 ± 69 particles L
1 over the course of the experiment by adding fresh particles when necessary (Fig. S2). However, there was a decline in concentrations after 20 weeks, probably due to removal of plastics through handling, cleaning, or deposition of particles in biofilms or within the corals. 2.4. Water chemistry and maintenance of corals Water parameters (alkalinity: 2.52 mmol L
1, Ca2þ: 410 mg L
1,


1
1 Mg2þ: 1230 mg L
1, PO3
4 : <0.03 mg L , NO3 : <0.02 mg L , NO2 :
1 <0.01 mg L
1, NHþ <0.025 mg L , salinity: 34) were monitored 4 weekly and, if necessary, adjusted by the addition of NaHCO3, CaCl2$2H2O, MgCl2$6H2O, deionized water or artificial seawater (Coral ocean plus, ATI, Germany). Corals received indirect feeding through the connected water system, which was supplied daily with frozen food (i.e., copepods, Mysis spp.). To avoid position effects within tanks, the nubbins were repositioned randomly every week. Coral nubbins were checked daily and cleaned from algae, if necessary. Accumulating detritus was removed from the tanks twice a week by siphoning. To minimize the impact of corallivorous flatworms (Amakusaplana acroporae), which prey on Acropora spp., all Acropora nubbins were treated with iodine solution and cleared from flatworm eggs four times during the experiment. 2.5. Coral growth and health assessment Growth parameters (i.e., changes in surface area, volume, and calcification rate) were assessed using 3D scanning and buoyant weight measurements. For doing so, coral nubbins were documented at the beginning of the six-month experiment and every six weeks thereafter using a handheld 3D scanner (Artec Spider 3D, Artec 3D, Luxembourg), according to previous studies (Reichert et al., 2016). Briefly, corals were placed on a rotating plate and scans were captured in air within 60e90 s. The resulting images were processed using the scanning software Artec Studio 10 (Artec 3D). Models were calculated with a final mesh size of 0.2 mm; for detailed settings see Table S2. During model calculation ‘outlier removal’ settings were adjusted to the underlying scan quality. For tissue surface area determination meshes were trimmed manually at the tissue border. All bleached and necrotic tissue was trimmed in each scan to assess only the healthy, photosynthetically active tissue surface area. For volume increase determination, meshes of the same individuals from all time points were aligned and trimmed horizontally. All meshes were exported as Wavefront ‘.obj’ files to MeshLab (Visual Computing Lab-ISTI-CNR, Italy; v1.3.4 beta), and surface area and volume were calculated using the ‘compute geometric measures’ tool. Coral growth rates were assessed based on increases in tissue surface area and skeletal volume. Growth rates were calculated for each six-week interval as increases in surface area and volume, normalized to surface area. Signs of compromised coral health (i.e., bleaching, tissue

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necrosis, feeding scars from parasites) were analyzed using the 3D models. Severe tissue reduction was defined as loss of more than 50% of healthy tissue (Marshall and Baird, 2000), over a six-week interval. Calcification rates were determined using the buoyant weight technique (Davies, 1989) every six weeks. Prior to 3D scanning, corals were placed into 12 L of artificial seawater (salinity: 34, temperature: 26  C) and weighed on a scale pan, attached to a hook under a balance (Kern-KB 360 3N, Kern & Sohn, Germany; precision: 0.001 g). Calcification rates were calculated for each six-week interval as total weight increase, normalized to surface area. 2.6. Photosynthetic activity of symbionts Photosynthetic activity of the photosymbionts was quantified with the pulse-amplitude modulated (PAM) fluorometry (Ralph and Gademann, 2005) at six-week intervals. Data were collected using a PAM-2500 fluorometer (Walz, Germany) with a 6 mm diameter fibre optic probe and a distance clip, which stabilised the probe at a distance of 5 mm above and at 60 to the tissue. PAM settings were adjusted to obtain stable maximum fluorescence yields for all species (Fm); for detailed settings see Table S3. Effective (DF/F'm) and maximum (Fv/Fm) photochemical efficiency were measured for each nubbin at three different positions. Effective photochemical efficiency was determined during daytime (measured after 3 h of light exposure). Maximum photochemical efficiency was measured after 40 min of dark incubation during daytime (incubated after 3 h of light exposure) and the maximum value was selected for subsequent analyses. In addition, rapid light curves (RLC) were generated from a subset of corals (n ¼ 6 nubbins per species and per colony) to assess the maximum relative electron transport rate (rETRmax), the minimum saturating irradiance (Ek), the efficiency of light capture (a), and photoinhibition (b). The RLCs consisted of successive measurements of the effective quantum yield (Fv/Fm) under ambient light, where light intensity increased in 10 steps (0, 1, 30, 100, 197, 362, 618, 980, 1385, and 2014 mmol photon m
2 s
1). Hyperbolic tangent functions (Jassby and Platt, 1976), which included an exponent for photoinhibition at high irradiances (Platt et al., 1980), were fitted to each curve and rETRmax, Ek, a, and b were extracted for statistical analyses. Due to refinements of the settings during the experiment, the first two time points (t0 and after 6 weeks) were excluded from all photosynthesis analyses. 2.7. Symbiont densities and chlorophyll concentrations Coral nubbins were snap-frozen in liquid nitrogen at the end of the experiment. Symbiont densities and chlorophyll concentrations were determined for a subset of corals (n ¼ 3 per species and per colony). Coral tissue was removed completely with an airbrush using 13e90 mL micro-filtered (0.22 mm) autoclaved artificial seawater. For each tissue-seawater suspension, symbiont concentrations were determined under a microscope using a haemocytometer slide (n ¼ 8 replicate counts per sample). Counts were normalized to tissue surface area as derived from 3D scanning. Subsequently, chlorophyll concentrations were determined in a subsample of 10 mL of the remaining tissue suspension. To do so, the sample was centrifuged at 1600 g for 10 min. The supernatant was discarded and the symbionts re-suspended in 5 mL 100% acetone. Chlorophyll was extracted for 24 h in darkness and subsequently centrifuged at 1600 g for 10 min. Concentrations of chlorophyll a (chl a) and chlorophyll c2 (chl c2) were determined in the supernatant with a spectrophotometer (BioMate 3, Thermo Fisher Scientific, USA) at 630 and 663 nm (n ¼ 3 replicate measurements per sample) and calculated according to Jeffrey and

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Humphrey (1975). Total chlorophyll content was determined (chl a þ chl c2) and normalized to surface area and number of symbiont cells. 2.8. Statistical analyses

respectively). The effects decreased for both species over time (linear-mixed-effects model, ‘treatment interaction with time’ effect, P ¼ 0.0146 and P ¼ 0.0005, respectively). Further, microplastic exposure caused elevated DF/F'm in A. muricata (linear-mixed-effects model, ‘treatment’ effect, P ¼ 0.0045) as well as higher rETRmax and Ek in P. verrucosa (linear-mixed-effects model, ‘treatment’ effect, P ¼ 0.0055 and P ¼ 0.0008, respectively). However, these effects decreased over time (linear-mixed-effects model, ‘treatment interaction with time’ effect; A. muricata: DF/F'm P ¼ 0.0172, P. verrucosa: rETRmax P ¼ 0.0382 and Ek P ¼ 0.0062). While differences in photosynthesis parameters were strongest after 12 weeks and decreased gradually in P. verrucosa, A. muricata exhibited maximum differences after 18 weeks, followed by a decrease after 24 weeks.

All statistical analyses and data plots were performed in the R statistical environment (v.3.4.1; R Development Core Team, 2017). The effect of microplastic exposure on growth rates (surface area, volume, and calcification rate), photosynthesis (DF/F'm, Fv/Fm, rETRmax, Ek, a, and b), symbiont densities (cells cm
2), and chlorophyll concentrations (total chlorophyll content per cm2 and total chlorophyll content per symbiont-cell) were analyzed using linear mixed-effect models, generalized linear mixed-effect models (family Poisson), or linear models. Individual models were constructed for each response variable and species. Treatment and time were always included as fixed interaction effects. As random effects, colony, tank, and the individual nubbin were included in the model if applicable. If necessary, data were transformed (scaled or logged) prior to the analysis to ensure that test assumptions are met. Residual structures were checked visually by a graphical residual analysis. The models were fitted using the lme4 package (v.1.1e14, Bates et al., 2015). Bootstrap confidence intervals were calculated and p-values were derived testing estimated coefficients using the multcomp package (v1.4-7, Hothorn et al., 2008). Detailed specifications on the individual model equations are provided in Tables S4eS7. The effect of microplastics on coral health was analyzed using mixed-effect Cox models, based on cumulative incidence analyses, comparing the occurrence of negative health effects in the control and the microplastic treatments (Cox, 1972; Ripatti and Palmgren, 2000; Therneau et al., 2003). One model was constructed for each species. Tanks and colonies were specified as random effects. The models were fitted using the coxme package (v.2.2e5, Therneau et al., 2003) and plots were generated using the survival package (v.2.41e3, Therneau, 1999).

Signs of compromised coral health (i.e., bleaching, tissue necrosis, feeding scars from parasites) were documented throughout the experiment. While P. lutea and H. coerulea remained unaffected (Fig. 3, Table S8), both A. muricata and P. verrucosa showed first signs of compromised health six weeks after the start of exposure. However, treatment effects were only significant for P. verrucosa (Mixed-effects Cox model, ‘treatment’ effect, P ¼ 0.0400), where 16% (9 of 45) of exposed corals were negatively affected in contrast to 2% (1 of 45) of the control corals.

3. Results

4. Discussion

3.1. Coral growth rates decrease under microplastic exposure

The chronic exposure to microplastics resulted in speciesspecific patterns of stress response in major hermatypic corals. Growth parameters were negatively impacted in A. muricata and H. coerulea when exposed to microplastics. Further, photosynthetic performance of the associated photosymbionts changed under microplastic exposure for A. muricata and P. verrucosa, while P. lutea and H. coerulea showed no differences. In parallel, coral health was compromised in A. muricata and P. verrucosa, while P. lutea and H. coerulea remained unaffected. The reduced growth of A. muricata and H. coerulea (Fig. 1) indicates a stress response of the coral host, potentially associated with a depletion of the corals' energy reserves. The most likely causes of this stress response are, first, that the interaction with microplastics might impede heterotrophic feeding as previously shown for cold-water corals (Chapron et al., 2018) and freshwater polyps (Murphy and Quinn, 2018). Second, interaction, ingestion, and subsequent egestion of microplastics involves energetically costly movement of tentacles, tissue contraction, and movement of cilia on the coral tissue surfaces, with probably no or little nutritional value being derived from the consumed microplastics or its biofilm. Third, fractions of the ingested particles might be retained and potentially simulate satiation or cause blockages of digestive cavities, as observed in other coral species (Allen et al., 2017; Hankins et al., 2018). Besides feeding interactions, corals also respond to microplastic exposure with cleaning reactions, such as increased mucus production (Reichert et al., 2018). Mucus, released during periods of stress, is known to account for up to 50% of the

Growth parameters (i.e., changes in surface area, volume, and calcification rate) were evaluated analyzing 3D scans and buoyant weight. Microplastics had significant negative impacts on growth parameters of A. muricata and H. coerulea (Fig. 1, Table S4). For A. muricata, growth in surface area was 33% lower compared to the control treatment regardless of exposure time (linear-mixed-effects model, ‘treatment’ effect, P ¼ 0.0170). For H. coerulea, calcification rates were reduced by 3% under microplastic exposure (linear-mixed-effects model, ‘treatment’ effect, P ¼ 0.0051) with a decrease in effect strength over time (linear-mixed-effects model, ‘treatment interaction with time’ effect, P ¼ 0.0032). 3.2. Photosynthetic performance changes under microplastic exposure Maximum (Fv/Fm) and effective (DF/F'm) photochemical efficiency, maximum relative electron transport rate (rETRmax), minimum saturating irradiance (Ek), efficiency of light capture (a), and photoinhibition (b) were assessed with pulse-amplitude modulated (PAM) fluorometry after 12, 18 and 24 weeks. Under microplastic exposure, photosynthesis parameters significantly increased for A. muricata and P. verrucosa, while P. lutea and H. coerulea remained unaffected (Fig. 2, Tables S5 and S6). Especially, Fv/Fm was up to 4% higher for both A. muricata and P. verrucosa (linearmixed-effects model, ‘treatment’ effect, P ¼ 0.0038 and P ¼ 0.0239,

3.3. Symbiont densities and chlorophyll concentrations do not change under microplastic exposure Symbiont densities and chlorophyll concentrations were determined at the end of the experiment. Neither symbiont densities nor chlorophyll concentrations changed in A. muricata, P. verrucosa, P. lutea, or H. coerulea under microplastic exposure (Linear-mixed-effects models, ‘treatment’ effect, P > 0.05, Table S7). 3.4. Microplastics compromise coral health

J. Reichert et al. / Environmental Pollution 254 (2019) 113074

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Fig. 1. Growth rates for Acropora muricata, Pocillopora verrucosa, Porites lutea, and Heliopora coerulea under microplastic exposure (yellow) and control conditions (blue). Cumulative increases in surface area, volume, and calcification rate are normalized to surface area. Data are displayed as box-and-whisker plots with raw data points and p-values (treatment and interaction effects <0.05) derived from mixed-effects models. For specifications of model equations and detailed results see Table S4. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

carbon assimilated by photosynthesis (Crossland et al., 1980; Wild et al., 2004) and might thus also contribute to higher energy demands. In addition, microplastics might cause reductions in energy levels through direct or indirect damage to the organisms. Microplastics may accumulate toxins (Bakir et al., 2012; Lee et al., 2014; Teuten et al., 2009) and opportunistic coral pathogens (e.g., Vibrio spp.) (Kirstein et al., 2016; Zettler et al., 2013) commonly present in low abundances in the water column. These pathogens or damageassociated molecular patterns might be transmitted to the corals (Rotjan et al., 2019) and induce, for example, energetically costly

immune upregulations (Palmer et al., 2010; Sheridan et al., 2014; Tang et al., 2018). Corals mediate energetic depletion through different compensation mechanisms. They may, for example, respond with augmented heterotrophic feeding efforts (Palardy et al., 2005) e though this might be a self-amplifying feedback loop, further increasing the negative effects of the microplastics. Direct measurements of potential changes in heterotrophic feeding behavior during microplastic exposure are required to test this hypothesis. Alternatively, photosynthetic performance may be adjusted

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Fig. 2. Effects of microplastic exposure on photosynthesis parameters of Acropora muricata, Pocillopora verrucosa, Porites lutea, and Heliopora coerulea. a) Composite rapid light curves for corals under microplastic exposure (yellow) and control conditions (blue) give the relative electron transport rate (rETR, arbitrary units (a.u.)) as a function of photosynthetically active radiation (PAR, mmol photons m
2 s
1), pooled over time. Rapid light curves are depicted as mean ± standard deviation and non-linear regressions fits (solid lines). b) Z-values of linear mixed-effects models estimating the effect of microplastic exposure (treatment effect) and its interaction with time (interaction effect) for the photosynthesis parameters DF/F'm, Fv/Fm, rETRmax, Ek, a, and b. For specifications on model equations and detailed results see Table S5. Significance levels: *P < 0.05, **P < 0.01, ***P < 0.001. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

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Fig. 3. Effects of microplastic exposure on coral health. a) Signs of compromised health in Acropora muricata, Pocillopora verrucosa, Porites lutea, and Heliopora coerulea over time under microplastic exposure (yellow) and control conditions (blue). Graphs give the number of coral nubbins (in percent) that showed >50% tissue loss (solid line) together with 95% confidence intervals (dashed line) and p-values (P < 0.05) derived from mixed-effects Cox models. b) Bleaching and necrosis occurred in A. muricata and P. verrucosa, shown in 3D models (top) and close-up photographs (bottom), white scale bar: 1 cm. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

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The compromised coral health observed (Fig. 3) also hints towards a depletion of the existing energy reserves, which likely increases the risk of adverse effects from other external stressors (Grottoli et al., 2006) and susceptibility to pathogens or parasites (Palmer et al., 2010). This might have fostered bleaching or necrosis in A. muricata and P. verrucosa. The observed effects of microplastic exposure were speciesspecific. Our findings suggest that A. muricata and P. verrucosa were most strongly affected as they were not able to compensate the increased energetic needs over a longer period of time. For H. coerulea the impact of the microplastics appeared to be less severe as the effects decreased over time and no changes in photochemistry or signs of compromised health were observed. This might indicate that H. coerulea has the capacity to buffer or even adapt to the negative impacts of microplastics. The underlying processes of these mechanisms need to be elucidated in further studies, addressing energy budgets, immune responses, and toxin or pathogens transmission. P. lutea in contrast remains apparently unaffected under microplastic exposure. The species-specific effects are likely driven by coral host characteristics and bio-physical feedbacks. Especially those species that are known to actively interact with microplastics (Reichert et al., 2018) were affected. Most likely, the microplastic particles are mistaken for food, and corals frequently catching and feeding on particulate matter might be particularly sensitive to microplastic exposure. This goes along with the observation that cold-water corals, which rely entirely on heterotrophic feeding, are strongly affected by microplastic exposure (Chapron et al., 2018), although the effects seem to be species-specific (Mouchi et al., 2019). Feeding on particulate matter is a common strategy of Acropora and Pocillopora spp. but rarely occurs in Porites spp. (Anthony, 1999; Palardy et al., 2005). Unfortunately, little is known about the feeding behavior of Heliopora sp., making it difficult to relate feeding strategy and stress response. Apart from that, the affected species share a higher structural complexity, which might increase both the probability of the corals to encounter particles and the capture efficiency of the polyps (Patterson, 1992). In addition, Porites spp. are known to exhibit effective cleaning mechanisms through the production of mucus layers (Coffroth, 1990; Edmunds and Davies, 1986), which might alleviate the direct impacts of the microplastics. In addition, tissue biomass and tissue thickness, which is higher in massive Porites spp., allow for higher photosymbiont densities and greater energy reserves (Loya et al., 2001; Putnam et al., 2017). This might also provide greater resistance against the impacts of microplastics. 5. Conclusions

(Anthony and Fabricius, 2000; Anthony and Larcombe, 2000) to increase energy acquisition in response to microplastic exposure, as seen in our study for A. muricata and P. verrucosa (Fig. 2). However, the changes in the photosynthetic parameters measured in the current study differed in direction and magnitude. For example, the increase in Fv/Fm indicates increased photosynthetic efficiency whereas the increase in Ek indicates decreased photosynthetic efficiency at low light levels. Remarkably, the effects on the photosynthetic activity decreased over time, suggesting that any upregulation of photosynthesis was transient, making this mechanism unlikely to boost coral energy acquisition in the long-term. This goes along with the observations, that symbiont properties, which often reflect changes in photochemistry (Philipp and Fabricius, 2003) and were studied at the end of the experiment, did not differ between treatments. Similar effects have been observed for corals under varying particle levels (Fabricius, 2005), potentially indicating limitations in nutrient levels, light, or heterotrophic food sources.

Overall, our findings provide evidence that microplastic exposure can have negative effects on important hermatypic corals. Although the impact observed during our six-month study was generally small and microplastic concentrations may still be lower in many natural systems (Reisser et al., 2013; Syakti et al., 2017), effects may cumulate over a coral's lifetime. In addition, the species-specific responses indicate that taxa known to feed on particulate food sources (i.e., Pocillopora spp. and Acropora spp.), and thus more frequently interact with microplastics (Reichert et al., 2018), were most affected by the exposure. This is of concern as these species are also particularly vulnerable to other stressors, such as increasing water temperatures, high light, and low water quality (Anthony and Kerswell, 2007; Putnam et al., 2017). In natural reef systems, microplastic pollution may add up to existing stressors and amplify the corals' susceptibility to bleaching and diseases. This might further drive corals towards a critical tipping point and foster community shifts in coral reef

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assemblages. Therefore, microplastic pollution should be considered as anthropogenic stressor in future environmental assessments and management actions for coral reefs. Likewise, immediate actions are needed to reduce plastic pollution by addressing the overuse of single-use plastics and improving wastewater treatment and recycling processes. Author contributions statement JR, AA, PS, and TW conceived the ideas and designed the methodology; JR and AA collected the data; JR, AA, and MH analyzed and interpreted the data; JR and TW led the writing of the manuscript. All authors contributed critically to the drafts and gave final approval for publication. Data availability The analyzed data and 3D meshes of the studied corals are made available at https://doi.org/10.6084/m9.figshare.9705263. Conflicts of interest The authors declare no competing interests. Acknowledgements This study was conducted as part of the ‘Ocean 2100’ global change simulation project of the Colombian-German Center of Excellence in Marine Sciences (CEMarin) funded by the German Academic Exchange Service (DAAD, project number 57480468). We thank all B.Sc. and M.Sc. students who helped with the experiment during their studies at Justus Liebig University Giessen. We also thank Christian R. Voolstra and Omar El Tall from King Abdullah University of Science and Technology for providing the opportunity and support to perform the FTIR analyses. We further thank Stefanie P. Glaeser from Justus Liebig University Giessen for the insights into the microbiological aspects of the experiment. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.envpol.2019.113074. References Allen, A.S., Seymour, A.C., Rittschof, D., 2017. Chemoreception drives plastic consumption in a hard coral. Mar. Pollut. Bull. 124, 198e205. https://doi.org/ 10.1016/j.marpolbul.2017.07.030. Andrady, A.L., 2017. The plastic in microplastics: a review. Mar. Pollut. Bull. 119, 12e22. https://doi.org/10.1016/j.marpolbul.2017.01.082. Andrady, A.L., 2011. Microplastics in the marine environment. Mar. Pollut. Bull. 62, 1596e1605. https://doi.org/10.1016/j.marpolbul.2011.05.030. Anthony, K.R.N., 1999. Coral suspension feeding on fine particulate matter. J. Exp. Mar. Biol. Ecol. 232, 85e106. https://doi.org/10.1016/S0022-0981(98)00099-9. Anthony, K.R.N., Fabricius, K.E., 2000. Shifting roles of heterotrophy and autotrophy in coral energetics under varying turbidity. J. Exp. Mar. Biol. Ecol. 252, 221e253. https://doi.org/10.1016/S0022-0981(00)00237-9. Anthony, K.R.N., Kerswell, A.P., 2007. Coral mortality following extreme low tides and high solar radiation. Mar. Biol. 151, 1623e1631. https://doi.org/10.1007/ s00227-006-0573-0. Anthony, K.R.N., Larcombe, P., 2000. Coral reefs in turbid waters: sediment-induced stresses in corals and likely mechanisms of adaptation. In: Proceedings of the 9th International Coral Reef Symposium, pp. 239e244. Bakir, A., Rowland, S.J., Thompson, R.C., 2012. Competitive sorption of persistent organic pollutants onto microplastics in the marine environment. Mar. Pollut. Bull. 64, 2782e2789. https://doi.org/10.1016/j.marpolbul.2012.09.010. Bates, D., M€ achler, M., Bolker, B., Walker, S., 2015. Fitting linear mixed-effects models using lme4. J. Stat. Softw. 67, 1e48. https://doi.org/10.18637/jss.v067.i01. Brocher, J., 2015. The BioVoxxel image processing and analysis toolbox. In: European BioImage Analysis Symposium. Paris, France. Browne, M.A., Chapman, M.G., Thompson, R.C., Amaral Zettler, L.A., Jambeck, J.,

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