Coral sampling

Sampling and imaging of corals were conducted at 540 m depth in the Lacaze-Duthiers canyon, northwestern Mediterranean Sea (42°32′72″N, 03°25′28″E) on the long-term site monitored since 2009 under the programme ‘Biodiversity, extreme marine environment and global change’ (LECOB) that had revealed large amounts of plastic debris in the canyon53,54 (Fig. 3). Coral branches were sampled from three distinct live colonies of L. pertusa in June 2016 using a Remotely Operated Vehicle (ROV) deployed from the R/V Janus II (COMEX company). On board, corals were transferred to aerated 30 L seawater tanks maintained at 13 °C. Once in the laboratory, corals were stabulated in thermoregulated room in the dark in a 80 L aquaria that received continuous flow (>1 renewal day−1) of filtered (5 μm) Mediterranean seawater pumped from 5 m depth55.

Figure 3 Representative views of plastic pollutions on Lophelia pertusa reefs in the Lacaze-Duthiers canyon, northwestern Mediterranean Sea. Different types of plastics are found in coral reefs, most of them are bags (A) and bottles (B). Dying (C) or dead (D) coral colonies covered by plastic bags. The distance between green dots is 6 cm. Fondation TOTAL/UPMC. Full size image

Plastics

Polyethylene is the most common plastic type used in the world and is found in oceans as micro- and macro-debris1,56,57. The macroplastics used in the experiment were 10 cm × 10 cm sized pieces of low density polyethylene (LDPE) meant to represent pieces of plastic bags. The macroplastic sizes were chosen to cover the entire height of the coral branch. The microplastics were composed of 500 μm LDPE microbeads (Santa Cruz Biotechnology Inc., Dallas, USA). The microplastic size corresponds to the size of the zooplankton commonly used to feed corals in aquaria58,59,60,61. Before the start of the experiment, both macro- and microplastics were incubated for two months in 5 L seawater tanks continuously supplied with filtered Mediterranean seawater. The goal of the pre-incubation was to allow colonization by environmental bacteria, a phenomenon observed in natural environments.

Experimental design

Experiments were conducted in recirculating flumes of 58 L each designed to maintain a regular and user defined rate of water circulation52. The flumes were maintained in the dark at 13 °C ± 0.5 °C in a thermoregulated room. A motor (Modelcraft) driven propeller maintained a constant flow of 2.5 cm s−1 in each flume. Each system was semi-closed with a continuous supply of 2.5 L h−1 of oxygenated, thermoregulated and filtered (5 μm) Mediterranean seawater (corresponding to one renewal per day). A 180 μm mesh at the spillway retained particles within the tank. Three different flumes were used to maintain three different experimental conditions for 2 months: one control, one with microplastics and one with macroplastics (Supplementary Fig. S1).

A total of 15 coral nubbins from the three distinct sampled colonies (each nubbin containing 4 to 33 living polyps) were randomly assigned to use within each flume. Nubbins were fixed to cement blocks using an aquatic epoxy resin62. Corals were acclimated in the flumes for four weeks before the addition of plastics. For microplastic conditions, the beads were added to the flume at a concentration of 350 beads L−1, which corresponds to the concentration of the zooplankton preys commonly used for regular feeding within the laboratory. The microplastic to zooplankton ratio that we use has been seen in the Mediterranean Sea surface waters with microplastic concentrations ranging from 0 to 2.28 mg L−1, and polyethylene being the most common plastic63,64. However, there are no precise quantifications of microplastics in the deep-sea in general, and in the Lacaze-Duthiers canyon in particular. Considering that not all the surface microplastics reach the sea floor, our experimental concentrations may be higher than in situ concentrations. Our results must thus be considered in the light of that caveat. For macroplastic conditions, the plastic pieces were placed in the flumes to partially cover (~50%) living polyps over the entire height of the coral branch, simulating conditions that we observed in the field (Supplementary Fig. S1) and that have been reported by others on deep sea corals in the Mediterranean Sea28,53,54. The macroplastics covered the living polyps that faced the current but not the polyps that were on the other side of the coral branch. Corals were fed three times a week alternately with freshly hatched Artemia salina nauplii (350 A. salina L−1) and with Marine Snow plankton diet (Two Little Fishies Inc, Miami Gardens, USA, 5 mL per flume), to provide a complete and diverse nutrient supply36,61.

Prey capture rate

Coral capture rate was measured 7, 20 and 47 days after the start of the experiment using a method published by Purser et al.52. Triplicate water samples (100 mL) were taken each hour after feeding over a 4-hour period and filtered on 55 μm mesh. Artemia salina nauplii were counted on the mesh with a binocular magnifier to calculate the concentration of suspended A. salina remaining in each flume. The number of zooplankton prey removed by coral capture was normalized against the number of living polyps present in the flume. Control measurements were also conducted in flumes containing no corals to quantify zooplankton sedimentation. The consumption of A. salina per polyp was corrected against control measurements. Most of the zooplankton consumption by L. pertusa occur during the first hour following prey delivery52 (Supplementary Fig. S2), and our results thus focus on this time period.

Polyp activity

Cold-water coral polyp behaviour can be studied by video monitoring65,66. We applied this approach to measure polyp activity using two-hour video captures. Two cameras (GoPro, San Mateo, USA) were installed in each flume. To maximize the total number of polyps observed, one camera filmed a lateral view and the other an aerial view of the corals. The activity rates were measured by analysing differences between consecutive images extracted from the videos67. Three subsamples of 20 images were extracted every 6 minutes from the video and combined into an image stack. Optimisation tests showed that the chosen time interval and number of images allowed to determine the highest number of active polyps for the duration of the sequence (Supplementary Fig. S3). The percentage of active polyps was calculated by counting the total number of moving polyps in each view using the Image J v1.51 software and by dividing this number by the total number of polyps.

Coral growth rates

Sclerochronological analysis was used to measure coral growth rates, after 2 months of exposure, on three replicate nubbins from each colony (each containing 5 or 6 living polyps) for each experimental condition. Polyps were labelled with a fluorescent calcein staining (150 mg L−1) at the start of the experiment as described in Lartaud et al.68. At the end of the experiment, the coral nubbins were cleaned for 12 hours in a hydrogen peroxide solution (H 2 O 2 , 4%) at 60 °C to remove organic tissues, and then rinsed in demineralized water. Each polyp calyx was then separated from the others, embedded into epoxy resin Sody33 for 24 hours and cut into slices with a Buehler Isomet low-speed saw. The coral sections were mounted on slides with Araldite© resin, abraded and then polished, for subsequent observation under an epifluorescence microscope (Olympus IX51) with an excitation at 495 nm. The new skeleton formed between the calcein marking (i.e., beginning of the experiment) and the summit of the septum of the calix (i.e., date of death) was measured using the ImageJ© software (repeated 5 times) by superposing the fluorescence pictures, which revealed the stain, and the optical light pictures showing the coral skeleton morphology68 (Supplementary Fig. S4).

To compare experimental and in situ growth rates, an analysis was also conducted on L. pertusa fragments that had been marked and redeployed in their natural habitat for ca. 2 months. The in situ corals used originated from an earlier study69 conducted at the same location in the Lacaze-Duthiers canyon and for the same duration.

Statistical analysis

Tests for normality of variance were performed using the Shapiro-Wilk test on R software (v3.4.3), which revealed that the distribution was not normal for feeding rates, percentages of active polyps or skeleton growth rates (p < 0.05). A multiple-comparison non-parametric Kruskal-Wallis test was thus used to analyse the differences between the three experimental conditions and for differences between 7, 20 and 47 days for each parameter investigated.