The corallivorous flatworm Amakusaplana acroporae: an invasive species threat to coral reefs?
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Fatal infestations of land-based Acropora cultures with so-called Acropora-eating flatworms (AEFWs) are a global phenomenon. We evaluate the hypothesis that AEFWs represent a risk to coral reefs by studying the biology and the invasive potential of an AEFW strain from the UK. Molecular analyses identified this strain as Amakusaplana acroporae, a new species described from two US aquaria and one natural location in Australia. Our molecular data together with life history strategies described here suggest that this species accounts for most reported cases of AEFW infestations. We show that local parasitic activity impairs the light-acclimation capacity of the whole host colony. A. acroporae acquires excellent camouflage by harbouring photosynthetically competent, host-derived zooxanthellae and pigments of the green-fluorescent protein family. It shows a preference for Acropora valida but accepts a broad host range. Parasite survival in isolation (5–7 d) potentially allows for an invasion when introduced as non-native species in coral reefs.
KeywordsAcropora-eating flatworms Amakusaplana acroporae Invasive species GFP-like fluorescent proteins Corallivory Photoprotection Green fluorescent protein Biomarker
Coral reefs, one of the most biodiverse and productive ecosystems in the world, are sensitive to a range of perturbations including the potentially devastating effects of corallivory, defined as the direct assimilation of live coral tissue (Hughes et al. 2007; Rotjan and Lewis 2008).
To judge the danger that AEFWs represent as potential invasive species for natural coral reefs, we studied the biology of an AEFW strain obtained from the ornamental trade in the UK. We applied molecular taxonomy approaches to identify the species, analysed host preference, feeding behaviour, association with zooxanthellae, effects on the host physiology and survival times in the absence of hosts.
Culture and aquarium experiments
An AEFW strain acquired from a local ornamental trader in Southampton (UK) was co-cultured for 6 months with a diverse range of acroporids and numerous other coral species (ESM Table 2) in a separated compartment of the experimental coral mesocosm of the Coral Reef Laboratory at the National Oceanography Centre Southampton (D’Angelo and Wiedenmann 2012). Host preference was judged using a classification scheme shown in ESM Table 2. For subsequent studies, AEFWs were dislodged from the host by a seawater jet. Survival in the absence of host corals was evaluated by maintaining ten isolated specimens in a partially shaded compartment without access to corals. To test the response of the AEFWs to increased temperatures, infested fragments of Acropora sp. were subjected to a heat stress treatment as described in (Wiedenmann et al. 2013). Horizontally growing Acropora millepora replicate colonies (infested and non-infested) were turned by 180°, and light acclimation was monitored through the fluorescence increase in the newly light exposed branch surface (D’Angelo et al. 2008).
Molecular identification of AEFW and zooxanthellae
Genomic DNA (gDNA) was prepared from a pool of 6 AEFWs collected from an A. millepora colony, the infested host colony itself and a co-cultured Acropora microphthalma colony using a protocol described in (Hume et al. 2013).
Phylotyping of zooxanthellae (Symbiodinium spp.) in the corals and in the AEFW was conducted by amplifying, cloning and sequencing of the ITS1-5.8S-ITS2 ribosomal DNA using zooxanthellae-specific primers (ESM Table 3) (Hume et al. 2013; Savage et al. 2002). Sequences were deposited in Genbank (accession numbers JN711475–JN711498).
The 18S rDNA region of the AEFW genomic DNA was amplified as two overlapping fragments, using primers designed against conserved regions of polyclad 18S sequences (ESM Table 3). A primer pair to amplify a 618-bp fragment of the 28S region was developed using polyclad 28S sequences available in GenBank (ESM Table 3). Sequences were deposited in Genbank (accession numbers JN711499–JN711500).
Phylogenetic analysis of 18S and ITS2 sequences was performed using MEGA 5 (Tamura et al. 2011). Sequences were aligned using ClustalW, and phylogenetic trees were constructed using maximum likelihood (ML) methods. Analysis was performed to infer an optimal nucleotide substitution method. For 18S analysis, a Tamura-Nei with gamma distribution and 5 rate categories were selected based on Akaike Information Criterion. ITS2 sequences were analysed using the Hasegawa–Kishino–Yano model with equal rates. The certainty of ML nodes was tested with bootstrap analysis (100 replications).
Photographic documentation, fluorescence measurements and microscopy
Coral fluorescence was imaged using a yellow longpass filter and a ~450-nm excitation light source (Nightsea, Andover, USA). Microscopic close-ups were obtained as described in (D’Angelo et al. 2012; Smith et al. 2013).
Photosynthesis efficiency of zooxanthellae was determined by pulse amplitude modification (PAM) fluorometry using a Diving-PAM (Walz) for a pool of 5 AEFW specimens.
Results and discussion
Interestingly, the AEFWs collected from A. pulchra show fluorescence patterns perfectly matching those of the host, with the coral fluorescence being derived from the cyan GFP-like protein apulFP583 (D’Angelo et al. 2008) and red chlorophyll fluorescence of zooxanthellae (Oswald et al. 2007) (Fig. 2). In contrast, depending on their developmental stage, eggs and embryos show only a weak blue or orange fluorescence under UV (365 nm) excitation (ESM Fig. 1c). The absence of fluorescence in the feeding marks in the coral tissues proposes that the parasites extract the fluorescent pigments from the host (Fig. 2f). The cyan fluorescent pigments appear to be evenly distributed in the parenchyma of the parasites, suggesting an incorporation of host pigments in their functional form to perfect the camouflage of the parasite (Fig. 2b, d). Such strategy could be facilitated by the high stability and slow turnover of coral GFP-like proteins (Leutenegger et al. 2007).
Sequence analyses of the 18S rDNA region demonstrated that the UK AEFW strain groups within the cotylean clade of polyclad flatworms (ESM Fig. 3a), ruling out the phenotypically similar Acropora-associated Waminoa sp. from Okinawa (Matsushima et al. 2010). Analysis of the 28S rDNA region identified the UK strain as the recently described A. acroporae, an AEFW from two US aquaria and from one natural location on Lizard Island (Rawlinson et al. 2011; Rawlinson and Stella 2012) (ESM Fig. 3b). Our 28S rDNA sequence showed a 100 % identity to the Virginia strain of A. acroporae, but could be distinguished in three shared residues from the New York and the Lizard Island strains. These nucleotide substitutions may indicate that at least two molecularly distinct A. acroporae strains are widely distributed in aquaria. Our global survey of reported AEFW infestations (Fig. 1, ESM Table 1) together with the results of the present paper revealed that many of the unidentified AEFWs show characteristics of A. acroporae such as exclusive preference for a diverse range of acroporids, excellent camouflage, preferred occurrence on the shaded branch sides, the typical ‘bite marks’ and egg clusters being deposited on dead skeletons (ESM Table 1). A. acroporae was considered responsible for the loss of Acropora colonies at Birch Aquarium (USA) (Nosratpour 2008; Rawlinson et al. 2011). Taken together, these data suggest that A. acroporae is globally distributed in land-based coral cultures, including regions close to natural coral reefs such as Florida, Thailand or Hong Kong (Fig. 1). We conclude that due to the broad range of accepted host coral species with a high preference for the cosmopolitan A. valida, the excellent camouflage and the ability to survive at least 5 days without host, A. acroporae has the potential to become a dangerous invasive species when released to an environment to which it is non-native. To better categorise this potential risk, further data on the biogeographic range of A. acroporae in reefs around the globe, on its natural predators and on its genetic diversity in captivity are urgently required.
We thank the reviewers of this manuscript for their helpful suggestions. Funding: NERC (NE/K00641X/1 to JW; studentship to BCCH/JW), the European Union’s Seventh Framework Programme (FP/2007-2013)/ERC Grant Agreement No. 311179 to JW.
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