Oyster larval microbiome and its manipulation for the improvement of shellfish aquaculture

Abstract

Aquaculture is the practice of farming aquatic species and contributes significantly to global seafood production. Aquaculture production has increased dramatically since the 1950’s, and with this, so has the production of molluscs. Oysters make up the most commonly produced group of molluscs by quantity, being produced for both food sustainability and ecological restoration goals. Importantly, the security and expansion of oyster farming is reliant upon reliable production of good quality seedstock from hatcheries. The production of such seedstock is, however, greatly hindered by large-scale disease mortality events that often occur during the larval stages of the oyster lifecycle. The causes of these disease mortality events are known to be often caused by dysbiosis of the larval microbiome (the collection of microorganisms including bacteria, viruses and fungi that live within the larvae and their environment). It is also is well accepted that the microbiome plays a key role in maintaining larval health. Research into the bacteria that reside within oyster larvae and their environments is therefore vital to understand larval health and improve the ways in which they are farmed. This research is also of benefit to aquaculture generally with methods of improving oyster aquaculture being possibly applicable to other key bivalve species such as mussels, scallops or clams. Within a broader context still, the improvement of oyster aquaculture is beneficial to restoration efforts, where increased oyster production allows for the improvement of water quality, substrate stabilisation and increased biodiversity in coastal ecosystems.

This thesis aims to improve the current understanding of the larval microbiome and investigate possible methods of microbial manipulation for improved larval heath within the hatchery setting. Chapter two addressed this with an environmental microbiome study of the European flat oyster (Ostrea edulis). This was done via the collection of water samples taken throughout a natural spawning event of O. edulis at Loch Ryan, UK. It was reported that microbial changes within this spawning event were most closely correlated with date and driven by the differences in abundance of certain bacterial taxa such as those belonging to the Rhodobacteraceae family. Inverse abundance profiles were also observed between bacteria belonging to the Rhodobacteraceae family and Vibrio genera. With species of the Vibrio genera being known to cause disease mortality events in larvae, this finding allowed for the formation of a hypothesis that Sulfitobacter and Jannaschia genera of Rhodobacteraceae may be suitable probiotic candidates for usage within oyster hatcheries. This hypothesis was tested through both bacterial plate assays and in vivo challenges in adult oysters throughout chapter five. Although preliminary, these challenges provided promising data. Most importantly, the addition of Rhodobacteraceae to seawater improved oyster survival rate when challenged with Vibrio aestuarianus. This finding has possible applications to improving oyster heath and cultivation. Chapter five also discusses future experimental work needed to replicate these findings within larger sample sets.

Similarly, chapter three was carried out with the ultimate aim of improving larval health within the hatchery environment for improved oyster production. This chapter again uses full-length 16S rRNA sequencing to provide baseline bacterial profiles throughout commercial production at two different UK hatcheries, covering species of both O. edulis (produced at Portsmouth) and Crassostrea gigas (C. gigas, produced at Morecambe Bay). The core larval microbiomes from each hatchery were identified and microbial species richness tended to decrease as the larvae developed. Microbial profiles of larvae were also found to be distinct to that of their surrounding water. This broad-scale microbial data also allowed for the microbial implications of hatchery practices to be discussed with water filtration techniques at both locations being successful in reducing bacterial diversity within the water samples. Generally, this chapter improves the current understanding of the larval microbiome and their hatchery environment whilst also fulfilling a knowledge gap by conducting this research throughout commercial production.

Chapter four builds upon previous baseline data by investigating possible methods of microbiome manipulation to improve larval health and negate the need for antibiotic usage within hatcheries. The growth rates of larvae which were reared up to 15-days post fertilisation under different diet and treatment groups were collected. Microbial comparisons were also reported via both culture dependent and independent techniques. Most notably, the microbiomes of larvae grown under two different dietary treatments were found to be distinct from one another, with differences being driven by the abundance of bacterial taxa such as Actinobacteria, Bdellovibrionia and Bacilli.

In summary, this thesis provides vital baseline data about the O. edulis and C. gigas larval microbiomes throughout a range of different spawning types and locations. This has allowed for an improved understanding of the larval microbiome which is vital in order to better inform hatchery practices and improve production. Chapters four and five build upon the previous broad-scale data collected in two and three by demonstrating possible methods of targeted microbial manipulation.