Hello everyone !

My name is Mark, a marine biologist from England who had the pleasure of working with the Marine Savers team from May to August 2016. I have always been passionate about marine life and conservation and since graduating from Bangor University (Wales) with a Masters in Marine Biology, I have been eager to gain as much practical experience in marine conservation as possible. After discovering Seamarc, Marine Savers and the work they do, I couldn’t wait to get involved.

During my time in the Maldives I was lucky enough to be able to spend time at both resorts – Four Seasons Kuda Huraa and Four Seasons Landaa Giraavaru – and had the opportunity to assist on a wide range of conservation projects as well as conducting the daily guest excursions. The teams at both resorts are a fantastic group of people and I enjoyed every moment working with them all.

Whilst working at Landaa Giraavaru I was also able to assist in the ornamental fish breeding lab; helping to spawn and rear the current clownfish residents. For me personally, marine ornamental aquaculture (culturing ornamental fish and invertebrates in captivity to supply the global aquarium industry) is an area of marine conservation that I am particularly interested in, having specialised in this field for my Master’s degree. I was therefore extremely grateful to be given the chance to work with the team on the various breeding projects in the lab. The following report highlights some of the work I was involved in (and you might also be interested in my earlier Marine Biology blog).

At home in England

At home in England my first ever job was working in a large public aquarium where I was surrounded by all sorts of amazing aquatic creatures on a daily basis. Although I loved the job, one thing that I was always conscious of is the fact that over 90% of the marine fish and invertebrates currently traded for the home/public aquarium industry are taken directly from the wild. It is estimated that between 1.5 to 2 million people keep marine ornamentals in home aquaria worldwide (Wabnitz et al., 2003) and it is predicted that this number will continue to rise as the cost of setting up and maintaining a marine aquarium becomes more accessible to many people. Due to such a huge reliance on wild stock to supply this industry, it has become a conservation concern as the vast majority of marine ornamentals are captured from coral reefs – ecosystems that are already under threat from human-related activities such as pollution, habitat destruction and global warming.

Marine ornamental aquaculture is now cited as the best potential solution for producing and supplying marine species for the aquarium trade. At present, the number of marine species that can be successfully cultured in captivity is relatively low, but thanks to constant advances in technology and greater knowledge of the requirements of certain species, this number is increasing. Not only will culturing marine ornamentals in captivity reduce the need to take them from the wild, it is well known that captive-bred animals have many advantages over their wild counterparts when kept in aquaria. This is because captive-bred animals are raised in tanks and so become used to ‘tank life’. Captive-bred animals also become more tolerant of changes in water chemistry and tend to live longer in aquaria.

In the Maldives, fishing for the aquarium trade is highly regulated in order to protect the reefs and safeguard tourism. These regulations are great for conserving the reefs, but they do limit the growth of a potentially highly profitable fishery. At the Marine Discovery Centre at Landaa Giraavaru, the ornamental Fish Lab was set up not only to help promote and educate people on marine conservation issues, but also to carry out further research into the breeding of ornamental species, with the aim of eventually establishing ornamental breeding facilities on neighbouring local islands. In the long term this would certainly benefit local people by providing an alternative livelihood to fishing.

Maldivian Clownfish (Amphiprion nigripes) with eggs

Clark’s clownfish (Amphiprion clarkii) in anemone

Clark’s clownfish (Amphiprion clarkii) in anemone

My Work in the Fish Lab

The Fish Lab is home to two different species of clownfish, the bold and attractive looking Clark’s clownfish (Amphiprion clarkii) and the Maldivian or ‘Black-Foot’ clownfish (Amphiprion nigripes), found only in the waters of the Maldives and Sri Lanka. The breeding projects have two main objectives:

1) To produce captive-raised fish that can eventually be sold on to supply the aquarium industry.
Clownfish are some of the most popular marine ornamentals kept in aquaria worldwide. However, if we continue to collect these fish from the wild they may eventually disappear from the reefs altogether. Luckily, clownfish will readily reproduce in captivity and in the Fish Lab the aim is to produce happy, healthy and hardy fish that can eventually be sold on to the aquarium market. Studying these fish in the lab also allows us to better understand their life-cycle and allows us to refine and improve culture techniques.

2) To produce captive-raised fish that can be released into the wild.
Individuals are released when they reach a certain size in order to maximize their chance of survival. In the Fish Lab we were also looking at ways to culture host anemones such as the Magnificent Anemone (Heteractis magnifica) to provide the clownfish with a home when released. This will also increase their chances of survival in the wild.

The Clark’s clownfish (also known as the yellowtail clownfish) is a widely distributed species that inhabits tropical lagoons and outer reef slopes. It reaches 15cm in length and with its attractive coloration and hardy nature, has become a firm aquarium favourite.

During my time in the lab, there were eleven tanks containing Clark’s clownfish, six tanks of adult broodstock pairs and five of fish of various ages. There were also an additional five tanks that were set up for larval rearing. Each day in the Fish Lab I assisted with all of the feeding, cleaning and general maintenance of all the tanks containing A. clarkii. Feeding occurred twice a day; once in the morning and again in the afternoon. All tanks were cleaned every couple of days in order to remove any uneaten food and fish waste.

Broodstock Pairs

The Clark’s clownfish is a monogamous species, with a dominant female pairing with a dominant male. There were 6 pairs of broodstock A. clarkii in the Fish Lab during June-July and several of the pairs were producing eggs regularly. The broodstock pairs were housed in separate rectangular 120 litre aquariums (60 x 47 x 50cm). Water inflow / outflow was constant and each tank was provided with aeration. Each tank also contained a host anemone of the species Heteractis magnifica (‘Magnificent anemone’) as well as a ceramic plant pot or tile to provide the hard surface which clownfish like to lay their eggs on. The broodstock pairs were fed twice a day, with their diets consisting of a blended shrimp paste and a dry pellet feed. Each day we would monitor fish behaviour and check for the presence of any eggs. In the days leading up to laying, adults were often observed cleaning and preparing the lay site.

On 23 June, the largest (and assumed oldest) pair of A. clarkii successfully spawned in the lab. The clutch size was fairly large and the eggs were attached to the side of their ceramic pot, which made the eggs very easy to spot for any visitors to the Fish Lab. Like others who have observed A. clarkii spawning in captivity (Krishna et al., 2015), the newly laid eggs were capsule-shaped and appeared to measure around 2-2.4mm in length. Once laid, the male was regularly observed fanning the eggs with his pectoral fins in order to ensure a continuous flow of aerated water over the developing embryos. He would also remove any damaged or infertile eggs so the clutch is kept healthy. The eggs also underwent several distinct colour changes during the following 6 days, changing from their initial reddish-orange colour to a striking silver. Once silver, the larvae’s eyes were clearly visible in the egg capsule and as in other studies (Swagat Ghosh et al., 2011) the eggs hatched within 12 hours of reaching this stage.

Hatching & Transfer of Larvae

Hatching took place on the evening of 29 June under ‘black-out’ conditions. That evening at 20:00h, all lights in the Fish Lab were switched off and black plastic covers were placed around the sides of the broodstock tank in order to make the tank as dark as possible. The water flow and aeration were temporarily switched off. After approximately 30 minutes under ‘black-out’ conditions the larvae began to hatch out. Once the majority of eggs appeared to have hatched, a fluorescent light was then hung in the corner of the broodstock tank. The new larvae are highly photopositive and so move towards this light, enabling us to carefully siphon them off into a large bucket and into a larval rearing tank. Once the larvae were removed the water flow and aeration was returned to the adult’s tank.

The larval rearing tank used in this study was the same size and volume as the adult broodstock tank. Two 20 litre buckets of broodstock tank water containing larvae was added to the rearing tank along with 5 litres of algae (Nannochloropsis) and 1 litre of cultured rotifers. No further water was added to the rearing tank and there was no water inflow/outflow. A lower water volume is necessary in the larval rearing tank as it condenses the food source, meaning the larvae have to travel less distance in order to capture their tiny prey. As the larvae are attracted to light, all sides of the rearing tank were covered with black or blue aquarium backing and the tank was only lit from above. The tank was also given very gentle aeration and was then covered and left overnight.

For the first 12 hours or so, the tiny larvae naturally fed on their own yolk sac and then moved onto rotifers once this was depleted. During this time the rotifers were busy consuming the algae, making them more nutritious and a perfect first feed.

Larval Husbandry

Each day the larval rearing tank was cleaned using a siphon and bucket. On the first day of my study no water was removed from the rearing tank as the larvae were still consuming the initial feed of rotifers from the night before. From the second day onwards a water change was done every morning and if needed, again in the afternoon (a water change was required in the afternoon if there was a large amount of larval mortality throughout the day). After each water change, the number of mortalities was recorded.

Initially, the larvae were fed with up to one litre of cultured rotifers enriched with SELCO every morning and evening. SELCO (Self-Emulsifying Liquid Concentrate) is a formula that was used to increase the levels of Highly Unsaturated Fatty Acids (HUFA) in the rotifers making them more nutritious for the larvae. The amount of food given to the larvae was adjusted depending on how much food was still visible in the tank from the previous feed; this was to prevent over-feeding which could affect water quality. From day 7 onwards the larvae were also fed with newly hatched Artemia shrimp. At this stage, the larvae were developed enough to consume these highly nutritious crustaceans, but to prevent any water quality problems, only a small amount of Artemia (50ml) was added 3 times a day (10am, 2pm and 5pm). The larvae were fed on both rotifers and Artemia for the entire length of my study and by the time I came to leave the Maldives they were also consuming crushed dry flake food.

Growth and Development Study

Once in the larval rearing tank, I began to study the larvae in order to witness how they grow and develop. Each day I would take 3 random individuals from the rearing tank and measured their body length under the microscope. I also observed the larvae’s development, making notes on how the body shape, colour pigmentation, eyes, mouth, fins and tail all developed over time.


My study of A. clarkii larvae ran for a total period of 30 days, with data on growth and development collected for the first 15 days of the study. This is because at around days 10-15 the larvae begin to undergo metamorphosis whereby they turn from their newly hatched form into miniature versions of their parents. Metamorphosis is very stressful for larvae and high mortality is often observed, so I decided to stop my growth study at the start of metamorphosis to prevent extra mortalities. Throughout the study period, the salinity and temperature of the larval rearing tank was recorded, with salinity maintained at 35ppm and the temperature range is shown in the graph below.

Larval growth is shown in the graph below. As mentioned, I measured 3 individual larvae each day and this graph shows the mean length for each day. In general, larvae showed steady growth and development over the study, with the early signs of impending metamorphosis (tail curling) first observed on day 9.

Although this early sign of metamorphosis was visible on day 9, the first larvae did not go through metamorphosis until day 15. From day 15 onwards, although I had stopped recording growth data, the larvae slowly went through metamorphosis and it was not until day 25 – 30 that all were completely metamorphosed. Although the majority of larvae did metamorphose shortly after day 15, I believe the reason it took some of the larvae so long was due to the gradual decrease in temperature from day 10 onwards.

From further reading, it was found that at a temperature of around 27 – 28 C the larvae would grow rapidly and go through metamorphosis in 8 – 12 days. At a lower temperature the larvae will grow and metamorphose at a slower rate. Therefore, I believe that if the temperature in the larval rearing tank had remained at 27 – 28 C, the larvae may have developed and gone through metamorphosis much faster than they actually did. However, as the water used in the Fish Lab is pumped in from the ocean, the temperature would fluctuate depending on weather conditions. Therefore, in future I would install an aquarium heater to keep the temperature in the larval rearing tank consistent.

When my time in the Maldives had come to an end I had successfully reared around 75 juvenile A. clarkii from tiny larvae into fully formed juvenile fish. As mentioned, ornamental aquaculture is a field of marine biology that I am very interested in and being able to work with the team on the fish breeding projects was a great experience. I would certainly love the opportunity to work on a project like this again in the future, and wish the Marine Savers teams at both Landaa Giraavaru and Kuda Huraa every success with their work.


  • Krishna M.V.R., Anil M.K. and Neethu Raj P. (2015) Studies on growth and development of hatchery produced juveniles of Amphiprion clarkii (Bennett, 1830). The International Journal of Science & Technoledge 3, 150-155.
  • Swagat Ghosh T., Ajith Kumar T., Nanthinidevi K. and Balasubrananian T. (2011) Hatchery production of Clark’s clownfish, Amphiprion clarkii (Bennett, 1830) using brackishwater. 2nd International Conference on Agriculture and Animal Science 22, 51-56.
  • Wabnitz C., Taylor M., Green E. and Razak T. (2003) From ocean to aquarium. UNEP-WCMC. Cambridge, UK.

Thanks for reading

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