WO2015149121A1

WO2015149121A1 - Method for the cultivation and production of macroalgae

Info

Publication number: WO2015149121A1
Application number: PCT/AU2015/050112
Authority: WO
Inventors: Christina CARL; Nicholas Paul; Rocky De Nys
Original assignee: James Cook University; Mbd Energy Limited
Priority date: 2014-03-31
Filing date: 2015-03-17
Publication date: 2015-10-08

Abstract

A method is provided for the cultivation and/or production of green macroalgae, such as Ulva spp. The green macroalgae are exposed to a temperature shock, such as a cold shock or a heat shock to induce reproduction, then maintained under suitable conditions. The method may further comprise fragmenting the temperature shocked green macroalgae. Green macroalgae may be produced according to the invention, such as for the production of biomass, wastewater treatment, bioremediation, human consumption, animal feed, and/or feedstock for the supply of pharmaceuticals, nutraceuticals, cosmeceuticals and/or biofuels. The method may be performed within a marine or brackish water environment, such as an aquaculture farm.

Description

TITLE

METHOD FOR THE CULTIVATION AND PRODUCTION OF MACRO ALGAE

TECHNICAL FIELD THIS INVENTION relates to a method of cultivating and producing green macroalgae. More particularly, the invention relates to an improved cultivation and production method for green macroalgae that may be particularly useful for the large scale production of algae biomass.

BACKGROUND

The application of macroalgal derived products is diverse, including for example, its use in human food products (Holdt and Kraan 2011), animals feeds (Soler-Vila et al. 2009), fertilisers (Bird et al. 2012) and as feedstock for the supply of pharmaceuticals (Tan et al. 2013), nutraceuticals (Zhang, Li, and Kim 2012), cosmeceuticals (Zubia et al. 2007), and biofuels (Gosch et al. 2012). Furthermore, large scale cultivation of macroalgae can be integrated into aquaculture production platforms, thereby providing value-adding processes (Cruz-Suarez et al. 2010) and environmental services by removing excess dissolved nutrients from waste waters (Neori et al. 2004).

A total of approximately 19 million tonnes of macroalgae were produced in 2010 with more than 95% of the total algae production from aquaculture (FAO 2012). The production of macroalgae is growing more than 7% per year (FAO 2012) and therefore, sustainable and economically feasible cultivation is essential.

Currently, long-line and net culture are the two major methods used for the cultivation of green marine macroalgae (seaweeds), including the species of Monostroma (Kida 1990; Ohno 1993) and Ulva (Ohno 1993). In order to attach algae to ropes and nets, fragments are either fastened on or pinched into ropes, or alternatively 'seeded' by spore settlement. Substrates can either be artificially seeded under controlled conditions or naturally by submerging substrates in calm areas with naturally high abundances of the desired species. Spores released from the natural populations then settle on the ropes and nets which are subsequently transferred to cultivation grounds (Dan et al. 2002). However, natural settlement is highly variable, and the current cultivation and production methods often produce poor yields.

Given the growing commercial interest in macroalgal derived products and their integration into aquaculture facilities for bioremediation, it is important to optimise the efficacy of cultivation and production techniques in order to satisfy the increasing global demand.

SUMMARY

The present inventors have determined an improved method of cultivating and/or producing green macroalgae.

Suitably, the invention is broadly directed to methods of cultivating and/or producing green macroalgae. In particular embodiments, the methods may be useful for the large scale production of macroalgae for the production of biomass, wastewater treatment, human consumption, animal feed, and/or feedstock for the supply of pharmaceuticals, nutraceuticals, cosmeceuticals and/or biofuels.

In a first aspect, the invention provides a method of cultivating green macroalgae, the method including;

exposing green macroalgae to a temperature shock, wherein the temperature shock induces the formation of reproductive bodies; and

maintaining the temperature shocked macroalgae under suitable conditions to thereby initiate production and release of the reproductive bodies.

In one embodiment the temperature shock is a cold shock. Suitably, the macroalgae are exposed to a cold shock of less than about 10°C.

Preferably, the green macroalgae are exposed to a cold shock of about 4 °C. In one embodiment the temperature shock is a heat shock. Suitably, the green macroalgae are exposed to a heat shock of about 15 °C to 35 °C.

Preferably, the green macroalgae are exposed to a heat shock of about 25 °C.

In one embodiment the green macroalgae are exposed to a temperature shock for a period of about 10 to about 20 minutes.

Preferably, the green macroalgae are exposed to a temperature shock for a period of about 10 minutes.

In a further embodiment the method may include the additional step of fragmenting the temperature shocked green macroalgae before maintaining the macroalge under suitable conditions. Suitably, the green macroalgae are fragmented for example by blending, cutting, chopping or tearing.

Suitably, the green macroalgae are fragmented to a size of about 1 to 50 mm.

In one embodiment, the temperature shocked macroalgae are maintained under suitable conditions for about 24 to 72 hours. Preferably, the temperature shocked macroalgae are maintained under suitable conditions for about 48 hours. In one embodiment, the temperature shocked macroalgae are maintained at a temperature from about 24°C to 26°C. Preferably, the temperature shocked macroalgae are maintained at a temperature of about 25°C.

In one embodiment, the temperature shocked macroalgae are exposed to a photoperiod comprising both light and dark periods during maintenance of the macroalgae.

Preferably, the temperature shocked macroalgae are maintained under suitable conditions for about 12 hours in the light and about 12 hours in the dark for every 24 hour period.

In a further embodiment the green macroalgae may comprise filamentous or blade species.

Preferably, the green macroalgae comprise filamentous species.

Suitably, the green macroalgae is of the Ulva genus.

Preferably, the macroalgae species are selected from one or more of the group consisting of: Ulva arasakii, Ulva armoricana, Ulva australis, Ulva bulbosa, Ulva californica, Ulva clathrata, Ulva clathratioides, Ulva compressa, Ulva enteromorpha, Ulva fasciata, Ulva fenestrate, Ulvaflexuosa, Ulva geminoidea, Ulva intestinalis, Ulva intestinaloides, Ulva laetevirens, Ulva lactuca, Ulva latissima, Ulva linza, Ulva lobata, Ulva muscoides, Ulva ohnoi, Ulva palmata, Ulva pertusa, Ulva procera, Ulva prolifera, Ulva proliferoides, Ulva pseudocurvata, Ulva purpurea, Ulva rigida, Ulva scandinavica, Ulva spinulosa, Ulva stenophylla, Ulva stenophylloides, Ulva stipitata, Ulva taeniata, Ulva tanneri, Ulva torta, Ulva sp. 3, Ulva sp. 2, Ulva sp. 4 and Ulva umbilicalia.

More preferably, the macroalgae is of the species Ulva sp. 3.

In a second aspect, the invention provides a method of producing green macroalgae, the method including:

producing reproductive bodies according to the first aspect;

seeding the reproductive bodies;

settling the reproductive bodies; and

growing the reproductive bodies to produce green macroalgae.

In one embodiment, reproductive bodies are seeded on to a growth structure.

Preferably, the growth structure is selected from the following non-limiting group comprising rope, net, sponge, balls and floats. More preferably, the growth structure is rope. Suitably, the reproductive bodies are seeded on to a growth structure at a density of 466,000 to 1,552,000 per metre of rope.

Preferably, the reproductive bodies are seeded on to rope at a density of about 621,000 reproductive bodies per metre of rope.

In one embodiment the reproductive bodies are seeded directly into an aqueous solution to form clusters. Suitably, the aqueous solution is sea water.

Preferably, the reproductive bodies are seeded into an aqueous solution at a density of about 10,000 to 20,000 reproductive bodies per mL.

More preferably, the reproductive bodies are seeded into an aqueous solution at a density of about 15,000 reproductive bodies per mL.

In one embodiment, the reproductive bodies are allowed to settle for about 4 to 5 days. Preferably, the reproductive bodies are settled for about 5 days.

In one embodiment, the reproductive bodies are settled at a temperature of about 24°C to about 26°C. Preferably, the reproductive bodies are settled at a temperature of about 25°C.

In particular embodiments, the reproductive bodies are exposed to a photo period capturing both light and dark periods during settlement. Preferably, the reproductive bodies are settled for about 12 hours in the light and about 12 hours in the dark.

In one embodiment, the settled reproductive bodies are grown for a period of about 10 to 16 days. Preferably, the settled reproductive bodies are grown for a period of about 14 days.

Suitably, the settled reproductive bodies may be grown in an open or closed system.

In one embodiment, the method according to the second aspect further comprises harvesting the green macroalgae.

Suitably, the green macroalgae is harvested about 18 to 25 days after exposing the macroalgae to a temperature shock.

Preferably, the green macroalgae is harvested about 21 days after exposing the green macroalgae to a temperature shock.

In one embodiment, about 50 to 90% of the weight of the green macroalgae is harvested.

Preferably, up to about 80% of the weight of the green macroalgae is harvested. Suitably, the macroalgae is harvested at two week intervals.

In particular embodiments, harvesting may be undertaken by cutting, trimming, tearing etc. Suitably, harvesting may be undertaken by mechanical or manual means.

In a third aspect, the invention provides a method of producing green macroalgae according to the method of the second aspect for the production of biomass, wastewater treatment, bioremediation, human consumption, animal feed, and/or feedstock for the supply of pharmaceuticals, nutraceuticals, cosmeceuticals and/or biofuels.

In one embodiment, the method according to any one of the aforementioned aspects is performed within a marine or brackish water environment.

Preferably, the marine environment is an aquaculture farm. Suitably, the aquaculture farm may be selected from any type of aquaculature farm, non-limiting examples of which include: fish, prawns, shell fish etc.

In one embodiment, wastewater is treated through removal of nutrients by the green macroalgae. Suitably, treated wastewater may be used for example for further cultivation of green macroalgae, further processes and/or for release into the environment.

In one embodiment the amount of nutrients that may be treated by the green macroalgae is about 3 kg of nitrogen and about 0.3 kg of phosphorus for about every 100 kg of green macroalgae produced.

In a fourth aspect, the invention provides green macroalgae produced according to the methods of the first, second and third aspects.

Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood by those of ordinary skill in the art to which the invention belongs. Although any methods and materials similar or equivalent to those described herein can be used in the practice or testing of the present invention, preferred methods and materials are described.

As used herein, except where the context requires otherwise, the term "comprise" and variations of the term, such as "comprising", "comprises" and "comprised", are not intended to exclude further additives, components, integers or steps.

It will be appreciated that the indefinite articles "a" and "an" are not to be read as singular indefinite articles or as otherwise excluding more than one or more than a single subject to which the indefinite article refers. For example, "a" alga includes one alga, one or more algae or a plurality of alga.

Any discussion of the prior art throughout the specification should in no way be considered as an admission that such prior art is widely known or forms part of the common general knowledge in the field.

BRIEF DESCRIPTION OF THE FIGURES

Figure 1. Reduced version of maximum likelihood tree of Ulva ITS sequences, showing the species used (shown in bold).

Figure 2. Transformation of vegetative cells into reproductive cells of Ulva sp. 3. (a) Vegetative cells, (b) formation of swarmers after 26 h, (c) reproductive cells with fully formed swarmers after 44 h, (d) reproductive cells with fully formed and discharged swarmers after 46 h. Scale bars 20 μπι.

Figure 3. Discharge of swarmers (%) after two and three days post collection and treatment. Samples were rinsed in dechlorinated tap water (DC) or filtered seawater (FSW) for 10 min prior to dehydration for 45 min (Dehydrated). Non- dehydrated filaments (Non-dehydrated) were used as a control. Filaments were either left whole (Whole) or segmented into pieces < 5 mm (Cut).

Figure 4. Discharge of swarmers (%) under photoperiods of (a) 12 h light: 12 h dark; (b) 18 h light:6 h dark; (c) constant light; and (d) constant darkness. Experiments were initiated at 1 pm and 7 pm on the day of collection (Day 1) and at 7 am, 1 pm, and 7 pm one day after collection (Day 2). Grey shaded background indicates duration of the dark period.

Figure 5. Discharge of swarmers (%) under a 12 h L: 12 h D photoperiod. Experiments were initiated at 1 pm and 7 pm on the day of collection (Day 1), and at 7 am, 1 pm, and 7 pm one day after collection (Day 2). Grey shaded background indicates the dark period.

Figure 6. Discharge of swarmers (%) two days after initiation of experiments at (a) 11 am and (b) 4 pm. Filaments of Ulva were exposed to 4°C and 25°C FSW and without pre-treatment as a control. After two days, the filaments were wrapped in moist paper, dried, or remained submersed from 7am for 4 h.

Figure 7. (a) Mean number of released swarmers over time (n = 3). (b) Number of released swarmers over time from three independently collected batches of algal biomass (n = 50). Algal batches were collected on 6/May/2013 (Batch 1), 7/May/2013 (Batch 2), and 14/May/2013 (Batch 3). Figure 8. Mean reproductive output (RO) of three independently collected algal batches (n = 50). Experiments were initiated on 6/May/2013 (Batch 1), 7/May/2013 (Batch 2), and 14/May/2013 (Batch 3).

Figure 9. Time course of induced sporulation of tropical Ulva sp 3. (a) Collection of algal samples and subsequent transportation to the laboratory, (b) Initiation of experiments at 1 pm by washing the thalli in FSW, subsequently chill Ulva for 10 min and then place thalli into autoclaved FSW under a 12 h L: 12 h D photoeriod at 25°C. (c) Induction of sporulation with visible formation.

Figure 10. Controlled cultivation stages of macroalgae.

Figure 11. Mean (± S.E.) biomass yield (g FW m^"1 rope) of Ulva seeded onto ropes at densities from 155 to 1552 ^χ 10³ swarmers m^"1 rope. Seeded ropes were maintained in a nursery for a period of (a) one, (b) five, and (c) ten days prior to transfer to outdoor cultivation. The fresh biomass yield was quantified after 7, 14, and 21 days of outdoor cultivation.

Figure 12. Mean (± S.E.) biomass yield (g DW m^"1 rope) of Ulva seeded onto ropes at densities ranging from 155 to 1,552 x 10³ swarmers m^"1 rope after 21 days of outdoor cultivation. The nursery periods of the seeded ropes ranged from one to ten days.

Figure 13. Mean (± S.E.) specific growth rate (SGR, % day^"1) of Ulva seeded onto ropes at densities ranging from 155 to 1,552 x 10³ swarmers m^"1 rope in the time period of: (a) 7 to 14 days; and (b) 14 to 21 days of outdoor cultivation.

Figure 14. Mean dry biomass yields and growth rates of Ulva seeded onto ropes over time. Experiments were run at an optimal seeding density of 621 x 10³ swarmers m^"1 rope and maintained for five days under nursery conditions, (a) Mean (± S.E.) dry biomass yield (g DW m^"1 rope) over time from three independently collected batches of algal biomass (n = 3 for each batch). Algal batches were collected on 11/09/2013 (batch 1), 18/09/2013 (batch 2), and 1/10/2013 (batch 3). (b) Mean (± S.E.) dry biomass yield (g DW m^"1 rope) over time (n = 3) for each batch, (c) Mean (± S.E.) specific growth rate (SGR, % day^"1) over time (n = 3).

Figure 15. Mean FW:DW ratios, filament width and ash content of Ulva seeded onto ropes over time. Experiments were run at an optimal seeding density of 621 x 10³ swarmers m^"1 rope and maintained for five days under nursery conditions, (a) Mean (± S.E.) FW:DW ratio (n = 3). (b) Mean (± S.E.) width of filaments of Ulva growing on ropes over time (n = 3). (c) Mean (± S.E.) ash content (%) of Ulva. Figure 16. Mean (± S.E.) weight (g DW m^"1 rope) of harvested Ulva biomass for successive harvests. (A) Mean harvested biomass over time (n = 3). (B) Mean harvested biomass (n = 13) of each batch for successive harvests. The ropes were independently seeded on 24 April 2014 (batch 1), 8 May 2014 (batch 2) and 9 July 2014 (batch 3).

Figure 17. Mean (± S.E.) specific growth rate (SGR, % day-1) of Ulva over time and after harvest (n = 3). The arrows indicate when the ropes were harvested (harvest 1 after 14 days of outdoor cultivation, harvest 2 after 28 days of outdoor cultivation, final harvest 3 after 42 days). Each cultivation cycle was 14 days long.

DETAILED DESCRIPTION

The present invention is broadly directed to improved methods for cultivating and/or producing green macroalgae. The inventors have established an improved method which considers the unique requirements of macroalgae cultivation and growth, to enable and enhance the production of high green macroalgae yields.

Controlled cultivation and production methods allow for the rapid development and growth of green macroalgae reducing the costs of production and providing a continuous supply of macroalgae biomass.

Method of Cultivating Macroalgae

In one aspect of the invention there is provided a method of cultivating green macroalgae, the method including;

maintaining the temperature shocked macroalgae under suitable conditions, to thereby initiate production and release of the reproductive bodies.

By "reproductive bodies" and "swarmers" is meant motile reproductive bodies, including gametes, zoospores, and zoids, which are capable of forming adult macroalgae plants.

In one embodiment, the temperature shock to which the green macroalgae are exposed may be a cold shock. Suitably, the green macroalgae are exposed to a cold shock at a temperature of less than about 10°C. More suitably, the green macroalgae are exposed to a cold shock of about 1, 2, 3, 4, 5, 6, 7, 8 or 9°C. Preferably, the macroalgae are exposed to a cold shock at a temperature of about 4°C.

In an alternate embodiment the temperature shock is a heat shock. Suitably, the green macroalgae are exposed to a heat shock at a temperature of about 15°C to 35°C. More suitably, the macroalgae are exposed to a heat shock at a temperature of about 15-20, 15-25, 15-30, 15-35, 20-25, 20-30, 20-35, 25-30, 25-35 or 30-35°C, or about 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34 or 35°C. Preferably, the green macroalgae are exposed to a heat shock at a temperature of about 25°C.

By abouf is meant a measured variation in the quantity, level, value, number, frequency, percentage, dimension, size, amount, weight or length that varies by as much 15, 10, 9, 8, 7, 6, 5, 4, 3, 2 or 1 % to a reference quantity, level, value, number, frequency, percentage, dimension, size, amount, weight or length.

Suitably, the green macroalgae are exposed to a temperature shock for a period of about 10 to 20 minutes. More suitably, the green macroalgae are exposed to a temperature shock for a period of about 10-12, 10-14, 10-16, 10-18, 12-14, 12- 16, 12-18, 12-20, 14-16, 14-18, 14-20, 16-18, 16-20 or 18-20 minutes or about 10, 11, 12, 13, 14, 15, 16, 17, 18, 19 or 20 minutes. Preferably, the macroalgae are exposed to a temperature shock for about 10 minutes or until the temperature shock induces the formation and release of reproductive bodies.

The temperature shock may be administered as a single treatment or as multiple treatments, depending on the requirements of the algal species, to induce the formation and release of reproductive bodies.

Suitably, the method of cultivating green macroalgae may further include fragmenting the temperature shocked macroalgae before maintaining the macroalgae under suitable conditions. The macroalgae may be fragmented by any method known in the art. Examples of fragmentation include without limitation such methods as blending, cutting, chopping, tearing, macerating, slicing and dicing.

The green macroalgae may be fragmented to any size, wherein reproductive bodies are capable of being produced. Preferably, the macroalgae is fragmented to a size of about 1 to 50 mm before maintaining the macroalgae under suitable conditions. Suitably, the macroalgae is fragmented to a size of about 1-5, 1-10, 1-15, 1-20, 1-25, 1-30, 1-35, 1-40, 1-45, 5-10, 5-15, 5-20, 5-25, 5-30, 5-35, 5-40, 5-45, 5- 50, 10-15, 10-20, 10-25, 10-30, 10-35, 10-40, 10-45, 10-50, 15-20, 15-25, 15-30, 15- 35, 15-40, 15-45, 15-50, 20-25, 20-30, 20-35, 20-40, 20-45, 20-50, 25-30, 25-35, 25- 40, 25-45, 25-50, 30-35, 30-40, 30-45, 30-50, 35-40, 35-45, 35-50, 40-45, 40-50 or 45-50 mm, or about 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47, 48, 49 or 50 mm.

Suitably, the temperature shocked green macroalgae are maintained under suitable conditions for about 24 to 72 hours or until reproductive bodies are produced. More suitably, the temperature shocked macroalgae are maintained under suitable conditions for about 24-28, 24-32, 24-36, 24-40, 24-44, 24-48, 24-52, 24-56, 24-60, 24-64, 24-68 or 24-72 hours or about 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47, 48, 49, 50, 51, 52, 53, 54, 55, 56, 57, 58, 59, 60, 61, 62, 63, 64, 65, 66, 67, 68, 69, 70, 71 or 72 hours. Preferably, the temperature shocked macroalgae are maintained under suitable conditions for about 48 hours.

In some embodiments, the temperature shocked green macroalgae are maintained at a temperature from about 24°C to 26°C. Suitably, the temperature may be about 24°C, 25°C or 26°C. Preferably, the temperature is about 25°C. The conditions under which the green macroalgae are maintained may be adapted to suit the specific macroalgae requirements, depending on the macroalgal species.

In some embodiments, the temperature shocked macroalgae are exposed to a photoperiod comprising both light and dark periods. Suitably, the macroalgae are exposed to a photoperiod for about 5 to 18 hours in the light and about 5 to 18 hours in the dark for every 24 hour period. Preferably, for at least about 12 hours in the light and about 12 hours in the dark for every 24 hour period, or under conditions suitable for reproductive bodies to be produced and released.

The green macroalgae may be maintained in any medium suitable to sustain the life of the macroalgae and macroalgae reproductive bodies and to allow for reproductive bodies to be produced and released. Examples of suitable medium include without limitation, seawater, saline ground water, artificial seawater media, or freshwater. The macroalgae may also be maintained in any means suitable for the maintenance of the macroalgae species including for example shallow containers capable of holding the medium.

In the general context of the invention, the macroalgae may comprise filamentous or blade species, including for example flat leaf and tubular leaf species.

Suitably, the green macroalgae may comprise Ulva genus.

Preferably, the macroalgae species are selected from one or more of the group consisting of Ulva arasakii, Ulva armoricana, Ulva australis, Ulva bulbosa, Ulva californica, Ulva clathrata, Ulva clathratioid.es, Ulva compressa, Ulva enteromorpha, Ulva fasciata, Ulva fenestrate, Ulvaflexuosa, Ulva geminoidea, Ulva intestinalis, Ulva intestinaloides, Ulva laetevirens, Ulva lactuca, Ulva latissima, Ulva linza, Ulva lobata, Ulva muscoides, Ulva ohnoi, Ulva palmata, Ulva pertusa, Ulva procera, Ulva prolifera, Ulva proliferoides, Ulva pseudocurvata, Ulva purpurea, Ulva rigida, Ulva scandinavica, Ulva spinulosa, Ulva stenophylla, Ulva stenophylloides, Ulva stipitata, Ulva taeniata, Ulva tanneri, Ulva torta, Ulva sp. 3, Ulva sp. 2, Ulva sp. 4 and Ulva umbilicalia.

Preferably, the macroalgae is filamentous green macroalgae.

More preferably, the algae is of the species Ulva sp. 3.

The macroalgae from which reproductive bodies are produced may be obtained directly from the ocean, estuarine and brackish environments, intertidal pools or streams or by farming (e.g., aquaculture ponds) or from any other known source.

It will be appreciated that the macroalgae for cultivation may be of any developmental stage that is capable of being induced to form and produce reproductive bodies.

Method of producing macroalgae

Green macroalgae may be grown using a number of steps, including reproduction, seeding, settling and growth of the adult macroalgae with specific production conditions. Figure 10 shows a non-limiting example of macroalgae production steps.

In one aspect of the invention there is provided a method for producing green macroalgae, the method including:

producing reproductive bodies according to the first aspect;

seeding reproductive bodies;

settling reproductive bodies; and

growing the reproductive bodies to produce macroalgae.

It will also be appreciated that settling is the process of allowing the reproductive bodies to form a stable attachment either to a physical substrate or to each other to form a cluster.

Suitably, in some embodiments, reproductive bodies are seeded on to a growth substrate. Suitably, the growth substrate may be selected from the following non limiting group comprising: rope, net, sponges, balls and floats. More preferably, the growth substrate is rope. The rope may be made of any material suitable for the settlement of algal reproductive bodies, an example of which includes polypropylene rope.

The reproductive bodies may be seeded on to a growth substrate at a density of about 466,000 to about 1,552,000 reproductive bodies per metre of rope. Suitably, the reproductive bodies may be seeded on to a growth substrate at a density of about 466,000-600,000, 466,000-700,000, 466,000-800,000, 466,000-900,000, 466,000- 1,000,000, 466,000-1, 100,000, 466,000-1,200,000, 466,000-1,300,000, 466,000- 1,400,000 or 466,000-1,500,000 reproductive bodies per metre of rope, or about 466,000, 500,000, 600,000, 700,000, 800,000, 900,000, 1,000,000, 1, 100,000, 1,200,000, 1,300,000, 1,400,000, 1,500,000 or 1,522,000 reproductive bodies per metre of rope.

Preferably, the reproductive bodies are seeded onto rope at a density of about

621,000 reproductive bodies per metre of rope.

The density of the reproductive bodies may be determined by any method known in the art, for example using a haemocytometer.

In some embodiments the reproductive bodies are seeded directly into an aqueous solution to form clusters. Clusters provide a suspended culture option that does not require a growth substrate for culture and can be cultured in shallow pond- based systems as clusters are not buoyant. Clusters may also be cultivated for example in tumble cultures in land-based ponds, tanks or trends or in High Rate

Algal Ponds (HRAPs)

The aqueous solution in which the reproductive bodies are seeded may be any aqueous solution capable of supporting growth of reproductive bodies. Non- limiting examples of an aqueous solution include sea water, sea water with nutrients, artificial seawater, artificial seawater with nutrients, freshwater, etc.

In some embodiments, the reproductive bodies are seeded into an aqueous solution at a density of about 10,000 to 20,000 reproductive bodies per mL. Suitably, the reproductive bodies are seeded into an aqueous solution at a density of about 10,000-12,000, 10,000-14,000, 10,000-16,000, 10,000-18,000, 12,000-14,000,

12,000-16,000, 12,000-18,000, 12,000-20,000, 14,000-16,000, 14,000-18,000,

14,000-20,000, 16,000-18,000, 16,000-20,000 or 18,000-20,000 reproductive bodies per raL or about 10,000, 11,000, 12,000, 13,000, 14,000, 15,000, 16,000, 17,000,

18,000, 19,000, 20,000 reproductive bodies per mL. Preferably, the reproductive bodies in an aqueous solution are at a density of about 15,000 reproductive bodies per mL.

The reproductive bodies may be settled for a period of about 4 to 5 days or until the reproductive bodies form an attachment to a growth substrate and/or the formation of clusters. Preferably, the reproductive bodies are settled for a period of about 5 days.

In some embodiments, the reproductive bodies are settled at a temperature of about 24 to 26°C. Suitably, the reproductive bodies are settled at a temperature of about 24°C, 25°C or 26°C. Preferably, the reproductive bodies are settled at a temperature of about 25°C.

In some embodiments, the reproductive bodies are exposed to a photo period capturing both light and dark periods during settlement. Suitably, the macroalgae are exposed to a photoperiod for about 5 to 18 hours in the light and about 5 to 18 hours in the dark for every 24 hour period. Preferably, for at least about 12 hours in the light and about 12 hours in the dark for every 24 hour period, or under conditions suitable for reproductive bodies to be produced and released.

The seeded and settled reproductive bodies may be grown for a period of about 10 to 16 days. Suitably, the settled reproductive bodies may be grown for a period of about 10-12, 10-14, 10-16, 12-14, 12-16, 14-16 days or about 10, 11, 12, 13, 14, 15 or 16 days. Preferably, the seeded and settled reproductive bodies are grown for a period of about 14 days or until the macroalgae reach the desired size, biomass, or growth stage.

Before growth of the reproductive bodies, the growth structures and/or clusters may be transferred to different growth systems or may be maintained in the same growth system, depending on the number of reproductive bodies.

Suitably, the macroalgae may be grown in open or closed water bodies, including open or closed pond growth systems and HRAPs. Preferably, the macroalgae are grown in flow through water bodies. The water bodies may comprise nutrients including for example nitrogen in the form of ammonium or nitrate and phosphorous in the form of phosphate. Suitably, for example, the nitrogen concentration may be above lmg L^"1 and the phosphorus concentration above 0.1 mg L-¹.

Suitably, any water containment means may be used that meets the macroalgae growth requirements, including natural or artificial means. In a further embodiment the method may further include harvesting the produced green macroalgae.

Suitably, the green macroalgae may be harvested about 18 to 25 days after exposing the macroalgae to a temperature shock. More suitably, the green macroalgae may be harvested about 18-20, 18-22, 18-24, 18-25, 20-22, 20-24, 20-25, 22-24, 22-25 days after exposing the macroalgae to a temperature shock, or about 18, 19, 20, 21, 22, 23, 24 or 25 days after exposing the macroalgae to a temperature shock. Preferably, the macroalgae is harvested about 21 days after exposing the macroalgae to a temperature shock.

When harvesting the macroalgae, about 50 to 90% of the weight of the macroalgae may be harvested. Preferably, up to about 90% of the weight of the macroalgae is removed and harvested.

In some embodiments, sequential harvests are performed.

In a particular embodiment no more than two (2) sequential harvests are performed.

Suitably, harvesting may include removing only a portion of the macroalgae plant, leaving the rooting portion. Suitably, harvesting may include separating the upper portion of the macroalge plant from the rooting portion, to allow the rooting portion to continue growing for subsequent harvest.

In some embodiments, the macroalgae is completely harvested.

Harvesting may take place about every two to three weeks. Preferably, the macroalgae is harvested at two week periods, providing continuous growth and harvest of the macroalgae. Any number of harvests may be undertaken, depending on the species of green macroalgae. Reseeding may be undertaken between harvests to further populate the growth structures or clusters.

Suitably, harvesting may be undertaken by any known process in the art, including for example mechanical and/or manual means. Methods of harvesting macroalgae include without limitation, cutting, tearing and ripping.

In one embodiment, the macroalgae may be dried prior to harvesting.

In one embodiment the macroalgae may comprise filamentous or blade leaf species, including for example tubular leaf and flat leaf species.

Suitably, the macroalgae may comprise Ulva genus.

Preferably, the macroalgae is filamentous green algae.

More preferably, the algae is of the species Ulva sp. 3.

The macroalgae produced by the methods of the invention may be used for a range of applications, non-limiting examples of which include the production of biomass, wastewater treatment, human consumption, and/or feedstock for the supply of pharmaceuticals, nutraceuticals, cosmeceuticals and/or biofuels. Large scale cultivation of macroalgae may be intergrated into aquaculture production platforms, removal of nutrients or toxic substances from water systems, including waste water.

Suitably, the method according to any one of the aforementioned aspects is performed within a marine or brackish water environment.

Preferably, the marine environment is an aquaculture farm (e.g., fish, prawns, shell fish etc.), wherein the macroalgae removes excess nutrients and produces biomass, which can be harvested.

Wastewater may also be treated through removal of nutrients such as phosphorous and nitrate by the macroalgae to produce clean wastewater for environmental use and/or for example for use in further macroalgae cultivation and/or production. In some embodiments, the amount of wastewater that may be treated by the macroalgae is about 3 kg of nitrogen and about 0.3 kg of phosphorous for about every 100 kg of macroalgae produced.

In some embodiments, the invention provides for the renewable production and use of macroalgae biomass. For example, the harvested macroalgae may be dried and processed for further use in macroalgae derived products and processes.

In an aspect, green macroalgae is produced according to the methods of the first, second and third aspects.

So that the invention may be readily understood and put into practical effect, the following non-limiting Examples are provided. EXAMPLES

EXAMPLE 1

Materials and Methods

Study species and collection

The genus Ulva has a worldwide distribution with a broad tolerance of salinity, with species occurring in hypersaline to freshwater environments (Shimada et al. 2008). The Ulva species used in this study was analysed using molecular barcoding. Barcoding compares short DNA sequences from a standardised region of the genome (the barcode) to a library of reference sequences derived from individuals of known identity (Hebert et al. 2003). The species was identified as Ulva sp. 3 (Shimada et al. 2008) using newly generated DNA sequences from the internal transcribed spacer (ITS) region of the ribosomal cistron (Genbank accession number KF534755). The species Ulva sp. 3, is characterised by flat tubular thalli. Phylogenetic tress constructed using these sequences show the Ulva species (Figure 1).

Ulva sp. 3 (hereafter Ulva) was collected by hand from a land-based aquaculture facility at Guthalungra (19° 55'S, 147° 50Έ), Queensland, Australia. Notably, reproductive patterns driven by lunar periodicity as found for temperate species of Ulva (Scardino et al. 2008) were not observed in the field populations collected over a 4 month period. Samples were placed in a 25 L container filled with pond water and then transported within 3 h to the laboratory at James Cook University (JCU) in Townsville, Australia. Subsequently, the samples were gently washed three times with filtered seawater (FSW; 0.2 μπι and UV sterilised) to remove debris, epiphytes and invertebrates.

Laboratory experiments

Effect of salinity, dehydration, and segmentation

To determine the effect of salinity, dehydration and segmentation on the formation and release of swarmers, these factors were manipulated in a fully factorial experiment under laboratory conditions using samples collected in the morning and gently cleaned with FSW as described above. Firstly, Ulva was exposed to a lower salinity by placing filaments in dechlorinated tap water (DC) for 10 min with FSW being used as a salinity control. Secondly, the effect of dehydration was tested for each salinity treatment. All filaments were dried using paper towels to remove excess water and subsequently either dehydrated by exposure to air for 45 min in the dark at 25 °C (dehydration treatments), or alternatively placed in FSW in the dark for 45 min (non-dehydrated control). Thirdly, the effect of segmentation was tested in the factorial design by cutting a section of approximately 50 - 70 mm from each filament (hereafter referred to as whole), or alternatively cutting this section further into pieces < 5 mm using a razor blade (hereafter referred to as cut). Both treatments had similar biomass per unit volume. Each of the whole and cut filaments were placed in individual Petri dishes (Iwaki; 1010-060) filled with 10 mL autoclaved FSW and sealed using Parafilm to prevent evaporation. The Petri dishes were then placed in a culture cabinet at 25°C at an irradiance of 125 μπιοΐ photon m^"2 s^"1 under a 12 h L: 12 h D photoperiod for three days. A total of three replicates was used for each treatment combination (n = 3 for each salinity x dehydration x segmentation).

The percentage of the total area of each filament (either whole or cut) that had released swarmers (hereafter referred to as discharge) was visually quantified daily at 3 pm using a dissecting microscope (Olympus SZ61) according to Nielsen and Nordby (1975). Vegetative cells of Ulva transform directly into reproductive cells (Figure 2). The colouration differs between vegetative (Figure 2a), reproductive (Figure 2c) and discharged cells (Figure 2d) and is classified by green, brown-ish and white colour, respectively (Han & Choi 2005). The discharge was quantified using only white cells that had unequivocally released swarmers (Figure 2d).

Effect of time of initiation of experiments and photoperiod

To determine the effect of time of initiation of experiments and photoperiod, on the formation and release of swarmers, samples were collected in the morning and gently cleaned with FSW as described above. Algae were subsequently maintained in an outdoor aquarium at the Marine & Aquaculture Research Facilities Unit at JCU until experiments were initiated by cutting a single section from each filament. Experiments were initiated at five times, two on the day of collection at 1 pm and 7 pm, and three on the day after collection at 7 am, 1 pm and 7 pm. The times of initiation affected the length of exposure to light and darkness at the start of the experiment (experiments initiated at 7am were initially exposed to 12 h light in contrast to experiments initiated at 7 pm where thalli were initially exposed to 12 h darkness).

For all replicates, a single section of approximately 50 - 70 mm was cut from each filament (see Results) and placed in Petri dishes (Techno Plas; S6014S10) filled with 10 mL autoclaved FSW and sealed with Parafilm. A total of 1080 replicate dishes were then placed in one of three culture cabinets at 25°C at photoperiods of a normal light period (12 h L: 12 h D), an extended light period (18 h L:6 h D), or constant light (24 h L). In addition, another 360 Petri dishes were wrapped in aluminium foil to provide constant darkness (24 h D) and these were split between the culture cabinets.

The discharge of swarmers was quantified at 7 pm on the day of collection and then daily at 7 am, 1 pm, and 7 pm for the following three days. At each sampling point, nine randomly selected dishes were destructively sampled from each treatment (n = 9 for each photoperiod x time of experimental initiation) and the discharge quantified as a percentage of the total area of each filament, as described above.

Effect of temperature shock and controlled release of swarmers

This experiment determined the effect of temperature shock on the formation of swarmers and assessed the potential to control their release by initially constraining and subsequently inducing the immediate release of swarmers in order to rapidly obtain a dense suspension of swarmers. Samples were collected in the morning and gently cleaned with FSW as described above. In the early afternoon, filaments of Ulva were placed in chilled (4°C) FSW and stored in the fridge for 10 min prior to placing a single section of approximately 50 - 70 mm from each filament (see Results) in Petri dishes (Techno Plas; S6014S10) filled with 10 mL autoclaved FSW. The use of a 10-min period was based on a pilot experiment where extended exposure periods to 4°C did not increase the formation and release of swarmers. As controls, filaments were either placed in 25°C FSW and stored in the dark for 10 min or immediately used in assays after the cleaning procedure without any further pre-treatment. The dishes were then placed in a culture cabinet at 25°C at an irradiance of 125 μπιοΐ photon m^"2 s^"1 under a 12 h L: 12 h D photoperiod.

After two days in the culture cabinet, prior to the expected onset of the release of swarmers, filaments were wrapped in moist paper towel at 7 am prior to the onset of the release of swarmers or exposed to air for 4 h by placing each filament individually on baking paper. After 4 h (11 am), each filament was placed in a new Petri dish filled with 10 mL autoclaved FSW. These methods were used in an attempt to constrain the release of swarmers to a short period of time in order to rapidly obtain a dense suspension of swarmers. A control remained submersed over this same period of time. The time of day selected reflected the onset of release in previous experiments (see Results). The discharge of swarmers was determined at 11 am within 5 min post re- submersion, and at 4 pm. A total of ten replicates was used for each treatment combination (n = 10 for temperature pre-treatment x restraining- treatment).

Reproductive output

The reproductive output (the number of swarmers released per unit area) of Ulva was estimated by quantifying the surface area of individual thalli and subsequently determining the percentage area of released swarmers, and the numbers of discharged swarmers over time for each thallus.

Samples of Ulva were collected in the morning and gently cleaned with FSW as previously described. The most successful combination of treatments tested above was used to induce sporulation and control the release of swarmers (see Results).

Briefly, the samples were chilled at 4°C for 10 min to maximise the formation and release of swarmers prior to cutting a section of approximately 50 - 70 mm from filaments (n = 50). Images of the filament pieces were captured using a camera (Olympus DP25) attached to a dissecting microscope (Olympus SZ61) for surface area measurements (Image J). Subsequently, these single filaments were placed in Petri dishes filled with 10 mL autoclaved FSW which were then placed in a culture cabinet at 25°C at an irradiance of 125 μπιοΐ photon m^"2 s^"1 under a 12 h L: 12 h D photoperiod.

To determine the number of released swarmers over time, two days after the initiation of experiments each filament was transferred into a new Petri dish filled with 10 mL autoclaved FSW every 90 min (from 7 am until 1 pm). This time period was the window of peak release for swarmers (see Results). To obtain accurate numbers of discharged swarmers in the water column, the dishes and water were changed at each measurement point (every 90 min) to minimise settlement. The water in the dishes was preserved in 1% Lugol's solution and subsequently transferred into sealable sample tubes. To increase the concentration of swarmers, the samples were centrifuged (400 g for 1 min; Thompson et al. 2007) and concentrated to a volume of 2.5 mL. Subsequently, the number of swarmers was determined using a haemocytometer. The reproductive output (RO) of each filament was calculated using the equation RO = NoS / (SA- D), where NoS is the number of released swarmers, SA the surface area of each filament and D is the discharge (percentage area of swarmer release). Finally, the number of flagella on released swarmers was counted using a compound microscope (Olympus BX53) to determine whether swarmers were biflagellate or quadriflagellate.

In addition, the viability and germling development of released swarmers was determined by inducing sporulation in a second cut section from each of the 50 filaments. These filaments were placed in individual Petri dishes filled with 10 mL autoclaved FSW and incubated as previously described to induce sporulation. After two days, the filaments were removed at 2 pm and nutrients were added to each dish (AlgaBoost lOOOx ill). Subsequently, the dishes were returned to the culture cabinet at 25°C under a 12 h L: 12 h D photoperiod. After a culture period of five days, the swarmers released by each filament were examined for germination using an inverted microscope (Olympus CKX41).

Experiments were conducted with a total of three independent collections of Ulva over an eight day period with 50 replicates from each collection time.

Phototactic behaviour of released swarmers

To determine the phototactic behavior of released swarmers, sporulation was induced by chilling samples and subsequently incubating an excised section of filaments (n = 40) in individual Petri dishes as previously described. After two days, the dishes were placed on a window sill at noon and any lights in the laboratory were turned off so that natural light was the primary light source. Released swarmers showed phototactic responses and consequently concentrated on the dark side of the dish facing away from the natural light when negatively phototactic or on the side with natural light source when possessing a positive phototaxis. Phototactic swarmers were removed and collected in sealable sample tubes over 90 min using a transfer pipette and preserved in 1% Lugol's solution. Subsequently, the number of flagella was counted to determine whether swarmers were bi- or quadriflagellate using a microscope (Olympus BX53). In addition, images of a total of 64 released swarmers were taken for size measurements (width and length).

Statistical analysis

Data were analysed by permutational analysis of variance (PERMANOVA) using PRIMER 6 (v. 6.1.13) and PERMANOVA+ (v. 1.0.3) (Clarke & Gorley 2006). The Bray-Curtis dissimilarity measure was used for all PERMANOVAs and /^-values were calculated using permutation of residuals under a reduced model with 9999 random permutations. If there was a significant difference, pair-wise a posteriori comparisons were made among the significant groups using the Bray- Curtis similarity measure (a = 0.05). All data are reported as mean ± 1 standard error (S.E.) unless stated otherwise.

The effects of salinity shock, dehydration, and segmentation on the discharge of swarmers were considered as fixed factors in the first experiment. Because the second experiment evaluated the interactive effects of different times of initiation and photoperiod, these data had to be analysed in two ways. A formal comparison using PERMANOVA was made between the effects of time of initiation and photoperiod by treating the day of sampling (day 3 and 4) and the sampling time of day (7 am, 1 pm, and 7 pm) as fixed factors because dishes were destructively sampled. An alternative plot of the effect of time of initiation (5 initiation times over 2 days past sampling) was then made for the key photoperiod (12 h L: 12 h D, see Results) by standardising the time of initiation at 0 h but was not formally analysed. The third experiment assessing the effect of temperature shock and treatments to control the release of swarmers, PERMANOVAs were run for each sampling point (11 am and 4 pm) with pre-treatment and restraining-treatment as fixed factors. To test for the effect of batch on the release of swarmers over time in the fourth experiment, a two-factor PERMANOVA was used with time as a fixed factor and batch as a random factor

Results

Effect of salinity, dehydration, and segmentation

There was no effect of salinity shock (three-factor PERMANOVA: F_(1: ιβ) = 0.86, p = 0.382), dehydration (F_(1: ιβ) = 0.55, p = 0.512), or segmentation (F_(1: ιβ) = 0.04, p = 0.948) on the discharge of swarmers among treatments after two and three days (Figure 3). No swarmers were released one day after initiation of experiments. After two days, the discharge of swarmers ranged from 23.3 ± 23.3% (FSW x dehydrated x segmented) to 76.7 ± 23.3% (FSW x non-dehydrated x whole) and was generally lower than after three days, where the discharge ranged from 23.3 ± 23.3 % (FSW x dehydrated ^χ segmented) to 90.0 ± 10.0% (FSW ^χ non-dehydrated ^χ whole, and also, dechlor ^χ dehydrated ^χ segmented). Therefore, the treatment combination of FSW x non-dehydrated x whole was chosen for the subsequent experiments on the basis of maximised discharge of swarmers with the least number of treatments. Effect of time of initiation of experiments and photoperiod The discharge of swarmers varied significantly between photoperiods (Figure 4), with the highest discharge in the 12 h L: 12 h D photoperiod (47.8 ± 10.4%). The discharge of swarmers decreased with extended light periods and was below 31% and 23%) at photoperiods of 18 h L:6 h D and 24 L, respectively. The lowest overall discharge occurred under constant darkness, with less than 12% discharge at any time (Figure 4). Photoperiod had a significant interactive effect with time of initiation (p < 0.001; Table 1), sampling day (p < 0.001) and time of sampling day (p = 0.001). Furthermore, there was a complex interactive effect of photoperiod, sampling day, and time of sampling day on the discharge of swarmers (p < 0.047; Table 1). This effect was driven by significant differences in the release of swarmers between photoperiods and higher overall discharge on day 4 in comparison to day 3. Regardless of the time of initiation, the values of released swarmers were similar at the end of the experiment for each photoperiod and ranged from 22.8 ± 7.4% to 44.4 ± 12.2%) under the normal photoperiod (12 h L: 12 h D), with generally large variations in the discharge between samples of the same treatment combinations.

Under the normal photoperiod (12 h L: 12 h D), the discharge generally peaked between 42 and 48 h after the initiation of experiments (two days post initiation), with the exception of the early initiation of 7am one day past collection where the discharge peaked after only 30 h (Figure 5). However, the same trend of an onset of release in the morning after the filaments were exposed to light occurred under normal photoperiod across all initiation times.

Effect of temperature shock and controlled release of swarmers

In general, the discharge of swarmers was higher for chilled filaments (4°C) than the other treatments (Figure 6), with the mean discharge of swarmers of temperature shock being nearly double (34.4 ± 4.6%>) that of both the 25°C pre- treatment (20.9 ± 7.9%)) and without any pre-treatment (18.5 ± 3.9%). However, the variance within the pre-treatments was relatively high (Figure 6) and pre-treatment was not a significant effect (two-factor PERMANOVA: F_{(2 )} = 1.88, p = 0.107). There was no clear effect of restraining-treatment combinations to manipulate the release of swarmers (F_(2,si) = 0.57, p = 0.718).

The discharge of swarmers was not constrained for filaments wrapped in moist paper towel or dried for 4 h (Figure 6a). In fact, treatments to constrain the discharge of swarmers resulted in slightly higher discharge at 1 1 am than continuously submersed filaments, with the exception of the 25 °C pre-treatment (submersed: 18.3 ± 7.8%; dried; 1.7 ± 0.8%; wrapped: 26.1 ± 10.0%). The unwrapping of filaments at 11 am had a high discharge of swarmers on the moist paper towel and these were clearly visible due to the change from white to green/ brown-ish colour of the paper towel. Furthermore, the unsuccessful constraint of discharge of swarmers by wrapping and drying was demonstrated by similar discharge between 11 am and 4 pm for those filaments (Figure 6a, b). Up to 33% of the biomass discharged swarmers while being wrapped (Figure 6a), whereas the discharge ranged from 18.9 ± 7.7% (control) to 43.3 ± 11.9% (4°C) after being re- submersed for 5 h (4 pm; Figure 6b). Similarly, the discharge of dried filaments was up to 23%) while exposed to air (Figure 6a) and increased marginally at 4 pm, ranging from 8.3 ±4.0 (25°C) to 30.6 ± 8.5% (4°C) (Figure 6b).

Reproductive output

The time of day had a significant effect on the number of swarmers released (two-factor PERMANOVA: F₍₄, 735) = 14.19, p = 0.003), with a peak of release at 11 :30 am with 842,708 ± 190, 123 swarmers released (mean of three batches ± 1 S.E.) (Figure 7a). The number of swarmers released was an order of a magnitude smaller at all other times and ranged from 34,375 ± 18,311 (7 am) to 94,791 ± 51,549 (1 pm). There was also a significant effect of batch on the number of swarmers released (F₍₂, 735) = 3.16, p = 0.025); however, the number of swarmers released showed a similar trend among batches and the highest release consistently occurred at 11 :30 am, regardless of batch (Figure 7b).

The reproductive output differed between batches and was nearly doubled for batch 1 and 2 in comparison to batch 3 with 2.3 ± 0.9, 2.4 ± 0.5 and 1.3 ± 0.4 l06 released swarmers per cm² respectively (Figure 8). Due to the low discharge of some thalli, only 59 samples could be used for counting the number of flagella on swarmers. The number of flagella on approximately 20 swarmers per sample was quantified in order to determine whether swarmers were bi- or quadriflagellate. Biflagellate swarmers were much more common (95%; 56 out of 59 analysed samples) than quadriflagellate swarmers (5%; 3 out of 59 analysed samples) in the samples.

Out of 150 thalli used to determine the viability and germling development of swarmers, a total of 147 thalli released swarmers. The released swarmers, both bi- and quadriflagellate, settled and germinated successfully in all 147 samples after 5 days. Phototactic behaviour of released swarmers

A total of 22 thalli released swarmers, all of which were biflagellate with a negative phototactic response. However, on one occasion, a small number of released biflagellate swarmers showed positive phototaxis, while the vast majority of swamers released from the same thallus were negatively phototactic. The average length and width of the biflagellate swarmers was 6.55 ± 0.85 (mean ± S.D.) and 3.75 ± 0.52 μπι, respectively.

Discussion

Photoperiod and temperature shock were successfully manipulated to enhance the formation and release of swarmers, while the effects of segmentation, dehydration, salinity, and time of initiation of experiments were negligible. The efficient manipulation of photoperiod and/or temperature shock is therefore the key in the reliable supply of swarmers with applications for fouling studies of tropical Ulva species and the seeding of nets for mass-cultivation. Therefore, one method - factoring in the maximum release and minimum practical effort and timing - is to collect Ulva in the morning and initiate the experiments in the early afternoon (at 1 pm) by washing the thalli in FSW, subsequently chilling the thalli for 10 min at 4°C and then placing them into autoclaved FSW under a l2 h L: 12 h D photoperiod at 25°C. Consequently, swarmers are released with peak after two days between 10:00 and 11 :30 am (Figure 9).

Notably, photoperiod had a significant effect on the formation and release of swarmers, with a discharge of up to 50% under normal photoperiod (12 h L: 12 h D). In contrast, extended light periods resulted in lower discharge and this is in agreement with temperate environments, where the day length plays a key role in the reproduction of seaweed (Forbord et al. 2012). While shorter day lengths result in a reduced growth rate and trigger reproduction, longer day length allows continuation of the vegetative growth phase for seaweeds in temperate systems. However, unlike previous studies on Ulva in temperate and cold waters where continuous light and dark cycles restrained the discharge of swarmers (Wichard & Oertel 2010), tropical Ulva sp. 3 continued to form and release swarmers under both extremes (up to 23% and 12%), respectively).

The present study highlights the importance of the dark phase for the formation and release of swarmers in tropical Ulva sp. 3, as discharge was halved when thalli were kept under continuous light. For Ulva sp. 3, the release of swarmers peaked at 4 ½ h after onset of light, around 11 :30 am.

A further factor affecting the formation and release of swarmers of Ulva sp. 3 was temperature shock. Sporulation was increased by approximately 10% when thalli were exposed to chilled (4°C) seawater for 10 min. The increased formation and release of swarmers following temperature shock may be a strategy to disperse under unfavourable conditions.

In general, stress treatments of segmentation (Dan et al. 2002), dehydration (Imchen 2012), and salinity (Dan et al. 2002) can initiate the formation and release of swarmers of Ulva (McArthur & Moss 1979). In contrast, these factors had no effect on the sporulation of Ulva sp. 3. This was unexpected, particularly that segmentation did not result in an increase of sporulation as shown for U. mutabilis, where sporulation increased by more than 60% when thalli were fragmented (Nordby 1977). The dehydration of filaments was also ineffective in increasing the sporulation of Ulva sp. 3. Previous studies have used this method to maximise the release of swarmers and dehydration times ranged from less than 1 h (Corradi et al. 2006) up to 12 h (Imchen 2012).

Therefore, the tested dehydration times of 45 min and 4 h are well within these timeframes and provide confidence that dehydration is ineffective for tropical Ulva sp. 3. In addition, neither dehydration nor wrapping in moist paper were effective constraining treatments for Ulva sp. 3 in the present study and swarmers were released during the desiccation process, indicating that once the reproductive cells are ready for release, this process might not be delayed.

The overall discharge of swarmers was generally low, at approximately 50% of previous studies, where sporulation reached up to 90 - 100 % (Mantri et al. 2011). In general, the discharge differs between sporophytes and gametophytes, with more than 90% in sporophytes, and only 40% for gametophytes (Hiraoka & Enomoto 1998). A total of 95% of thalli in the present study released biflagellate swarmers. The possession of a majority of negative phototaxis and the ability to germinate and grow within 5 days without fusing with complementary gametes suggests a simple asexual life history via biflagellate zoids (Hiraoka, Dan, et al. 2003). However, this requires confirmation through the cultivation of successive generations. In general, gametes are positively phototactic (Kuwano et al. 2012), while asexual biflagellate zoids are negatively phototactic (Hiraoka, Dan, et al. 2003), as are quadriflagellate zoospores (Mantri et al. 2011) and zygotes (Kuwano et al. 2012). The negative phototaxis guides swarmers to suitable surfaces for settlement and attachment (Callow & Callow 2006). Quadriflagellate swarmers were also released in the present study, although only from 5% of the thalli, and it remains unknown as to whether these are zoospores or asexual zoids (Hiraoka, Shimada, Ohno, et al. 2003). In general, Ulva species with a simple asexual life history produce either exclusively bi- or quadriflagellate zoids (Hiraoka, Dan, et al. 2003). It is possible that both bi- and quadriflgallate swarmers were found in this study because used samples could have been from different Ulva sepcies due to morphological similarity between filamentous Ulva species; alternatively Ulva sp. 3 may have several life histories as reported for U. prolifera (formerly Enteromorpha prolifera; Hiraoka, Dan, et al. 2003). Overall, the simplicity of the reproduction of Ulva sp. 3 may be a trait of tropical green macroalgae species that has genetic basis.

In conclusion, a method to induce the sporulation in tropical Ulva sp. 3 within 30 to 48 h after initiation was established. Sporulation was enhanced by temperature shocking thalli prior to incubating at for example a photoperiod of 12 h L: 12 h D. Swarmers were released two days after initiating experiments with a maximum release between 10:00 and 11 :30 am. More than 95% of the collected field population released biflagellate swarmers with negative phototaxis. These swarmers also had the ability to settle and germinate without crossing with complementary gametes. The findings of this study have application in laboratory antifouling assays for the tropics and in the seeding of nets for the mass-cultivation of seaweed. EXAMPLE 2

Materials and Methods

Collection of algal biomass and preparation of reproductive material

The species used in this study was Ulva sp. 3 (Shimada et al. 2008) and identified using molecular barcoding (Lawton et al. 2013). Ulva sp. 3 (hereafter Ulva) was characterised by flat tubular thalli and collected in the morning by hand from a land-based aquaculture facility at Guthalungra (19° 55'S, 147° 50Έ), Queensland, Australia. Permission was obtained from owners to collect algae from this site. Samples were placed in a 25 L container filled with pond water and then transported within 3 h to the laboratory at James Cook University in Townsville, Australia. Subsequently, the samples were gently washed three times with filtered seawater (FSW; 0.2 μπι and UV sterilised) to remove debris, epiphytes and invertebrates. To induce the release of swarmers, Ulva was shocked at a temperature of 4°C for 10 min and subsequently chopped using a blender in the early afternoon on the day of collection. The chopped filaments were retained with a sieve (120 μπι), washed with autoclaved FSW and subsequently placed in a crystallising dish filled with approximately 200 mL autoclaved FSW. Approximately 0.5 - 1 mL of this suspension was then transferred in Petri dishes filled with 10 mL autoclaved FSW using a transfer pipette and subsequently placed in a temperature controlled culture cabinet (Sanyo MLR-351) at 25°C at an irradiance of 125 μπιοΐ photon m^"2 s^"1 under a 12 h L: 12 h D photopenod to maximise the release of swarmers. The release of swarmers peaked after two days between 10:00 and 11 :30 am. The Petri dishes were then emptied in a beaker through a sieve (120 μπι) to filter out the chopped filaments, while the released swarmers were collected with the seawater in the beaker. The density of the swarmers was calculated using a haemocytometer.

Optimal seeding density and nursery period

To determine which seeding density and nursery period resulted in the highest biomass yield and growth, swarmers were seeded onto polypropylene ropes (0 4 mm, Syneco) at densities of 155, 310, 466, 621, and 1,552 10³ swarmers m^"1 rope. A total of 45 pieces of rope were seeded, each with a length of 580 mm. These were attached to the bottom of individual Petri dishes (0 60 mm, Techno Plas; S6014S10) as a flat spiral coil using the pressure-sensitive adhesive Ezy Tack (Selleys). The seeding density was altered by manipulating the density of swarmers to 5, 10, 15, 20, and 50 x 10³ swarmers mL^"1 and then adding 18 mL of this swarmer suspension to each Petri dish with a piece of rope using a syringe. The tested nursery periods were 1, 5, and 10 days and the Petri dishes were kept in a culture cabinet (Sanyo MLR-351) in a 12 h L: 12 h D photoperiod at 25°C during this period. The water was changed, where applicable, after 3, 5, and 8 days. There were three replicates (n = 3) for each seeding density for each tested nursery period. Subsequent to the time in the nursery, the seeded ropes were carefully removed from the Petri dishes using tweezers, uncoiled and transferred to aerated flow-through outdoor tanks under ambient light at the Marine & Aquaculture Research Facilities Unit (MARFU).

A total of 5 seeded ropes (one from each density) were attached to a weighted frame (380 x 500 mm) using cable ties. Each frame was placed on the bottom of a tank (n = 9 replicate tanks) so that the ropes were immersed horizontally in the water at a depth of approximately 100 mm below the surface. To minimise temperature fluctuations, the tanks holding the ropes were placed in a circulating water bath (see cultivation conditions following). After 7, 14, and 21 days of outdoor cultivation, the tanks, frames and air lines were cleaned and the seeded ropes were repeatedly sampled to measure the fresh weight (FW) of Ulva. The ropes were spun dry and weighed (FW) using a digital balance (Kern KB 3600-2N). After 21 days of outdoor cultivation, the seeded ropes were dried in an oven at 65°C for at least 7 days to determine the dry weight (DW) of Ulva. The fresh and dry biomass yield was calculated as FW (g FW m^"1 rope) and DW (g DW m^"1 rope) per linear meter of rope, respectively. Individual FW:DW ratios were calculated for each replicate. Specific growth rate was calculated for each replicate using the equation SGR (% day^"1) = In (B₂ / Bi) / (t₂ - ti)^■ 100, where Bi and B₂ are the biomasses (g FW) at time ti and t₂ of outdoor cultivation (days).

Experiments were conducted during the Australian winter with an average water temperature of 25°C and the seeded ropes received on average photo synthetically active radiation of 29.2 (± 6.4 S.D.) mol photons m^"2 day^"1. The tanks were on a recirculating system with an average nitrate-N and phosphorous concentration of 2.09 (± 1.47 S.D.) mg N0₃-N L^"1 and 0.37 (± 0.22 S.D.) mg P L^"1, respectively. Each tank held approximately 28 L of FSW (1 μπι) and had a flow rate of 0.5 L min^"1. The average salinity was 31.4 (± 1.4 S.D.) ppt.

Biomass yield and growth of Ulva seeded at optimal seeding density and nursery period

To determine the dry biomass yield, growth, FW:DW ratio, ash content and change in morphology over time, ropes were seeded at an optimal density of 621,000 swarmers m^"1 rope and maintained for five days under nursery conditions (see results Optimal seeding density and nursery period'). The arrangement of ropes in Petri dishes, and the seeding and nursery procedure, followed the methods described in the previous experiment. After five days under nursery conditions, the seeded ropes were removed from the Petri dishes and transferred to aerated flow-through outdoor tanks at MARFU as described above. Each seeded rope was individually attached to a weighted frame placed on the bottom of a tank holding approximately 28 L of FSW (1 μπι and UV sterilised) for 25 days. Holding tanks, frames and air lines were cleaned after 7, 13, and 19 days. The seeded ropes were destructively sampled (n = 3) after 7, 10, 13, 16, 19, 22, and 25 days of outdoor cultivation and photographed with a digital camera (Canon PowerShot D20) to determine the width of filaments (n = 50 per samples) using Image J freeware (www.nih.giv). Subsequently, samples were spun dry and weighed to determine the growth (SGR) using the equation above where Bi and B₂ are the average biomasses (g FW). Thereafter, the samples were dried in an oven at 65°C for at least 7 days to quantify the DW and to calculate the dry biomass yield as DW per linear meter of rope (g DW m^"1 rope). FW:DW ratios were calculated as described above. The ash content of each replicate was quantified by heating a 1.5 g homogenised subsample of dried biomass at 110°C in a moisture balance until a constant dry weight was reached and then combusting at 550°C in a muffle furnace for 24 h until a constant weight was reached.

Experiments were conducted with a total of three independent sample collections of Ulva as initial parent generation used as a source of swarmers to seed the ropes during the Australian spring (hereafter referred to as 'batch'). The water temperatures in the tanks ranged between a night time minimum of 21.7°C and a daytime maximum of 36.4°C during the trial, with an average water temperature of 27.8 (± 3.0 S.D.)°C. The average salinity was 32.7 (± 0.9 S.D.) ppt and the seeded ropes received an average photosynthetically active radiation of 47.0 (± 6.8 S.D.) mol photons m^"2 day^"1 during the experiment. The tanks had a flow rate of 0.5 L min^" ¹ and were on a recirculating system with an average concentration of nitrogen as nitrate of 1.50 (± 1.89 S.D.) mg NO₃-N L^"1 and an average concentration of phosphorous of 0.40 (± 0.25 S.D.) mg P L^"1.

Statistical analysis

To formally test the effect of seeding density on the biomass yields (fresh and dry) after 21 days of outdoor cultivation for each nursery period in the first experiment, density was considered as a fixed factor and tank as an unreplicated blocking factor as seeded ropes in one tank were not independent. To determine the effect of nursery period and sampling day on the fresh biomass yields of seeded ropes, the optimal seeding density of 621,000 swarmers m^"1 (see results) was selected for PERMANOVA analysis and nursery was considered as fixed and sampling day as random factors. To analyse the effect nursery period on the dry biomass yield after 21 days of outdoor cultivation of ropes seeded at 621,000 swarmers m^"1, data was analysed using an one-factor PERMANOVA. To test for the effect of sample collection on the dry biomass yield, FW:DW ratio, SGR, ash content, and width of filaments over time in the second experiment, two-factor PERMANOVAs were used with time as a fixed factor (with destructively sampled replicates) and sample collection as a random factor. FW:DW ratios for both experiments were correlated with dry biomass yield for each replicate using the software Statistica (v.12).

Results

Optimal seeding density and nursery period

The highest fresh biomass yields were obtained for ropes maintained for five days in the nursery, regardless of seeding density, and had approximately two to six times the biomass than ropes maintained under shorter and longer nursery periods (Figure 11). The highest fresh biomass yield was obtained for the seeding density of 621,000 swarmers m^"1 rope after the nursery period of five days, and this density was nearly double the yield obtained at any other seeding density and nursery period (Figure 11). Therefore, this seeding density in combination with a five day nursery period was chosen for the subsequent experiment on the basis of biomass yield. When specifically analysing the fresh biomass yield of ropes seeded at 621,000 swarmers m^"1 rope to identify differences between nursery periods over time within a single seeding density, there was a significant interactive effect between nursery and days of outdoor cultivation, driven by significant differences between nursery periods and lower biomass yields overall after day 7 in comparison to day 14 and 21 (Table 2).

After the shortest nursery period (one day), the average fresh biomass yields increased for all seeded ropes over 21 days and also increased with increasing seeding densities (Figure 11a). The lowest biomass yields were observed after seven days and were below 0.4 g FW m^"1 rope for all seeding densities. In the following days, the yields increased more than 10-fold and ranged from 5.5 ± 1.4 g FW m^"1 rope to 27.5 ± 7.1 g FW m^"1 rope after 14 days and from 12.1 ± 4.3 g FW m^"1 rope to 48.5 ± 17.1 g FW m^"1 rope after 21 days (Figure 1 la). The highest seeding density of 1,552 x 10³ swarmers m^"1 rope had a greater biomass yield than lower seeding densities after a nursery period of one day, despite some variability between tanks in the analysis (Table 3).

After the nursery period of five days, the biomass yields were generally low after seven days and increased approximately 16-fold during the following days, ranging from 30.8 ± 10.9 g FW m^"1 rope to 72.5 ± 6.6 g FW m^"1 rope at day 14. Subsequently, the yields decreased after 21 days and ranged from 17.7 ± 5.8 g FW m^"1 rope to 42.9 ± 7.9 g FW m^"1 rope (Figure 1 lb).

After the longest nursery period (ten days), the biomass yields ranged from

10.1 ± 1.9 g FW m^"1 rope to 18.2 ± 1.3 g FW m^"1 rope after seven days (Figure 11c) and were generally higher than for shorter nursery periods. However, the biomass yields overall were low for the following days and below 23 g FW m^"1 rope at all times, which is less than half the yield than that of shorter nursery periods.

Similar to the fresh biomass yields, the dry biomass yields overall were generally higher for ropes maintained at a nursery period of five days (4.2 ± 0.5 g DW m^"1 rope) compared to shorter (3.1 ± 0.6 g DW m^"1 rope) and longer nursery stages (1.8 ± 0.3 g DW m^"1 rope) (Figure 12). The average dry biomass yields ranged from 2.4 ± 0.8 g DW m^"1 rope to 5.6 ± 1.2 g DW m^"1 rope for a nursery period of five days, and were halved for a longer nursery period of ten days where yields ranged from 0.8 ± 0.1 g DW m^"1 rope to 2.7 ± 1.0 g DW m^"1 rope. Notably, the dry biomass yields were measured after 21 days of outdoor cultivation when the biomass on ropes maintained under a nursery period of five days had already degraded, on contrast to the shorter nursery period of one day which continued to increase. Nursery period had no significant effect when specifically analysing the dry biomass yield of ropes seeded at a single seeding density (Table 2). The dry biomass yield increased nearly 3-fold from the lowest seeding density of 155,000 swarmers m^"1 rope (1.6 ± 0.4 g DW m^"1 rope) to the highest seeding density of 1,552 x 10³ swarmers m^"1 rope (4.3 ± 0.9 g DW m^"1 rope) (Figure 12) with significant differences in dry biomass yield between seeding densities for nursery periods of one and ten days (Table 3). The average FW:DW ratios were highly variable ranging from 4.7 ± 1.0 to 8.6 ± 0.3 and generally decreased with increasing nursery periods with no consistent trend between FW:DW ratio and seeding density. There was no correlation between the FW:DW ratio and the dry biomass yield (r = 0.185, p = 0.224 ).

In contrast to fresh and dry biomass yields, specific growth rates were highest for the shortest nursery period of one day across all seeding densities because of the low initial biomass for each measurement (Figure 13). The average growth rates for

Ulva maintained at a nursery period of one day ranged from 46.4 ± 1.7 % day^"1 to

64.4 ± 1.0 % day^"1 between 7 and 14 days of outdoor cultivation, while those maintained under the longest nursery period of ten days ranged from 2.4 ± 2.6 % day^"1 to 9.5 ± 1.6 % day^"1 (Figure 13a). The growth rates between 14 and 21 days of outdoor cultivation had substantially lower growth rates with the shortest nursery period being the only treatment with positive values (Figure 13b).

Biomass yield and growth o Ulva seeded at optimal seeding density and nursery period

The average dry biomass yield increased more than 10-fold from day 7 (1.8 ± 0.5 g DW m^"1 rope) to day 10 (10.1 ± 4.9 g DW m^"1 rope) and then day 13 (23.0 ± 8.8 g DW m^"1 rope) (Figure 14a). Subsequently, the biomass yield decreased to below 20.4 g DW m^"1 rope for extended time periods of outdoor cultivation. There was a significant interaction of batch and days of outdoor cultivation (Table 4), driven by differences in the time when the highest biomass yield was attained among batches (Figure 14b). While the biomass yield peaked at day 13 for the first batch (40.0 ± 3.3 g DW m^"1 rope) and third batch (19.0 ± 2.8 g DW m^"1 rope), the second batch had the highest biomass yield at day 16 (16.2 ± 0.4 g DW m^"1 rope). Furthermore, there were striking differences in the biomass yield between batches; however, batches showed a similar trend of an initial increase and subsequent decrease in dry biomass yield over time (Figure 14b). Notably, the biomass matured and was reproductive during the course of the experiment and the release of swarmers occurred after 13 days of outdoor cultivation. Average specific growth rates decreased significantly over time (Figure 14c, Table 4) with the highest growth rate between 7 and 10 days of outdoor cultivation (67.1 ± 12.2 % day^"1). There was a subsequent reduction in growth rate between 10 and 13 days (29.9 ± 10.9 % day^"1). After 13 days of outdoor cultivation, growth rates were generally lower and ranged from -13.4 ± 16.4 % day^"1 (days 13 to 16) to 3.5 ± 9.8 % day^"1 (days 16 to 19).

Average FW:DW ratios ranged from 5.9 ± 0.5 (day 7) to 8.9 ± 1.3 (day 13) (Figure 15a) and overall patterns were generally variable between batches, reflected by a significant interaction between batch and days of outdoor cultivation (Table 4). In addition, FW:DW ratios were positively correlated with an increase in dry biomass yield (r = 0.788, p < 0.001). The average width of filaments increased more than 7-fold from day 7 (86.7 ± 7.0 μιη) to day 19 (644.3 ± 158.4 μιη) (Figure 15b). Subsequently, the average width decreased to 390.3 ± 73.7 μπι and 390.6 ± 63.0 μπι at days 22 and 25, respectively. The ash content ranged from 25.9 ± 0.6 % (day 7) to 36.2 ± 4.9 % (day 13) (Figure 5c).

Discussion

This study demonstrates the importance of seeding density and nursery period on the biomass yield and growth rate of Ulva sp. 3 when artificially seeded onto ropes. Under certain cultivation conditions high biomass yields were achieved at a seeding density of 621,000 swarmers m^"1 rope and a nursery period of five days. Seeding density was a key factor affecting both growth rate and biomass yield of Ulva sp. 3 confirming previous studies, where the biomass and size of seaweeds was related to their settlement density (Zhang et al. 2012). In general, the density of germlings is negatively related to growth and survival due to intraspecific competition and shading effects (Steen and Scrosati 2004), yet low seeding densities on ropes can result in higher growth of epiphytes and thus high seeding densities are also a strategy to control biofouling on cultured seaweeds (Liining and Pang 2003). However, this might be less critical for fast-growing and opportunistic species such as filamentous Ulva, meaning that the selection of seeding density can be based on controlled cultivation conditions. In addition, the seeding density for Ulva of 621,000 individuals m^"1 rope, as quantified in this study, is generally much higher than for other species, where common seeding densities are 2,000 individuals m^"1 rope for the brown seaweeds Undaria pinnatifida (Peteiro and Freire 2012) and Sargassum fulvellum (Hwang, Park, and Baek 2006), 2,000 to 3,000 individuals m^"1 rope for Saccharina latissima (Peteiro and Freire 2013), and 20,000 individuals m^"1 rope for the red seaweed Gracilaria chilensis (Alveal et al. 1997). Furthermore, brown and red seaweeds typically have longer nursery and culture cycles with correspondingly higher productivities expected per linear metre (~7 kg FW m^"1 rope after 5 months for G. chilensis; Alveal et al. 1997). This highlights the differences in the morphology and cultivation techniques of commercial brown and red seaweeds compared to filamentous Ulva, with clear distinction in the length of production cycle. A further key factor affecting growth and biomass yield of Ulva was the nursery period prior to grow-out. This period is important for the success of viable mass-cultivation of Ulva as an increased contact time acts to minimise detachment and loss of germlings due to hydrodynamic forces (Zhang et al. 2012). This effect is consistent with the difference in fresh biomass yields for seeded ropes maintained under nursery conditions for one and five days where values were lower for the shorter nursery period, most probably due to higher detachment of settled germlings after only one day. Interestingly, an extended nursery period of ten days had adverse effects on the growth of Ulva with the lowest biomass yields and growth rates overall among nursery periods. This implies that resources may be limited under static nursery conditions (Hurd 2000; Kregting et al. 2008) resulting in poor growth performance of the germlings.

The trend of initially increasing and then decreasing biomass yield, coupled with decreasing growth rates over time, is likely due to a general decrease of light availability with increasing biomass on the ropes (Demetropoulos and Langdon 2004). Furthermore, the decline in growth of Ulva sp. 3 also reflects the maturation of biomass and the reproductive events after 13 days of outdoor cultivation. This short life cycle is characteristic of opportunistic species. Vegetative cells of Ulva transform directly into reproductive cells and, therefore, maturation and reproduction results in the degradation of filaments once the swarmers have been released. As such, the timing of harvest is important to maximise yields. Furthermore, it is also important if mature populations on the ropes are to be used as parental stock for future generations through artificial seeding.

The species Ulva sp. 3 has a high biomass yield and growth rate as shown in this study, and an ability to integrate with existing aquaculture facilities (Lawton et al. 2013), which together promise to make algal cultivation for biomass applications more cost-effective. The specific growth rate was high at more than 65% day^"1 which is higher than recorded in a previous laboratory study (-30% day^"1; Lawton et al. 2013). Overall, the growth rate was higher than for other species of Ulva and macroalgae, such as U. reticulata (-4% day^"1; Msuya, Kyewalyanga, and Salum 2006), U. ohnoi (-38% day^_1;Yokoyama and Ishihi 2010), Cladophora and Chaetomorpha (-44% day^"1; de Paula Silva et al. 2008), Saccharina latissima (-5% day^"1; Handa et al. 2013), and Porphyra linearis (-16% day^"1; Kim et al. 2007). Furthermore, Ulva sp. 3 was a robust species in this study tolerating varying environmental conditions across the experimental period. Water temperatures reached up to 36.4°C, without any signs of degradation of biomass, and other species of filamentous Ulva can be grown in water temperatures up to 40°C (Moll and Deikman 1995). In addition, filamentous species of Ulva generally have a broad tolerance towards a wide range of salinities, including as low as 5 psu over 7 days, without a significant decrease in the viability of cells (Ichihara et al. 2013). As such, Ulva sp. 3 may be used for year-around cultivation in tropical environments which are typically characterised by strongly seasonal rainfall providing a challenge for algal species. Furthermore, Ulva sp. 3 is a native species common at land-based aquaculture facilities in Eastern Australia and could therefore also be considered for bioremediation of waste waters from land-based aquaculture (Lawton et al. 2013).

In conclusion, this study highlights the importance of controlling seeding densities in culture with large effects of doubling of the biomass yield at an optimal range per length of rope or potentially any unit of culture infrastructure. Therefore, ropes artificially seeded at an optimal density under controlled conditions will importantly maximise yields with an efficient use of the resource inputs and this provides a fundamental step in the cultivation of filamentous Ulva at scale. A reliable source of biomass for year-round production through optimised hatchery and culture cycles, in line with maximising biomass production per unit effort, allows for a reliable bioremediation strategy and further permits the development and management of selected breeding lines (genotypes). This study also revealed the need to select genotypes with high productivity for the cultivation of this species and to test the potential interactive effects with environmental conditions. Finally, populations seeded onto ropes can be used as parental plants for seedling production, which in turn reduces the need to harvest wild stocks.

EXAMPLE 3

Material and Methods

Algal biomass and preparation of reproductive material

The species used in this study was Ulva sp. 3 (Shimada et al. 2008) identified using molecular barcoding (Genbank accession number KM406999). Biomass of Ulva sp. 3 was maintained in culture for more than four months in a temperature and light controlled laboratory (23°C, 12 h light: 12 h dark cycle, 50 μπιοΐ m^"2 s^"1) at James Cook University (JCU) in Townsville, Australia. To obtain seedlings for artificial seeding, the release of swarmers was induced using a temperature shock at 4°C for 10 min (Carl et al. 2014) and the filaments were subsequently cut using a blender (Carl, de Nys, and Paul 2014). The release of swarmers peaked after two days between 10:00 and 11 :30 am and the density of swarmers was calculated using a haemocytometer.

Experimental set-up

To determine the effect of multiple harvests on the yield and quality of the harvested biomass, the released swarmers were artificially seeded onto ropes at a density of 621,000 swarmers m^"1 rope (n = 13 for each batch; see below) and maintained for five days under nursery conditions with a water change after three days. Subsequent to the time in the nursery, the seeded ropes were transferred to aerated flow-through outdoor tanks holding approximately 28 L of FSW (1 μπι and UV sterilised) under ambient light at the Marine & Aquaculture Research Facilities Unit at JCU. Each seeded rope was individually attached to a weighted frame (380 x 500 mm) using cable ties and each frame was placed on the bottom of a tank so that each rope was immersed horizontally in the water at a depth of approximately 100 mm below the water surface. To minimise temperature fluctuations (see 'Cultivation conditions' below), the tanks holding the ropes were placed in a circulating water bath. Holding tanks, frames and air lines were cleaned weekly. The ropes were repeatedly sampled weekly, spun to remove excess water and weighed to determine the fresh weight (FW) and growth of the biomass for each replicate. The algal FW for each replicate was calculated by subtracting the weight of the moist rope from the total weight of the seeded rope with the algal biomass. The specific growth rate (SGR) for each replicate was calculated using the equation SGR (% day^"1) = In (B₂/Bi) / (t₂-ti) · 100, where Bi and B₂ are the biomasses (g FW) at time ti and t₂ (days).

After 14 days of outdoor cultivation, the biomass was harvested (harvest 1) by cutting off the seaweed using scissors and leaving approximately 1 cm of biomass on each rope. Ropes with the remaining biomass were then returned to the outdoor tanks under the previous conditions. The harvested biomass of all replicate ropes for each harvest and batch was pooled and then freeze-dried for 24 h (Vitris benchtop 2K, VWR, Australia). The dried biomass was then homogenised using a coffee grinder (Breville, CG2B) and maintained in the dark at -20°C in airtight containers until further processing (see 'Yield and quality of harvested biomass' below). For the subsequent second (harvest 2) and third harvest (harvest 3), the biomass on the ropes were harvested as described above at day 28 and 42 of outdoor cultivation, respectively. This corresponds to a total of three cycles of culture and harvest at 14 days for each independent production cycle. Experiments were conducted at different times with three batches of swarmers from three independent reproductive events (hereafter referred to as 'batch') resulting in three independent production cycles.

Cultivation conditions

Experiments were conducted from April to August 2014. The water temperatures in the outdoor tanks ranged between 15.9°C and 32.6°C during the trial, with an average water temperature of 25.0 (± 1.3 S.D.)°C. The average salinity was 32.6 (± 1.1 S.D.) ppt and the seeded ropes received an average photo synthetically active radiation of 27.5 (± 7.6 S.D.) mol photons m^"2 day^"1 during the experiment. The tanks had a flow rate of 0.5 L min^"1 and were on a recirculating system. The nutrient concentration was measured three times per week and the concentration of nitrogen as nitrate was adjusted to 1 - 3 mg NO₃-N L^"1. The average concentration of nitrogen as nitrate and phosphorous was 2.6 mg NO₃-N L^"1 (± 0.2 S.D.) and 0.5 (± 0.1 S.D.) mg P L^"1, respectively.

Yield and quality of harvested biomass

To determine the yield of the cultivated biomass for each harvest, the biomass from all replicate ropes within each harvest in each batch was pooled, freeze-dried and weighed. The dry biomass yield was calculated as dry weight (DW) per linear meter of rope (g DW m^"1 rope). Given that only two successive harvests are feasible (see Results), only the biomass of the first and second harvest were characterised and analysed to determine the quality of the cultivated biomass. The quality was defined by biomass morphology, colour, ash, elemental composition and mineral content. In addition, amino acids (protein), lipids, carbohydrates and fibre were analysed for batch 1 and 3, given their high biomass yield, and not for batch 2 as there was insufficient biomass for these additional analyses (see Results).

To quantify morphology (n = 50 per sample), the fresh biomass of each rope was photographed upon harvest using a digital camera (Canon PowerShot D20) and the width of filaments was then analysed using Image J freeware (www.nih.gov). The colour of the homogenised dried biomass of each harvest for each batch was matched by visual comparison to the Pantone swatch book reference (GP1301XR, The Plus Series; Solid Uncoated) using two assessors. The ash content of the dried biomass of each harvest for each batch was quantified by heating a 2 g homogenised subsample of dried biomass at 110°C in a moisture balance until a constant dry weight was reached. The sample was then split into triplicates and subsequently combusted at 550°C in a muffle furnace for 24 h until a constant weight was reached. Furthermore, a homogenised subsample of the dried biomass was analysed for carbon (C), hydrogen (H), oxygen (O), nitrogen (N), sulphur (S) and iodine (I) content by ultimate analysis of each harvest for each batch. The samples were analysed by the OEA Laboratories (Cornwall, UK). In addition, a homogenised subsample of the dried biomass was analysed for a total of 23 minerals (Al, As, B, Ba, Ca, Cd, Co, Cr, Cu, Fe, Hg, K, Mg, Mn, Mo, Na, Ni, P, Pb, Se, Sr, V and Zn) following Roberts et al. (2013). Analyses were conducted by the Advanced Analytical Centre at JCU (Townsville, Australia).

For dried biomass samples of batch 1 and 3 (batch 2 did not provide sufficient biomass; see Results), amino acids (protein), lipids, carbohydrates, and fibre were determined. Amino acid profiles were quantified by hydrolysis following analysis using the Waters AccQTag Ultra chemistry on a Waters ACQUITY UPLC at the Australian Proteome Analysis Facility (Sydney, Australia). The theoretical protein content was calculated as the sum of all amino acids. The content of essential amino acids was calculated as the sum of histidine, isoleucine, leucine, lysine, methiodine, phenylalanine, threonine and valine. The total lipid content was analysed following Folch et al. (1957). The total carbohydrate content was determined by difference using the equation Carbohydrates (%) = 100% - (ash + moisture + total lipids + protein content). The total, insoluble and soluble fibre content was analysed using the enzymatic-gravimetric method and analyses were conducted by Grain Growers Ltd (North Ryde, Australia).

Statistical analysis

The data was statistically analysed by permutational analysis of variance (PERMANOVA) using PRIMER 6 (v. 6.1.13) and PERMANOVA+ (v. 1.0.3) (Clarke and Gorley 2006). The Bray-Curtis dissimilarity measure was used for all PERMANOVAs on square root transformed data and /^-values were calculated using permutation of residuals under a reduced model with 9999 random permutations (Anderson et al. 2008). All data are reported as mean ± 1 standard error (S.E.) unless stated otherwise. To formally test the effect of multiple harvests on the dry biomass yield, two- factor PERMANOVAs were used with harvest as a fixed factor and batch as a random factor. To determine changes of the specific growth rate over time, batch and time (days) were considered as random and fixed factor, respectively.

Results

Yield of harvested biomass

The average dry biomass yield decreased with the increasing number of harvests for all batches (Figure 16a) with a similar biomass yield for the first (7.0 ± 3.2 g DW m^"1 rope) and second harvest (6.6 ± 3.4 g DW m^"1 rope), followed by a more than five-fold decrease for the third harvest (1.2 ± 0.6 g DW m^"1 rope). The average biomass yield varied significantly among batches (Figure 16b) and there was a significant interaction between 'batch' and 'harvest' (Table 5a). The difference between batches was most pronounced in the first two harvests for batches 1 and 3 with a yield more than eight-fold higher than batch 2 (Figure 16b).

The average specific growth rates decreased over time (Figure 17) with the highest growth rate between days 7 and 14 (31.91 ± 3.0% day^"1). Subsequently, the growth rate halved between days 14 and 21 (15.4 ± 7.0% day^"1) and decreased further to below 9% day^"1 after day 21 with negative growth rates between days 35 and 42 (-3.6 ± 3.3% day^"1). There was a significant interaction between days of cultivation and batch, and highly significant effects for both of the main effects (Table 5b).

Quality of the harvested biomass

The quality parameters of ash content, elemental composition, mineral, colour, lipid and carbohydrate content were consistent between harvests, whereas the width of filaments, protein and fibre content increased from the first to the second harvest. The average width of filaments increased from 140.3 ± 28.3 μπι for the first harvest to 215.8 ± 39.6 μπι for the second harvest. The colour of the dried biomass was dark green and similar between harvests and batches.

The ash and moisture contents of Ulva sp. 3 were consistent with ash ranging from 29.7 ± 0.9% for the first harvest to 26.8 ± 1.4% for the second harvest. The moisture content ranged from 6.4 ± 0.9% for the first harvest to 5.8 ± 0.5% for the second harvest. The elemental composition of Ulva sp. 3 was also relatively consistent between harvests. Carbon (30-31%) and oxygen (21-23%) were the major elements characterised by ultimate analysis with sulphur being the lowest (3%). Potassium (K) and sodium (Na) were the main minerals in the harvested biomass, followed by magnesium (Mg), calcium (Ca) and phosphorous (P). The content of the 24 elements measured in the harvested biomass was relatively consistent between harvests with the exception of nickel (Ni), iodine (I) and potassium. The relative content of nickel was low (< 1 mg 100 g^"1 dried biomass) but more than doubled from the first to the second harvest. Similarly, the relative content of iodine was low (< 10 mg 100 g^"1 dried biomass) but also doubled from the first to the second harvest. In contrast, the potassium content (> 4,300 mg 100 g^"1 dried biomass in the first harvest) approximately halved in the second harvest.

Carbohydrates were the main biochemical component of Ulva sp. 3 and made up 46% of the dried biomass. Lipids were the smallest component at less than 2%. Notably, the content of carbohydrate and lipid was similar between harvests. In contrast, the protein content (the sum of all amino acids) increased by more than 25% from the first to the second harvest (Table 6) and this proportional increase occurred for all amino acids. Aspartic acid and glumatic acid, including their respective amides, were the main amino acids making up 29% of the total amino acid content. The quantity of the essential amino acids lysine and methionine, as a proportion of total amino acids, decreased by approximately 10% from the first to the second harvest. This decrease was more pronounced for lysine which decreased from 5.5 to 4.8% of the total amino acid content. Overall, the proportion of essential and non-essential amino acids remained consistent between harvests.

Dietary fibre was the main component of carbohydrates comprising between 55%) and 65%> of total carbohydrate. The total dietary fibre content increased by more than 20% from the first to the second harvest with insoluble fibre increasing by 35%). In contrast, the soluble fibre content decreased by approximately 5%.

Discussion

This study demonstrates that Ulva sp. 3 can be harvested multiple times during a production cycle to optimise cultivation. Notably, the biomass yield was similar for the first two harvests, while the third harvest resulted in a more than five- fold decrease in yield coupled with decreased growth rates and an increase of fouling. Therefore, a maximum of two harvests may be beneficial for Ulva sp. 3, at least under the conditions assessed here, resulting in approximately double the total biomass yield of that for ropes only harvested once. Multiple harvests generally result in greater total production (Ohno 2006) coupled with minimised operational inputs in the seeding and nursery stage (Carl, de Nys, and Paul 2014). This facilitates effective cultivation with two harvests in one production cycle reducing the time period required to yield equivalent biomass from two harvests from six to five weeks.

The biomass yield was highly variable between batches and ranged from 1 to

13 g DW m^"1 rope for the first two harvests with an average of 7 g DW m^"1 rope. Previously studies have also shown highly variable yields of Ulva sp. 3 (Carl, de Nys, and Paul 2014). Notably, previous studies harvested the entire biomass after a cultivation cycle of 14 days, while the present study only partially harvested the biomass on the ropes. Consequently, biomass remained on the ropes when only partially harvested resulting in a comparatively reduced yield. Other filamentous species of Ulva are grown on nets with yields ranging from 60 to 485 g DW m² (Ohno 1993), however it is difficult to make quantitative comparisons based on the different cultivation techniques of nets versus ropes. Other commercial brown and red seaweeds are cultivated on ropes with longer production cycles (> 6 months) and consequently higher overall yields (Alveal et al. 1997; Hwang et al. 2006; Hwang et al. 2010; Dring et al. 2013).

It is important to note that the number of harvests during the production cycle depends on the species of seaweed being cultured. For example, the brown seaweed Cladosiphon is harvested up to ten times during a cultivation period of three to six months (Ohno 2006), while the green seaweed Monostroma is harvested two to four times over an eight month cultivation period (Kida 1990; Ohno 1993; Ohno 2006). Filamentous species of Ulva, including U. prolifera, U. compressa and U. intestinalis, are harvested two to three times during a two to three months culture period (Ohno 1993; Ohno 2006).

The biochemical characteristics of Ulva sp. 3, analysed for the first time, show that this species has high nutritional value as a food product. The main biochemical component was carbohydrates, which made up nearly 50% of the biomass. The carbohydrates of Ulva are mostly cell-wall polysaccharides which are also referred to as dietary fibre and the consumption of these are health promoting (Elleuch et al. 2011). The second largest component of Ulva sp. 3 was ash (dry inorganic content) which is similar to other filamentous species of Ulva ranging from 21% to 29% (McDermid and Stuercke 2003; Zhuang et al. 2012). The ash content is primarily made up of minerals (Holdt and Kraan 2011) which are differentiated as trace elements and macro -minerals depending on the quantity required by the body.

In general, the ash content of seaweeds is much higher than terrestrial crops (Ross et al. 2008) with the ash content of most vegetables being < 1.1% on dry weight basis (Hanif et al. 2006; de Souza Araiijo et al. 2014). The main minerals of Ulva sp. 3 were potassium, sodium, magnesium, calcium and phosphorous, all of which are essential macro-minerals for human (Suter et al. 2002) and animal health (Suttle 2010). The mean sodium and potassium content of Ulva sp. 3 (Na: -4.8%; K: -3.1%)) was similar to other species of Ulva (Kumar et al. 2011). However, Ulva sp. 3 has a low Na:K ratio similar to other seaweeds (Ruperez 2002) and is therefore of interest in food applications to reduce their sodium content (Gupta and Abu- Ghannam 2011).

Furthermore, the addition of Ulva sp. 3 in food can increase the content of magnesium and calcium which is currently under consumed in the Western diet. Ulva sp. 3 has a high magnesium content of up to 1.4%. This is in agreement with other studies where Ulva has a higher magnesium content than other seaweeds (Hwang et al. 2008; Kuda and Ikemori 2009; Kumar et al. 2011). In contrast, the iodine content of Ulva sp. 3 (-0.003 - 0.009%>) is relatively low compared to other seaweeds (Lee et al. 1994; Teas et al. 2004; Holdt and Kraan 2011). However, the iodine content of Ulva sp. 3 is approximately an order of a magnitude higher than for most terrestrial crops (Mahesh et al. 1992). Consequently, it is a promising food and feed supplement to correct nutritional deficiencies in both human and animal diets with its high content of essential trace elements and macro-minerals.

Protein (sum of all amino acids) was the third largest component of Ulva sp. 3 with an average of 19.1% of the dried biomass which is in the range for Ulva (7 - 29%; Wong and Cheung 2000; Ortiz et al. 2006; Marsham et al. 2007; van der Wal et al. 2013; Neveux et al. 2014). The quality of the protein was consistent between harvests and the ratio of essential to non-essential amino acids was approximately 60:40.

Overall, multiple harvests during the production cycle of Ulva sp. 3 resulted in an increase in total production, while maintaining desirable nutritional composition. Ulva sp. 3 represents a product high in fibre, essential minerals and proteins with use as a functional food and feed ingredient. Throughout the specification the aim has been to describe the preferred embodiments of the invention without limiting the invention to any one embodiment or specific collection of features. It will therefore be appreciated by those of skill in the art that, in light of the instant disclosure, various modifications and changes can be made in the particular embodiments exemplified without departing from the scope of the present invention.

All computer programs, algorithms, patent and scientific literature referred to herein is incorporated herein by reference.

Table 1: PERMANOVA output testing the effect of photoperiod (Ph; 12 h L: 12 h D, 18 h L:6 h D, 24 h L, and 24 h D), time of initiation (In; 1pm and 7pm on collection day, 7am, 1pm, and 7pm one day after collection), sampling day (Day; 3 and 4 days past collection), and time of sampling day (sTime; 7am, 1 pm, and 7 pm) (all fixed factors) on the discharge of swarmers.

Source df F P

Ph 3 68.05 < 0.001

In 4 5.01 < 0.001

Day 1 68.66 < 0.001

sTime 2 12.89 < 0.001

Ph In 12 2.67 < 0.001

Ph Day 3 5.33 < 0.001

Ph x sTime 6 2.81 0.001

In x Day 4 1.72 0.101

In x sTime 8 0.90 0.549

Day x sTime 2 6.83 < 0.001

Ph x In x sTime 12 1.08 0.356

Ph x In x sTime 24 0.89 0.688

Ph x Day ^χ sTime 6 1.43 0.140

In x Day ^x sTime 8 1.70 0.047

Ph x In x Day ^χ 24 0.75 0.886

sTime

Residual 960

Table 2: PERMANOVA analysis on Bray-Curtis distances testing the effects of nursery period (Nursery, fixed factor) and days of outdoor cultivation (Day, random factor) on the fresh biomass yield of ropes seeded at 621 ^χ 10³ swarmers m^"1 rope; and the effect of nursery on the dry biomass yield after 21 days outdoor cultivation. Mean square (MS), pseudo-F (F and P values are presented, significant terms shown in bold.

Biomass yield per metre rope

Fresh biomass Dry biomass

Source d MS F P MS F P f

Nursery 2 5552 1.26 0.363 1015 2.61 0.106

Day 2 7760 26.58 < 0.001 No test

Nursery x Day 4 4423 15.15 < 0.001 No test

Table 3: PERMANOVA analysis on Bray-Curtis distances testing the effects of seeding density (Density, fixed factor) on the (a) fresh (g FW m^"1 rope) and (b) dry biomass yield (g DW m^"1 rope) of seeded ropes at each nursery period in an unreplicated blocked design (Tank: blocked factor) after 21 days outdoor cultivation. Pseudo-F (F) and P values are presented, significant terms shown in bold.

Biomass yield per metre rope

1 day nursery 5 days nursery 10 days nursery

Source df F P F P F P

(a) Fresh biomass

Density 4 2.35 0.053 2.64 0.086 3.00 0.075

Tank 2 5.91 0.003 0.67 0.576 1.24 0.341

(b) Dry biomass

Density 4 2.77 0.030 2.16 0.130 4.01 0.030

Tank 2 4.85 0.006 0.41 0.757 0.60 0.620

Table 4: PERMANOVA analysis on Bray-Curtis distances testing the effects of days of outdoor cultivation (Day, fixed factor) and batch (Batch; random factor) on the biomass yield per metre rope (Fresh biomass; Dry biomass), FW:DW ratios (FW:DW), width of filaments (Width), ash content (Ash) and specific growth rate (SGR) of Ulva. Pseudo-F (F) and P values are presented.

Dry biomass FW: DW Width Ash

yield

Source df F P F P F P F P

Day 6 5.5 2.6 16.69 < 3.3 0.025

0.001 0.650 0.001

Batch 2 34.2 < 51.1 < 51.20 < 39.42 <

0.001 0.001 0.001 0.001

Day x 12 5.0 < 6.1 < 18.75 < 2.5 0.006

Batch 0.001 0.001 0.001 SGR

Source df F P

Day^" 5 7.47 0.005

Batch 2 0.19 0.828

Day x 10 No test

Batch

Table 5: PERMANOVA analysis on Bray-Curtis distances testing the effects of (a) multiple harvests (Harvest, fixed factor) and batch (Batch, random factor) on the dry biomass yield of Ulva sp. 3 cultivated in outdoor cultivation and (b) days of outdoor cultivation (Day, fixed factor) and batch (Batch, random factor) on the specific growth rate. Degrees of freedom (df), pseudo-F (F) and p values (p) are presented, significant terms shown in bold.

Source df F p

(a) Biomass yield

Harvest 2 3.43 0.140

Batch 2 96.67 < 0.001

Harvest x Batch 4 22.17 < 0.001

(b) Specific growth rate

Day 4 5.65 0.018

Batch 2 17.05 < 0.001

Day x Batch 8 11.31 < 0.001

Table 6: Biochemical analysis of Ulva sp. 3. Data are mean (± S.E.; n = 2, batch 1, 3) values (weight % on a dry basis) for the first and second harvest. Carbohydrate content was determined by difference. Protein content equals total amino acid contents (sum of all analysed amino acids).

Harvest 1 Harvest 2

Carbohydrate 46.62 ± 1.8 46.26 ± 1.4

Lipid 1.7 ± 0.3 1.5 ± 0.2

Protein 16.8 ± 0.2 21.3 ± 1.1

REFERENCES

Alveal K, Romo H, Werlinger C, Oliveira EC. 1997. Mass cultivation of the agar- producing alga Gracilaria chilensis (Rhodophyta) from spores. Aquaculture 148:77- 83.

Alves A, Pinho ED, Neves NM, Sousa RA, Reis RL. 2012. Processing ulvan into 2D structures: cross-linked ulvan membranes as new biomaterials for drug delivery applications. Int. J. Pharm. 426:76-81.

Alves A, Sousa RA, Reis RL. 2013a. Processing of degradable ulvan 3D porous structures for biomedical applications. J. Biomed. Mater. Res. Part A 101 A: 998- 1006.

Alves A, Sousa RA, Reis RL. 2013b. A practical perspective on ulvan extracted from green algae. J. Appl. Phycol. 25:407-424.

Anderson MJ, Gorley R, Clarke K. 2008. PERMANOVA + for PRIMER: Guide to software and statistical methods. Plymouth, UK: Primer-E. 214 p.

Bird MI, Wurster CM, De Paula Silva PH, Paul NA, de Nys R. 2012. Algal biochar: effects and applications. GCB Bioenergy 4:61-69.

Bobin-Dubigeon C, Lahaye M, Guillon F, Barr J-L, Gallant DJ. 1997. Factors limiting the biodegradation of Ulva sp cell-wall polysaccharides. J. Sci. Food Agric. 75:341-351.

Callow ME, Callow JA, Pickett-Heaps JD, Wetherbee R. 1997. Primary adhesion of Enteromorpha (Chlorophyta, Ulvales) propagules: quantitative settlement studies and video microscopy. J. Phycol. 33 :938-947.

Carl C, de Nys R, Lawton RJ, Paul NA. 2014. Methods for the induction of reproduction in a tropical species of filamentous Ulva. PLoS One:9(5):e97396.

Carl C, de Nys R, Paul NA. 2014. The seeding and cultivation of a tropical species of filamentous Ulva for algal biomass production. PLoS One 9(6):e98700.

Clarke K, Gorley R. 2006. PRIMER v6: User manual/tutorial. Plymouth, UK:

Primer-E. 190 p. Corradi MG, Gorbi G, Zanni C. 2006. Hypoxia and sulphide influence gamete production in Ulva sp. Aquat Bot 84: 144-150.

Cox S, Abu-Ghannam N. 2013. Enhancement of the phytochemical and fibre content of beef patties with Himanthalia elongata seaweed. Int. J. Food Sci. Technol.

48:2239-2249. Cruz-Suarez LE, Leon A, Pena-Rodriguez A, Rodriguez-Pena G, Moll B, Ricque- Marie D. 2010. Shrimp/ Ulva co-culture: A sustainable alternative to diminish the need for artificial feed and improve shrimp quality. Aquaculture 301 :64-68. Dan A, Hiraoka M, Ohno M, Critchley AT. 2002. Observations on the effect of salinity and photon fluence rate on the induction of sporulation and rhizoid formation in the green alga Enteromorpha prolifera (Miiller) J. Agardh (Chlorophyta, Ulvales). Fish. Sci. 68: 1182-1188. Demetropoulos CL, Langdon CJ. 2004. Enhanced production of Pacific dulse (Palmaria mollis) for co-culture with abalone in a land-based system: effects of stocking density, light, salinity, and temperature. Aquaculture 235:471-488. de Paula Silva PH, McBride S, de Nys R, Paul NA. 2008. Integrating filamentous "green tide" algae into tropical pond-based aquaculture. Aquaculture 284:74-80. de Souza Araujo DF, da Silva AMRB, de Andrade Lima LL, da Silva Vasconcelos MA, Andrade SAC, Asfora Sarubbo L. 2014. The concentration of minerals and physicochemical contaminants in conventional and organic vegetables. Food Control 44:242-248.

Dring M, Edwards M, Watson L. 2013. Development and demonstration of viable hatchery and ongrowing methodologies for seaweed species with identified commercial potential. 31 p.

Du S, Neiman A, Batis C, Wang H, Zhang B, Zhang J, Popkin BM. 2014.

Understanding the patterns and trends of sodium intake, potassium intake, and sodium to potassium ratio and their effect on hypertension in China. Am. J. Clin. Nutr. 99:334-343.

Elleuch M, Bedigian D, Roiseux O, Besbes S, Blecker C, Attia H. 2011. Dietary fibre and fibre-rich by-products of food processing: Characterisation, technological functionality and commercial applications: A review. Food Chem. 124:411-421.

Englyst FIN, Hudson GJ. 1996. The classification and measurement of dietary carbohydrates. Food Chem. 57: 15-21.

Folch J, Lees M, Stanley GHS. 1957. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chemsitry 226:497-509.

Forbord S, Skjermo J, Arff J, Handa A, Reitan KI, Bjerregaard R, Liining K. 2012. Development of Saccharina latissima (Phaeophyceae) kelp hatcheries with year- round production of zoospores and juvenile sporophytes on culture ropes for kelp aquaculture. J. Appl. Phycol. 24:393-399.

Gosch BJ, Magnusson M, Paul NA, de Nys R. 2012. Total lipid and fatty acid composition of seaweeds for the selection of species for oil-based biofuel and bioproducts. GCB Bioenergy 4:919-930. Gupta S, Abu-Ghannam N. 2011. Recent developments in the application of seaweeds or seaweed extracts as a means for enhancing the safety and quality attributes of foods. Innov. Food Sci. Emerg. Technol. 12:600-609.

Handa A, Forbord S, Wang X, Broch OJ, Dahle SW, St0rseth TR, Reitan KI, Olsen Y, Skjermo J. 2013. Seasonal- and depth-dependent growth of cultivated kelp (Saccharina latissimd) in close proximity to salmon (Salmo solar) aquaculture in Norway. Aquaculture 414-415: 191-201.

Hanif R, Iqbal Z, Iqbal M, Hanif S, Rasheed M. 2006. Use of vegetables as nutritional food: role in human health. J. Agric. Biol. Sci. 1 : 18-22. Hebert PDN, Ratnasingham S, de Waard JR. 2003. Barcoding animal life:

cytochrome c oxidase subunit 1 divergences among closely related species. Proc Biol Sci. 270:S96-S99.

Hiraoka M, Enomoto S. 1998. The induction of reproductive cell formation of Ulva pertusa Kjellman (Ulvales, Ulvophyceae). Phycol Res 46: 199-203. Hiraoka M, Shimada S, Ohno M, Serisawa Y. 2003. Asexual life history by quadriflagellate swarmers of Ulva spinulosa (Ulvales, Ulvophyceae). Phycol. Res. 51 :29-34.

Hiraoka M, Shimada S, Uenosono M, Masuda M. 2003. A new green-tide-forming alga, Ulva ohnoi Hiraoka et Shimada sp. nov. (Ulvales, Ulvophyceae) from Japan. Phycol. Res. 51 : 17-29.

Holdt SL, Kraan S. 2011. Bioactive compounds in seaweed: functional food applications and legislation. J. Appl. Phycol. 23 :543-597.

Hurd CL. 2000. Water motion, marine macroalgal physiology, and production. J. Phycol. 36:453-472. Hwang EK, Park CS, Baek JM. 2006. Artificial seed production and cultivation of the edible brown alga, Sargassum fulvellum (Turner) C. Agardh: Developing a new species for seaweed cultivation in Korea. J. Appl. Phycol. 18:251-257.

Hwang EK, Gong YG, Park CS. 2010. Cultivation of a brown alga, Undariopsis peterseniana (Kjellman) Miyabe and Okamura, as a warm-temperature species by artificial seed production in Korea. J. Appl. Phycol. 23 :449-455.

Ichihara K, Miyaji K, Shimada S. 2013. Comparing the low-salinity tolerance of Ulva species distributed in different environments. Phycol. Res. 61 :52-57.

Imchen T. 2012. Recruitment potential of a green alga Ulvaflexuosa Wulfen dark preserved zoospore and its development. PLoS One 7(3): e32651. Kida W. 1990. Culture of seaweeds Monostroma. Mar. Behav. Physiol. 16: 109-131.

Kim JK, Kraemer GP, Neefus CD, Chung IK, Yarish C. 2007. Effects of temperature and ammonium on growth, pigment production and nitrogen uptake by four species of Porphyra (Bangiales, Rhodophyta) native to the New England coast. J. Appl. Phycol. 19:431-440.

Kregting LT, Hurd CL, Pilditch CA, Stevens CL. 2008. The relative importance of water motion on nitrogen uptake by the subtidal macroalga Adamsiella chauvinii (Rhodophyta) in winter and summer. J. Phycol. 44:320-330. Kuda T, Ikemori T. 2009. Minerals, polysaccharides and antioxidant properties of aqueous solutions obtained from macroalgal beach-casts in the Noto Peninsula, Ishikawa, Japan. Food Chem. 112:575-581.

Kumar M, Kumari P, Trivedi N, Shukla MK, Gupta V, Reddy CRK, Jha B. 2011. Minerals, PUFAs and antioxidant properties of some tropical seaweeds from

Saurashtra coast of India. J. Appl. Phycol. 23 :797-810.

Kuwano K, Hashioka T, Nishihara GN, lima M. 2012. Durations of gamete motility and conjugation ability of Ulva compressa (Ulvophyceae). J Phycol 48: 394-400.

Lahaye M, Robic A. 2007. Structure and functional properties of ulvan, a

polysaccharide from green seaweeds. Biomacromolecules 8: 1765-74.

Lawton RJ, Mata L, de Nys R, Paul NA. 2013. Algal bioremediation of waste waters from land-based aquaculture using Ulva: selecting target species and strains. PLoS One 8(10):e77344.

Lee SM, Lewis J, Buss DH, Holcombe GD, Lawrance PR. 1994. Iodine in British foods and diets. Br. J. Nutr. 72:435-446.

Liining K, Pang S. 2003. Mass cultivation of seaweeds: current aspects and approaches. J. Appl. Phycol. 15: 115-119.

Mahesh DL, Deosthale YG, Rao BSN. 1992. A sensitive kinetic assay for the determination of iodine in foodstuffs. Food Chem. 43 :51-56. Mantri VA, Singh RP, Bijo AJ, Kumari P, Reddy CRK, et al. 2011. Differential response of varying salinity and temperature on zoospore induction, regeneration and daily growth rate in Ulva fasciata (Chlorophyta, Ulvales). J Appl Phycol 23 : 243- 250.

Marsham S, Scott GW, Tobin ML. 2007. Comparison of nutritive chemistry of a range of temperate seaweeds. Food Chem. 100: 1331-1336.

McArthur DM, Moss BL. 1979. Gametogenesis and gamete structure of

Enteromorpha intestinalis (L.) Link. Br Phycol J 14: 43-57.

McDermid KJ, Stuercke B. 2003. Nutritional composition of edible Hawaiian seaweeds. J. Appl. Phycol. 15:513-524. Moll B, Deikman J. 1995. Enteromorpha clathrata: a potential seawater-irrigated crop. Bioresour. Technol. 52:255-260.

Msuya FE, Kyewalyanga MS, Salum D. 2006. The performance of the seaweed Ulva reticulata as a biofilter in a low-tech, low-cost, gravity generated water flow regime in Zanzibar, Tanzania. Aquaculture 254:284-292.

Neori A, Chopin T, Troell M, Buschmann AH, Kraemer GP, Hailing C, Shpigel M, Yarish C. 2004. Integrated aquaculture: rationale, evolution and state of the art emphasizing seaweed biofiltration in modern mariculture. Aquaculture 231 :361-391. Neori A, Msuya FE, Shauli L, Schuenhoff A, Kopel F, Shpigel M. 2003. A novel three-stage seaweed (Ulva lactuca) biofilter design for integrated mariculture. J. Appl. Phycol. 15:543-553.

Neveux N, Magnusson M, Maschmeyer T, de Nys R, Paul NA. 2014. Comparing the potential production and value of high-energy liquid fuels and protein from marine and freshwater macroalgae. GCB Bioenergy. DOI: 10.1111/gcbb.12171.

Nordby 0. 1977. Optimal conditions for meiotic spore formation in Ulva mutabilis F0yn. Bot Mar 20: 19-28.

Ohno M. 1993. Cultivation of the green algae, Monostroma and Enteromorpha "Aonori." In: Ohno M, Critchley AT, editors. Seaweed cultivation and marine ranching, pp. 7-16.

Ohno M. 2006. Recent developments in the seaweed cultivation and industry in Japan. In: Advances in seaweed cultivation and utilisation in Asia: proceedings of a workshop held in conjunction with the 7^th Asian Fisheries Forum, Penang. Kuala Lumpur: University of Malaya, pp. 1-20.

Ortiz J, Romero N, Robert P, Araya J, Lopez-Hernandez J, Bozzo C, Navarrete E, Osorio A, Rios A. 2006. Dietary fiber, amino acid, fatty acid and tocopherol contents of the edible seaweeds Ulva lactuca and Durvillaea antarctica. Food Chem. 99:98- 104. Peteiro C, Freire O. 2012. Outplanting time and methodologies related to mariculture of the edible kelp Undaria pinnatifida in the Atlantic coast of Spain. J. Appl. Phycol. 24: 1361-1372.

Peteiro C, Freire O. 2013. Biomass yield and morphological features of the seaweed Saccharina latissima cultivated at two different sites in a coastal bay in the Atlantic coast of Spain. J. Appl. Phycol. 25:205-213.

Roberts DA, de Nys R, Paul NA. 2013. The effect of C0₂ on algal growth in industrial waste water for bioenergy and bioremediation applications. PLoS One 8(l l):e816.

Ross AB, Jones JM, Kubacki ML, Bridgeman T. 2008. Classification of macroalgae as fuel and its thermochemical behaviour. Bioresour. Technol. 99:6494-504.

Ruperez P. 2002. Mineral content of edible marine seaweeds. Food Chem. 79:23-26.

Scardino AJ, Guenther J, de Nys R. 2008. Attachment point theory revisited: the fouling response to a microtextured matrix. Biofouling 24: 45-53.

Shimada S, Yokoyama N, Arai S, Hiraoka M. 2008. Phylogeography of the genus Ulva (Ulvophyceae, Chlorophyta), with special reference to the Japanese freshwater and brackish taxa. J. Appl. Phycol. 20:979-989. Soler-Vila A, Coughlan S, Guiry MD, Kraan S. 2009. The red alga Porphyra dioica as a fish-feed ingredient for rainbow trout (Oncorhynchus mykiss): effects on growth, feed efficiency, and carcass composition. J. Appl. Phycol. 21 :617-624.

Steen H, Scrosati R. 2004. Intraspecific competition in Fucus serratus and F.

evanescens (Phaeophyceae: Fucales) germlings: effects of settlement density, nutrient concentration, and temperature. Mar. Biol. 144:61-70.

Suter PM, Sierro C, Vetter W. 2002. Nutritional factors in the control of blood pressure and hypertension. Nutr. Clin. Care 5:9-19.

Suttle NF. 2010. Mineral nutrition of livestock, 4^th edition. Wallingford, UK: CABI. 579 p.

Tan SP, McLoughlin P, O' Sullivan L, Prieto ML, Gardiner GE, Lawlor PG, Hughes H. 2013. Development of a novel antimicrobial seaweed extract-based hydrogel wound dressing. Int. J. Pharm. 456: 10-20.

Teas J, Pino S, Critchley A, Braverman LE. 2004. Variability of iodine content in common commercially available edible seaweeds. Thyroid Off. J. Am. Thyroid Assoc. 14:836-841. van der Wal H, Sperber BL, Houweling-Tan B, Bakker RR, Brandenburg W, Lopez- Contreras AM. 2013. Production of acetone, butanol, and ethanol from biomass of the green seaweed Ulva lactuca. Bioresour. Technol. 128:431-7. Wichard T, Oertel W. 2010. Gametogenesis and gamete release of Ulva mutabilis and Ulva lactuca (Chlorophyta): regulatory effects and chemical characterization of the "swarming inhibitor." J. Phycol. 46: 248-259.

Wong KH, Cheung PCK. 2000. Nutritional evaluation of some subtropical red and green seaweeds. Part I - proximate composition, amino acid profiles and some physico-chemical properties. Food Chem. 71 :475-482.

Yokoyama H, Ishihi Y. 2010. Bioindicator and biofilter function of Ulva spp.

(Chlorophyta) for dissolved inorganic nitrogen discharged from a coastal fish farm — potential role in integrated multi-trophic aquaculture. Aquaculture 310:74-83.

Zhang C, Li X, Kim S. 2012. Application of marine biomaterials for nutraceuticals and functional foods. Food Sci. Biotechnol. 21 :625-631.

Zhang QS, Tang YZ, Liu SK, Zhang SB, Lu ZC, Cu SH, Yu YQ. 2012. Zygote- derived seedling production of Sargassum thunbergii: focus on two frequently experienced constraints in tank culture of seaweed. J. Appl. Phycol. 24:707-714.

Zhuang Y, Guo J, Chen L, Li D, Liu J, Ye N. 2012. Microwave-assisted direct liquefaction of Ulva prolifera for bio-oil production by acid catalysis. Bioresour. Technol. 116: 133-139. Zubia M, Robledo D, Freile-Pelegrin Y. 2007. Antioxidant activities in tropical marine macroalgae from the Yucatan Peninsula, Mexico. J. Appl. Phycol. 19:449- 458.

Claims

CLAIMS:

1. A method of cultivating green macroalgae, the method including;

2. The method according to claim 1, wherein the temperature shock is a cold shock.

3. The method according to claim 2, wherein the macroalgae are exposed to a cold shock of less than about 10°C.

4. The method according to claim 1, wherein the temperature shock is a heat shock.

5. The method according to claim 4, wherein macroalgae are exposed to a heat shock of about 15 to 35°C.

6. The method according to any one of claims 1 to 5, wherein the macroalgae is exposed to the temperature shock for a period of about 10 to 20 minutes.

7. The method according to any one of claims 1 to 6, further comprising fragmenting the temperature shocked macroalgae before maintaining the macroalgae under suitable conditions.

8. The method according to any one of claims 1 to 7, wherein the temperature shocked macroalgae are maintained under suitable conditions for about 24 to 72 hours.

9. The method according to any one of claims 1 to 8, wherein the temperature shocked macroalgae are maintained at a temperature from about 24 to 26°C.

10. The method according to any one of claims 1 to 9, wherein the temperature shocked macroalgae are maintained for about 12 hours in the light and about 12 hours in the dark for every 24 hour period.

11. The method according to any one of claims 1 to 10, wherein the macroalgae comprise filamentous and/or blade leaf species.

12. The method according to any one of claims 1 to 11, wherein the macroalgae is of the genus Ulva.

13. The method according to claim 12, wherein the Ulva species are selected from one or more of the group consisting of: Ulva arasakii, Ulva armoricana, Ulva australis, Ulva bulbosa, Ulva californica, Ulva clathrata, Ulva clathratioides, Ulva compressa, Ulva enteromorpha, Ulva fasciata, Ulva fenestrate, Ulva flexuosa, Ulva geminoidea, Ulva intestinalis, Ulva intestinaloides, Ulva laetevirens, Ulva lactuca, Ulva latissima, Ulva linza, Ulva lobata, Ulva muscoides, Ulva ohnoi, Ulva palmata, Ulva pertusa, Ulva procera, Ulva prolifera, Ulva proliferoides, Ulva pseudocurvata, Ulva purpurea, Ulva rigida, Ulva scandinavica, Ulva spinulosa, Ulva stenophylla, Ulva stenophylloides, Ulva stipitata, Ulva taeniata, Ulva tanneri, Ulva torta, Ulva sp. 3, Ulva sp. 2, Ulva sp. 4 and Ulva umbilicalia.

14. The method according to claim 13, wherein the green macroalgae is of the species Ulva sp. 3.

15. A method of producing green macroalgae, the method including:

producing reproductive bodies according to the method of any one of claims 1 to 14;

seeding the reproductive bodies;

settling the reproductive bodies; and

growing the reproductive bodies to produce macroalgae.

16. The method according to claim 15, wherein the reproductive bodies are seeded on to a growth structure.

17. The method according to claim 16, wherein the growth structure is selected from the group comprising: rope, net, sponge, balls and floats.

18. The method according any one of claims 15 to 17, wherein the reproductive bodies are seeded on to a growth surface at a density of about 466,000 to 1,552,000 per metre of rope.

19. The method according to any one of claims 15 to 17, wherein the reproductive bodies are seeded directly into an aqueous solution.

20. The method according to claim 19, wherein the reproductive bodies are seeded at a density of about 10,000 to 20,000 reproductive bodies per mL.

21. The method according to any one of claims 15 to 20, wherein the reproductive bodies are allowed to settle for about 4-5 days.

22. The method according to any one of claims 15 to 21, wherein the reproductive bodies are allowed to settle at a temperature of about 24 to 26°C.

23. The method according to any one of claims 15 to 22, wherein the reproductive bodies are settled for about 12 hours in the light and about 12 hours in the dark.

24. The method according to any one of claims 15 to 23, wherein the settled reproductive bodies are grown for a period of about 10 to 16 days.

25. The method of any one of claims 15 to 24, wherein the reproductive bodies are grown in an open or closed system.

26. The method according to any one of claims 15 to 25, further comprising harvesting the grown green macroalgae.

27. The method according to claim 26, wherein the macroalgae is harvested about 18 to 25 days after exposing the macroalgae to a temperature shock.

28. The method according to claim 26 or claim 27, wherein about 50 to 90% of the weight of the algae is harvested.

29. The method of claim 28, wherein no more than two (2) sequential harvests are performed.

30. A method of producing macroalgae according to any one of claims 15 to 29 for the production of biomass, wastewater treatment, human consumption, and/or feedstock for the supply of pharmaceuticals, nutraceuticals, cosmeceuticals and/or biofuels.

31. Green macroalgae produced according to the method of any one of claims 15 to 30.