There are two primary cultivation systems that are currently used to grow microal­gae: (1) open-air system and (2) photobioreactors [4] . Table 2 compares the two cultivation systems in terms of important cultivation parameters. The selection of cultivation systems depends on several factors: the microalgal species, the avail­ability of sunlight and water, the cost of land, the type of desired final product, and

the supply of CO2 [4, 53]. The amount of nutrients and certain metals (i. e., iron and magnesium) in the cultivation water is also important as it directly affects the growth rate and the CO2 fixation efficiency of microalgal biomass. Research from an out­door mass culture in New Mexico (US) shows that CO2 fixation by microalgae increases with optimal nutrient level [58]. Susceptibility of the microalgal species to

contamination by other microorganisms also needs to be considered when selecting cultivation systems. For example, Tetraselmis, Skeletonem, and Isochrysis are easily invaded by other local microorganisms and, as a result, are preferably cultivated in a closed photobioreactor system [4].

There are four different types of open-air systems: shallow big ponds, tanks, circular ponds, and raceway ponds [4]. Even though open-air systems are less com­plex to construct and cheaper to operate than photobioreactors, they suffer from numerous disadvantages, such as slower CO2 diffusion, requirement for a large land area, substantial evaporative losses, poor light utilization, and difficulty in control­ling cultivation conditions [55]. The depth of the open system also needs to be care­fully calculated. The pond needs to be shallow enough for the penetrating sunlight to reach all the microalgal cells yet, at the same time, deep enough for sufficient mixing of the culture to occur. Maximum culture concentration that can be achieved in an open system varies between 0.1 and 0.5 g dried microalgae/L [4].

Among the open-air systems, the most common design is the raceway pond (Fig. 3) [56]. The raceway pond is made up of a closed loop recirculation channel with a paddle wheel that provides circulation and mixing. Baffles are also placed in the channel to guide the culture flow. To ensure uniform light penetration, the race­way pond is only 0.3 m in depth. The ponds are made from concrete or compacted earth that can be lined with plastic [9]. Companies, such as New Zealand’s Aquaflow Bionomics and the US’ Live Fuels Inc., have been using the raceway ponds to cul­tivate their microalgae.

Since photobioreactors have a “closed” design which allows users to tightly con­trol cultivation conditions (temperature, degree of mixing, degree of illumination) as well as level of sterility, they are able to achieve significantly higher biomass pro­ductivity and CO2 fixation rate for most microalgae than raceway ponds. There are

four different designs of photobioreactors: flat-plate, tubular, bubble-sparged verti­cal column, and airlift vertical column. Of these designs, tubular photobioreactors are the most popular [55]. Tubular photobioreactors are composed of an array of clear straight tubes made of thin glass or plastic (usually less than 0.1 m in diameter) to allow for maximum light penetration. The tubes are arranged either horizontally where they are placed parallel to each other flat on the ground or vertically where they are configured to form fence-like structures. Figure 4 shows the two different configurations of tubular photobioreactors [9].

Apart from the four primary designs, there are other types of photobioreactor systems that are currently used. The basics of the systems are the same with some modifications to maximize its efficiency. One such example is a 450-ft-long by 50-ft-wide photobioreactor made up of twin transparent plastic algal waterbeds pat­ented by a company called A2BE Carbon Capture LLC. Another company, Green Shift Corporation based in New York, has produced a pilot-scale photobioreactor that is incorporated with an ethanol-producing facility to capture the CO2 emitted from power plants [33] .

Algae contain large amounts of polysaccharides, notably cell wall structural poly­saccharides that are extruded by the hydrocolloid industry: alginate from brown algae, carrageenans, and agar from red algae. Edible algae contain 33-50% total fibers, which is higher than the levels found in higher plants. Other minor polysac­charides are found in the cell wall: fucoidans (from brown algae), xylans (from certain red and green algae), ulvans (from green algae), and cellulose (which occur in all genera, but at lower levels than found in higher plants). Algae also contain storage polysaccharides, notably laminarin (b-1,3 glucan) in brown algae and floridean starch (amylopectin-like glucan) in red algae ] 16]. Most of these polysaccharides are not digested by humans and can be regarded as dietary fibers.

Water soluble and water insoluble fibers have different physiological effects associated. Insoluble fiber primarily promotes the movement of material through the digestive system, thereby improving laxation. Therefore, insoluble fiber can increase feelings of satiety [178]. The majority of insoluble fiber is fermented in the large intestine, supporting the growth of intestinal microflora, including probiotic species. Soluble fiber can help to lower blood cholesterol and regulate blood glucose levels [190]. The insoluble fibers include cellulose, hemicellulose, and lignin; the soluble fibers include the oligosaccharides, pectins, b-glucans, and galactomanan gums.

Table 3 shows, for comparison, the dietary fiber content in some sea vegetables, seaweed by-products and plants [95]. As can be seen, algae contain slightly more fiber than cabbage, although the amounts consumed in the diet would be lower. The red alga Kappaphycus shows the highest levels of total fiber (70.7% dry weight).

Algae contain sulfated polysaccharides which possesses important functional properties. For instance, fucoidans (soluble fiber), polysaccharides containing sub­stantial percentages of L-fucose and sulfate ester groups, are constituents of brown algae. For the past decade, fucoidans isolated from different brown algae have been extensively studied due to their varied biological activities, including anticoagulant and antithrombotic, antiviral, antitumoral and immunomodulatory, anti-inflammatory, blood lipids reducing, antioxidant and anticomplementary properties, activity against hepatopathy, uropathy, and renalpathy, gastric protective effects, and therapeutic potential in surgery [94]. Compared to other sulfated polysaccharides, fucoidans are widely available from various kinds of cheap sources, so more and more fucoidans have been investigated in recent years as natural sources of drugs or functional ingre­dients. Fucoidans had been isolated of different brown algae, such as, U. pinnatifida [92], Laminaria angustata [83], Sargassum stenophyllum [29], H. fusiforme [93], Adenocytis utricularis [146], and Cystoseira canariensis [149].

Red algae contain water soluble sulfated polysaccharide galactan, agar, and car­rageenans. One of the most studied marine-sulfated homopolysaccharides class, together with fucoidans, are the sulfated galactans. In general, the sulfated galactans are polymers of a-L — and a-D — or b-D-galactopyranosyl units. Unrelated to their natural biological roles as components of the biological wall, the sulfated galactans show important and potent pharmacological actions. These include antiviral, antitu­moral, immunomodulation, antiangiogenic, anti-inflammatory, anticoagulant, and antithrombotic properties [144]. Their beneficial effects on the cardiovascular sys­tem are the most studied and exploited clinical actions, especially due to the serious need for new antithrombotic drugs as a consequence of the continuously increasing incidence of thromboembolic diseases [ 145]. Sulfated galactans have been identified in several red algae, among others, Grateloupia elliptica, Sinkoraena lancifolia, Halymenia dilatata, Grateloupia lanceolata, Lomentaria catenata, Martensia den- ticulata, Schizymenia dubyi, and C. crispus [91].

Agar extracted from species, such as Gracilaria and Gelidium, is composed of a mixture of the sulfated galactans D — galactose and 3,6-anhydro-a-L-lactose. The term agarose and agaropectin represent an oversimplification of the agar structure.

Carrageenan is a generic name for a group of linear-sulfated galactans, obtained by extraction from numerous species of marine red algae. These carbohydrates consist of a linear structure of alternating disaccharide repeat units containing 3-linked P-d- galactopyranose and 5-linked a-D-galactopyranose.

Porphyrans, the sulfated polysaccharides making up the hot-water soluble por­tion of the cell wall, are the main components of Porphyra. Structurally, they have a linear backbone of alternating 3-linked b-D-galactosyl units and 4-linked a-L — galactosyl 6-sulfate or 3,6-anhydro-a-L-galactosyl units. In a former study, the con­tent of ester sulfate in porphyran extracted from Porphyra haitanensis was measured ranging from 16 to 19% and showing generic antioxidant activity [202]. Several investigations of the structure and function of porphyrans isolated from different species have been undertaken [122, 201]. Although the chemical components and structures show great variation, porphyrans have also been shown to have immuno — regulatory and antitumor activities [128, 135].

On the other hand, green algae, such as those of the genera Ulva and Enteromorpha, contain sulfated heteropolysaccharides in their mucilaginous matrix [72]. Sulfated polysaccharides extracted from the green algae belonging to Ulvales (Ulva and Enteromorpha) are ulvan. Ulvan is a heteropolysaccharide, mainly composed of rhamnose, xylose, glucose, glucuronic acid, iduronic acid, and sulfate, with smaller amounts of mannose, arabinose, and galactose. The mainly repeating disaccharide units are (b-D-Glcp A-(1 ^ 4)-a-L-Rhap 3S) and (a-L-ldop A-(1 ^ 4)-a-L-Rhap 3S) [139]. Most of the recent work on Ulvales cell wall polysaccharides focused on ulvan as it display several physicochemical and biological features of potential interest for food, pharmaceutical, agricultural, and chemical applications. Ulvans have been shown to have antioxidant [148], antitumor [ 104], and antihyperlipidemic [139] activities.

In general, biological production of methane from algae or cyanobacteria is a two — step process. The first step is biomass production or capturing and conversion of sun light energy into new algal cells. The second step is a transformation of energy stored as biomass into a more applicable form, such as methane gas, through the ADP. Methane is easily stored, transported, and used for the production of heat or electricity. Methane can also be used as a motor fuel. The efficiency of methane production from sunlight energy relies on the performance of these coupled steps.

5.1 Algal Biomass Improvement

The performance of the biomass production step can be described by productivity per acre but the algal methane potential is controlled by algal biochemical composi­tion. In this section, we review factors that control and limit algal productivity and methane potential. We also describe methods used for the improvement of methane production from algae.

Fertilizers and inorganic chemicals are the major costs associated with intensive algal production systems. As example, producing 1 kg of biodiesel in fresh water requires 3,726 kg water, 0.33 kg nitrogen, and 0.71 kg phosphate [449]. On the other hand, algae and cyanobacteria play a role in self-purification of water bodies and in wastewater treatment by direct assimilation of simple organic compounds [294,450] and nutrients [451-456], removal ofheavy metals [388,453,457-459], and finally providing oxygen for organic matter oxidation by heterotrophic bacteria. Wastewater stabilization ponds with naturally occurring algal flora are widely used in developing countries and local sewage systems [460, 461]. Closed coastal areas, such as bays, fjords, and lagoons near urban and agricultural runoffs. are potential systems for cultivation, harvesting, and utilization of macroalgae for biomethane production [177, 412]. Golueke and Oswald first suggested the combination of wastewater treatment with production of algae to yield a biofuel [109, 110, 462, 463]. Coupling the treatment of nutrient-rich wastewater with algal growth followed by conversion to methane represents a potentially attractive biofuel production pro­cess with reduced impact on the environment [464-468]. Moreover, mixotrophic microalgal growth is attractive due to induction of lipids accumulation in algal cells.

Of the three main dissociation methods, depressurization appears best suited to the conditions of Class 1 deposits because of its simplicity, technical, and economic effectiveness, and the fast response of hydrates to the rapidly propagating pressure wave [129, 125]. Hong and Pooladi-Darvish [68] applied constant-P depressuriza­tion at a well at the center of a GH reservoir, and analyzed the sensitivity of the continuously declining production to various properties and operational conditions. They reported that, at the end of the first year of production, about 48% of the pro­duced gas had been replenished by hydrate-originated CH4 , thus confirming the technical feasibility of production from hydrates using conventional technology.

Moridis et al. [129] conducted a long-term (10-30 years) study of constant-rate (=0.81944 ST m3/s = 2.5 MMSCFD) production from Class 1 hydrate deposits. To describe the contribution of gas released from hydrate to the production stream, they introduced the concepts of Rate Replenishment Ratio (RRR) and Volume Replenishment Ratio (VRR). RRR is defined as the fraction of the gas production rate at the well(s) that is replenished by CH4 released from hydrate dissociation. VRR is defined as the fraction of the cumulative gas volume produced at the well(s) that is replenished by hydrate-originating CH4. During the 30-year production period, the VRR increases continuously to a maximum of about 0.74, and the corresponding RRR is 0.54 (Fig. 11 — [129]). The desirability and the great production potential of such deposits are obvious. The evolution of the SH distribution over time is shown in Fig. 12. Production from these Class 1 deposits (a) involved conventional technolo­gies and (b) necessitated continuous heating of the wellbore to prevent hydrate for­mation and plugging [129]. Note that the use of horizontal wells does not confer any practical advantages to gas production from Class 1 deposits [137].

Because there are currently no unconventional developments, oil or gas, in the fron­tier areas where hydrates occur, and because these areas also contain significant amounts of developed and undeveloped conventional gas resources with no access to markets, GHs will have to compete with frontier conventional gas developments. This puts GHs at a distinct disadvantage compared to other unconventional gas resources (such as the booming shale-gas production in the United States) for access to the larger North American gas market. While a local market use of gas from gas hydrates may be feasible at some point, this situation appears likely to defer the tim­ing of GH developments until sometime in the not-too-near future. Offshore GH developments may proceed sooner if the premium price required is not onerous when there is no conventional gas competition, and where security of supply is a major consideration.

The studies of Hancock et al. [59], Hancock [58], and the review study of Moridis et al. [127] were the first in-depth analyses of the economics of gas production from hydrates, and their results, while preliminary, have been encouraging. Assuming 2009 prices, it appears that (a) for onshore gas hydrates, stand-alone developments could be economic with a gas price in the upper range of historical North American prices, and (b) for offshore gas hydrates, stand-alone developments could be eco­nomic with a gas price in the upper range of what India has been paying for liquefied natural gas imports on the spot market. Thus, using the admittedly limited data from the numerical predictions of gas production in the literature [58, 59], it appears that a reasonable rate of return (i. e., 15%) can be achieved with prices in the order of $6.00-$12.00/MSCF for offshore and onshore projects, respectively (Figs. 18 and 19). However, considering the various risks and uncertainties associated with such production (well performance, geological uncertainty, reservoir characteristics,

gas-in-place, thermodynamic conditions, the absence of a long-term field test of production from hydrates, proximity to infrastructure, access to markets, uncer­tainty in forecasting gas prices, etc.), sustained gas contract prices in the range of $10.00-$ 16.00/MSCF for offshore and onshore projects respectively may be required before GH projects will proceed [127]. Given the current gas price reali­ties, it appears that production from GH may be delayed, although unique circum­stances may allow production of onshore gas hydrates for local community or industrial use, especially where there is some underlying gas. Fundamental changes in the North American gas market supply picture, as well as advances in technology may also have a significant impact on the price range required for GH development, and will inevitably affect the timing of commercial gas production from HBS.

The strain that is being used for metabolic engineering is R. eutropha H16 (cur­rently classified as Cuparividus necator), which is a facultative chemolithoauto — troph that has been extensively studied for both PHB production and chemolithoautotrophic metabolism [16]. The genome of R. eutropha H16 has been sequenced, demonstrating that it consists of three circular replicons: two chromo­somes (4.0 and 2.9 MB) and a megaplasmid (0.45 MB), termed pHG1 [15]. R. eutro­pha H16 is capable of growing either heterotrophically on reduced carbon substrates under aerobic conditions or growing autotrophically under an atmosphere of H2/CO2/O2.

A critical element in the success of this project will be the development of genetic tools for metabolic engineering of R. eutropha. Engineering of R. eutropha will entail successful implementation of the following elements: (1) heterologous DNA delivery; (2) compatible plasmid system for rapid construction and testing of heterologous pathways; (3) chromosomal manipulation comprising insertions and knockouts, and (4) deployment of tunable parts such as promoters, ribosomal bind­ing sites, spacers, and terminators for precise control of heterologous genetic cir­cuits. To ensure expression compatibility, we will perform in silico analyses of codon usage and transcript stability of all heterologous genes and parts described in this project and if necessary, commercially synthesize optimized sequences.

1.2 Solvent-Based Extraction Systems

The yields of linear olefins obtained through traditional organic solvents extraction (n-hexane, chloroform, and methanol) and SPS-based systems (DBU/octanol and DBU/ethanol) are compared in Fig. 16.

The SPS DBU/octanol is the best solvent in the extraction of freeze-dried sam­ples (16 ± 2% total hydrocarbons yield on dry weight basis), followed by the SPS DBU/ethanol (12 ± 2% yield). The two SPS are both better than the extraction sys­tem based on traditional organic solvents that gives the lowest yields (7.8 ± 3%) after an extraction process performed at the same temperature and after the same time interval (60°C and 4 h).

Also, in the direct extraction of algal cultures, the SPS DBU/octanol affords bet­ter results than traditional organic solvents, under both the speed and time condi­tions we checked. It is important to underline that the SPS DBU/octanol gives an extractive yield of 8.2 ± 1% after 24 h at 300 rpm, which is slightly higher than that obtainable from freeze-dried sample with traditional organic solvents under reflux (7.8% yield). This is an indication that the extraction process can be improved from energetic, economic, and sustainability points of view, without the need of dewater­ing biomass, consuming energy to evaporate the solvent, and adopting special safety measures to reduce the risk for operators.

The culture velocity within the solar receiver is very important, as the cells must be evenly distributed throughout the tubing to avoid extended periods within the dark zones located at the centre of the tubing. The maximum velocity obtained within the system is dependent on the size of micro-eddies in comparison to the algae cell dimension. Acien Fernandez et al. [1] found that the maximum velocity in an exter­nal loop reactor (ELR) for Phaeodactylum tricornutum strain was 1 m/s. This veloc­ity was obtained by a specific power input of 170 W/m3. However, the actual velocity used in the ELR was 0.5 m/s due to issues associated with the mechanical properties of the solar receiver. The velocity must be high enough to ensure turbulence, thus preventing bio-sedimentation. However, the liquid velocity cannot be applied at gratuitous speeds, as this could potentially cause damage to the algal cells. Generally the liquid velocity must comply with two constraints: a turbulent Reynolds number and a micro eddy length that is significantly larger than the cell dimensions [1].

1.3.3 pH

The pH of the cultivation system increases as the algal cells photosynthesise. The consumption of carbon dioxide and the production of dissolved oxygen from pho­tosynthesis can significantly alter the pH thereby impeding growth. The cultivation system requires a relatively neutral environment, usually maintained at pH ~8 [34]. To prevent variations in culture pH, appropriate control systems are incorporated to monitor the pH. Another technique to control variations in pH is to employ carbon dioxide injection points along the tube run. This prevents excessive culture pH and any carbon limitation that may occur [23]. However, this is not economically viable when considering large algal plants.

1.7.1 Cultivation Economics

The economic model, as shown in Fig. 9, identified the raceway pond as the cheap­est production system ($2.77/kg) followed by the HTR ($9.91/kg). The ELR was the most expensive option ($12.98). The greater complexity of the reactor-style systems was found to require a much greater level of FCI, $2.7 billion for the HTR system and $3.6 billion for the ELR system compared with only $0.73 billion for the raceway ponds. This FCI was represented in annual cost terms as the deprecia­tion of the cultivation system, expressed in Fig. 9, by the black equipment cost por­tion of the graph. Figure 9 shows these higher equipment costs are the major contributors to the greater overall production cost. Furthermore, the magnitude of FCI required to build the reactor-style systems makes investment in these alterna­tives unlikely, at least on such a vast scale.

The running cost of each cultivation system is represented by the grey segment in Fig. 9 and is examined in greater detail in Fig. 10, which divides the costs into specific components. Notably, the major contributors to the annual running costs shown in Fig. 10 were found to vary greatly between the raceway pond and the reactor-style systems. Major contributors to the annual running costs of the raceway

Raceway Pond HTR ELR

E Electricity H Culture Medium H Wastewater Treatment

□ Maintenance DD Other Expenses

Fig. 10 Breakdown of annual running costs for different cultivation systems pond were found to be the culture medium and the wastewater treatment, while in the reactor-style systems’ electricity consumption and maintenance costs were the great­est contributors to running costs. The larger volume of fluid processed in the raceway pond system, due to its lower volumetric productivity, led to greater culture medium and wastewater treatment costs. In contrast, the larger maintenance costs of the HTR and ELR systems resulted from the greater complexity of the system operation.

The considerably larger electricity consumption of the reactor-style systems could be attributed to the use of an airlift pump to mix the culture, which used significantly larger amounts of energy to operate than the simple paddle wheel used

H Capital Costs □ Electricity Costs □ Flocculant Cost

Fig. 11 Biomass dewatering costs for raceway pond (RP) in the raceway pond system. Despite the lower production costs of the raceway pond system, it is necessary to account for the risk of additional costs resulting from con­tamination of the algal culture. This risk is a significant drawback in the use of raceway ponds for cultivation compared to the use of reactor-style systems. Contamination results from a lack of control and exposure to the external environ­ment, and can lead to lower growth rates of biomass unsuitable for downstream.