The physiological requirements of microalgae determine the basic design of a photobioreactor. Phototrophic microorganisms capture sunlight or artificial light and transform light energy into chemical energy in the form of ATP and reduced NADPH which are essential for carbon fixation. A photobioreactor needs to supply cells with sufficient light and at the same time with enough carbon dioxide to build up carbon hydrates for anabolism and storage purpose. The generation of oxygen is stoichiometrically linked with carbon dioxide consumption. Accordingly, excess oxygen needs to be removed from the system. Furthermore, microalgae need inor­ganic nutrients and trace elements for growth. The stoichiometric demand of nitro­gen and phosphorous, for example, can be deduced from the elemental composition of the microalgae.

In terms of hydrodynamics, a photobioreactor represents a three phase system with the liquid system providing the inorganic nutrients which are dissolved in the broth. The gaseous phase supplies carbon dioxide and excess oxygen is removed from the system via gas bubbles. Eventually, the solid phase consists of cells. A fourth interacting component is the superimposed light radiation field.

The major challenge, and the function of a bioreactor, is to provide favorable conditions which allow for high productivities and avoidance of inhibiting or limiting effects. However, considerable gradients of CO2 and O2 can affect growth. Light that impinges on the reactor surface (I0) is absorbed and scattered and thus not all cells in the reactor receive light with the same intensity. Even local differences in concen­trations of inorganic nutrients can occur (Fig. 1). No photobioreactor concept provides optimal mass transfer and light distribution in a manner that the occur­rence of gradients is completely avoided.

Improvements of photobioreactor designs mainly focus on three key areas, namely light transfer, reaction (related to aeration), and hydrodynamics. It is not sufficient to address these aspects all separated from each other but there are significant interferences which mainly influence physiology, growth, and productivity in photobioreactors (Fig. 2). Yet in order to reduce complexity, experimental scale — down approaches try to separate interdependent influences to unravel underlying mechanisms influencing productivity [37].

These key aspects and their interdependency are further addressed in the follow­ing sections.

Sample Collection and Preparation: Both filtration and centrifugation were used to obtain highly concentrated samples of algae from various sources. Algae were col­lected from the effluent of the sewage treatment plant Virginia Initiative Plant (VIP), close to Old Dominion University (ODU), Norfolk, VA, from May, 2007 to February, 2008. The dominant algae at the VIP were Centric diatoms (10-30 mm), Scenedesmus/Desmodesmus spp., and Chlorella sp. Water from the wastewater facil­ity with entrained algae was passed through a tangential flow ultrafiltration system (Millipore Pellicon) with 0.2 mm filter, and the retentate, as condensed algal concen­trate, was collected. Total cell concentration and algae specification for each sample was examined under microscope [21] . To remove salts, the algal concentrate was centrifuged (6,500 RPM, 250 mL rotor, 30 min), and the supernatant was decanted and replaced with MilliQ water. This procedure was repeated twice. The final con­centrate was freeze-dried before chemical treatment. Algae were also collected from the ODU Algae Farm located in Spring Grove, VA. This farm, established in 2008, is an open pond design in which mainly Scenedesmus/Desmodesmus spp. algae are being grown and harvested. Algae at the ODU algal farm were concentrated by use of a continuous flow centrifuge (Lavin Centrifuge Model 12-413 V). The concen­trated algae, collected as a paste, were removed and lyophilized to dryness.

Conversion of Algae to Biodiesel in a Prototype Reactor: The TMAH method was adapted for the conversion of algae to biodiesel in a horizontal tube furnace reactor [16,17,26]. Approximately 1-2 g of algae sample was used in the prototype reactor (Fig. 1) . Algae samples were placed in a 30-mL glass culture tube with 1-2 mL of a TMAH in methanol solution (25 wt%). The methanol was evaporated to dryness under N2 and placed in the Thermo Scientific Lind-berg® Blue M reactor tube oven. The temperature was programmed to increase from 25°C to the desired temperature (250°C up to 550°C) over 15 min. The sample was held at final tem­perature for 45 min and the reactor was cooled to room temperature. An oxygen — free environment was created by using nitrogen gas flowing at a rate of approximately 100 mL/min. Gaseous vapors were collected and condensed in an ice bath. These condensed vapors were analyzed for FAME content by GC-MS.

Condenser

Biodiesel

Collected

Fig. 1 Schematic diagram of the prototype reactor. Reactor is designed for use with TMAH in a batch mode

I dentification by GC-MS: Samples were analyzed by an Agilent 6890 gas chromatograph interfaced to a Leco III mass spectrometer. The temperature was programmed from 50 to 300°C at a rate of 15°C/min. The mass spectrometer was repeatedly scanned in the low-resolution mode from 45 to 500 mass (u) at a rate of 20 spectra/s. Compounds were identified by their mass spectra. Most peaks were identified by comparison with the Wiley/NBS library and some were confirmed by comparison with standards. Quantitative measurements of the concentrations of individual peaks were made using an internal standard л-tetracosane as well as with external fatty acid methyl ester (FAME) standards. FAME standards were GLC-40, 50, and 90 standard mixtures (Supelco Analytical) containing fatty acid mixtures of

C16:0’ C180 C20:0, and C22:0; C16:1’ C18:1 ’ C20:T and C22:1; and C13:0, C150 C17:0, C19:0, and

C210; respectively.

Elemental Carbon (C) and Nitrogen (N) Analysis of Algae Residue: The carbon (C) and nitrogen (N) contents of algae residues were determined by a Thermo Finnigan Flash 1112 Elemental Analyzer using a nicotinamide standard for calibra­tion. Approximately 1-2 mg of each solid sample was placed in a 3.3 x 5-mm tin capsule for combustion. The method used for analysis was a furnace at 900°C, oven at 75°C, and carrier gas helium at 91 mL/min.

Solid-state 13C NMR: The NMR cross-polarization magic angle spinning (CPMAS) 13C experiments, with a recycle delay time of 1.0 s and 2,048 scans, were performed using a Bruker Avance II 400 spectrometer operating at 100 MHz for 13C and 400 MHz for 1H [18]. All the experiments were conducted using a 4-mm triple resonance probe. Dry algae and conversion residues were packed into a 5-mm Zirconia NMR rotor fitted with a Kel-F cap for analysis.

Molina Grima et al. [24] have shown that lipid extraction from P tricornutum can be achieved on wet biomass. The study achieved 90% fatty acid yield on wet bio­mass in comparison to a 96% fatty acid yield from lyophilised biomass. The wet biomass was extracted at approximately 20% (w/v). The wet biomass leaving the dewatering stage from this study is at 50% (w/v). Therefore, the extraction method described by Molina Grima et al. [24] should provide enhanced interaction between the solvent and the cells. However, this extraction method involved three steps: (1) direct saponification of biomass oil, (2) extraction of unsaponifiable lipids and (3) extraction of purified free fatty acids (FFA) [24]; hence, it was compared to a simi­lar method conducted by Ramirez Fajardo et al. [27], which was a two-step solvent extraction process. The two-step method involved lipid extraction using ethanol and lipid purification using a two-phase system of hexane and water. The crude lipid extraction involved a reaction between ethanol (96% v/v) and the biomass in an agitated tank for ~20 h. The crude lipid contains both saponifiable and unsaponifiable lipids. The saponifiable lipids include glycolipids and acylglycerols which are use­ful for biodiesel, whereas unsaponifiable lipids contain components such as amino acids and chlorophyll pigments which must be removed from the extract [27]. Water is added to the ethanol/crude-extract mixture to create a hydroalcoholic solution, and hexane is added to create a biphasic extraction system that separates the exploit­able and unusable lipids thus purifying the extract. The majority of the saponifiable lipid will be found in the hexane phase, whereas the more polar components will remain in the hydroalcoholic phase [ 24] . It is assumed that hexane purification reaches 80% recovery after four continuous extractions.

For biodiesel production, a transesterification process is used to convert the extracted and purified lipids into FAME. The process involves different unit operations. However, the emission sources can be categorised based on either steam or electric­ity usage. Both emissions are Scope 2. For the modelled facility, 1,834 MJ/year of steam and 470,716.3 kWh/year electricity are required. For this study, it was assumed that a natural gas boiler is used to produce the required steam and the emis­sions are calculated based on the amount of natural gas needed to produce the required energy. Net emissions from the production of biodiesel are 668.42 tonnes of CO2-e/year. 85% of the emissions are due to power requirements in running the pumps and the remaining 15% is due to the production of steam. The extraction design chosen has a significant amount of pumping and filtration units, all of which are energy-intensive processes.

Global warming can bring about extreme weather occurrences, rise in sea levels, extinction of species, retreat of glaciers, and many other calamities. The rise in global temperature is attributed to the high amount of carbon dioxide (CO2) gases in the atmosphere [5, 13, 56]. CO2 is emitted from the burning of fossil fuels for elec­tricity, transport, and industrial processes [3]. The Kyoto Protocol in 1997 proposed a reduction of greenhouse gases by 5.2% based on the emissions in 1990.

Different CO2 mitigation options have been considered to meet the proposed tar­get [5]. The various strategies can be classified as either chemical-reaction-based approaches or biological mitigation. Chemical-reaction-based strategy captures CO2 by reaction with other chemical compounds before the CO2 is released into the atmo­sphere. The disadvantage of this method is that the chemical reactions can be very energy-intensive and costly. Furthermore, the wasted chemical compounds will need to be disposed of. On the other hand, biological mitigation seems more favourable as it not only captures CO2 but also generates energy through photosynthesis [56].

Photosynthesis is carried out by all plants and any photosynthetic microorganism. Even though the use of plants to capture CO2 is viable, it is inefficient due to their low photosynthetic rates. In contrast, owing to their structural and functional sim­plicities, microalgae are able to photosynthesize and hence capture CO2 with an efficiency up to 10-50 times greater than that of higher-order plants [56]. Microalgae include both prokaryotic cyanobacteria and eukaryotic unicellular algal species [5]. In addition to CO2 and sunlight, microalgae need nutrients, trace metals, and water to grow [2, 38, 46, 58]. In short, microalgal biomass is produced based on the fol­lowing reaction:

CO2 + H2O + nutrients + light energy ^ biomass + O2

Unlike plants, microalgae can be grown with waste or brackish water as their high adaptability enables them to survive in a hostile environment contaminated with excess nitrogen, excess phosphorous, and heavy metals. In fact, microalgae can directly metabolize the excess nitrogen and phosphorous in waste water as nutrients for their cultivation. As such, microalgal cultivation does not interfere with use of fresh water, a limited resource in many parts of the world [2, 34, 38, 46, 58].

Among the 30,000 species of microalgae on Earth, many of them are known to contain a variety of high-value bio-products that can be commercially harnessed, such as biodiesel-convertible neutral lipids, different isomers of carotenoids, poly­saccharides, polyunsaturated fatty acids, and phycobiliproteins [10]. In addition to being a CO2 bio-sequester, commercial applications of microalgal biomass also include: (1) biodiesel through transesterification of its neutral lipids; (2) bio-ethanol through fermentation of its carbohydrates; (3) nutritional supplements for humans; (4) natural food colourants; (5) natural food source for many aquacultural species; (6) natural colourants in cosmetics; (7) bio-fertilizers through pyrolysis; (8) protein feed for farm animals [10, 14, 52]. Some of these applications require specific compo­nents of the biomass to be recovered while others utilize the entire cellular biomass.

Algae are important sources of various bioactive compounds with different physi­ological effects (toxic or curative) on human health. Many of them possess antioxi­dant, antimicrobial, and antiviral activities that are important for the protection of algal cells against stress conditions. The discovery of new analytical methods and techniques is important for the study of metabolites in algae and similar organisms with respect to their applications in pharmacology and the food industry [132].

4.1 Carotenoids

Carotenoids are prominent for their distribution, structural diversity, and various functions. More than 600 different naturally occurring carotenoids are now known, not including cis and trans isomers, all derived from the same basic C40 isoprenoid skeleton by modifications, such as cyclization, substitution, elimination, addition, and rearrangement. The different carotenoids have been isolated and characterized from natural sources as plants [43, 187], algae [142,143], bacteria [183,191], yeast [119], and fungi [70].

Carotenoids play a key role in oxygenic photosynthesis, as accessory pigments for harvesting light or as structural molecules that stabilize protein folding in the photosynthetic apparatus. Carotenoids are powerful antioxidants. The beneficial effects of carotenoids have been well documented from the numerous clinical and epidemiological studies in various populations. Due to its high antioxidant activity, carotenoids have been proposed as cancer prevention agents [173], potential life extenders [88], and inhibitors of ulcer [74], heart attack, and coronary artery disease [151, 194].

All photosynthetic eukaryotes are able to synthesize lycopene, a C40 polyene, which is the precursor of two different carotenoid synthesis pathways, the b, e-carotene and the b, b-carotene pathways [1694. Xanthophylls are oxidation products of carotenes; diversification of xanthophylls increases by the inclusion of allene or acetylene groups. Allenic and acetylenic carotenoids are highly represented in algae, and at least 30 different carotenoids have been identified in this group [169]. The distribution of carotenoids having different molecular structures or the presence of specific biosynthesis pathways can be an index for algae classification. For example, the major carotenoids that occur in seaweeds (Fig. 1) include b-carotene, lutein, violax-
anthin, neoxanthin, and zeaxanthin in green algae (Chlorophytes); a — and b-caro — tene, lutein, and zeaxanthin in red seaweeds (Rhodophytes) and b-carotene, violaxanthin, and fucoxanthin in brown algae (Phaeophytes).

Carotenoid composition of algae can present great variations mainly related to environmental factors, such as water temperature, salinity, light, and nutrients avail­able. Most of the environmental parameters vary according to season, and the changes in ecological conditions can stimulate or inhibit the biosynthesis of several nutrients, such as carotenoids. For example, D. salina is a green microalga, well known for being one of the main natural sources of b-carotene. Under particular conditions, this microalga is able to produce b-carotene up to 14% of its dry weight. Moreover, the particular growing conditions able to maximize the production of b-carotene at industrial scale have been investigated [48-50, 84, 198, 118, 206]. Because b-carotene may play important roles in preventing degenerative diseases due to its associated antioxidant activity, different procedures have been studied, not only for the production of this compound but also for its extraction and isolation [66, 97, 106, 118]. The most widely employed technique has probably been SFE. The low polarity characteristics of the supercritical CO2 make this solvent appropri­ate for the b-carotene extraction from this microalga [66, 97, 106, 118].

Other example is the green microalgae H. pluvialis that produces chlorophylls a and b and primary carotenoids, namely, b-carotene, lutein, violaxanthin, neoxan — thin, and zeaxanthin, while it has the ability to accumulate, under stress conditions, large quantities of astaxanthin, up to 2-3% on a dry weight basis [150]. Using this carotenogenesis process, it undergoes different changes in cell physiology and morphology, giving as a result large red palmelloid cells [76, 204] . Astaxanthin is present in lipid globules outside the chloroplast, its functions in the cell include protection against light-related damage by reducing the amount of light available to the light-harvesting pigmented protein complexes. These pigments possess powerful biological activities, including antioxidant capacity [19], ulcer preven­tion [74] as well as immunomodulation and cancer prevention [130]. In fact, the extraction of astaxanthin has been thoroughly investigated. Different methods have been tested, including neat supercritical CO2 [189] or supercritical CO2 with differ­ent cosolvents [127], PLE [25, 67], MAE [203], direct extraction with vegetable oils [76] or solvents [75], or even treating cells with various solvents and organic acids at 70°C before acetone extraction, with the aim to facilitate the astaxanthin extraction from the thick cell wall without affecting the original astaxanthin esters profile [166].

Fucoxanthin is the most characteristic pigment of brown algae, and is also one of the most abundant carotenoids in nature [61] , accounting for more than 10% of estimated total natural production of carotenoids [103]. Fucoxanthin is an oxygen­ated carotenoid that is very effective in inhibiting cell growth and inducing apopto­sis in human cancer cells [60, 87]; it also has anti-inflammatory [172], antioxidant [158], antiobesity [99], and antidiabetic [100] properties.

A summary of the methane yield and volumetric production rate during algal diges­tion in continuous reactors appears in Fig. 11. The curves drawn through the data points on each figure represent general trends for easy comparison of the nonuni­form data.

Based on these data, algae as a substrate for ADP can be classified into three groups. The first group consists of algae with the methane yield larger than 0.3 L/gVS and includes brown macroalga M. pyrifera with high mannitol content, cyanobac­terium Arthrospira. green microalgae Chlorella and Scenedesmus. The second group has methane yield about 0.2 L/g VS and includes brown macroalga Laminaria and green macroalgae Ulva, Cladophora. and Chaetomorpha. Lastly, the third group has methane yield lower than 0.15 L/g VS and includes brown macroalga Sargassum, red macroalga Gracilaria, and green macroalga Enteromorpha.

Another important conclusion is that AD of M. pyrifera is stable at values of OLR up to 10 gVS/L-day. High values of the methane volumetric production rate are achieved 2.7 L(CH.)/L(digester)-day, reducing the required volume and the capital costs of the digester.

Finally, several methods such as application of advanced digestion reactors, co­digestion of algae with other substrates, algal hydrolysis and extraction of cellular liquids, and digestion of alginate extraction residues, significantly enhance methane yield, production rate, and the overall process efficiency.

The ADP is widely used for processing a variety of organic residues and wastes. The AD can be integrated into many potential pathways for algal biofuels production as shown in Fig. 19 [433]. Coupling biodiesel and green diesel production processes with the ADP (Fig. 20) can improve the overall efficiency of energy recovery and reduce the final cost of biofuel [434]. Based on the theoretical calculations, the ADP can recover approximately 55-85% of the biomass energy content in a coupled biodiesel-ADP, depending on algal lipid content [263]. Similar results were achieved by the calculation of theoretical energy output from conversion of T. suecica [435].

Chlorella residue from the biodiesel production process is a feasible substrate for methane production [436, 437]. Either 1-butanol as a solvent for lipid extraction or acid catalyzed in situ transesterification was recommended because application of the normal chloroform/methanol mixture inhibited methane production. The observed methane yield from algal residues was approximately 52-63% from fresh algae. Addition of glycerol as a co-substrate slightly increased the methane yield by

4- 7% possibly due to a more favorable N to C balance [437].

The ability to numerically simulate the behavior of geologic hydrate reservoirs has improved substantially over the past 5 years in terms of both code availability and capabilities [8]. There are currently several numerical models that can simulate the system behavior in hydrate-bearing geologic media (e. g., [30,68,94,130,143,152, 153, 188]). Several of these codes were calibrated against the Mallik 2002 produc­tion test data, and the data from the Mt. Elbert MDT test [2] . A code-comparison study [1, 214] indicated that most of the participating codes appear capable of simu­lating the behavior of hydrates and reservoir fluids during common dissociation scenarios. The current consensus is that the models generally account for the impor­tant physics of the problem, and that validation and calibration (rather than ade­quacy of the numerical code capabilities) will be a constraining factor in the assessment of hydrates as an energy resource [214, 2] . Additionally, while uncer­tainties exist in the description of properties and processes involved in numerical simulators (e. g., thermal properties of composite GH-bearing systems, relative per­meability and capillary pressure, geomechanical properties related to subsidence after the dissociation of the cementing GH from the porous media, etc.), these knowledge gaps are being addressed [125, 127].

Such challenges address the basic conceptual framework upon which theoretical and laboratory studies on the thermodynamics and flow properties of the GH sys­tems are based.

5.2.4 Development of Universal Standards for Hydrate Sample Creation

This is an important challenge for fundamental physico-chemical hydrate research, and it first involves the establishment of a protocol to fabricate artificial hydrate samples in sediment. The second part of this challenge is to ensure that the sample is a reasonable replicate of nature. Currently the method of Spangenberg et al. [181] appears to be dominant; however, months of sample preparation time are required, and there is an urgent need for a more time-effective techniques.

5.2.5 Thermodynamic Knowledge Gaps and Time-Dependence Issues

Gibbs energy minimization methods are currently the most effective tools in deter­mining the behavior of complex hydrates involving more than CH4 (a subject that needs to be tackled, as such hydrates are likely in GH systems), and form the basis of the statistical thermodynamics approach in the description of the properties and behavior of such systems. Because of serious experimental difficulties, the predic­tions of these methods currently cannot be verified in a large part of the P-T-X spectrum of composite hydrates, and especially in the presence of more than one inhibitors. Additionally, time-dependent measurements are required, to establish kinetic phenomena, which are currently confounded by the addition of heat and mass transfer.