The strain of microalgae considered for the design is P. tricornutum, which is a type of marine algae from the class Bacillariophyceae [31]. Figure 2 shows the proximate biochemical composition of P tricornutum. The design basis for the cultivation sys­tem is 50,000 tonnes of dry biomass per year. The cultivation system will operate 330 days per year with the number of batches dependent on the type of cultivation

system employed, as well as the specific yields and productivities. Seawater is used as the major source of water due to the marine nature of the algal specie. The life span of the plant is fixed at 10 years. It is assumed that 80% of the biomass is removed from the cultivation system at the end of each batch. After 330 days of cultivation, the facility will shutdown for major maintenance. Dominant strains of algae and unwanted parasites can often enter the reactor and destroy the culture [5]; thus it is pivotal that the cultivation system is shutdown for scheduled maintenance periodically. The cultivation system will rely on carbon dioxide from a power sta­tion, and this will be the only source of carbon dioxide for photosynthesis. The car­bon dioxide from the power station is assumed as a mixture with compressed air such that the mass fraction of carbon dioxide entering the cultivation system is 10%.

The major processes investigated as dewatering alternatives in this study include single-stage dewatering using centrifugation, chamber filtration, vacuum filtration, suction filtration and a dual-stage process using flocculation followed by centrifuga­tion. In comparing the dewatering of different cultivation options, the raceway pond was approximately 15 times more expensive to dewater using a single-stage process than the reactor-style systems. This is primarily due to a combination of the significantly larger volume of the raceway pond and its much lower concentration of biomass, which requires the dewatering equipment to run for a greater length of time to process comparable amounts of dry biomass, leading to exorbitant energy costs. The costs of dewatering biomass from the raceway pond cultivation system using the four options investigated can be seen in Fig. 11. The capital costs neces­sary in the dewatering stage were found to be very low, whereas the contribution of running costs was found to be significantly larger. Of the alternatives shown in Fig. 11, the chamber and suction filtration options were found to be significantly cheaper than the single and dual-stage centrifuge systems, chiefly due to their lower electricity consumption.

However, the large consumption of electricity demanded by single-stage centrifugal recovery has other major environmental and economic impacts, which makes the dual-stage dewatering process the preferred option. As previously noted, the costs in dewatering biomass from the reactor-style cultivation systems were considerably less than the cost in dewatering raceway pond culture. In Fig. 12 filtration again appeared to be the cheapest dewatering option; however, there were a number of aforementioned hidden costs associated with fouling which were not included in the model. Significantly, as shown by the dot shaded bar in Fig. 12, the higher capital costs relative to running costs made dual-stage dewatering uncom­petitive at this smaller reactor volume and higher concentration of algae. Thus, due to its greater reliability and cost effectiveness, a single-stage centrifugal dewatering process would be the optimal production selection in the dewatering of reactor-style cultivation systems.

Based on this discussion, there are at least four metrics that should be included in life cycle studies of algae-to-energy technologies:

• Net energy

• GWP

• Land use

• Water use

Net energy is important because efforts to use algae for fuel production are predi­cated on the assumption of a positive net energy balance. Similarly, GWP is impor­tant because of the expectation that algae-to-energy systems will be no more carbon intensive than conventional fossil fuels. In addition, land use and water use should be considered because of algae’s high productivity relative to terrestrial crops and its unique requirements for water that set it apart from other sources of bioenergy.

Today, there is a wide range of classical or conventional extraction techniques that have been traditionally employed for the extraction of interesting compounds from natural matrices, such as algae. In this group, techniques such as Soxhlet, liquid-liquid extraction (LLE), solid-liquid extraction (SLE), and other techniques based on the use of organic solvents are included. Although these techniques are routinely used, they have several well-known drawbacks; they are time consuming, laborious, they lack of automation and therefore are more prone to present low reproducibility, have low selectivity and/or provide low extraction yields. These shortcomings can be partially or completely overcome by using the newly developed advanced extraction techniques. This new kind of extraction techniques are characterized by being faster, more selective towards the compounds to be extracted, and also very important nowadays, these techniques are more environmentally friendly. In fact, by using the considered advanced extraction techniques, the use of toxic solvents is highly lim­ited. In the next sections, the most important advanced extraction techniques that have been employed to extract bioactive compounds from algae are briefly described and commented.

Chlorophyll a and b are the dominant pigments in Chlorophyta and are the source of the second name of these organisms—Green algae. The secondary pigments are carotenoids (b-carotene, prasinoxanthin, siphonaxanthin, astaxanthin) which some­times give algae their yellowish-green and red-green colors [44]. The major habitat for green algae is freshwater although they are also found in sea or brackish water, and in soil [38, 39, 45, 46]. Chlorophyta species are unicellular or colonial motile and non-motile, filamentous, coccoid, parenchimatous, and siphonous [37, 47] .

Table 8 Chlorophyta species major organic matter characteristics

Characteristic Description References

Nutrient reserves Chlorophycean: mix of amylose [511,533-540]

(a-1,4-linkage) and amylopectine (a-1,4 and a-1,6 linkage) inside of chloroplast Lipids

Polyphosphate granules

Cell wall organization Mostly two layered [161,162,170,541-549]

Outer mucilage or capsule Structural component—crystalline

cellulose (Cladophorales), amorphous cellulose (Ulvales, Oedogoniales, coccoid algae), xylose or mannose (Caulerpales, Codiaceae,

Polyphysaceae) in hemicellulose, glycoproteins (Volvocales)

Several microalgae (e. g., Chlorellaceae,

Scenedesmaceae, Hydrodictyaceae families) have resistant trilaminar structure containing nonhydrolysable biopolymer—algaenan Some marine siphonous species are

______________________ calcified with CaCO3___________________________________

Halophilic microalga Dunaliella is widely cultivated for the production of b-carotene and other human nutritional products [48]. Species from Ulvophyceae group are mostly marine macroscopic algae that are used as food in coastal regions and can be used for nitrogen removal during wastewater treatment [49] (Tables 8-10).

Heterotrophically cultivated cells use organic substrate(s) for carbon and energy while mixotrophically grown cells can use light and organic carbon for energy and use either organic or inorganic carbon for biomass synthesis [180, 181, 284]. Many algae can utilize various organic carbon substrates: sugars (glucose, mannose, fructose, lactose, etc.), VFA (acetate), glycerol, molasses, and organic carbon from wastewater. Under mixotrophic conditions, algae exhibit a five — to tenfold higher growth rate compared to photoautotrophic growth [285]. Species that grow mixotrophically include:

• Green algae: C. reinhardtii [286-288]; Chlorella sp. [289-293]; Scenedesmus [294-296]; Tetraselmis suecica [297]; Platymonas subcordiformis [298]; Botryococcus braunii [299]; Micractiniumpusillum [300]; Haematococcusplu — vialis [301-303]; Haematococcus lacustris [304, 305].

• Red algae: Porphyridium cruentum [306, 307]; Galdieria sulphuraria [308].

• Diatoms: Phaeodactylum tricornutum [256,285,309-312]; Nannochloropsis sp. [313-315]; Navicula saprophila [316]; Nitzschia [316, 317].

• Cyanobacteria: Synechococcus [318, 319]; Arthrospira [320-323]; Nostoc flagelliforme [324]; Anabaena variabilis [325].

The main disadvantage of mixotrophic growth is the high cost of organic carbon sources. Main advantages of mixotrophic growth are the elimination of light pene­tration limitation that allows high concentration of algae (opportunity to reduce harvesting cost), better process control, increased growth rate and production of lipids, and potential to use waste streams as an organic carbon source. One attractive solution is the coupling of wastewater treatment and algal production [326]. Another possible source of organic carbon is acetate from a modified AD system where the major products of anaerobic fermentation are acetate and hydrogen. The drawback of systems grown on waste streams is bacterial and viral contamination.

Light Conditions

Light characteristics have a significant impact on pigment content, photosynthetic activity, and lipid content [327, 328]. Generally, high irradiation inhibits the forma­tion of polar lipids, but stimulates synthesis of storage carbohydrates and neutral lipids (usually triglycerids). Light limitation favors the production of total proteins and structural polar lipids associated with chloroplasts [258, 329-332]. B. braunii cultivated under continuous illumination gave the highest yield of exopolysaccha­rides. A light-dark cycle of 16-8 h resulted in the highest hydrocarbon yield [333]. Blue light was found to promote protein synthesis, while red light stimulated carbo­hydrate production [334].

Gas hydrates (GH) are solid crystalline compounds in which gas molecules (referred to as guests) occupy the lattices of ice-like crystal structures called hosts. Under suitable conditions of low temperature T and high pressure P, the hydration reaction of a gas G is described by the general equation

G + NHH2O = G • NHH2O, (1)

where NH is the hydration number. GH deposits occur in two distinctly different geographic settings: in the permafrost and in deep ocean sediments [91].

Naturally occurring hydrocarbon gas hydrates contain CH4 in overwhelming abundance. Simple CH4 hydrates concentrate methane volumetrically by a factor of 164 when compared to standard P and T conditions (STP). Such hydrates have 5.77 <NH< 7.4, with NH = 6 being the average value and NH = 5.75 the maximum one 4 1784 • Natural gas hydrates can also contain other hydrocarbons (alkanes CH2v+2, v = 2-4), but may also comprise lesser amounts of other gases (mainly CO2, H2S, or N2).

Although there has been no systematic effort to map and evaluate this resource and current estimates of the in-place amounts vary widely, the consensus is that the world­wide quantity of hydrocarbon GH is vast [80, 120, 178]. Given the magnitude of the resource, the ever-increasing global energy demand, and finite conventional fossil fuel reserves, the potential of GH as an energy source demands technical and eco­nomic evaluation. The attractiveness of GH is further enhanced by the environmental desirability of natural gas, as it is an energy resource with significantly lower carbon intensity than coal, oil, or other solid and liquid fuels.

The past decade has seen a marked acceleration in GH research and development (R&D). Among the most important developments are the increasing focus of research on gas hydrate-bearing sediments (HBSs) rather than crystalline hydrate, the improvements in tools available for sample collection and analysis, the emer­gence of robust numerical simulation capabilities, and the transition of GH resource assessment from in-place estimates to potential recoverability [8]. A fuller under­standing of the complexities of GH geological systems has emerged, including new insights into the effects of solubility, salinity and heat flow, reservoir lithology, and rates and migration pathways of both gas and H2O [151,167]. Additionally, critical data gaps, such as information on the mechanical and hydraulic properties of HBS, are being addressed. Significant inroads are also being made into our understanding of hydrate response under different production scenarios.

GH are often compared to coalbed gas, which was also considered an uneco­nomic resource in the not too distant past [21] • However, once the resource was geologically understood, the reservoir properties defined, and the production chal­lenges addressed, coalbed gas became a viable fuel in its own right and an important part of the energy mix in the United States, where it accounts for almost 10% of the natural gas production. Past experience with other unconventional energy resources shows that the evolution of GH into a producible source of energy will require a significant and sustained R&D effort. Here we discuss the current state of this effort and of the corresponding knowledge status.

An ideal well-test interpretation solution should be comprehensive enough to incor­porate the important mechanisms, and sufficiently simple to allow estimation of the controlling properties and parameters through inversion. This has been possible in the case of hydrate-capped gas reservoir [49]. Similar solutions for more prevalent GH accumulations, i. e., those that are within the hydrate stability zone and coexist with formation water, have yet to be developed. It is known whether such a complex and nonlinear problem can be simplified sufficiently to yield analytical solutions. Further complexities make the application of such solutions (even if they are pos­sible) problematic. Thus, the strong nonlinearity of the hydrate problem renders the principle of superposition (upon which a large part of well-testing techniques are founded) inapplicable. Additionally, the concept of radius of influence may be inap­plicable to GH reservoirs [127].

5.2.1 Practical Challenges

Actual well-test data from GH deposits are scarce. It is expected that our ability to interpret these tests will improve as we acquire more well-test data of longer duration [127].

Autotrophic production of PHB in R. eutropha has been studied previously, and the fermentation parameters to maximize culture density and mitigate explosion risk
have been studied in detail [14-16]. Also, stoichiometric formulae for autotrophic biomass and PHB production have been calculated, with the latter being [14]:

4CO2 + 33H2 + 12O2 ^ C4H6O2 + 30H2O (1)

Adapting this stoichiometry for IBT production, the overall mass balance on the gaseous inputs and liquid products would be:

4CO2 + 36H2 + 12O2 ^ C4H10O + 31H2O (2)

The standard free energy (AG°) for this overall reaction is -5.0 MJ/mol IBT, which demonstrates that IBT production is not in violation of thermodynamic laws. Assuming that 50% of the electrical energy used for H2O electrolysis (AG° = 0.237 MJ/ mol H2O) is lost as heat, approximately ((0.237 MJ/mol H2) x (36 mol H2)/ (0.5)) = 17.1 MJ of electrical energy are needed to produce 1 mol of IBT (C4H10O). As the approximate heat of combustion of IBT is 2.4 MJ/mol, the IBT production process involving a recombinant R. eutropha strain discussed here should be able to convert approximately 10% of the input electrical energy to transportation fuel energy. The required input H2:O2 ratio of 3:1 can easily be achieved via H2O elec­trolysis, which produces separate H2 and O2 product streams that can be fed to IBT — producing R. eutropha cultures in any desired ratio. However, integrating the fermentation with in situ generation of H2 and O2 presents a reactor design chal­lenge, which is discussed below (see Sects. 4.1 and 4.2).

When grown autotrophically, R. eutropha generates the energy and reducing equivalents required to drive carbon fixation by the oxidation of H2 gas, catalyzed by hydrogenase enzymes. There are three types of hydrogenases present in R. eutro­pha. Two of these are energy conserving hydrogenases (Fig. 2): a membrane bound hydrogenase (MBH, Fig. 2a) and a soluble hydrogenase (SH, Fig. 2b). The third is a regulatory hydrogenase (RH), which serves as a hydrogen sensor [17].

Electrons generated by the MBH are directed into the respiratory chain, provid­ing the reducing equivalents for reduction of O2 to H2O and the proton gradient for ATP synthesis [18] . The cytoplasmic SH directly reduces NAD+ to NADH [19], which is necessary to drive carbon fixation via the Calvin-Benson-Bassham (CBB) cycle [20]. Because all three hydrogenases in R. eutropha are resistant to inhibition by ambient oxygen concentrations, H2 oxidation can be coupled to the reduction of O2, a rare property among microorganisms [21]. Despite the presence of a hydrogen sensor, expression of both energy-conserving hydrogenases is linked to global energy level, not the amount of available hydrogen [22]. The combination of consti­tutive expression and oxygen resistance allows use of the hydrogenases in aerobic processes, such as the autotrophic production of IBT.

For design of an IBT production pathway in R. eutropha. carbon must first be diverted from PHB biosynthesis. A deletion of the PHA synthase gene, phaC, abol­ishes PHB production [23]. However, to maximize carbon flow to IBT, the b-keto — thiolase and acetoacetyl-CoA reductase genes (phaA and phaB, respectively) have also been deleted. Further optimization of carbon flow requires that many of the

genes and enzymes needed for the IBT biosynthesis pathway (Fig. 1), most which are native to R. eutropha, be expressed concomitantly and that the temporal expres­sion of the pathway is highest during “carbon storage” conditions.

The CBB cycle is used by a vast majority of autotrophic microorganisms for CO2 assimilation, and is often coupled with photosynthesis. Ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) is the key enzyme of the cycle. RuBisCO is a bifunctional enzyme that is involved both in photosynthesis and photorespiration in photosynthetic organisms. The enzyme catalyzes the initial step in the fixation of CO2. In this reaction, one molecule of CO2 is added to ribulose-1,5-bisphosphate, yielding two molecules of 3-phosphoglycerate (3-PGA) which can be metabolized to pyruvate or other central metabolites. When R. eutropha is growing autotrophi- cally, the reducing equivalents required for the CBB cycle are generated by the oxidation of hydrogen, rather than photosynthesis. Figure 3 shows a schematic of the R. eutropha CBB pathway.

To enhance CO2 fixation, organisms have developed efficient methods to acquire inorganic carbon. Carbonic anhydrase (carbonate dehydratase) enzymes play important

Fructose-6-Р

Fig-3 Schematic diagram of the Calvin-Benson-Bassham (CBB) cycle in R. eutropha. Ribulose- 5-phosphate is phosphorylated by the enzyme phosphoribulose kinase (CbbP). The resulting com­pound, ribulose-1,5-bisphosphate is then carboxylated by ribulose-1,5-bisphosphate carboxylase/ oxygenase (RuBisCO) (CbbL and CbbS) The outcome of this carboxylation are two molecules of 3-phosphoglycerate (3-PGA). 3-PGA is phosphorylated by phosphoglycerate kinase (CbbK) to yield 1,3-bisphosphoglycerate (1,3-BP). 1,3-Bisphosphoglycerate is reduced by NADPH to yield NADP+ and glyceraldehyde-3-phosphate (GAP) by glyceraldehyde-3-phosphate dehydrogenase (CbbG). GAP is then converted fructose-6-phosphate (F6P) by aldolase (CbbA) and fructose bis — phosphatase (CbbF). The reversible reactions of the reductive pentose phosphate cycle involving erythrose-4-phosphate, fructose-6P, sedoheptulose-7P, xylulose-5P, and ribose-5-P are catalyzed by the enzymes: Transketolase (CbbT), fructose-bisphosphate aldolase (CbbA), fructose/sedohep — tulose bisphosphatase (CbbF), ribulose-5-epimerase (CbbE), and triosephosphate isomerase (TpiA). Ribose-5P is isomerized by ribose-5-phosphate isomerase (RpiA) to yield ribulose-5P, which can then be put back into the cycle [27, 96] roles in this process [24]. Carbonic anhydrases (CAs) are zinc-containing enzymes that catalyze the reversible formation of bicarbonate (HCO3-) from water and carbon diox­ide [25]. These enzymes are important to many physiological processes as well, such as respiration, photosynthesis, transport, and autotrophic fixation of CO2 as well as HCO3- or H+ coupled ion transport, pH regulation, or carboxylation reactions. Recent work has shown that CAs are present in a wide range of metabolically diverse species from both Archaea and Bacteria, indicating that the enzyme has a more extensive and fundamental role in prokaryotic biology than previously recognized [24, 26]. Maintenance of the optimal CO2 concentration in the R. eutropha cell during auto­trophic fermentation avoids CO2 limitation during carbon fixation by the CBB cycle and, in consequence, ensures optimal IBT production. The action of CAs could play a central role in this process.

The 3-PGA produced in the CBB cycle is converted into pyruvate [27] , which can be utilized by the branched-chain amino acid (BCAA) production pathway to produce the intermediate ketoisovalerate (KIV) (Fig. 1). The BCAA valine, leucine, and isoleucine are synthesized by plants, algae, fungi, Bacteria, and Archaea through

Fig. 4 Schematic diagram of BCAA metabolism in R. eutropha. Pyruvate is the common precur­sor, which is reacted by acetohydroxyacid synthase (AHAS). AHAS can also incorporate 2-keto — butyrate, allowing a branch point to isoleucine biosynthesis. Ketoacid reductoisomerase (KARI) and dihydroxyacid dehydratase (DHAD) then produce the key intermediate 2-ketoisovalerate (KIV). A transaminase (TA) produces valine from 2-KIV. The IBT production pathway competes with TA for 2-KIV. Other enzymes: IPMS isopropylmalate synthase; IPMD isopropylmalate syn­thase; IPMDH isopropylmalate dehydrogenase

a common pathway [28] . The common enzymes in BCAA biosynthesis pathways (Fig. 4) are acetohydroxyacid synthase (AHAS), ketoacid reductoisomerase (KARI), dihydroxyacid dehydratase (DHAD), and transaminase (TA). These enzymes are involved in synthesis of all three BCAAs, and their expression and activity are tightly regulated through tRNABCAA repression, substrate specificity, and feedback inhibition. R. eutropha BCAA biosynthesis enzymes have not been studied previ­ously. However, sequence alignment with other characterized BCAA biosynthesis enzymes from E. coli, Corynebacterium glutamicum, Bacillus subtilis, and Streptomyces avermitilis revealed that the R. eutropha enzymes are most similar to the ones from E. coli on the level of primary sequence.

The KIV produced in the BCAA pathway is a key intermediate in IBT production (Fig. 4). To decarboxylate KIV to isobutyraldehyde, the precursor of IBT, a heter­ologous enzyme must be expressed in R. eutropha. To accomplish this, the ketois — ovalerate decarboxylase (kivd) gene from Lactococcus lactis is expressed, either on a plasmid or inserted into the R. eutropha genome, to allow production of isobutyr — aldehyde from 2-KIV (data not shown). Subsequently, an alcohol dehydrogenase (Adh) converts isobutyraldehyde to IBT. Initial assays for Adh activity in cell extracts of R. eutropha using isobutyraldehyde as the substrate yielded no detectable activity. Multifunctional Adh activity is repressed in R. eutropha during growth under ambient O2 concentrations [29-31]. Thus, a native, constitutively expressed Adh or a heterologous Adh with substrate specificity for isobutyraldehyde is needed for the final step of IBT production. A “short-chain alcohol dehydrogenase” has
been described in R. eutropha, but in wild-type cells is only expressed under anaerobic conditions [31]. Thus, the enzyme must be expressed under the right conditions for use in IBT production. Additionally, heterologous Adh enzymes, such as YqhD, have shown promise for converting isobutyraldehyde to IBT. The yqhD gene from

E. coli [32] has been used in previous iterations of IBT production pathways in heterotrophic organisms [4, 33].

Amongst the many microorganisms used for bioethanol production, Saccharomyces sp. remains the prime species. Currently, both alcoholic beverages and ethanol fuels are produced through fermentation performed by Saccharomyces sp. The species are resistant to high temperatures and provide a high ethanol tolerance level, allow­ing fermentation to continue at ethanol concentrations of 16-17% (v/v) [8]. Bacteria, particularly Zymomonas mobilis and Escherichia coli, have been successfully used to ferment biomass for bioethanol production [34]. However, bacteria are less robust than yeast and their growth requires a narrow pH range (6.0-8.0), thus less prefer­able to be used in the fermentation process. Table 6 shows some wild-type of micro­organisms commonly used for industrial ethanol production. Most of the listed microorganisms fail to ferment xylose even though it is one of the sugars obtained from the hydrolysis process. In order to overcome the hurdle, genetically-modified microorganisms have been cultivated and tested to ferment xylose [25, 65]. To date, the successful use of genetically-modified strains for fermentation has been reported only at a laboratory scale.

The fermentation of simple sugars into bioethanol involves a glycolytic pathway which occurs in two major stages. The first stage is the conversion of the various sugar molecules to a common intermediate, glucose-6-phosphate. The second phase is the metabolism of each molecule of glucose-6-phosphate to yield two molecules of pyruvate [38]. The products from the glycolysis steps are further metabolized to complete the breakdown of glucose. Under anaerobic conditions, the pyruvate is further reduced to ethanol with a simultaneous release of CO2 as a by-product.