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.

Figure 7 illustrates typical extraction curves obtained in all experiments using either acetone or methanol as an extracting solvent. Even though the specific chlorophyll yields were different between the various extractions, the trends in which the yield evolved throughout experimental duration were similar from one extraction to another. From the two curves presented, it can be observed that chlorophyll yield increased most rapidly in the beginning and that the rate of extraction decreased with experimental time. For extraction using methanol, the majority of extraction (more than 80%) was achieved within the first 2 h and extending the extraction time

beyond this point hardly affected the final chlorophyll yield. This asymptotic behav­iour advocates for the diffusion-driven nature of chlorophyll extraction, where extraction rate is directly proportional to the amount of unextracted intracellular chlorophyll [37] .

Table 6 shows the final chlorophyll yields for the various extractions in the study. The final chlorophyll yields (between 0.020 and 0.045 g/g microalgae) are slightly higher than the typical amount expected from the species (~0.020 g/g dried microal­gae) [53]. It is noted that chlorophyll contents of a particular microalgal species are highly dependent on the culture conditions adopted and are not always comparable from one study to another. The highest chlorophyll yield (0.045 g/g dried microal­gae) in the study was obtained when extraction was performed using acetone at ambient temperature directly on wet microalgal paste that had previously been stored at 4°C (exp no. 3).

The ratio of chlorophyll a : chlorophyll b in the extract remained similar for most extractions (between 2.0 and 2.5) with the exception of experiment no. 3 where extraction was performed directly from wet paste (ratio of chlorophyll a:b = 1.3). All of the chlorophyll ratios reported in Table 6 fall within the normal range of green algae (between 0.64 and 5) [53] and verify chlorophyll a as the principal chlorophyll in this microalgal biomass. The decrease in chlorophyll ratio during wet extraction can be explained by the difference in polarity between chlorophyll a and chlorophyll b. The presence of a formyl group (-CHO) instead of a methyl group (-CH3) in ring II position of its cyclic molecule makes chlorophyll b slightly more polar than chlorophyll a. During extraction from wet paste, the water molecules in the paste dissolved in the acetone and increased the solvent’s polarity. As such, the

Yields are represented as: average of three replicates ± maximum error. Final chlorophyll yield = final chlorophyll a yield + final chlorophyll b yield. Amount of water in powder=0 wt.%, amount of water in paste = 77.6 wt.%

solvent was able to interact more favourably towards chlorophyll b, resulting in its increased co-extraction and the decreased chlorophyll ratio.

A decrease in the storage temperature of microalgal paste prior to chlorophyll extraction (exp no. 2 compared to exp no. 1) was observed to increase final chloro­phyll extraction yields (both a and b). The combination of freezing and thawing actions which occurred only when paste was stored at -20°C was expected to rup­ture a portion of the microalgal cell membranes. The partial cell disruption would then liberate intracellular chlorophyll molecules directly into the extracting solvent, thereby enabling more rapid solvent-analyte interaction and producing the increased fi nal chlorophyll yield.

Acetone extraction from wet microalgal paste produced more than twice the final chlorophyll yield of the same extraction from dried microalgal powder (exp no. 3 compared to exp no. 2). Several potential reasons can be attributed to this significant yield discrepancy. The prior exposure of the powder to thermal drying (65°C for 16 h) oxidized some the more susceptible chlorophyll molecules and depleted the biomass of chlorophyll contents. On the other hand, the presence of water in the paste allowed the biomass to form a homogeneous mixture with the extracting sol­vent and increased chlorophyll extraction through enhanced solvent-analyte interac­tion. As previously discussed, the water in the paste also acted as a co-solvent and increased the selective extraction of chlorophyll b.

Figure 7 shows extraction with methanol to be significantly more efficient than extraction with acetone. Methanol extraction (exp no. 4) produced almost twice the chlorophyll yield of an equivalent acetone extraction (exp no. 2). Despite a similar finding from previous studies [ 24, 49] , acetone is still the preferred solvent for majority of chlorophyll extraction works due to its known propensity to reduce the co-extraction of chlorophyllase enzyme responsible for chlorophyll degradation [11, 24]. The continuous decrease in chlorophyll yield of methanol extraction after 4 h of operating time (Fig. 7) indicates the onset of chlorophyll degradation in that solvent. In the future, the co-extracted chlorophyllase enzyme should be inactivated by spiking the methanol with a small quantity of metal carbonate (sodium, calcium, and magnesium) throughout the extraction [11].

From experiment 5, 2, and 6, chlorophyll yield is observed to increase with extraction temperature. Operating the extraction at 40°C rather than at ambient tem­perature (approximately 20°C) increased chlorophyll yield by more than 100%. Increasing the temperature of chlorophyll extraction results in two simultaneous competing effects. Rapid thermal degradation depletes the overall chlorophyll con­tent of the biomass and can potentially reduce extraction yield, while the increase of chlorophyll solubility in the extracting solvent enhances mass transfer and can potentially improve extraction yield. In our case, the temperature rise seemed to lead to an increase in chlorophyll solubility that more than offset the decrease in chlorophyll content due to thermal degradation and resulted in a higher final chlo­rophyll yield. A previous study by Sartory and Grobbelaar [44] found that chloro­phyll extraction from microalgal biomass reached its optimum efficiency when the extraction was carried out at an elevated temperature near the boiling point of the extracting solvent.

The Rhodophyta is a relatively well-defined group of about 6,000 algal species with several features that differentiate them from other algal divisions, such as the pres­ence of accessory phycobilin pigments, the absence of flagella and centrioles [36]. The vast majority of red algae are marine multicellular, macroscopic species, which account for the majority of the so-called seaweeds [37]. The main habitats are near­shore and offshore zones (down to 40-60 m) in tropical and temperate climate regions while the presence of accessory pigments allow algae to grow at depths down to 200-250 m. Species with calcified cell walls are important for the estab­lishment and support of coral reef formation. Red algae are also found in brackish and fresh water, as well as in soil [38, 39].

Porphyra species are an important food source for humans in the Asia region [40]. Several Rhodophyta species (Gelidium, Gracilaria) are an important source of agar and agarose [41] . These polysaccharides are used in many laboratories for preparing culture media and separating nucleic acids [42]. Carrageenan is widely used in the food industry as a gel forming substance and stabilizer [43] (Tables 5-7). Structural, biochemical characteristics and productivity of selected red algae species are presented in Tables 5-7.

Many researchers have studied the response of algae to nitrogen and phosphorus limitation. The general trend for most green algae and diatoms is decreasing synthe­sis of proteins, polyunsaturated fatty acids, and structural polar lipids (phospholip­ids, glycolipids, and sulfolipids). An increasing production of carbon storage products (carbohydrates and triglycerides) is observed under nitrogen limitation [266-272] .

Growth with phosphorus depletion can decrease the algal growth rate and stimu­late an increase in the lipid to protein and carbohydrate to protein ratios [267, 273­277]. Phosphorus-starvation conditions negatively affect synthesis of long-chain polyunsaturated phospholipids [266] .

Silicon-Deficient Conditions

Silicon is an important element for lipid metabolism in diatoms. Silicon-deficient growth conditions stimulate synthesis of more carbohydrates and triglycerides with higher ratio of saturated and mono-unsaturated fatty acids compared to cells grown in silicon rich conditions [275, 278-280[ . After 4 h of silicon deficient growth, acetyl-CoA carboxylase activity increased by 100%, whereas activity of

P-(1^3)-glucan-P-3-glucosyltransferase (chrysolaminarin synthase) was reduced by 31% during the same period [275, 278-280].

Trace Metals Availability

Trace elements are important for optimal cell metabolism. Dunaliella tertiolecta deprivation in iron and cobalt significantly affected growth rate but simultaneously caused rapid accumulation of lipids [281]. Botryococcus spp. harvested during stationary phase and exposed to an iron-enriched medium stimulated lipids accumulation from 5-18 to 16-36% [282]. One Botryococcus species showed the highest lipid content after an exposure in nitrogen-free, but iron-enriched media with high light intensity. Chlorella vulgaris harvested in late-exponential growth phase and resuspended in iron-free media for 2 days followed by iron addition had a lipid content up to 57% by dry weight with 1.2 x 10-5 mol/L chelated FeCl3 or three to seven fold larger compared to cells resuspended in lower iron concentrations [283] .

George J. Moridis, Timothy S. Collett, Ray Boswell, Stephen Hancock, Jonny Rutqvist, Carlos Santamarina, Timoth Kneafsey, Matthew T. Reagan, Mehran Pooladi-Darvish, Michael Kowalsky, Edward D. Sloan, and Carolyn Coh

Abstract Gas hydrates are a vast energy resource with global distribution in the permafrost and in the oceans, and its sheer size demands evaluation as a potential energy source. Here we discuss the distribution of natural gas hydrate (GH) accu­mulations, the status of the international R&D programs. We review well-characterized GH accumulations that appear to be models for future gas production, and we analyze the role of numerical simulation in the assessment of their production potential. We discuss the productivity from different GH types, and consistent indi­cations of the possibility for production at high rates over long periods using con­ventional technologies. We identify (a) features, conditions, geology, and techniques that are desirable in production targets, (b) methods to maximize production, and

(c) some of the conditions and characteristics that render GH deposits undesirable. Finally, we review the remaining technical, economic, and environmental chal­lenges and uncertainties facing gas production from hydrates. [12] [13]

Symbols, Abbreviations, and Nomenclature

DP Pressure depletion (Pa)

GH Gas hydrate

HBL Hydrate-bearing layer

HBS Hydrate-bearing sediment

k Intrinsic permeability (m2)

keff Effective permeability (m2)

mbsf Meters below sea floor

MMSCF 106 of standard ft3 MMSCFD 103 of standard ft3/day MSCF 103of standard ft3

NH Hydration number

P Pressure (Pa)

Pcap Capillary pressure (Pa)

Pe Hydrate equilibrium pressure at a given T (Pa)

SH Hydrate saturation

STP Standard pressure and temperature

QM Mass production rate (kg/s)

QP Gas production rate (ST m3/s)

Qavg Average gas production rate (ST m3/s)

t Time (days)

T Temperature (K or °C)

RRR Rate replenishment ratio

TCF 1012 STP ft3 of gas

VRR Volume replenishment ratio

WZ Water zone

1 Introduction

Well testing is a key technique that is widely employed for reservoir characterization. Well-testing and pressure transient analysis (PTA) techniques are complementary to other characterization techniques as (1) they fill a gap between the small-scale characterization based on cores and logs, and large-scale characterization based on geophysical measurements, and (2) they provide a measure of flow capacity, in contrast to static properties determined from other techniques.

In hydrate reservoirs, there are at least three reported cases of pressure transient tests that along with other techniques have been used for reservoir characterization. There has been an evolution in the techniques available for interpretation of the test results. Initially, techniques developed for conventional reservoirs were applied, ignoring temperature and phase change effects associated with hydrate dissociation. Kurihara et al. [92] determined that there is a reasonable agreement between the effec­tive permeability keff found from application of the conventional PTA techniques, and the average keff over the dissociation zone obtained from a simulation model, but cau­tioned that the application of conventional PTA techniques to hydrate reservoirs is not straightforward. Gullapalli et al. [55] suggested that estimates of keff from conven­tional analysis could not be used because of significant uncertainties, and supported the use of hydrate-specific numerical simulators. Gerami and Pooladi-Darvish [49] developed a semi-analytical PTA technique for the interpretation of the flow (draw­down) data and parameter estimation for hydrate reservoirs that are underlain by a free-gas zone, which they verified against data from numerical simulators.

Despite the steady progress in testing and interpretation of the well-test data, significant challenges remain.

Despite the initial successes in IBT biosynthesis with E. coli [4] and yeast [8] using glucose as the main carbon source, the lone published attempt to establish IBT produc­tion in a host organism that can utilize CO. as a carbon source, the photosynthetic cyanobacterium Synechococcus elongates, resulted in an IBT titer of less than 1 g/L [4]. The facultatively autotrophic Gram-negative bacterium Ralstonia eutropha is capable of reducing CO2 in the presence of O2. It has also been observed to utilize H2, both exoge­nously delivered to culture vessels and produced in situ via electrolysis [9]. R. eutropha is a metabolically versatile bacterium that can also utilize sugars, fatty acids, amino acids, and triacylglycerols as carbon sources [10, 11]. When available in excess, carbon is typically stored by R. eutropha in the form of polyhydroxybutyrate (PHB), a natural polyester present in intracellular inclusion bodies [12-14]. Designing a recombinant strain of R. eutropha to produce IBT as opposed to PHB would require a redirection of carbon flux via the pathway shown in Fig. 1. A well-developed collection of R. eutropha basic biology and strain engineering tools, enumerated in Table 1, exists allowing facile

Fig. 1 Schematic diagram of the engineered isobutanol (IBT) production pathway in Ralstonia eutropha. Shown in red are the steps of the pathway that direct carbon flux away from branched chain amino acid (BCAA) biosynthesis towards isobutyraldehyde and IBT synthesis. IlvB aceto — hydroxyacid synthase; IlvC ketoacid reductoisomerase; IlvD dihydroxyacid dehydratase; IlvE transaminase; Kivd ketoisovalerate decarboxylase; Adh alcohol dehydrogenase; 3PGA 3-phospho — glycerate; 2PGA 2-phosphoglycerate; PEP phosphoenol pyruvate. All enzymes present in this figure are discussed in the text

modification of the R. eutropha genome. These factors make R. eutropha an ideal organ­ism for metabolic engineering to produce IBT from CO2, H2, and O2.

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.

In situ transesterification is another possibility that simplifies or even eliminates the need to perform the pre-processing of feedstocks, in particular the oil extraction and refining steps. The transesterification reaction is directly performed in the macer­ated oil seeds, such as soybeans flakes or animal fats containing the lipidic material [24,40, 70, 88, 102].

Although the in situ transesterification was proposed some years ago, it has not yet been used extensively. Reasons for this may be the large molar ratios between oil and alcohol that are necessary to obtain full oil conversion and the dependence on the seeds characteristics and on its oil content [ 88] . This process may seem simple, but it is still not fully worked out for practical applications and it is not economically efficient.

Haas et al. [47] investigated biodiesel production by in situ transesterification using as feedstocks corn dried grains (a by-product of ethanol production) and meat and bone meal (a by-product of animal rendering). As a result, these authors achieved almost the maximum theoretical transesterification conversion (91.1%) at ambient pressure and 35°C of temperature. For a higher temperature of 55°C, no significant increase in the conversion was achieved. Partial drying of the corn grains contrib­uted to reduce the methanol requirements to achieve a high degree of transesterification. For meat and bone meal, drying was not required to achieve a high degree (93.3%) of transesterification.

Unit operations such as centrifugation and microfiltration may be preceded by flocculation to improve the efficiency of recovery [21]. Flocculants can improve dewatering characteristics during centrifugation and fil tration because of their binding capabilities. Flocculants can help to maintain cellular properties when the culture experiences high shear forces during processes such as centrifugation [3]. Multi-stage dewatering processes have the potential to significantly reduce the energy consumption involved with large-volume cultures. It is estimated that bio­mass recovery contributes up to 20-30% of the total biomass production cost [12]. Therefore, multi-stage dewatering processes have the potential to reduce the eco­nomics involved with biomass production.