Gas can be produced from GH by inducing dissociation, which also releases large amounts of H2O (1). The three main methods of hydrate dissociation are (1) depres­surization, in which the pressure P is lowered to a level lower than the hydration pressure Pe at the prevailing temperature T, (2) thermal stimulation, in which T is raised above the hydration temperature Te at the prevailing P, and (3) the use of inhibitors (such as salts and alcohols), which shifts the Pe-Te equilibrium through competition for guest and host molecules [111] . Long-term production strategies often involve combinations of the three main dissociation methods [ 131, 132] . Another production method involves CH4 exchange with another hydrate-forming gas (e. g., CO2) through a thermodynamically favorable reaction [52, 213].

2 Occurrence, Research Activities and Priorities, and Prospective Production Targets

Dedicated GH production wells will have to both drill and produce through and in HBS. This presents a number of unique design challenges which must be consid­ered for both onshore and offshore GH wells including reservoir subsidence, loss of mechanical strength of the HBS along the wellbore, and development of high exter­nal pressures along the wellbore [46, 168]

Reservoir compaction greater than 5% is a consistent indicator for potential cas­ing failures. Casing shear is the dominant failure mechanism, typically located in the overburden up to several 100 ft above the reservoir. There is typically little that can be done to prevent casing shear, other than strategic well placement. Field development economics should include a suitable budget for future well replace­ments if casing shear is expected. Reservoir subsidence can also result in tensile failures of the casing above the reservoir, and buckling failure within the reservoir. Tensile failures may be prevented through the use of slip joints or length expandable casing joints, placed strategically in the wellbore. Casing failures due to column buckling in the reservoir interval can be prevented by selecting heavy wall casings and by employing good cementing and solids control practices.

Carbonic anhydrase (CA) catalyzes the reversible hydration of dissolved CO2 via a two-step mechanism that involves an attack of zinc-bound OH- on a CO2 molecule loosely bound in a hydrophobic pocket. The resulting zinc-coordinated HCO3- ion is displaced from the metal ion by H2O [25]. The overall reaction is as follows:

CO2 + H2O о H+ + HCO3- (3)

Although this reaction occurs without CA, the uncatalyzed hydration and dehy­dration reactions are slow. Therefore, CA is important when the availability of CO2 or HCO3- becomes limiting to a metabolic reaction [43]. Because the reaction shown in (3) involves protons, the equilibrium ratio of the two carbon forms is a function of pH. At physiological ionic strengths, CO2 predominates at a pH of less than 6.4, and HCO3- is the dominant form in the pH range of 6.4 and 10.3.

Three main evolutionarily distinct families of CAs were initially identified (a, b, and g-CAs), and recently two more CA classes, the 5 and Z classes, have been found in marine diatoms [44]. All CA families require zinc at the active site to activate the water molecule, although the g-CAs have been shown to use iron at the active site under anaerobic conditions. However, there is no significant sequence homology between families, and they appear to be examples of convergent evolution of cata­lytic function.

2.2.1 Types of CA Present in R. eutropha

a-CA: The best-studied group of the CAs is the a-class, first described and mostly studied in mammals, but also found in other organisms [24, 26]. Most a-CAs are monomeric enzymes, with an active site zinc ion coordinated by three histidine resi­dues. Members of this family may be involved in maintaining pH balance, in facili­tating transport of carbon dioxide or bicarbonate, or in sensing carbon dioxide levels in the environment [25, 45].

b-CA: The b-class has been identified in plants, Bacteria, red and green algae, fungi, andArchaea [46,47]. Formanyorganisms, including R. eutropha, b-CAis essential for growth at atmospheric concentrations of CO2 [47, 48]. The fundamental struc­ture of b-CA, a dimer, is the only one known to exhibit allosteric regulation by bicarbonate ion [46] .

y-CA: The g-CAs, which are predominantly found in Archaea, have strikingly dif­ferent sequence features than the a — and b-CAs, The g-CAs. are trimeric enzymes that contain a zinc in the active site. In an anaerobic environment, the zinc is replaced by iron as the physiologically relevant active site metal [45, 49]. R. eutropha is capable of growth under anaerobic conditions, and may utilize a g-CA (see below) under these conditions.

Although the primary structure and number of subunits of the various classes of CAs are strikingly different, the metal — (in most cases zinc) coordinating site is remarkably similar at the structural level [45, 46].

Four putative CA genes were identified in the genome sequence of R. eutropha strain H16. Two are located on chromosome 1, and two on chromosome 2. H16_ A1192 encodes a g-like-CA/acetyltransferase, can (locus tag H16_A0169) and can2 (locus tag H16_B2270) encode b-CA enzymes, and caa (locus tag H16_B2403) encodes a periplasmic a-CA. The presence of genes for multiple carbonic anhy — drases in R. eutropha suggests that these enzymes play a major role in its physiology and that the function of the different types is complementary [24] . However, the exact roles of all four CA enzymes are still largely unknown. The only R. eutropha CA gene studied to date is can [48], which was identified as being essential for growth under atmospheric concentrations of CO2. Either the presence of the other three CAs was not sufficient to support growth under ambient CO2 concentrations, or these other CA genes were not expressed [48]. The metabolic processes in which the activity of the other three CAs plays a role remain to be identified.

Since CO2 and HCO3- are both involved in a wide range of cellular processes, CAs can have different physiological roles. One role is to increase the supply of CO2 or HCO3- for metabolic reactions. For example, during the carboxylation reaction of RuBisCO, which uses only CO2, competition between CO2 and O2 for the active site is attenuated when the concentration of CO2 is higher. It is mainly for this purpose that CA activity, and enhancement thereof, may be crucial for autotrophic IBT pro­duction in R. eutropha. Other roles CA may play include delivery of carbon to the correct location within the cell and retention of CO2. Both of these roles are impor­tant, as CO2 can readily pass through biological membranes and quickly leak out of the cell. The role of CAs in transportation and retention of CO2 is shown through the carboxysomes, or CO2 concentration mechanism (CCM), which are important in cyanobacteria and algal carbon fixation [50, 51] . The action of CAs may also be important in pH homeostasis [43]. The roles of CA enzymes in IBT production are likely to prove important, both for concentrating CO2 to drive the RuBisCO reaction towards carbon fixation and for increasing CO2 tolerance.

In order to obtain high-purity bioethanol, solids and other aqueous components associated with the bioethanol need to be removed by clarification and distillation respectively. This separation process, however, has not yet been demonstrated for microalgal-bioethanol broth. The residual biomass produced after the separation process can theoretically be concentrated and converted to other products, such as animal feeds or fertilizers. The purity of bioethanol must satisfy international stan­dards for fuel specifications, ASTM D5798—09B. The produced bioethanol can be either blended with gasoline to form E10 (10% bioethanol) and E85 (85% bioetha­nol) or used directly in vehicles as a substitute for gasoline. Each of the blends has its own specifications which vary from one country to another. The overall cost of bioethanol production from microalgae should be made low enough to compete with existing commercial fuels. Due to the lack of any existing pilot-scale produc­tion facility of bioethanol from microalgae, practical information on operating and production costs is not readily available.

Acknowledgement This work was supported by an Australian Research Council (ARC) Linkage grant between Bio-Fuel Pty Ltd (Victoria, Australia) and Monash University Department of Chemical Engineering (Victoria, Australia).

A variant of thermal cracking is the catalytic cracking, extensively used in the petrochemical industry to produce a significant percentage of the fossil-derived fuel currently used. This possibility has also been pursued for the production of biofuels from a wide variety of feedstocks, especially from low-value triglyceride-based biomass. The reaction takes place in fluid catalytic cracking (FCC) units where triglyceride mol­ecules are transformed into water, CO2, CO, and a mixture of hydrocarbons, some of the aromatic type [68]. The employment of a catalyst permits the utilization of milder con­ditions of temperature and pressure, with a better control of the final products [27, 92].

Hua et al. [50] studied the catalytic cracking transformation of vegetable oils and animal fats in the laboratory. The results show that they can be used as FCC feed singly or co-feeding with vacuum gas oil, which can give high yield (by mass) of liquefied petroleum gas (LPG), C2-C4 olefins, for example, 45% LPG, 47% C2-C4 olefins, and 77.6% total liquid yield produced with palm oil cracking. Co-feeding with vacuum gas oil gives a high yield of LPG (39.1%) and propylene (18.1%).

Different combinations of reactors and catalysts can be used, as for example pil­lared clays, alumina metal-supported catalysts, zeolites, among others. Also, the huge experience gathered in the petrochemical industry can be relevant in the devel­opment and implementation of cracking processes for biodiesel production.

Miguel Herrero, Jose A. Mendiola, Merichel Plaza, and Elena Ibanez

Abstract At present, functional foods are seen as a good alternative to maintain or even improve human health, mainly for the well-known correlation between diet and health. This fact has brought about a great interest for seeking new bioactive products of natural origin to be used as functional ingredients, being, nowadays, one of the main areas of research in Food Science and Technology. Among the different sources that can be used to extract bioactives, algae have become one of the most promising. Algae have an enormous biodiversity and can be seen as natural factories for producing bioactive compounds since either by growing techniques or by genetic engineering approaches, they can improve their natural content of certain valuable compounds. In this book chapter, a revision about the different types of bioactives that have been described in algae is presented including compounds, such as lipids, carotenoids, proteins, phenolics, vitamins, polysaccharides, etc. Also, the modern green techniques used to achieve the selective extraction of such bioactives are presented and the methods for fast screening of bioactivity described.

Two dewatering cases have been analysed in this section. The first study assumes that all cultivated biomass would either be produced solely by centrifugation or filtration and the second analysis considers a two-stage dewatering technique based on flocculation followed by centrifugation or filtration. The use of an initial flocculation step reduces the power usage in the subsequent centrifugation or filtration stage.

For a single-stage process, the calculations indicate that the dewatering of the RP culture is more energy intensive than that of the HTR or ELR. This trend is consis­tent across the majority of the centrifugation and filtration options. This is due to the significant volume of culture produced per batch in the RP system, and the greater degree of processing required in concentrating the RP culture as opposed to that of HTR or ELR. For the dewatering of the HTR and ELR cultures, the emissions pro­duced are comparable for both centrifugation and filtration, as the total culture vol­ume produced in these two cultivation options are about the same.

The second analysis was based on a two-step dewatering process: flocculation first step and a second step consisting of either centrifugation or filtration. A number of different flocculants were considered. The use of a two-step process significantly reduces the overall emissions compared with a one-step process as represented by the HTR cultivation system in Table 5. This trend is consistent for other types of cultiva­tion systems. The percentage emission reduction from a one-step process to a two — step process is around 90%, which is a significant drop in emissions. The study clearly shows that the combination of chitosan 1 (95% efficiency of water removal) with a second-stage decanter bowl centrifugation has the greatest drop in emissions: 99.5% reduction for the HTR, 99.6% reduction for the ELR and 99.7% for the RP. The lowest net emission for the dewatering of the HTR culture was 1,690.73 tonnes CO2-e/year which was for the dewatering process combination of chitosan 1 with a secondary vacuum-suction filtration method. The same combination of chitosan 1 and a suction filter produced the lowest emission of 2,686.37 tonnes CO2-e/year for the dewatering of the ELR culture and 10,781.52 tonnes CO2-e/year for the RP culture. Thus, the most desired dewatering combination per NGER regulations would be a first-stage chitosan 1 flocculation and a secondary stage suction filter. However, it is interesting to note that in terms of percentage reduction due to the addition of a flocculant, this combination has the lowest percentage reduction (only 9% for the HTR and 14.9% for the ELR culture). This is due to the low-energy consumption of the suction filter com­pared with its concentration factor. To concentrate a sample by 80 times, the energy input is only 0.1 kWh/m3; thus even though the volume is reduced, the energy required to dewater the culture by a single or double stage process is insignificant.

NGER regulations consider both Scope 1 and 2 emissions, but there are no Scope 1 emissions associated with the dewatering stage. Thus with respect to CPRS requirements, the selection of dewatering options could only be reliant on economic and design considerations. The combinations which were studied in detail are the two flocculants of chitosan 2 and LT-25/NaOH with a secondary dewatering stage of either: centrifugation (disc stack or nozzle discharge) and fi2tration (chamber, suction, drum filtration or TFF). The data for the HTR, ELR and RP are given in Table 6. The results clearly indicate that chitosan 2 is the better option, as it creates

the least amount of emissions. This is obviously due to the higher dewatering efficiency of chitosan 2: 90% compared with 30% for LT-25/NaOH. This reduces the energy requirements needed for the secondary centrifugation/filtration stage.

Ronald Halim and Michael K. Danquah

Abstract Chlorophyll, a green pigment found abundantly in plants, algae, and cyanobacteria, plays a critical role in sustaining life on earth and has found many applications in pharmaceutical, food, as well as cosmetic industries. Because of their high intracellular chlorophyll accumulations (up to 10% of cell dry weight), green microalgae are recognized as promising alternative chlorophyll sources. Successful co-production of a high value product such as chlorophyll in a microal­gal bio-refinery is desirable as it will alleviate the overall cost of producing microal­gal biodiesel. This chapter evaluates the bioprocess engineering required to recover and to purify chlorophyll molecules from microalgae. The use of organic solvents and supercritical fluids to extract microalgal chlorophyll on a commercial scale is examined. The use of chromatographic techniques to purify the recovered chloro­phylls is also reviewed. Finally, the chapter ends by presenting a case study which investigates the use of organic solvents (acetone and methanol) to extract chloro­phyll from Tetraselmis suecica on a laboratory scale.

1 Introduction

Microalgae are microscopic unicellular organisms capable to convert solar energy to chemical energy via photosynthesis. They require carbon dioxide, light, water, and other nutrients which facilitate the photosynthetic process to grow and large — scale production of microalgal biomass normally uses open-ponds. Growing con­cern over the widespread exploitation of food crops for biodiesel production has

R. Halim • M. K. Danquah (H)

Department of Chemical Engineering, Bio Engineering Laboratory (BEL), Monash University, Melbourne, VIC 3800, Australia

e-mail: michael. danquah@eng. monash. edu. au; kobinadanquah@yahoo. com

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_34, 807

© Springer Science+Business Media New York 2013 sparked an unprecedented level of commercial interests in exploring the use of microalgae as alternative biodiesel feedstocks. In addition to having high intracel­lular contents of biodiesel-convertible neutral lipids, microalgae contain numerous high-value bio-products that can also be harnessed such as different isomers of car­otenoids, polysaccharides, polyunsaturated fatty acids, and phycobiliproteins [40]. These bio-products have extensive applications in food and pharmaceutical indus­tries and their successful co-production from microalgae will significantly alleviate the high cost associated with microalgal biodiesel production. Unfortunately, lack of process understanding for the recovery and purification of these bio-products from the microalgal biomass has prevented the realization of their full potential applications.

Chlorophyll is one of the high-value bio-products that can be extracted from microalgae. As a pigment that selectively absorbs light in the red and blue regions, chlorophyll emits a green colour [24] . It is used as a natural food colouring agent and has antioxidant as well as antimutagenic properties [24]. The process of produc­ing chlorophyll from microalgae begins with concentrating the highly dilute microalgal culture (biomass concentration=0.1-1% w/v). The resulting paste is then dried to form powder before the cells are exposed to either an organic solvent or a supercritical fluid during chlorophyll extraction. These steps are explained more thoroughly in Sect. 4.

Chlorophyll is present abundantly in nature (plants, algae, and cyanobacteria) and plays a vital role in photosynthesis due to its “light harvesting” propensity. The specific functions that chlorophyll performs during photosynthesis can be viewed elsewhere [11, 20, 21]. Photosynthesis is a process which uses harvested light energy together with water and carbon dioxide to produce oxygen and carbohy­drates; as such, it converts solar energy into chemical energy. The direct product from this chemical process, carbohydrate, is used as the primary building block for plants and eventually all living organisms [21]. The importance of photosynthesis for life on earth is further highlighted by plants forming the basis of all food chains. It is estimated that 1.2 billion tons of chlorophyll/year are produced globally by both terrestrial and aquatic organisms [21].

There are two types of chlorophyll, chlorophyll a and chlorophyll b. However, these bio-molecules are highly susceptible to oxidation and their exposure to weak acids, oxygen, or light rapidly results in the formation of numerous degradation products [11, 20, 24]. Figure 1 shows the structures making up chlorophyll mole­cules [21, 45, 51]. The skeleton of chlorophyll molecule is the porphyrin macrocy­cle (Fig. 1a] , which comprises of four pyrrole rings. An attachment of a single isocyclic ring to one of the pyrrole rings gives rise to the phorbin structure (Fig. 1b). Each pyrrole ring contains four carbon atoms and one nitrogen atom. All of the nitrogen atoms face inward to create a central hole where an Mg2+ metal ion easily binds to form the chlorophyll structure (Fig. 1c). In chlorophyll b, the methyl group in ring II of chlorophyll a is replaced by a formyl group. This structural difference results in chlorophyll a being a blue/green pigment with maximum red absorbance at 660-665 nm and chlorophyll b being a green/yellow with maximum red absor­bance at 642-652 nm [20]. Chlorophyll in plant cells is confined within chloroplasts

where it is not only complexed with phospholipids, polypeptides, and tocopherols but also protected by a hydrophobic membrane [21].

When chlorophyll is removed from this original environment, its magnesium ion becomes unstable and may easily be displaced by a weak acid. This reaction leads to a series of reactions which eventually degrade the entire molecule. To increase stability, the magnesium ion in a free-standing chlorophyll molecule is often artificially substituted with a copper ion [21, 54].

Inflammatory processes are related with several cardiovascular diseases and oxidative stress, therefore its study is of high interest. Among anti-inflammatory compounds from algal sources astaxanthine, terpenes, sterols, indols, and shikimate- derivatives have been described [105] . There is a huge amount of enzymes and secondary metabolites involved in inflammatory processes, but the general trend is to measure the expression of some of those metabolites and/or enzymes when cells involved in the inflammatory response are “activated.” Leukocytes are among the most studied models; leukocyte migration has been shown to be one of the first steps in the initiation of an inflammatory/immune response and is essential for accumulation of active immune cells at sites of inflammation. The chemotaxis assay used to analyze the test material is designed to assess the ability of a test material to inhibit the migration of polymorphonuclear leukocytes (PMNs) toward a known chemotactic agent.

For example, polysaccharides from red microalga primarily inhibited the migration of PMNs toward a standard chemoattractant molecule and also partially blocked adhesion of PMNs to endothelial cells [101].

Although the biochemical composition of algae is very different among algal groups, cellulose is a common material among many algal species. The process of cellulose biological degradation has been extensively studied in recent years. The mechanism of cellulose enzymatic hydrolysis by anaerobic bacteria is quite different from the mechanism of aerobic organisms. Anaerobic bacteria have a large multienzyme complex—cellulosome, which is attached to the cell envelope and consists of up to 11 different catalytic enzymes carried by scaffold-proteins [113, 114]. The enzy­matic hydrolysis of algal cellulose is relatively slow and can be inhibited by the close association with other structural materials, such as polyphenols, fucoidan, protein, and alginate. Therefore, other specie-specific sulphonated, methylated or carboxylated polysaccharides, mannitol, proteins, and lipids usually determine the more readily biodegradable fraction of algal biomass.