Lignocellulosic biomass is a mixture of cellulose, hemicelluloses, and lignin which are held together by covalent bonding, various intermolecular bridges, and van der Waals forces forming a complex structure. Several studies have been conducted using model compounds such as cellulose, hemicelluloses, and lignin in sub — and supercritical water to establish the reaction pathways of these compounds in hydro­thermal medium. The chemistry behind the reactions of the individual components of biomass under hydrothermal conditions is well understood. The generalized individual reaction pathways of the major components of biomass (cellulose, hemicelluloses, and lignin) are discussed in the following section.

The objective of this approach is to convert feedstock into FT liquid as much as possible. The heat and particularly electricity generations are of secondary impor­tance. This approach requires a large production capacity. Since FT process has a high fixed cost, the economy scale is important to produce FT liquid competitively with other biofuels such as bioethanol, biodiesel, as well as FT liquids derived from fossil fuels [46]. This means gasifiers of the size at least greater than 1,000 MWth are required. Fundamental assumptions and other elements of this approach are:

1. FT off-gas is recycled to the biomass gasifier to achieve maximum syngas con­version. Electricity is produced as a “by-product” from the relatively small recy­cle bleed stream.

2. The yield of syngas H2 + CO in the FT feed gas must be as high as possible.

3. Since significant energy remains in CH4, C2H4, BTX, and tar, the process requires tar cracker and shift reactor or a reformer to convert all hydrocarbons into CO and H2. The increase in feed H2 + CO and increase in their yields result in much higher production of FT liquids and wax and higher overall efficiency of 63.5%.

4. Both gasifier and tar cracker must be oxygen blown to reduce dilution of recycle stream by N2 and other inert materials.

5. Unlike in the front-end approach, the part of gas conditioning system, CO2 removal step is essential.

CO2-TPD was used to study Fe catalysts promoted by different contents of Zn, K, and Cu. Figure 5 is the results of desorbed CO2 from these catalysts. There is much CO2 desorbed from the catalysts with 2 mass% K. However, the amount of CO2 decreases evidently for the catalysts promoted by 1 mass% K. Potassium is a main factor to determine the adsorption of CO2 on Fe catalyst [16, 30, 49]. For the catalysts with same content of K, Zn, and Cu show their influence. The amount of desorbed CO2 decreases firstly when Zn and Cu contents increase from 2 to 4

293 493 693

Temperature (K)

mass%, and then increase with the further raise of Zn and Cu content to 6 mass%. There is a slight increase for the 1 mass% K-promoted Fe catalysts after Zn and Cu contents are changed from 2 to 4 mass%.

Figure 6 illustrates the desorbed CO during CO2-TPD. The peak around 763 K is related with the loading content of K which is similar to desorbed CO2 from these catalysts. A sharp peak exists at this range for 2 mass% K-promoted catalysts, whereas it decreases greatly with the Fe catalysts promoted by 1 mass% K. On the contrary, 1 mass% K-added catalysts show the maximum of CO desorption around 930 K. In this temperature range, only Z6K2C6/Fe which belongs to the series of 2 mass% K-promoted catalysts has a clear desorption peak. The effect of K on des­orption is displayed by the difference around 1,040 K, too. Much CO is desorbed from 2 mass% K-promoted catalysts, but catalysts Z4K1C4/Fe and Z2K1C2/Fe with low content of K do not emit CO in this temperature range.

Therefore, it can be concluded that K is the main component to control the adsorp­tion and evolution of CO2 based on the results revealed by TPD. Although Zn and Cu are inferior components relative to K, their action become larger in the case of higher ratio of Zn/K. Z6K2C6/Fe has a desorption peak at ~930 K contrasting with the flat pattern of Z4K2C4/Fe and decline of Z2K2C2/Fe in this range. Considering the results of mono-promoted Fe catalysts in Fig. 4, the CO peak around 930 K in Fig. 6 is related to the influence of Zn other than Cu on Fe catalyst structure.

Production of butanol in a fed-batch reactor is another system where hydrolysis, fermentation, and recovery can be combined (SSFR). Using such a system, butanol was produced from wheat straw [58]. The reactor was loaded with 86 g/L pretreated wheat straw, enzymes, and the butanol producing culture. The system was operated for 533 h at a pH 5.0 and temperature 35°C. During the initial period of 120 h, wheat straw was hydrolyzed completely by the added hydrolytic enzymes. In order to ascertain that the culture was not deficient in sugar, a mixture containing glucose, xylose, arabinose, galactose, and mannose was fed to the reactor to mimic their proportion in wheat straw. In a 1 L reaction mixture a total (including sugar present in WS) of 430 g sugar was used thus producing 192 g total ABE with a yield of 0.44 (Table 4). In this reactor, a productivity of 0.36 g/L h was achieved which is 20% higher than achieved (0.30 g/L h) in a control reactor. One of the major problems associated with this fermentation was that the culture had difficulty utilizing xylose in the later part of fermentation. In this system, gas stripping was used to agitate the treated biomass and recover ABE.

2.1.2 Genetic Manipulation of Cellulose Characteristics

One of the central aims of cell wall scientists today is to find a formula for manipu­lating cellulose polymer structure without impairing plant growth. It has been pro­posed that altering the degree of crystallization and polymerization in the cell wall can result in cellulose polymers more responsive to hydrolysis [60]. The highly organized crystalline domains of the cell walls are interrupted with less crystalline, “amorphous” regions. There is an inverse correlation between the degree of cellu­lose crystallinity and the initial rate of cellulose hydrolysis [ 64]. At least three CesA genes are required for assembly and function of a single cellulose synthase complex [119]. Genes, such as CesA and korrigan (kor), the membrane-localized b-1,4-glucanase, are directly involved in cellulose synthesis and modification [65, 85]. The Arabidopsis mutant ixr1-2, which bears a mutation within the conserved C-terminal transmembrane region of CesA3, has lower cellulose crystallinity. The cell walls of the transgenic plants had higher conversion rates to fermentable sugars than wild type, with only a slight decrease in growth rate [54]. It is suggested that the generation of the same amino acid change in CesA subunits (CesA4, 7 and 8) that modify the secondary cell wall may further increase the efficiency of biomass conversion in biofuel feedstocks [54] .

Korrigan is part of the cellulose synthase complex and is a key membrane-bound endoglucanase required for cellulose biosynthesis [119] . Korrigan is believed to cleave stacking glucan chains of microfibrils at the beginning of their synthesis. In this way, it promotes accurate synthesis [122]. Overexpression of Arabidopsis kor­rigan (AtKOR) in hybrid poplar resulted in reduced crystallinity without any significant effect on cell wall composition and plant growth [77] .

Cellulose crystalline structure can be further modified post-synthesis in the cell wall. For example, the extracellular fungal enzyme, cellobiose dehydrogenase (CDH) [136], enhances cellulose degradation in the presence of cellulase mixtures directly [7] by preventing recondensation of glycosidic bonds of cellulose chains nicked by endoglucanases [5], or indirectly by generating hydroxyl radicals in a Fenton type reaction [58]. CDH binding to cellulose is highly specific [59] and can lead to oxidation of highly crystalline as well as amorphous cellulose [ 16]. We hypothesized that the expression of CDH in plants cell walls can result in the dis­ruption of the microcrystalline lattice through the modification of cellobiose to cellobiono-(1,5)-lactone via reduction of hydrogen bonds responsible for the highly crystalline structure of cellulose [1]. It is still unclear if and how plant fitness and growth are affected by reductions in cellulose crystallinity.

James Weifu Lee

Abstract This chapter reports two inventions: designer proton-channel algae (US Patent No. 7,932,437 B2) and designer switchable photosystem-II algae (US Patent No. 7,642,405 B2), for more efficient and robust photobiological production of hydrogen (H2) from water. Use of these inventions could eliminate the following six technical problems that severely limit the yield of algal H2 production: (1) restric­tion of photosynthetic H2 production by accumulation of a proton gradient, (2) com­petitive inhibition of photosynthetic H2 production by CO2 , (3) requirement of bicarbonate binding at photosystem-II (PSII) for efficient photosynthetic activity, (4) competitive drainage of electrons by O2 in algal H2 production, (5) oxygen sensitivity of algal hydrogenase, and (6) the H2-O2 gas separation and safety issue. By eliminating these six technical problems that currently challenge those in the field, the designer algae approach could enhance photobiological production of hydrogen with a yield likely over ten times better than that of the wild-type.

1 Introduction

Photoautotrophic H2-producing microorganisms, including microalgae such as Chlamydomonas reinhardtii, have the potential to be a clean energy resource. In the algal system, H2 can be produced through hydrogenase-catalyzed reduction of pro­tons by the electrons generated from photosynthetic oxidation of water using sunlight energy, as illustrated in Fig. 1. The net result is photoevolution of H2 and

J. W. Lee (*)

Department of Chemistry and Biochemistry, Old Dominion University,

Physical Sciences Building, Room 3100B, 4402 Elkhorn Avenue, Norfolk, VA 23529, USA

Johns Hopkins University, Whiting School of Engineering, 118 Latrobe Hall,

Baltimore, MD 21218, USA

e-mail: jwlee@ODU. edu; JLee349@JHU. edu

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_20, 371

© Springer Science+Business Media New York 2013

O2 from H2O. Recently, there were a few interesting algal H2 production research efforts as mentioned in some of the recent review articles [1, 2]. For example, sulfur — deprivation with C. reinhardtii was explored in trying to improve algal H2 produc­tion [3, 4]. More recently, certain phylogenetic and molecular analyses were performed in H2-producing green algae [5, 6]. However, so far, there was essentially no truly tangible fundamental improvement on the rate and the yield of algal photo­biological H2 production. Algal H2 production remains impractical with very lim­ited yield; and the solar-to-hydrogen energy conversion currently is still less than

0. 1% [7], which clearly is not commercially viable.

According to a recent analysis, the most urgent and challenging technical barri­ers are the four trans-thylakoidal proton gradient-associated physiological problems

[8] that seriously limit the rate and the yield of algal photobiological H2 production: (1) accumulation of a proton gradient across the algal thylakoid membrane, (2) competition from carbon dioxide fixation, (3) requirement for bicarbonate binding at photosystem-II (PSII) for efficient photosynthetic activity, and (4) competitive drainage of electrons by molecular oxygen.

This chapter reports two inventions: designer proton-channel algae (US Patent No. 7,932,437 B2) and designer switchable photosystem-II algae (U. S. Patent Number: US 7,642,405 B2), which may enable efficient and robust photobiological production of hydrogen with an enhanced yield likely more than ten times better than that of the wild-type. The first invention (designer proton-channel algae [9] ) uses a highly innovative “one stone killing four birds” approach to simultaneously eliminate all the four — rans-thylakoidal proton gradient-associated physiological problems by genetic insertion of a polypeptide proton channel into algal thylakoid membrane using synthetic biology; whereas, the second invention [10] is on creat­ing designer switchable photosystem-II algae for robust photobiological production of hydrogen from water splitting, which can eliminate all the following three molec­ular oxygen (O2)-associated technical problems: (4) competitive drainage of elec­trons by molecular oxygen, (5) oxygen sensitivity of algal hydrogenase, and (6) the H2-O2 gas separation and safety issue.

The following describe the four t rans — thylakoidal proton gradient-associated physiological problems that may be solved by the use of the designer proton-chan­nel algae invention for enhanced photoautotrophic hydrogen production:

1. Accumulation ofaproton gradient across the algal thylakoid membrane. For each hydrogen molecule (H2) produced by photosynthesis, six protons are trans­located across the algal thylakoid membrane (Fig. 1). Since the membrane has a limited permeability to protons, photosynthetic electron transport will result in accumulation of protons inside the lumen of the thylakoids, with no mechanism for dissipation. The resulting static back-proton gradient seriously impedes the electron transport, thus limiting the rate of H2 production. This phenomenon can be explained by the difference between the Calvin-cycle CO — — fixation process and the ferredoxin (Fd)/hydrogenase H2 production pathway. In photosynthetic CO2 fixation, this proton gradient across the thylakoid membrane is used by the coupling factor CF0CF1 complex to drive the formation of ATP that is required by the Calvin cycle. However, the Fd/hydrogenase H2 production pathway does not consume ATP. Consequently, under the conditions of H2 photoevolution where the consumption of ATP by the Calvin cycle stops due to the absence of CO2, the CF0CF1-mediated conduction of protons from the lumen to the stroma will quickly become limiting because of the accumulation of ATP and the shutdown of ATP synthase activity. As a result, photosynthetic H2 production quickly results in an increased proton gradient across the thylakoid membrane that has no mechanism for dissipation. The static back-proton gradient seriously impedes the electron transport, thus limiting the rate of H2 production, since electron transport from water through photosystem-II (PSII), plastoquinone (PQ), the cytochrome b/f (Cyt b/f) complex, plastocyanin (PC), and photosystem I (PSI) to the Fd/hydrogenase pathway is coupled with proton translocation across the thylakoid membrane, as illustrated in Fig. 1.

2. Competitionfrom carbon dioxide fixation. Carbon dioxide (CO2) fixation by the Calvin cycle can compete with the Fd/hydrogenase H2 production pathway for photosynthetic electrons. This has been demonstrated in experimental studies [11]. Steady state H2 photoevolution was inhibited to nearly zero by injecting 58 ppm of CO2 . Therefore, increasing the photosynthetic H2 production would require a nearly CO2-free environment. However, lowering the CO2 concentra­tion would countermand the following factor.

3. Requirementfor bicarbonate binding at PSllfor efficientphotosynthetic activity. Experimental studies [12] have demonstrated that adding bicarbonate (HCO3-) to depleted samples can result in a six to sevenfold stimulation of PSII electron — transport activity. Because CO2 and HCO3- are interchangeable in aqueous medium, removal of CO2 can lead to depletion of HCO3 -, and thereby reduce PSII electron-transport activity. This presents a dilemma when one tries to lower the CO2 concentration in order to increase H2 production.

4. Competitive drainage ofelectrons by molecular oxygen. In a previous study [13] performed by the author, a new molecular oxygen (O2 ) sensitivity in algal H2 production was discovered that is distinct from the O2 sensitivity of hydrogenase per se and more significant. This is likely due to the background O2 which appar­ently serves as a terminal electron acceptor through RuBisco’s oxygenase activ­ity at the Calvin cycle, in competition with the H. — production pathway. Our experimental studies demonstrated that photosynthetic H2 production can be inhibited by an O2 concentration as low as about 500-1,000 ppm, while the algal hydrogenase is still active at an O2 concentration as high as 5,000 ppm. This indicates that the drainage of electrons by O2 in competition with the Fd/hydro — genase H2 production pathway is a serious problem that must be solved in order for H2 production to work efficiently.

According to one of the various embodiments, a designer Calvin-cycle-channeled pathway is created that takes the Calvin-cycle intermediate product, 3-phospho — glycerate, and converts it into 3-methyl-1-pentanol, 4-methyl-1-hexanol, and/or

5- methyl-1-heptanol by using, for example, a set of enzymes consisting of (as shown with the numerical labels 34, 35, 03-05, 36-39, 53-55, 39′-43′, 39′-43′, and 39"-43" in Fig. 9): NADPH-dependent glyceraldehyde-3-phosphate dehydrogenase 34, NAD-dependent glyceraldehyde-3-phosphate dehydrogenase 35, phosphoglycerate mutase 03, enolase 04, pyruvate kinase 05, citramalate synthase 36, 2-methylmalate dehydratase 37, 3-isopropylmalate dehydratase 38, 3-isopropylmalate dehydroge­nase 39, acetolactate synthase 53, ketol-acid reductoisomerase 54, dihydroxy-acid dehydratase 55 , designer isopropylmalate synthase 40′, designer isopropylmalate isomerase 41′, designer 3-isopropylmalate dehydrogenase 39′, designer 2-keto acid decarboxylase 42′, short-chain alcohol dehydrogenase 43′, designer isopropylmalate synthase 40", designer isopropylmalate isomerase 41", designer 3-isopropylmalate dehydrogenase 39", designer 2-keto acid decarboxylase 42", and designer short — chain alcohol dehydrogenase 43".

According to another embodiment, a designer Calvin-cycle-channeled pathway is created that takes the intermediate product, 3-phosphoglycerate, and converts it into 3-methyl-1-pentanol, 4-methyl-1-hexanol, and/or 5-methyl-1-heptanol by using, for example, a set of enzymes consisting of (as shown with the numerical labels 34, 35, 03, 04, 45-55, 39′-43′, 39′-43′, and 39"-43" in Fig. 9): NADPH — dependent glyceraldehyde-3-phosphate dehydrogenase 34, NAD-dependent glycer — aldehyde-3-phosphate dehydrogenase 35, phosphoglycerate mutase 03, enolase 04, phosphoenolpyruvate carboxylase 45, aspartate aminotransferase 46, aspartokinase 47, aspartate-semialdehyde dehydrogenase 48, homoserine dehydrogenase 49, homoserine kinase 50 , threonine synthase 51 , threonine ammonia-lyase 52, ace — tolactate synthase 53, ketol-acid reductoisomerase 54, dihydroxy-acid dehydratase 55, designer isopropylmalate synthase 40′, designer isopropylmalate isomerase 41′, designer 3-isopropylmalate dehydrogenase 39′, designer 2-keto acid decarboxylase 42′, short-chain alcohol dehydrogenase 43′, designer isopropylmalate synthase 40 , designer isopropylmalate isomerase 41", designer 3-isopropylmalate dehydroge­nase 39", designer 2-keto acid decarboxylase 42", and designer short-chain alcohol dehydrogenase 43".

These pathways (Fig. 9) are similar to those of Fig. 8, except they use acetolac — tate synthase 53, ketol-acid reductoisomerase 54, dihydroxy-acid dehydratase 55 as part of the pathways for production of 3-methyl-1-pentanol, 4-methyl-1-hexanol, and/or 5-methyl-1-heptanol. They all share a common feature of using an NADPH- dependent glyceraldehyde-3-phosphate dehydrogenase 34 and an NAD-dependent glyceraldehyde-3-phosphate dehydrogenase 35 as a mechanism for NADPH/NADH conversion to drive production of 3-methyl-1-pentanol, 4-methyl-1-hexanol, and/or

5- methyl-1-heptanol through a designer Calvin-cycle-channeled pathway in combi­nation with a hydrocarbon chain elongation pathway (40′, 41′, 39′). This embodi­ment also takes the advantage of the broad substrate specificity (promiscuity) of

2- isopropylmalate synthase 40 , isopropylmalate isomerase 41, 3-isopropylmalate dehydrogenase 39, 2-keto acid decarboxylase 42, and short-chain alcohol dehydro­genase 43 so that they can also serve as: designer isopropylmalate synthase 40′, designer isopropylmalate isomerase 41′, designer 3-isopropylmalate dehydrogenase 39′, designer 2-keto acid decarboxylase 42′, and short-chain alcohol dehydrogenase 43′, designer isopropylmalate synthase 40", designer isopropylmalate isomerase 41", designer 3-isopropylmalate dehydrogenase 39", designer 2-keto acid decar­boxylase 42", and designer short-chain alcohol dehydrogenase 43".

The net results of the designer photosynthetic NADPH-enhanced pathways (Fig. 9) in working with the Calvin cycle are production of 3-methyl-1-pentanol (CH3CH2CH(CH3)CH2CH2OH), 4-methyl-1-hexanol (CH3CH2CH(CH3)CH2CH2 CH2OH), and 5-methyl-1 — heptanol (CH3CH2CH(CH3)CH2CH2CH2CH2OH) from carbon dioxide (CO2) and water (H2O) using photosynthetically generated ATP and NADPH according to the following process reactions:

6CO2 + 7H2O ^ CH3CH2CH(CH3)CH2CH2OH + 9O2 (14)

14CO2 + 16H2O ^ 2CH3CH2CH(CH3)CH2CH2CH2OH + 21O2 (15)

8CO2 + 9H2O ^ CH3CH2CH(CH3)CH2CH2CH2CH2OH + 12O2 (16)

The proportion of greenhouse gas (CHG) emissions from each material and energy used is shown in Fig. 5. The main contribution came from transesterification reac­tion for J. curcas and oil extraction for oil palm.

Biodiesel production from palm oil has bigger GHG emission than that from Jatropha oil.

Infrastucture, energy, raw materials, auxialary materials

Fig. 4 The system boundary of Jatropha biodiesel production

0 0.5 1.0 1.5 2.0 2.5 (kg CO2)

Fig. 5 Comparison of life cycle GHG emissions of biodiesel production from palm oil and Jatropha oil based on material and energy used in each steps of life cycle [59]

Oil Extraction

Land Preparation and Cultivation

0 0.05 0.1 0.15 (POINT)

With roughly 61% of global bioethanol production, sugar crops such as sugar beet, sugar cane and molasses are the most widely used feedstocks for fermentation (Fig. 4). Corn comes second with roughly 20% of global production. However, production of bioethanol from these sources competes with the limited agricultural lands needed for food production [58]. Also both crops require more herbicides or nitrogen-based fertilizers than any other crop, thus have a high soil erosion potential [49]. High costs of pre-treatment, harvesting and transportation of these materials are some of the problems encountered with their utilization as raw materials for bioethanol production.

Lignocellulosic materials are alternative feedstocks that are being intensely explored for bioethanol production currently. The three major components of ligno — cellulose are cellulose, hemicellulose and lignin. Cellulose is a long-chain highly — uniform polymer of glucose monomers joined by b — linkages. Hemicellulose is another polysaccharide composed of many different sugar monomers, including xylose and mannose. Unlike cellulose, hemicellulose is not uniform in structure and tends to be more reactive. Both cellulose and hemicellulose can be fermented once their sugar monomers have been released through a process known as saccarification [52]. On the other hand, lignin is a polymer of phenylpropylene subunits and cannot be fermented. Lignin forms a crystalline protective structure around cellulose and

hemicellulose in the biomass and is very difficult to be biologically degraded. Such formation interferes with the saccarification, and hence fermentation, of cellulose and hemicellulose [36]. Therefore, lignocellulosic materials are more challenging to be used in fermentation. Moreover, the cost of ethanol production from these mate­rials has been reported to be relatively higher compared to other feedstocks [58].

Microalgal biomass is currently being explored as a potential feedstock for bio­ethanol production. Microalgal cells contain carbohydrates that can be used as car­bon sources for bioethanol production during fermentation, and this varies from strain to strain. Selection of the right strain is necessary in order to achieve the high­est bioethanol yield. Table 1 shows the carbohydrates and protein contents of differ­ent microalgal strains. The production of bioethanol from microalgae provides many advantages: (1) Microalgal cells have a very short harvesting cycle (~1-10 days) compared to other feedstocks (harvest once or twice a year) [54], thus poten­tially providing enough supplies to meet bioethanol production demands in the future. (2) Compared with microalgal biodiesel, the process for producing microal­gal bioethanol is more energy effective due to the utilization of simpler equipment or machinery to start up. (3) No delignification process is required since microalgae contain no lignin, thus reducing the production cost of bioethanol [13]. (4) The CO2 by-product after the fermentation process can be recycled to the microalgal cultiva­tion system, thus emitting minimal amount of carbon dioxide to the environment. (5) The leftover biomass (containing protein residue) after the fermentation process can be utilized as a fertilizer or an animal feed [27]. As such, the process of bioetha­nol production from microalgae is sustainable, environmentally friendly and cost efficient. The production of bioethanol from microalgae consists of four main stages: pre-treatment of microalgal biomass, hydrolysis, fermentation and product recovery. A more detailed outline of each stage is discussed in Sect. 6.

The various embodiments further teach how the designer organisms may be used with a photobioreactor and an ethanol-separation-harvesting system for photosyn­thetic production of ethanol (CH3CH2OH) and O2 directly from CO2 and H2O using sunlight (Figs. 9-11). There are a number of embodiments on how the designer organisms may be used for photobiological ethanol production. One of the preferred embodiments is to use the designer organisms for direct photosynthetic ethanol pro­duction from CO2 and H2O with a photobiological reactor and ethanol-harvesting (distillation) system (Fig. 9), which includes a specific operational process described as a series of the following steps: (a) Growing a designer transgenic organism pho- toautotrophically in minimal culture medium using air CO2 as the carbon source under aerobic (normal) conditions before inducing the expression of the designer ethanol-production-pathway genes; (b) When the designer organism culture is grown and ready for ethanol production, sealing or placing the culture into a specific condition, such as an anaerobic condition that can be generated by removal of O2 from the photobiological reactor, to induce the expression of designer ethanol- production genes; (c) When the designer ethanol-production-pathway enzymes are expressed and inserted into the stroma region of the designer organism’s chloroplast,

supplying visible light energy such as sunlight for the designer-genes-expressed cells to work as the catalysts for photosynthetic ethanol production from CO2 and H2O; (d) Harvesting the ethanol product by any method known to those skilled in the art. For example, harvesting the ethanol product from the photobiological reactor by a combination of membrane filtration and ethanol-distillation techniques and flexibly collecting the O2 gas product from the reactor.

The above process to use the designer organisms for photosynthetic CH3CH2OH and O2 production from CO2 and H2O with a biological reactor and ethanol-harvesting (distillation) and gas product separation and collection system can be repeated for a plurality of operational cycles to achieve more desirable results. Any of the steps (a) through (d) of this process described above can also be adjusted in accordance of the invention to suit for certain specific conditions. In practice, any of the steps (a) through (d) of the process can be applied in full or in part, and/or in any adjusted combination as well for enhanced photobiological ethanol production in accordance of this invention.

The sources of CO2 that can be used in this process include, but not limited to, industrial CO2, (bi)carbonates, and atmospheric CO2. For an example, flue-gas CO2 from fossil fuel-fired and/or biomass-fired industrial facilities can be fed through a pipeline into a photobiological reactor in this process as illustrated in Fig. 9.

The industrial facilities that can generate CO2 supplies for the designer photosynthetic ethanol-production process include (but not limited to): coal-fired power plants, iron and steelmaking industries, cement-manufacturing plants, petroleum refinery facilities, chemical fertilizer production factories, biomass-fired and/or fossil fuel-fired ethanol distillation/separation facilities, biomass-pyrolysis processes, smokestacks, fermentation bioreactors, biofuel-refinery facilities, and combinations thereof.

Alternatively, this designer photobiological ethanol-production process can also use the CO2 in the environment and from the atmosphere (Figs. 10 and 11) as well. Gaseous COr, dissolved COr, bicarbonate, and carbonates can all be used by the designer-organism photobiological ethanol-production technology.

This embodiment is illustrated in more details here using designer algae as an example. As described above, designer algae of the present invention, such as the one that contains a set of designer HydA1 promoter-controlled designer ethanol-produc­tion-pathway genes, can grow normally under aerobic conditions by autotrophic pho­tosynthesis using air CO2 in a manner similar to that of a wild-type alga. The designer algae can grow also photoheterotrophically using an organic substrate as well.

In a preferred embodiment, a designer alga is grown photoautotrophically using air CO2 as the carbon source under the aerobic conditions as shown in Fig. 4a in a minimal medium that contains the essential mineral (inorganic) nutrients. No organic substrate such as acetate is required to grow a designer alga under the normal conditions before the designer photosynthetic ethanol-production genes are expressed. Most of the algae can grow rapidly in water through autotrophic photosynthesis using air CO2 as long as there are sufficient mineral nutrients. The nutrient elements that are commonly required for algal growth are: N, P, and K at the concentrations of about 1-10 mM, and Mg, Ca, S, and Cl at the concentrations of about 0.5-1.0 mM, plus some trace elements Mn, Fe, Cu, Zn, B, Co, Mo among others at pM concen­tration levels. All of the mineral nutrients can be supplied in an aqueous minimal medium that can be made with well-established recipes of algal culture media using water and relatively small of inexpensive fertilizers and mineral salts such as ammo­nium bicarbonate (NH4HCO3) (or ammonium nitrate, urea, ammonium chloride), potassium phosphates (K2HPO4 and KH2PO44 , magnesium sulfate heptahydrate (MgSO47H2O), calcium chloride (CaCl2), zinc sulfate heptahydrate (ZnSO47H2O), iron (II) sulfate heptahydrate (FeSO47H2O), and boric acid (H3BO3), among others. That is, large amounts of designer algae cells can be inexpensively grown in a short period of time because, under aerobic conditions such as in an open pond, the designer algae can photoautotrophically grow by themselves using air CO2 as rapidly as their wild-type parental strains. This is a significant feature (benefit) of the invention that could provide a cost-effective solution in generation of photoactive biocatalysts (the designer photosynthetic ethanol-producing algae) for renewable solar energy production.

When the algal culture is grown and ready for ethanol production, the grown algal culture is sealed or placed into certain specific conditions, such as anaerobic conditions that can be generated by removal of O2 from the sealed photobiological reactor (Fig. 9) , to induce the expression of designer photosynthetic ethanol — production-pathway genes. When the designer ethanol-production-pathway enzymes are expressed and inserted into the stroma region of algal chloroplast, visible light energy such as sunlight is supplied for the designer-genes-expressing algal cells to work as the catalysts for photosynthetic ethanol production from CO2 and H) O. When the designer genes are expressed, the algal cells can essentially become efficient and robust “green machines” that are perfect for photosynthetic production of ethanol (CH3CH2OH) and O2 from CO2 and H2O. The ethanol product from the algal photobiological rector can be harvested by a combination of membrane filtration and ethanol-distillation techniques.

Photosynthetic production of CH3CH2OH and O2 directly from CO2 and H2O in accordance with the present invention can, in principle, have high quantum yield. Theoretically, it requires only 24 photons to produce a CH)CH2OH and 3O) from water and carbon dioxide by this mechanism. The maximal theoretical sunlight-to — ethanol energy efficiency by the process of direct photosynthetic ethanol production from CO2 and H2O is about 10%, which is the highest possible among all the biological approaches. Consequently, this approach has great potential when implemented properly with an algal reactor and ethanol-oxygen-harvesting system (Fig. 9).

The above process to use the designer algae for photosynthetic production of CH3CH2OH and O2 from CO2 and H2O with an algal reactor and an ethanol-harvesting (distillation) and gas product separation and collection system (Fig. 9) can be repeated for a plurality of operational cycles to achieve more desirable results.

Another feature is that the designer switchable ethanol-production organism provides the capability for repeated cycles of photoautotrophic culture growth under normal aerobic conditions with a manner similar to that of a wild type (Fig. 1) and efficient photobiological production of ethanol (Fig. 2) when the designer ethanol — production pathway is switched on by an inducible promoter (such as hydrogenase promoter) at certain specific inducing conditions (such as under anaerobic conditions) in a bioreactor (Fig. 9). For example, the switchable designer alga with designer hydrogenase promoter-controlled ethanol-production genes contains normal mito­chondria, which uses the reducing power (NADH) from organic reserves (and/or exogenous substrates, such as acetate) to power the cell immediately after its return to aerobic conditions. Therefore, when the algal cell is returned to aerobic condi­tions after its use under anaerobic conditions for production of ethanol, the cell will stop producing ethanol-production-pathway enzymes and start to restore its normal photoautotrophic capability by synthesizing normal functional chloroplast. Consequently, it is possible to use this type of genetically transformed organism for repeated cycles of photoautotrophic culture growth under normal aerobic conditions (Fig. 1) and efficient production of ethanol under anaerobic conditions (Fig. 2) in an anaerobic reactor (Fig. 9). That is, this photobiological ethanol-production technology can be operated for a plurality of operational cycles by rejuvenating the used culture under aerobic conditions and recyclably using the rejuvenated algal culture under ethanol-producing conditions to achieve more desirable results. Optionally, this photobiological ethanol-production technology is operated continuously by circulating rejuvenated algal culture from an aerobic reactor into the anaerobic reactor while circulating the used algal culture from the anaerobic reactor (after its use for ethanol production) into the aerobic reactor for rejuvenation by synthesizing normal functional chloroplasts through photosynthetic CO2 fixation and photoautotrophic growth.

Some of the designer organisms could grow photoautotrophically even with the ethanol-production pathway(s) switched on. Whether or how fast a designer organ­ism could grow under the ethanol-producing conditions may depend on its genetic background and how much of the Calvin cycle products are still available for cell growth after use by the designer ethanol-production pathway(s). Designer organisms that can, under the ethanol-producing conditions, maintain essential cellular functions with an appropriate growth rate can also be used for continuous photobiological production of CH3CH2OH and O2 from CO2 and H2O with a bioreactor and an ethanol­harvesting (distillation) system (Figs. 9 and 10).

There are additional ways that the switchable designer organisms can be used. For example, the used designer algal culture from a photobiological ethanol — production reactor does not have to be circulated back to a culture-growth reactor. Instead, the used algal culture is taken out to be used as fertilizers or biomass feed stocks for other processing because the photoautotrophic growth of the switchable designer alga in a culture-growth reactor (Fig. 11, left) is capable of continuously supplying algal cells to a photobiological ethanol-production reactor for the biofuel production (Fig. 11, right). This embodiment is, especially, helpful to using some of the designer organisms that can grow photoautotrophically only before but not after the ethanol-production pathway(s) is switched on. For example, as illustrated in Fig. 11, by keeping a continuously growing culture of a designer alga (that can grow photoautotrophically only before the ethanol-production pathway(s) is switched on) in a culture-growth reactor, it can provide continuous supplies of grown algal cells for use in a photobiological ethanol-production reactor. This approach makes it pos­sible to use those designer organisms that can grow only before the ethanol-production pathway(s) is switched on for photobiological ethanol production as well.

Because of various reasons, some of the designer ethanol-production organisms could grow only photohetrotrophically or photomixotrophically but not photoauto — trophically. Use of a culture-growth reactor as illustrated in Fig. 11 can also grow this type of designer ethanol-production organisms photohetrotrophically or photo — mixotrophically using organic substrates including, but not limited to, sucrose, glucose, acetate, ethanol, methanol, propanol, butanol, acetone, starch, hemicellulose, cellu­lose, lipids, proteins, organic acids, biomass materials, and combination thereof. The so-grown culture can also be supplied to a photobiological ethanol-production reactor for induction of the designer pathways for ethanol production as illustrated in Fig. 11. This modified embodiment on culture growth makes it possible to use those designer organisms that can grow only photohetrotrophically, or photomixotrophically also for photobiological ethanol production as well.

For certain specific designer organisms with designer nitrate reductase (Nial) promoter-controlled ethanol-production-pathway genes, the above photobiological reactor process may be further adjusted to achieve more beneficial results. For example, a designer alga that contains Nial promoter-controlled ethanol-production­pathway genes can grow normally in a culture medium with ammonium (but no nitrate) by autotrophic photosynthesis using air CO2 in a manner similar to that of a wild-type alga. This is because the expression of the ethanol-production-pathway genes in this designer organism will be turned on only in the presence of nitrate as desired owning to the use of nitrate reductase (Nial) promoter in controlling the designer pathway expression. A significant feature of the designer organisms with Nial promoter-controlled ethanol-production-pathway genes is that the expression of the designer ethanol-production pathways can be induced by manipulating the concentration levels of nitrate (NO3-) relative to that of ammonium (NH4+) in the culture medium without requiring any anaerobic conditions. That is, the expression of the designer ethanol-production pathway(s) can be induced under both aerobic and anaerobic conditions. This enables the designer photobiological ethanol — production process to operate even under aerobic conditions using atmospheric CO2 (Fig. 10). Likewise, this type of designer organisms with Nial promoter-controlled ethanol-production-pathway genes can grow photoautotrophically both under aero­bic and anaerobic conditions as well. Therefore, as a further embodiment, the operational process of using designer organism with nitrate reductase (Nial) promoter-controlled ethanol-production-pathway genes is adjusted to the following:

(a) Growing a designer transgenic organism photoautotrophically in minimal culture medium in the presence of ammonium (NH4+) but no nitrate (NO3-) before inducing the expression of the designer ethanol-production-pathway genes; (b) When the designer organism culture is grown and ready for ethanol production, adding nitrate (NO3-) fertilizer into the culture medium to raise the concentration of nitrate (NO3-) relative to that of ammonium (NH4+) to induce the expression of designer ethanol — production-pathway genes; (c) When the designer ethanol-production-pathway enzymes are expressed and inserted into the stroma region of the designer organism’s chloroplast, supplying visible light energy such as sunlight for the designer-genes — expressed cells to work as the catalysts for photosynthetic ethanol production from CO2 and H2O; (d) Harvesting the ethanol product from the photobiological reactor by a combination of membrane filtration and ethanol-distillation techniques.

Furthermore, the harvesting of ethanol product from photobiological liquid culture media can be achieved also through the use of another invention: greenhouse distil­lation that is disclosed in details in PCT International Patent Application Publication No. WO 2009/105714 A2. Briefly, when sunlight is used to drive photobiological ethanol production, it also generates waste heat in the liquid culture medium. Through combined use of a designer photosynthetic organism culture with a green­house distillation system, the waste solar heat associated with the photobiological ethanol-production process can be utilized in vaporizing the product ethanol (and water) for harvesting ethanol and producing freshwater by fractional greenhouse distillation. Consequently, this approach can help improve the total process energy efficiency with minimal cost and high sunlight utilization efficiency [14].

In addition to ethanol production, it is also possible to use a designer organism or part of its designer ethanol-production pathway(s) to produce certain intermediate products including: acetaldehyde, pyruvate, phosphoenolpyruvate, 2-phosphoglyc- erate, 1,3-diphosphogly cerate, glyceraldehye-3-phosphate, dihydroxyacetone phosphate, fructose-1,6-diphosphate, fructose-6-phosphate, glucose-6-phosphate, and glucose-1-phosphate. Therefore, a further embodiment comprises an additional step of harvesting the intermediate products that can be produced also from an induced transgenic designer organism. The production of an intermediate product can be selectively enhanced by switching off a designer-enzyme activity that catalyzes its consumption in the designer pathways. The production of a said inter­mediate product can be enhanced also by using a designer organism with one or some of designer enzymes omitted from the designer ethanol-production pathways. For example, a designer organism with the alcohol dehydrogenase or pyruvate decarboxylase omitted from the designer pathway of Fig. 4b may be used to produce acetaldehyde or pyruvate, respectively.

Acknowledgments This work was supported in part by a Seed Money project at Oak Ridge National Laboratory (ORNL) which is managed by UT-Battelle, LLC, for DOE under contract no. DE-AC05-00OR22725. The author wishes to thank also the ORNL/Technology Transfer Office and the Laboratory for the nice support.