The life cycle energy analysis of Jatropha biodiesel production was conducted by evaluating direct energy input (such as electricity, diesel, gasoline, fuel oil, palm fiber, palm shell, etc.) and indirect energy input (energy accumulated in fertilizers, agrochemicals, and chemical production, excluding equipment and machinery used in the processes). The net energy value (NEV) and the net energy ratio (NER) can be estimated. The NEV is a measure of the energy gain or loss from the biodiesel used, which is defined as the energy content of the biodiesel minus the nonrenew­able energy used in the life cycle of the biodiesel production [63]. The NER is a ratio of energy output to total energy input for the life cycle of the product [64].

Prueksakorn and Gheewala [64] calculated the energy consumption in every pro­cess in producing 1 kg of Jatropha biodiesel. The energy analysis results of the present situation of Jatropha biodiesel production compared to palm oil methyl ester is shown in Table 5. The results show that the selected biodiesel production process determines energy efficiency and environmental impacts.

9 Conclusions

High cost of biodiesel production is the major impediment to its large-scale com­mercialization. Methods to reduce the production cost of Jatropha biodiesel must be developed. One way to reduce production costs is to increase the added values of protein-rich Jatropha seedcakes, by-product of oil extraction, through detoxification process. The development of integrated biodiesel production process and the detoxification process results in two products, namely biodiesel and protein-rich seedcakes that can be used for animal feed. Assuming that an average seed yield on land of 5 tons/ha/year (2 tons/acre/year) could be achieved, the estimated theoretical maximum yield of biodiesel would be 750 kg/acre/year and seedcake products would be 500 kg/acre/year.

Since the cost and efficiency of the selected process will be closely correlated with the production for a long time and affect the capital and operating costs and the environmental load of the product, selecting an appropriate process for the biodiesel production is a critical decision. There are still many future potential improvement of biodiesel production of J. curcas. These include (1) development of better and cheaper oil extraction and postreaction processing methods; (2) development of better and cheaper catalysts; (3) improvements in current technology for producing high-quality biodiesel with cheaper cost production; (4) development of technology to use methanol/ethanol in in situ extraction and transesterification; (5) develop­ment of technique to improve fuel stability of Jatropha biodiesel; (6) conversion of by-products, such as glycerol and seedcake to useful and value-added products, such as methanol and ethanol or glycerol tert-butyl ether (GTBE); and (7) develop­ment of integrated Jatropha biodiesel processing and detoxification process.

LCA has become an important decision-making tool for promoting alternative fuels because it can systematically analyze the fuel life cycle in terms of energy efficiency and environmental impacts. LCA analysis shows that the selected biodie­sel production process determines energy efficiency and environmental impacts of Jatropha biodiesel production.

5.1.1 Basic Principles

During lipid extraction, microalgal cells (either as a wet paste or dried biomass) are exposed to a non-polar organic solvent, such as hexane or chloroform, which inter­acts with the neutral lipid molecules and overcomes their weak hydrophobic inter­actions with other biomolecules. As a result, the neutral lipid molecules desorb from their cellular matrix and dissolve in the solvent [30, 39]. The remaining cell residue is then separated from the solvent via solid-liquid separation methods. The solvent is evaporated to yield dry lipid extract.

However, some neutral lipids are strongly linked via hydrogen or electrostatic bonds to other biomolecules, such as proteins and polar lipids, in the cell membrane. These bonds are too tough to be disrupted by the non-polar solvent alone and require the presence of a more reactive solvent for their destruction [30, 39]. A small amount of polar solvent (such as methanol or isopropanol) is thus added together with the non-polar solvent (such as hexane or chloroform) to facilitate the extraction of these membrane-associated neutral lipids. Unfortunately, the addition of a polar co-solvent also leads to the increased co-extraction of undesirable polar lipids.

When using a non-polar/polar solvent mixture (such as hexane/isopropanol or chloroform/methanol), both solvents are added to the microalgal cells (either as a wet paste or dried biomass) in the desired ratio at the same time. Upon removal of cell debris via solid-liquid separation, biphasic separation of the solvent mixture is induced by equivolume addition of the non-polar solvent (hexane for hexane/isopro­panol mixture and chloroform for chloroform/methanol mixture) and water. Once the solvent mixture separates into two layers, the lipids (both neutral and polar frac­tions) will partition in the organic phase (a mixture of non-polar solvent and polar solvent), while the aqueous phase (a mixture of water and polar solvent) will contain co-extracted non-lipid contaminants (proteins and carbohydrates) [30, 39] . The organic phase is collected and the solvent is evaporated to obtain dry lipid extract.

5.1.2 Selection of Organic Solvents

In addition to satisfying the aforementioned criteria for the lipid extraction method, the selected organic solvents must be relatively cheap, relatively non-toxic, able to form two-phase with water for the removal of co-extracted non-lipids and volatile for recovery from the lipid extract [22, 39] . Furthermore, the solvent molecules must be sufficiently small so that permeation through the microalgal cell wall can be easily achieved.

Chloroform/methanol (2:1 v/v) is the most commonly used solvent mixture for laboratory-scale lipid extraction from any living tissue. The system uses residual endogenous water as a ternary component to enhance the extraction of polar lipids and does not require cells to be completely dried. Upon removal of the cell debris, the solvent mixture equilibrates with the dissolved lipids and separates into 2 phases upon addition of chloroform and water. The lower organic phase (chloroform with some methanol) contains most of the lipids while the upper aqueous phase (water with some methanol) contains most of the non-lipids [39]. Even though the extrac­tion process using this solvent mixture is fast and quantitative, chloroform is a highly toxic solvent whose large-scale usage is environmentally unattractive. Since the method was originally developed by Folch et al. [23] for the isolation of total lipids from brain tissues, its effectiveness in extracting lipids from microalgal cells still needs further assessment.

Hexane/isopropanol (3:2 v/v) works in a similar fashion as the chloroform/meth — anol mixture. Upon biphasic separation, the upper organic phase (hexane with some isopropanol) contains most of the lipids while the lower aqueous phase (water with some isopropanol) contains most of the non-lipids. This solvent mixture is quickly replacing chloroform/methanol as a laboratory favourite due to its lower toxicity and its higher selectivity towards neutral lipids [26, 33, 46]. However, the hexane/ isopropanol mixture has been reported to yield a lower lipid recovery than the chlo — roform/methanol mixture when applied to microalgal cells [33] .

Even though alcohols, such as butanol and ethanol, are cheap and highly volatile, their use as pure lipid extraction solvents is limited due to their lower affinities towards neutral lipids. In a study conducted by Lee et al. [33], the performances of five different solvent mixtures during lipid extraction from bead-beaten Botryococcus braunii were evaluated (Fig. 7) . Chloroform/methanol was found to be the best solvent mixture with a lipid yield of ~0.29 g/g dried microalgae, while all dichloro — ethane-based mixtures seemed rather ineffective.

Sandy soils in the coastal plain of South Carolina have inherently low soil fertility and a meager capacity to retain nutrients. Increased levels of SOC are regarded as an important determent to improve their fertility. Organic carbon compounds

returned as crop residues to sandy soils are temporal; a longer lasting solution is need. Therefore, it would be sensible to supplement sandy soils with biochar. This is not a new concept, but has been practiced by Amerindian populations for a long period of time [63]. In fact, it is arguably the starkest example of improving impov­erished Amazonian soils. In this region, the inhabitants stock piled char-like mate­rial on red-colored, infertile soils to convert them into a dark earth colored soil called “terra preta do Indio” [45, 95, 100] . Today, large amounts of C supplied through biochar additions to the Terra Preta soils have lasted for thousands of years after they were deserted [61] . In fact, Glaser et al. [44] reported that as much as 250 Mg C ha-1 has been sequestered in the Terra Preta as compared to 100 Mg C ha-1 typically measured in surrounding untreated soils. The message is apparent that biochars applied to the Terra Preta soils improved their fertility while also supplying C in recalcitrant forms that have lasted for several thousand years.

Building on the fertility gains by applying biochars to Terra Preta soils, let us establish the Norfolk’s Ap low fertility as the target issue to improve (Table 4). The next step would be selection of a feedstock and pyrolysis conditions (Fig. 11) that produces a biochar with properties chosen to compensate for these targeted (pH, SOC, N, P, etc.) problems. Among the biochar properties shown in Table 1, peanut hulls and poultry litter biochar contain greater N, P contents, and would act as a liming agent because of their alkaline pH. Biochars produced from the remainder of the feedstocks contain lesser amounts of nutrients or are not as alkaline. So, a logi­cal choice would be to use peanut hull and poultry litter feedstock and the prefer­ence of pyrolysis temperature could be selected based on the desired biochars nutrient concentration or by its alkalinity. If more nutrients and a better liming agent are desired, then the biochars should be produced using a higher pyrolysis tempera­ture (>500°C; Table 1).

The biochar application rates to this example Norfolk Ap, however, should be carefully chosen to avoid causing excessive alkaline or macronutrient imbalances.

Because this Norfolk Ap sample has a low buffer capacity, gross chemical changes occur with high biochar applications. For example, Novak et al. [74] reported that intensive application of poultry litter biochar (40 Mg ha-1) to a Norfolk Ap resulted in high soil pH values (8-9.7) and excessively high Mehlich-1 extractable P concen­trations (1,280-1,812 kg ha-1). Under these conditions, the Norfolk Ap contained plant available P concentrations that were grossly in excess of soil plant P sufficiency levels [51]. These disproportionate P concentrations, if moved off-site, poses surface and ground water quality issues [17, 47]. Crops may also experience micronutrient deficiencies; micronutrients have low solubility at elevated soil pH levels [102].

Unwanted soil pH increases may be avoided by employing alternate feedstocks such as peanut hulls, pecan shells, hardwood, or pine chips because they contain modest N-P-K ratios and are not as alkaline (Table 1). It should be understandable that when applying biochar to soil, it is important to not create an additional prob­lem while attempting to solve the target soil problem. The impact of biochars pro­duced from these alternate feedstocks and under different pyrolysis conditions (high vs. low temperature) on the fertility of a Norfolk Ap was shown in Table 4. Both peanut hull (400 and 500°C) and hardwood biochars were added at 2% (w w“ 1, 40-44 Mg ha-1). The treatments were laboratory incubated for 4 months and were then leached monthly with water to simulate loss of nutrients due to rainfall and/or irrigation. All biochar treatments after 120 days of incubation significantly raised soil pH, SOC, and TN contents, CEC had mixed results, when compared to the control (Table 4). After 120 days of incubation, the CEC increases were not particu­larly large (<0.6 cmolc kg-1), but some of the increases were still significant. The Norfolk Ap fertility was increased because Mehlich 1 extractable P, K, Ca, and Mg were all significantly higher than the control (Table 4). Both peanut hull bio­chars caused the greatest increases in OC, TN, and K relative to the control. Both OC and Ca concentrations were increased after applying hardwood biochar; minimal improvement occurred in pH, TN, P, K, and Mg concentrations. These results imply that hardwood would be an appropriate feedstock for a biochar designed to improve SOC and Ca levels alone, without causing large upward shifts in soil pH. Unfortunately, hardwood did not improve other soil problems such as low N and P contents. If OC and N improvements were the target soil fertility issue, then peanut hull would be an appropriate feedstock and either pyrolysis temperature.

Water leaching of the treatments resulted in loss of K and some P, whereas mixed results were obtained for the other nutrients. Leaching of K is not unexpected in sandy soils; its monovalent charge causes it to be less attracted to cation exchange sites [102]. This Norfolk’s Ap fertility status was improved by employing a biochar with appropriately designed characteristics. We avoided using an ill-suited biochar (poultry litter) in this situation because prior laboratory soil incubation showed that would cause a negative soil legacy (e. g., excessive nutrient concentrations, alkaline pH values, etc. [75]) potentially resulting in crop productivity declines. Poultry lit­ter biochar has special chemical properties, such as high P and alkalinity, which may be useful as a fertilizer and lime source if their concentrations are diluted through blending with benign biochar (see Sect. 6).

Biochars applications are not just limited to infertile soils, but the technology can be applied to fertile, mid-western soils as a supplement for increased C sequestra­tion and to replace nutrients lost through plant uptake, erosion and leaching. Laird et al. [59] incubated a hardwood biochar produced by slow pyrolysis in an Iowa Mollisol (Typic Hapludoll) and reported significant increases in total N, OC and Mehlich 3 extractable P, K, Mg, and Ca concentrations. In a similar study, Laird et al. [60] reported that the same biochar reduced total N and dissolved P leaching from swine manure applied to this Mollisol. These results imply that hardwood biochar additions to a mid-western Mollisol can be an effective agricultural and environmental management option by improving fertility and minimizing nutrient leaching. The authors did not choose to investigate if other feedstocks and different pyrolysis temperatures could have resulted in biochars with designed characteristics to improve the biochars performance at modifying fertility and nutrient leaching.

Biomass fast pyrolysis is the summation of its major components fast pyrolysis. In addition to the cellulose, hemicellulose, and lignin, biomass usually contains some extractives which would also decompose during fast pyrolysis, making the pyrolytic products more complex.

The biomass fast pyrolytic pathways and subsequent product distribution will be influenced by many factors, including biomass composition, feedstock property, pyrolysis temperature, heating rate, pressure, pyrolysis reactor configuration, and a combination of these variables. The detailed effects of these factors can be found in previous studies, and will not be shown here.

Lignin is a complex and high molecular weight polymer of phenylpropane derivatives (p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol). The density of hydrothermal medium is found to be a key parameter in lignin decomposition. In hydrothermal reaction medium, most of the hemicelluloses and part of the lignin are solubilized below 200°C. Lignin fragments have high chemical reactivity. Part of these fragments again cross-links and re-condenses to form high molecular weight water insoluble products [9]. Recently, Fang et al. have proposed the reaction pathway (Fig. 10) for lignin in supercritical water [29] .

The reaction steps consist of four phases: oil phase, aqueous phase, gas phase, and a solid residue phase [6, 29]. Their study concluded that lignin can be completely dissolved and undergoes homogeneous hydrolysis and pyrolysis preventing further re-polymerization.

The following sections discuss about application of sub — and supercritical water/ hydrothermal medium’s properties for bioethanol, biocrude, biochar, and gaseous fuels production.

While significant progress have been made (particularly in Europe) to commercial­ize BTL plants to generate green fuel, there are several drawbacks in the use of biomass alone as a feedstock for either generation of power or the production of synthetic oil. There is no consistent supply of biomass to operate a commercial sized power or fuel plant. For fuel production, the maximum advantage of the econ­omy of scale requires 500,000 Bbl (barrels)/day of oil production. At the present time, the largest BTL (biomass-to-liquids) plant that may be built will most likely

Table 10 Advantages and disadvantages of use of coal and biomass mixture feed for gasification

[50-55]___________________________________________________________________

Advantages Disadvantages

Expensive feed preparation

Complex feed systems for co-gasification

Two separate feed injectors, versus single feed injector, may affect the gasifier performance Negative impact of the slagging behavior of the combined ash in the gasifier Additional complication to gas cleaning system More tar and oil formation in raw syngas can be a problem depending on the gasification technology

produce 50,000 Bbl/day [47-49]. Also, as discussed earlier, biomass is difficult to store, feed, and transport and requires significant and costly pretreatment. One method that is presently being pursued is to obtain syngas by the gasification of coal and biomass mixture. Once the syngas (with appropriate methane content) from such a mixture is produced, it can be easily used either for power generation or producing a variety of transportation fuel by means of FT synthesis.

The advantages and disadvantages of the co-gasification of coal and biomass mixture are briefly presented in Table 10 [50-55]. As given in this table, co-gasification provides many advantages to the production of the syngas. Pure bio­mass gasification process is limited to small scale, has high capital (fixed) cost, has lower thermal efficiency, and carries shut down risk. All of these are alleviated by the use of coal. The mixture of coal and biomass provides a stable and reliable feed supply that generates less carbon dioxide. Coal can be considered as the “fly wheel” that allows a continuous plant operation when biomass feedstock is not easily avail­able. Co-gasification reduces the cost associated with fossil fuel consumption although some types of biomass can add significant cost to the overall fuel production. These advantages provide more security and less risks for project financiers than the use of pure biomass and are likely to engender more positive public attitudes towards the use of coal for fuel generation.

Table 10 also presents some disadvantages of co-gasification process. As given in this table, feed preparation and complex feed systems can be expensive. Two sepa­rate feed injectors, versus a single feed injector, may affect the gasifier performance. The slagging behavior of combined ash of coal and biomass in the gasifier may have a negative impact [17, 50-55] . The gas cleaning system may have the additional complications due to a mixture of organic and inorganic impurities. Also, depend­ing on the gasification technology, generation of more tar or oil in the product gas may be problematic [34, 36-43].

In summary, when coal and biomass are fed together as a mixture into a gasifier, the process of feeding should be uniform, consistent, and one that allows easy
fluidization in any types of gasifier. New short residence time gasifiers prefer dry feeding because it allows maximum flexibility in the allowable operating conditions of the gasifiers. The dry feeding also allows some flexibility in varying the nature of the coal-biomass mixture feedstock. Biomass also needs to be prepared such that it forms a homogenous mixture with coal. A successful CBTL process will truly open up new possibilities for fuel production that can use a wide variety of coal and bio­mass, our two sources of energy that can clearly replace our demands on oil.

The results of CO2 selectivity shown in Fig. 7 only indicate that the Fe catalysts with high Zn/K ratio or low K content possess the possibility to use CO2 contained in syngas for hydrocarbon synthesis at low temperature because the exact conversion from CO2 to hydrocarbons cannot be decided in our experiments. Therefore, Z4K1C4/Fe and Z4K2C4/Fe were assessed under the reactants of CO2 and H2 in order to investigate the influence of Zn/K ratio on CO2 hydrogenation [42].

Z4K1C4/Fe produces more C2+ hydrocarbons than Z4K2C4/Fe, it indicates that the catalyst with higher Zn/K ratio is more active to hydrogenate CO2 indirectly [9, 11, 18] or directly.

Furthermore, Z4K1C4/Fe shows similar activity and selectivity to those studied under 523 K [56], 538 K [30], or 623 K [33]. Based on the experimental results, we pointed out that it is possible to use CO2 contained in syngas for hydrocarbon synthesis at low temperature by the help of suitable promoter(s) [42].

1.2 Conclusions

CO2-TPD indicates that adsorbed CO2 on promoted Fe catalysts can partly be converted into CO. Promoter K or Zn is able to increase the adsorption of CO2 because more CO2 and CO are desorbed from K2/Fe and Z4/Fe than that from unpromoted Fe catalyst. K mainly increases CO2 adsorption and is inferior to Zn in producing CO.

Cu has no strong effect on CO2-TPD. For the Fe catalysts copromoted by Zn, K, and Cu, the desorbed amount of CO2 closely relies on the content of K although the content of Zn and Cu has effect on the amount of desorbed CO2. The desorbed CO2 from the tri-promoted catalysts with 2 mass% K is higher than the mono-promoted Fe catalyst by Zn or K, whereas the corresponding peak diminishes with the K content decreased to 1 mass%.

The combined promoters changes the CO desorption pattern evidently. The cata­lysts with high Zn/K ratio or low K content possess desorbed CO peak around 930 K. This peak reflects the possibility whether CO2 added into syngas can influence CO2 selectivity in FT synthesis. For the catalysts showing desorbed CO around 930 K, their CO2 selectivity based on converted CO is decreased due to the added CO2 in syngas.

The Fe catalyst with high Zn/K ratio shows high C2+ hydrocarbon selectivity for CO2 hydrogenation. It indicates that the CO2 contained in syngas is able to be activated by suitable promoter(s) for hydrocarbon synthesis at low temperature.

Jeffrey G. Linger and Al Darzins

Abstract The production of ethanol and other biofuels through the biochemical conversion of lignocellulosic biomass represents a promising path towards sustainably achieving the immense global demand for liquid transportation fuels. While numerous cellulosic ethanol production process configurations exist, the one known as Consolidated Bioprocessing (CBP) stands alone in combining all biologically medi­ated events into the action of a single organism (i. e., production and secretion of saccharolytic enzymes, hydrolysis of cellulose and hemicellulose, and fermentation of six-carbon and five-carbon sugars into biofuels such as ethanol). We discuss here the major issues with developing CBP technologies including the promises and challenges, the two prominently pursued routes to achieve this technology and several of the most promising candidate organisms. CBP represents a low-risk, high-reward proposition and its pursuit by researchers is most certainly warranted as we look to the future.

1 Introduction

The biological conversion of lignocellulosic biomass (biomass) to fuels such as etha­nol or butanol is broadly viewed as a very achievable means to producing large quantities of liquid transportation fuel from renewable resources in the shadow of a

J. G. Linger (*)

National Bioenergy Center, National Renewable Energy Laboratory, 1617 Cole Blvd, Golden, CO 80401, USA e-mail: Jeffrey. Linger@nrel. gov

A. Darzins

National Bioenergy Center, National Renewable Energy Laboratory, 1617 Cole Blvd, Golden, CO 80401, USA

DuPont Central Research and Development, 200 Powder Mill Road, Wilmington, DE 19803, USA e-mail: Al. Darzins@usa. dupont. com

J. W. Lee (ed.), Advanced Biofuels and Bioproducts, DOI 10.1007/978-1-4614-3348-4_16, 267

© Springer Science+Business Media New York 2013

waning supply of fossil fuels and an ever-changing energy marketplace. The bio­chemical conversion of cellulosic biomass to ethanol, for example, represents a major untapped potential fuel source, with minimal environmental impacts [1]. The challenge has always been to make the ethanol production process financially com­petitive in the current fuel market [2, 3]. While the commercial conversion of starches and monomeric sugars to fuel is a relatively straightforward process, the utilization of biomass as a starting feedstock is rather complex owing in large part to the recal­citrance of biomass, and to the heterogeneity of the substrate. Biomass recalcitrance is a general term which refers to the resistance of plant cell walls to enzymatic decon­struction [2], while heterogeneity refers to the complex mixture of lignin, cellulose and hemicellulose consisting of numerous sugar polymers, and the various chemical side-chain modifications that exist in different amounts and whose ratios vary among biomass feedstocks. The particularly difficult nature of using lignocellulosic bio­mass as the feedstock for the biological production of ethanol has led to the develop­ment of numerous process configurations that can be used to break down biomass in distinct, yet nontrivial steps. These steps include chemical pretreatment, enzymatic deconstruction and depolymerization of cellulose and hemicellulose, fermentation of hexose sugars, and fermentation of the pentose sugars [4, 5]. The detailed discus­sion of biomass chemical pretreatment is beyond the scope of our focus, but we mention it here because it is an absolutely essential component of current process configurations and still represents a significant research effort within the field of biofuels production. While numerous process configurations exist that separate the different steps mentioned above, perhaps no other configuration has garnered as much attention or caught the imagination of researchers as much as the one known as “Consolidated Bioprocessing,” also referred to as “CBP” [4, 6-12] (Fig. 1).

The concept of CBP refers to combining the four biologically mediated processing steps into the action of a single microorganism, or a consortium of microorganisms. As most CBP research has focused on single organisms, we limit our discussion to these candidates in this review. Essentially, an organism must produce and secrete multiple glycoside hydrolase enzymes to depolymerize the cellulose and hemicel — lulose within the pretreated biomass, uptake the newly released monomeric sugars, and metabolize both the five-carbon sugars and the six-carbon sugars to produce ethanol. While the concept is simple to describe, the intricacies of generating a viable CBP organism are extremely complex. In this review, we briefly discuss the promises and challenges of achieving commercially relevant CBP, the general strat­egies employed, and further discuss the advantages and disadvantages of candidate CBP organisms.

We proposed an additional novel cell wall remodeling approach by introducing algal and viral soluble polysaccharides to behave as “Trojan Horses” within the cell wall [1]. This new approach inserts novel polysaccharides into the plant cell wall, which may behave normally during plant growth, but allow enhanced solubility during processing. The “Trojan Horse” concept aims to reduce pretreatment input require­ments by the intercalating soluble polymers into the cell wall, enabling channel formation, and permitting rapid solvent and enzyme penetration and cell wall disas­sembly during and after the pretreatment stage. Such polysaccharides can be formed via exogenous expression of one or more enzymes capable of exploiting natural plant building blocks for their synthesis. The polysaccharides can be designed to be secreted or produced during cell wall development and to intercalate between cel­lulose fibers or serve as soluble “hemicellulose-like” polymers, creating soluble “pockets.” Polymers of the algae cell wall or bacterial exopolysaccharides can pro­vide a rich source of soluble polysaccharides, such as alginate, carrageenan, acetan, hyaluronan, chitosan, and levan. Most of the metabolic and genetic pathways for the synthesis of these polysaccharides are complicated and only a few have been fully elucidated.

A very similar biological process occurs naturally in algae-virus host-pathogen interactions. The paramecium bursaria Chlorella virus (PBCV-1) encodes multiple enzymes involved in extracellular hyaluronan synthesis [31]. PBCV-1-infected chlo­rella algae produce hyaluronan within their cell walls. As a result, the cell walls of the virus-infected algae are more porous when compared to that of the uninfected algae [48]. We have introduced genes for hyaluronan synthase together with appropriate targeting sequences into tobacco plants. Preliminary results have demonstrated enhanced cellulose hydrolysis upon acidic pretreatment in transgenic tobacco when compared to wild type plants (Abramson et al., unpublished results). We postulate that expression of hyaluronan synthase in plants can result in a reduction or modification of pectin levels due to competition for UDP-glucuronic acid, a substrate for pectin synthesis. This may result in a change in the mechanical properties and ultrastructure of the cell wall. Thus, ideally, hyaluronan synthase expression should be placed under control of a specific developmental promoter. Finally, as in the case of other cell wall modifying genes, the consequences of expression of each gene must be calculated to minimize side effects detrimental to plant growth, fitness, and physi­cal character.

3 Concluding Remarks

The secure supply of bio-energy will depend on the integration of various crop plat­forms as well as multiple technologies. These technologies will need to act upstream for plant modification and selection and downstream for processing to ensure the economic viability and environmental sustainability of future energy supplies.

Biotechnological tools can be utilized to develop and improve specific feed­stocks as dedicated crops for biofuel production. Appropriate modification of plant cell walls by genetic engineering approaches could produce new plant varieties with significantly enhanced properties for the efficient exploitation of biomass for bio­fuel production. We believe that advanced technology solutions will require the manipulation of combinations of genes and metabolic pathways, acting on different feedstocks, each suited to specific geographic environments.

Furthermore, these developments which are essential for the biofuels industry will also be beneficial for the pulp and paper industry by minimizing the use of energy and chemicals during the paper making process and by increasing pulping efficiency.

Interdisciplinary research combining agronomy, plant molecular biology, genet­ics, microbiology, mechanical and chemical engineering will be essential to advance the breakthroughs required for the development of economical, applicable solu­tions. All of these new approaches will need to be carefully evaluated for their effect on overall plant fitness and development and environmental sustainability.

Acknowledgments We are grateful to Professor Jonathan Gressel for his critical review, comments, and corrections on earlier drafts of the manuscript.

The designer proton-channel alga could be used with bioreactor systems for efficient photobiological H2 production (Fig. 7). As explained previously, the designer pro­ton-channel algae, such as the one that contains a hydrogenase-promoter-controlled designer proton-channel gene, can grow normally under aerobic conditions (Fig. 3a) by autotrophic photosynthesis using air CO2 in a manner similar to that of a wild — type organism. Therefore, to receive the maximal benefit by fully using the potential capabilities of the inducible proton-channel designer algae, it is a preferred practice

to grow the designer algae photoautotrophically using air CO2 as the carbon source under the aerobic conditions in a minimal medium that contains the essential min­eral (inorganic) nutrients. No organic substrate such as acetate is required to grow the designer algae under the normal conditions before the designer proton-channel gene is expressed.

Most of the algae 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, etc. at micromolar concentra­tion levels. All of the mineral nutrients are supplied in an aqueous minimal medium that is made with well-established recipes [23] of algal culture media using some water and relatively small amounts of inexpensive fertilizers and mineral salts, such as ammonium bicarbonate (NH4HCO3) (or ammonium nitrate, urea, ammonium

chloride), potassium phosphates (K2HPO4 and KH2PO4), magnesium sulfate hepta — hydrate (MgSO4 7H2O), calcium chloride (CaCl2), zinc sulfate heptahydrate (ZnSO47H2O), iron (II) sulfate heptahydrate (FeSO47H2O), and boric acid (H3BO3), etc. That is, large amounts of designer algae cells (biocatalysts) can be inexpen­sively grown in a short period of time because, under aerobic conditions such as in an open pond, the designer algae photoautotrophically grow by themselves using air CO2 as rapidly as their wild-type parental strains. This is a feature (benefit) that provides a cost-effective solution in generation of photoactive biocatalysts (the designer proton-channel algae) that are alternative to the silicone-photovoltaic — based technologies for renewable solar energy production.

When the algal culture is grown and ready for H2 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 algal reactor, to induce the expression of designer proton channels. When the designer proton-channel gene is expressed simultaneously with the induction of the hydrogenase enzyme under anaerobic conditions that can be achieved by removal of O2 from the culture, the algal cells are essentially turned into efficient and robust “green machines” that are perfect for photoevolution of H2 and O2 by water splitting (Fig. 2b). Production of H2 and O2 by direct photosynthetic water splitting can, in principle, have high quantum yield. Theoretically, it requires only four photons to produce a H2 molecule and VO2 from water by this mechanism. The maximal theoretical sunlight-to-H2 energy efficiency by the process of direct photosynthetic water splitting is about 10%, which is the highest possible among all the biological approaches. Application of the designer proton-channel algae maximally realizes the potential of this photosynthetic water-splitting process for H2 production because all the four pro­ton gradient-related physiological problems (illustrated in Fig. 1) that limit the rate of photobiological H2 production from water are eliminated by use of the designer alga (Fig. 2a, b) . Consequently, this approach has great potential when imple­mented properly with an algal H2-production reactor and a gas-separation/utiliza — tion system (Fig. 7).

Figure 7 illustrates an algal reactor and gas-separation-utilization system for simultaneous photosynthetic production of H2 and O2 from water (H2O) with effec­tive harvesting and utilization of the gas products including (but not limited to) fuel-cell application for electricity generation. An algal reactor contains a quantity of the designer alga that is exposed to light, such as sunlight. The H2 and O2 pro­duced in the algal reactor are pulled through a H2 separation membrane by vacuum pumps. The O2 on one side of the membrane is transferred to an O2 storage tank and a fuel cell. The H2 on the opposite side of the membrane is transferred to an H2 stor­age tank and the fuel cell.

It is worthwhile to note that the removal of O2 from algal culture can be achieved by a number of techniques, such as the use of a vacuum pump after the grown algal culture is sealed from atmospheric air O2. Because the production of H2 and O2 by direct photosynthetic water occurs in the same bioreactor volume, an effective gas — products separation process, such as the nanometer membrane technology shown in Fig. 7 for effective separation of H2 and O2 from the gas mixture, is necessary. Furthermore, consideration must be taken for safe handling of the H2 and O2 mix­ture, which is potentially explosive in case that the concentration of gas mixture reaches the explosion limits. Innovative application of a vacuo-photosynthetic reac­tor and gas-separation system also helps address the safety issue by strictly main­taining the concentration of H2 and O2 well below the explosion limits by effectively removing the gas products from the bioreactor system. In addition, use of a fuel cell that effectively consumes H2 and O2 for electricity generation could also help to remove the gas products from the bioreactor and gas-separation system. Furthermore, use of certain hydrogen storage materials, such as metal hydrides that can effec­tively adsorb H2, may also help to remove the H2 gas product from the bioreactor and gas-separation system. Certain engineering technology, such as use of argon or hydrofluorocarbon gas as an inert retardant in the bioreactor and gas-separation system, also improves the safe handling of the H2 and O2 mixture.

More information on how to use the designer proton channel algae for hydrogen production can be found in PCT patent application WO 2007/134340 [24]. Furthermore, the next section of this chapter, which reports the technology of designer switchable photosystem-II algae, will provide another solution to the O2 associated issues as well.

5 Designer Switchable-Photosystem-II Algae

According to the technology concept of designer switchable photosystem-II (PSII) algae, the designer transgenic algae comprise at least two transgenes for enhanced photobiological H2 production. The first said transgene serves as a genetic switch that controls PSII oxygen evolution and the second one encodes for creation of free proton channels in algal photosynthetic membranes, such as thylakoid membrane. The switchable PSII designer algae, combined with the benefits of programmable creation of free proton channels in algal photosynthetic membranes, provides an effective new solution for efficient and robust photobiological H2 production, by solving all the six major problems that currently challenge those in the field of pho­tosynthetic H2 production.

Accordingly, the switchable PSII designer alga is created by transformation of a host alga with at least one DNA construct as shown in Fig. 8a that contains a designer PSII suppressor gene linked with an externally inducible promoter such as a redox- condition-sensitive hydrogenase promoter serving as a genetic switch. According to this embodiment, the designer PSII suppressor is a PSII interfering RNA (iRNA) that can specifically suppress PSII oxygen evolution activity. The general design of the PSII suppressor gene is shown in its DNA construct, which comprises the fol­lowing components: (a) a PCR FD primer; (b) an externally inducible promoter; (c) a PSII-iRNA sequence; (d) a transcription terminator; and (e) a PCR RE primer.

A second transgene, illustrated in Fig. 8b, that encodes an inducible CF1-iRNA is added also into genome of the designer alga to create free CF0 proton channels in

algal photosynthetic membranes by suppressing the expression of CF1 along with the suppression of PSII and the induction of the hydrogenase under certain specific conditions, such as the anaerobic conditions. The general design of the DNA con­struct for the CF1 suppressor gene is shown in Fig. 8b, which includes the following components: (a) a PCR FD primer; (b) an externally inducible promoter; (c) a CFj — iRNA sequence; (d) a transcription terminator; and (e) a PCR RE primer.

Hydrogenase promoter is an anaerobic inducible promoter that serves as a genetic switch to desirably control the expression of the designer transgenes (Fig. 8a-d). Therefore, the designer transgenes can be expressed only under certain specific con­ditions, such as the anaerobic conditions. Under aerobic conditions, the designer transgenes are normally not expressed, except for the designer PSII-producing gene illustrated in Fig. 8e. Consequently, as illustrated in Fig. 9a, the switchable PSII designer alga performs autotrophic photosynthesis using ambient-air CO2 as the carbon source and grows normally, just like a wild-type organism under aerobic conditions, such as in an open pond. When the algal culture is grown and ready for H2 production (when sufficient amounts of starch are accumulated through the nor­mal oxygenic photosynthetic fixation of CO2), the algal cells are placed under anaer­obic conditions to express the designer transgenes simultaneously with the induction of the hydrogenase enzyme because of the use of the hydrogenase promoter.

As illustrated in Fig. 9b, the expression of the PSII inhibitor gene (as illustrated in Fig. 8a) will shut off PSII O2 evolution so that the alga will produce H2 by the metabolic (dark) and PSI-driven (light) H2 production pathways without production of O2 . The source of electrons for the metabolic and PSI-driven H2 production is organic reserves, such as starch (as illustrated in Fig. 9b) that are made in the previ­ous cycle of oxygenic photosynthetic CO2 fixation (Fig. 9a). Since the gas product in the embodiment illustrated in Fig. 9b is essentially pure H2 and CO2 without O2, the problems of H2 and O2 mixture (including problems 1-3: drainage of electrons by O2 , poisoning of the hydrogenase enzyme by O2 , and the gas-separation and safety issues) are eliminated with the designer algae.

According to one embodiment, use of a proton channel in the algal photosyn­thetic thylakoid membrane also enhances this mode of PSI-driven H2 production from organic reserves. As illustrated in Fig. 9b, operation of the PSI-driven H2 — production mechanism results in translocation of protons across the thylakoid mem­brane from the stroma into the lumen as the reducing power (NADH, NADPH, FADH) from degradation of organic reserves enters the PQ pool near the Q. site of Cyt b/f complex and is then oxidized at the Qo site. If a free proton-conducting chan­nel is not present in the thylakoid membrane, protons could accumulate at the lumen (inside the thylakoids) and impede the electron transport for this PSI-driven H2 pro­duction. Furthermore, the presence of a free proton-conducting channel in algal thylakoid membrane will ensure the inactivation of the Calvin-cycle activity that could compete with the Fd/hydrogenase H2 -production pathway for the electrons derived from the PSI-driven decomposition of the organic reserves. Therefore, it is a preferred practice to incorporate the benefits of a programmable thylakoid proton channel into the switchable PSII designer alga. This embodiment is achieved by delivery of a CF1-suppresor transgene, as illustrated in Fig. 8b, also into the genome

of the designer alga to create free CF0 proton channels in algal photosynthetic membranes by inhibiting the expression of CF 0 with a CF о о iRNA that the CF 0 — suppresor transgene can produce under the anaerobic conditions upon the induction of the hydrogenase along with the suppression of PSII oxygen-evolving activity.

Therefore, the coexpression of the PSII suppressor, the CFo suppressor, and hydrogenase makes this alga a more efficient and robust system for production of H2. This organism contains normal mitochondria, which can use 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 H2, the cell will stop generating nonfunctional PSII in thyla — koid membranes and start to restore its normal photoautotrophic capability by synthesizing functional thylakoids. Consequently, it is possible to use this type of genetically transformed organism for repeated cycles of photoautotrophic culture growth (starch accumulation) under normal aerobic conditions (Fig. 9a) and efficient production of H2 under anaerobic conditions (Fig. 9b).

One aspect is the innovative application of a genetic switch to control the expres­sion of PSII activity for production of H2 without the three O2 problems (illustrated in Fig. 9a, b). This switchability is accomplished through application of an externally inducible promoter such as a hydrogenase promoter. To function as intended, the designer transgenes (Fig. 8a-d) must be inducibly expressed under anaerobic condi­tions. In one embodiment, an algal hydrogenase promoter, such as the promoter of the hydrogenase gene (Hydl) of C. reinhardtii, is used as an effective genetic switch to control the expression of the designer genes to the exact time and conditions where they are needed for enhanced photobiological H2 production as illustrated in Fig. 9b. That is, the designer transgenes will be expressed only at the time when the hydroge — nase is induced and ready for H2 production under anaerobic conditions. Therefore, the hydrogenase promoter can be employed as an inducible promoter for each of the DNA constructs to serve as a genetic switch to control the expression of the designer genes. The reason that the designer alga can perform autotrophic photosynthesis using COo as the carbon source under aerobic condition (illustrated in Fig. 9a) is because the designer genes (illustrated in Fig. 8a-d) are not expressed under aerobic conditions owing to the use of a hydrogenase promoter, which can be turned on only under the anaerobic conditions when needed for photobiological H2 production.

In addition to the hydrogenase promoter, there are other promoters that can also be used to construct the desired genetic switch for designer genes (Fig. 8a-d). For example, the Nial promoter [25] which is induced by growth in nitrate medium and repressed in nitrate-deficient but ammonium-containing medium can be used to con­trol the expression of certain designer genes, such as the designer PSII-producing gene illustrated in Fig. 8e, according to the concentration levels of nitrate in a culture medium as well. Therefore, inducible promoters that can be used and/or modified to serve for this purpose include, but are not limited to, hydrogenase promoters, Cyc6 promoter, Nial promoter, CabII-1 promoter, Cal promoter, Ca2 promoter, coprogen oxidase promoter, and/or their analogs and modified designer sequences. Use of these externally inducible promoters can also create varieties of designer algae.

The PSII-iRNA DNA sequence (Fig. 8a) and the CF,-iRNA DNA fragment (Fig. 8b) are artificially designed based on the principle of the emerging RNA inter­ference technique. The RNA interference technique uses a piece of iRNA that can specifically bind with the mRNA of a particular gene, thus inhibiting (suppressing) the translation of the gene-specific mRNA to protein. Inactivation of PSII oxygen evolution activity is achieved by suppressing the expression of any key PSII compo­nents including its OEE1, D1, CP47, CP43, or D2 protein subunits. Therefore, the envisioned PSII iRNA further includes, but is not limited to, OEE1 iRNA, D1 iRNA, CP47 iRNA, CP43 iRNA, and/or D2 iRNA. For example, the OEE1 subunit [26] is a key component of the PSII oxygen-evolving complex (OEC) that is directly responsible for the oxygen evolution process. It is conceivable that deletion (sup­pression by an OEE1-specific iRNA) of the OEE1 subunit results in an OEE1- deficient PSII that is no longer able to evolve molecular oxygen by oxidation of water. The gene that encodes for the OEE1 subunit is PsbO (also known as Psbl), which is a nuclear DNA [ 27] that can be used with the hydrogenase promoter. Therefore, an anti-sense OEE1 DNA can be designed for generation of an anti-sense mRNA (iRNA) that can inhibit the synthesis of the OEE1 subunit by binding with the normal OEE1 mRNA. Unlike in the mammalian systems, where 21-nt probes are often used to target mRNA for inhibition, the iRNA in microbes (including the host organism C. reinhardtii) can be longer segments to be more gene-specific. It is a preferred practice to consider the following three criteria in designing the DNA sequence of the PSII suppressor (such as OEE1 iRNA): (a) the iRNA is a contigu­ous segment of the PSII subunit (e. g., OEE1) mRNA, typically with a length range of about 20-1,200 base pairs; (b) the iRNA is specific to the PSII subunit (e. g., OEE1), i. e., any significant portion of the iRNA does not have high sequence iden­tity to part of an RNA of any other genes in the host organism such as C. reinhardtii; (c) the iRNA does not have significant self-complimentarity to form stable second­ary structure that prevents hybridization with the mRNA of PSII subunit (e. g., OEE1). Because such a designed DNA fragment does not have any significant sequence similarity to other known genes, it (when transformed into the host alga) is able to produce an iRNA that specifically binds with normal mRNA of OEE1, thus selectively inhibiting the translation of the OEE1 mRNA into its protein. The CF1-iRNA sequence is designed with similar principles. Therefore, the iRNA tech­nique can be applied in conjunction with an inducible promoter, such as the hydro- genase promoter, as a genetic switch to create a desirable switchable PSII designer alga for improved photobiological H2 production.

)n the embodiments illustrated in Fig. 8a, b, each of the designer-suppressor DNA constructs also contains a terminator after the designer-suppressor encoding sequence. The terminator DNA sequence, which is designed based on the sequences of natural gene terminators, is to ensure that the transcription of said designer gene is properly terminated to produce an exact designer iRNA as desired.

In various embodiments, the host organisms for transformation of the designer genes (Fig. 8a-d) to create the transgenic designer photosynthetic organism are selected from the group that includes green algae, brown algae, red algae, blue — green algae, marine algae, freshwater algae, cold-tolerant algal strains, heat-tolerant algal strains, H2-consuming-activity-deleted algal strains, uptake hydrogenase-deleted algal strains, PSII-deficient algal strains, and combinations thereof. C. reinhardtii is a green alga that has had its genome sequenced. Therefore, it is a good model organ­ism, although the technology is applicable to any of the algae mentioned above for enhanced photobiological H2 production. Proper selection of host organisms for their genetic backgrounds and certain special features is also beneficial. For exam­ple, the switchable PSII designer alga created from a cold-tolerant host strain, such as Chlamydomonas cold strain CCMG1619 that has been characterized to produce H2 as cold as 4°C, enables the use even in cold seasons or regions such as Canada. Meanwhile, the switchable PSII designer alga created from a thermophilic photo­synthetic organism, such as Synechococcus bigranulatus, enables use into the hot seasons or areas such as Mexico and the Southwestern region of the United States including Nevada, California, Arizona, New Mexico, and Texas where the weather temperature can often be high. Furthermore, the switchable PSII designer alga cre­ated from a marine alga, such as Platymonas subcordiformis, enables using seawa­ter, while the designer alga created from a freshwater alga, such as C. reinhardtii, uses freshwater.

Another feature is that various embodiments of the designer algae are devoid of any H2-consuming activity that is not desirable for net H2 production. That is, the switchable PSII designer algae will not consume any H2 that it can produce. This feature, which further enhances the net efficiency for photobiological H2 produc­tion, is incorporated by genetic inactivation or deletion of the H2-consuming activity. The uptake hydrogenase activity is generally responsible for the H2 — consuming activity in algae. Therefore, this additional feature is incorporated by creating the designer algae from a host alga that has its uptake hydrogenase activity genetically deleted. Additional optional features of the designer algae include the benefits of reduced chlorophyll antenna size which has been demonstrated to provide higher photosynthetic productivity and O2-tolerant hydrogenase like the [NiFe] hydroge — nases of Ralstonia eutropha, which can function under aerobic conditions. In vari­ous embodiments, these optional features are incorporated into the designer algae also by use of an O 11 tolerant hydrogenase and/or chlorophyll antenna-deficient mutant (e. g., C. reinhardtii DS521) as a host organism for gene transformation with the designer DNA constructs (Fig. 8a-e).

There are a number of embodiments for constructing the switchable PSII designer algae by creating and/or using a genetically transmittable factor that can inducibly or programmably control PSII activity. That is, to create and/or use a factor that can be genetically encoded to programmably control PSII activity to solve the O2 — related problems in photobiological H2 production. An additional example on how to create the switchable PSII designer algae is described as follows. This example utilizes a hydrogenase promoter-linked streptomycin-production gene (Fig. 8c) to also create a switchable PSII designer alga for production of H2 without the three oxygen-related problems. It has been demonstrated that streptomycin is an inhibitor to the synthesis of a key PSII component, the D1 protein [28]. Without this D1 protein, PSII will no longer be able to perform photosynthetic water splitting to produce O2. The gene for streptomycin biosynthesis has already been cloned [29, 30].

Therefore, it is now possible by application of synthetic biology techniques to use this streptomycin-production gene in conjunction with the hydrogenase promoter as a programmable switch to control PSII activity in photosynthetic organisms. Figure 8c presents a DNA construct that can be genetically transferred into a host alga to create a switchable PSII designer alga with a phenotype similar to that illus­trated in Fig. 9a, b. That is, this designer alga containing the DNA construct can also grow and accumulate organic reserves (such as starch) by autotrophic photosynthe­sis using CO2 under aerobic conditions, such as in an open pond (Fig. 9a). When the algal culture is grown and ready for H2 production, the streptomycin-production gene is then expressed simultaneously with the induction of the hydrogenase enzyme under anaerobic conditions. The expression of the streptomycin-production gene inhibits the synthesis of the D1 protein, thus leading to gradual inactivation of PSII O2 evolution and activation of H2 production without O2 production (Fig. 9b).

Another aspect is the combination of switchable PSII with programmable proton channel and additional features. In one embodiment, it is a preferred practice to apply the feature of the switchable PSII in combination with the benefits of creating a free proton channel in algal photosynthetic thylakoid membrane (Fig. 9b). Proton channels can be created not only by selectively suppressing the expression of CF1 (Fig. 9b) using a designer CF1-iRNA gene (Fig. 8b), but also, in another embodi­ment, by targeted genetic insertion of proton channels such as polypeptide (or pro­tein) pores into algal photosynthetic thylakoid membrane (Fig. 9c) using a designer proton-channel gene (Fig. 8d). The detailed arts for targeted genetic insertion of proton channels into algal photosynthetic membrane have now been reported in the previous section of this chapter. The benefit s of a proton channel created with a designer proton-channel producing gene (Fig. 8d) are similar to those created with a designer CF1-iRNA gene while the features of the switchable PSII created with a designer PSII-iRNA gene are also similar to those created with a designer strepto­mycin (PSII inhibitor) producing gene. Therefore, these four types of designer genes (Fig. 8a-d) can be applied in various combinations to create the switchable PSII designer algae that also combine with the benefits of a free proton channel in algal photosynthetic thylakoid membranes under H2-producing conditions as shown in Fig. 9b, c. This combination of benefits is also achieved by algal hybridization of a switchable PSII designer alga with another transformed alga that contains a designer proton-channel gene (Fig. 8b or d), in addition to transformation of a designer proton-channel gene using a switchable PSII designer alga as a host organism.

As mentioned previously, operation of the PSI-driven H2-production mechanism results in translocation of protons across thylakoid membrane from the stroma into the lumen as the reducing power (NADH, NADPH, and FADH) from degradation of organic reserves enters the PQ pool near the Qr site of Cyt b/f complex and is then oxidized at the Qo site. If a free proton-conducting channel does not exist, protons could accumulate inside the thylakoids and impede the electron transport for this PSI-driven H2 production. Use of a free proton-conducting channel eliminates the restriction of photosynthetic H2 production by proton accumulation (problem 1). Furthermore, the presence of a free proton-conducting channel in algal thylakoid
membrane also ensures the needed inactivation of the Calvin-cycle activity (which is associated with problems 2-4) so that CO2 and/or O2 no longer act as a terminal electron acceptor through the Calvin-cycle activity to compete with the Fd/hydroge — nase H2-production pathway for the electrons derived from the PSI-driven decom­position of the organic reserves. Consequently, incorporation of a proton channel helps eliminate the four proton-gradient-related problems (1-4), while use of the switchable PSII solves the three O2-related problems (4-6). Therefore, the combina­tion of a switchable PSII with a proton channel in the switchable PSII designer alga (Fig. 9b, c) ensures to eliminate all the six technical problems that currently chal­lenge those in the field of photobiological H2 production.