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CHAPTER 3 Metabolic Engineering and Molecular Biotechnology of Microalgae for Fuel Production Su-Chiung Fang Biotechnology Center in Southern Taiwan, Academia Sinica Agricultural Biotechnology Research Center, Academia Sinica Tainan, Taiwan R.O.C. 3.1 INTRODUCTION Compared to other biofuel feedstocks, microalgae are the preferred option for many reasons: 1. They grow extremely fast and hence produce high biomass yield quickly. 2. Microalgae-based fuels do not compete with the food supply and hence present no food security concerns. 3. Biofuels generated from microalgae are renewable and can be carbon-reducing [generation of 100 tons of algal biomass is equivalent to removing roughly 183 tons of carbon dioxide from the atmosphere (Chisti, 2008)]. 4. Microalgal farming does not require arable land and can utilize industrial flue gas as a carbon source. 5. Selected oleaginous microalgae do not require fresh water and can grow in seawater, brackish water, or waste water. 6. Biodiesel fuels derived from microalgae can be integrated into the current transportation infrastructure. During the past few years, there have been significant advances in uncovering molecular components required for production of fuel molecules in microalgae. The availability of ge- nomic sequences in the model green alga Chlamydomonas reinhardtii has accelerated forward 47 Biofuels from Algae # 2014 Elsevier B.V. All rights reserved.

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Biofuels from Algae

C H A P T E R

3

Metabolic Engineering and MolecularBiotechnology of Microalgae for Fuel

ProductionSu-Chiung Fang

Biotechnology Center in Southern Taiwan, Academia Sinica

Agricultural Biotechnology Research Center, Academia Sinica

Tainan, Taiwan R.O.C.

3.1 INTRODUCTION

Compared to other biofuel feedstocks, microalgae are the preferred option for manyreasons:

1. They grow extremely fast and hence produce high biomass yield quickly.2. Microalgae-based fuels do not compete with the food supply and hence present no food

security concerns.3. Biofuels generated frommicroalgae are renewable and can be carbon-reducing [generation

of 100 tons of algal biomass is equivalent to removing roughly 183 tons of carbon dioxidefrom the atmosphere (Chisti, 2008)].

4. Microalgal farming does not require arable land and can utilize industrial flue gas as acarbon source.

5. Selected oleaginous microalgae do not require fresh water and can grow in seawater,brackish water, or waste water.

6. Biodiesel fuels derived from microalgae can be integrated into the current transportationinfrastructure.

During the past few years, there have been significant advances in uncovering molecularcomponents required for production of fuel molecules in microalgae. The availability of ge-nomic sequences in the model green alga Chlamydomonas reinhardtii has accelerated forward

47 # 2014 Elsevier B.V. All rights reserved.

48 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

genetic analysis and allowed for the use of reverse genetic approaches to uncover molecularmechanisms associated with fuel production (Merchant et al., 2007). Moreover, tran-scriptomics, proteomics, and metabolomics studies have provided new insights into generegulation networks and coordinated cellular activities governing physiological flexibilityand metabolic adaptation of microalgae. Understanding the basis of microalgal biology isimportant in laying the foundation for innovative strategies and for ultimate developmentof fuel surrogates. This review summarizes recent progress in elucidating molecular andcellular mechanisms of cellular physiology that are relevant to fuel production in microalgalsystems, with an emphasis on developing metabolic engineering strategies to increase fuelproduction.

3.2 BIODIESEL

Among the various fuel categories derived from microalgae, biodiesel receives the mostattention because it shares similar chemical characteristics with petrol diesel and can bedirectly channeled into the current transportation infrastructure without major alterationsof existing technology and fuel pipelines. Oleaginous green microalgae species that havethe capacity to accumulate oil in the form of triacylglycerols, or TAGs (Chisti, 2007; Sheehanet al., 1998), have been isolated and possess great potential as a feedstock for biodiesel fuels(Converti et al., 2009; Liu et al., 2008; Xu et al., 2006). However, microalgae-based biodiesel isfar from being commercially feasible, because it is not economically practical at present. Froma biological point of view, one of the obvious solutions is to increase oil content. Mostmicroalgae do not accumulate large amounts of lipid during a normal growth period. Cellsbegin to accumulate significant amounts of storage lipids after encountering stress conditionssuch as light and nutrient starvation (Hu et al., 2008; Sheehan et al., 1998). Nutrient starvation,however, slows cell proliferation and therefore limits biomass and overall lipid productivity.Despite the continuous interest in and enthusiasm about microalgal oil-to-biodiesel potential,the molecular mechanisms underlying the cellular, physiological, and metabolic networksconnecting to lipid and TAG biosynthesis remain largely unknown. Recent progress intranscriptomics, proteomics, metabolomics, and lipidomics studies have started to unravelthe complex molecular mechanisms and regulatory networks involved in lipid and TAGbiosynthesis in microalgae.

Current efforts to isolate and characterize the repertoire of genes required for lipid andTAG biosynthesis and accumulation in microalgae have focused on a model microalga:Chlamydomonas reinhardtii (Li et al., 2008; Miller et al., 2010; Msanne et al., 2012). With com-plete genome information, many enzymes required for lipid and TAG biosynthesis and me-tabolism have been identified based on in silico predictions of orthologous genes from otherorganisms (Riekhof et al., 2005). Similar to oilseed crops, the most common fatty acids inmicroalgae are 16- and 18-carbon fatty acids (Hu et al., 2008). Genome comparison and geneprediction analyses have shown that the pathways of fatty acid and lipid biosynthesis arelargely conserved between plants and green algae (Riekhof et al., 2005). In plants, de novo syn-thesis of fatty acids occurs in the plastid (Ohlrogge and Browse, 1995). The synthesized fattyacids are used as building blocks for synthesis of membrane lipids and storage lipids. AcetylCoA serves as the basic unit for fatty acid biosynthesis. It is converted tomalonyl CoA by acetyl

493.2 BIODIESEL

CoA carboxylase in the rate-limiting step for fatty acid biosynthesis. In green alga, acetyl CoApools are probably derived mainly from glycolysis (Hu et al., 2008). The malonyl moietyin malonyl CoA is transferred to a small acyl carrier protein (ACP) by malonyl-CoA:ACPtransacylase (MCT). The putative ortholog of MCT has been identified in Chlamydomonas(Lemaire et al., 2004). Following the synthesis of malonyl-ACP by MCT, fatty acid synthesiscontinues by the action of a type II fatty acid synthase composed of multiple proteins(White et al., 2005). The first condensation of malonyl-ACP and acyl-ACP is catalyzed by3-ketoacyl-ACP synthase. The step is followed by reduction catalyzed by 3-ketoacyl-ACPreductase and dehydration catalyzed by 3-hydroxylacyl-ACP dehydratase. Another roundof reduction is catalyzed by enoyl-ACP reductase. These serial reactions result in the additionof two methylene carbons to the growing acyl chain, and the cycle is repeated so that the acylchain reaches 16 or 18 carbons. Finally, chain elongation is terminated by fatty acyl-ACPthioesterases. Free fatty acids then leave the plastids and are converted into acyl-CoA byacyl-CoA synthetase, as shown in Figure 3.1 (Riekhof et al., 2005; Stern and Harris, 2009).

Because TAGs are derived from either acylation of diacylglycerol (DAG) via a de novo path-way (acyl CoA-dependent pathway) or recycling of membrane lipids (acyl CoA-independentpathway), most of the metabolic engineering strategies have been designed to manipulate therate-limiting steps of these two pathways in the hope of increasing themetabolic flux for TAGproduction. For the de novo pathway, fatty acids produced in the chloroplast are transferred inthe form of acyl-CoA to positions 1 and 2 of glycerol-3-phosphate by glycerol-3-phosphateacyltransferase (GPAT) and lysophosphatidic acid acyltransferase (LPAAT),respectively. The enzymatic reactions result in the formation of phosphatidic acid (PA)(Ohlrogge and Browse, 1995). Dephosphorylation of PA by phosphatidic acid phosphatasegenerates DAG. In plants, DAG is a common precursor to both membrane lipid and storageTAG (Ohlrogge and Browse, 1995). Diacylglycerol acyltransferases (DGAT) utilizes acyl-CoAas an acyl donor and catalyzes acylation of DAG for TAG production. Isolation andidentification of the DGATs that are able to enhance metabolic flux TAG production willbe important for increasing oil content in algal cells.

In addition to the de novo pathway, TAG is likely to be synthesized from an acylCoA-independent pathway in microalgae. In plants and budding yeast, phospholipids canbe used as acyl donors for the synthesis of TAG from DAG catalyzed by phospholipid:diacylglycerolacyltransferases, or PDATs (Dahlqvist et al., 2000). It is already known thatstorage lipids are accumulated under nutrient stress conditions concomitant with reorgani-zation of membrane lipids in microalgae (Fan et al., 2011). It is therefore reasonable to hypo-thesize that membrane lipids can be used as acyl donors for TAG biosynthesis under thesestress conditions (Figure 3.1).

Nitrogen deficiency is by far the predominant way to induce lipid body formation invarious microalgae (Hu et al., 2008). Detailed analyses have shown that nitrogen deprivationleads to a major redirection of carbon metabolism, decreased photosynthetic carbon fixation,and increased fatty acid biosynthesis (Miller et al., 2010). To explore the molecular events thatlink nitrogen deprivation to TAG synthesis, RNA transcriptome analysis has been employedto examine genome-wide transcript abundance and identify potential regulatory componentsregulating TAGmetabolism inC. reinhardtii (Boyle et al., 2012;Miller et al., 2010). As expected,mRNAs of some of the genes involved in lipid and TAGs metabolism are increasedunder N-deprived conditions. Among them are mRNAs of two DGATs, diacylglycerol

Chloroplast

Free fatty acids

Glucose

Glucose-6-phosphate

Pyruvate

Acetyl CoA

Malonyl ACP

Oil body

Cytosol

DHAP

Glycerol-3-phosphate

Oil body

3

GPAT

LPAAT

PA phosphatase

DGAT

ER

Acyl-ACP 3-Ketoacyl-ACP

3-Hydroxyacyl-ACP

Trans-Enoyl-ACP

Malonyl CoA

TAG

G3PDH

CoA

fatty acidsynthesis

Plasma membrane lipids

2

4

5

LCS

Acyl CoA

Phosphatidic acid

Diacylglycerol

TAG1

Lysophosphatidic acidAcyl CoA

FIGURE 3.1 Schematic representation of the pathways and presumed subcellular localizations of lipid synthesisand TAG assembly that are known or hypothesized to occur in microalgae. Free fatty acids are synthesized in thechloroplast. TAG-containing oil bodies are detected in both cytosol and chloroplast. The lipid biosynthesis pathwaythat leads to TAG assembly is depicted in black arrows. The unknown pathways used for TAG biosynthesisare depicted in arrows with dashed lines and labeled in numbers based on the origin of the acyl group. Acyl groupsfor TAG biosynthesis are potentially derived from 1-plasma membrane lipids, 2-chloroplast envelope membranes,3-fatty acids synthesized in the chloroplast, 4-acyl-ACP synthesized in the chloroplast, and 5-thylakoid membranes.G3PDH, glycerol-3-phosphate dehydrogenase; GPAT, glycerol-3-phosphate acyltransferase; LPAAT, lyso-phosphatidic acid acyltransferase; DGAT, diacylglycerol acyltransferase; LCS, long-chain acyl-CoA synthetase.

50 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

acyltransferase type-2 1 (DGTT1) and diacylglycerol acyltransferase 1 (DGAT1, type-1DGAT), which are increased under nitrogen-deprivation conditions (Boyle et al., 2012;Msanne et al., 2012) and likely play a role in controlling TAG synthesis in response tonitrogen-limited stress. Although inferences onmetabolism based on transcriptional analysesmay not be accurate, studies of orthologous genes in plants suggest their potential applica-tions. For example, DGATs catalyze the final step of the TAGs biosynthesis pathway, andoverexpression of arabidopsis DGAT in Brassica napus has been shown to increase oil contentin seeds (Post-Beittenmiller et al., 1992). Overexpression of DGATs in Chlamydomonas hasbeen investigated. However, this strategy does not enhance cellular TAG accumulation undereither nutrient-repleted or nutrient-depleted conditions (La Russa et al., 2012). This is not to-tally unexpected, because manipulation of single steps in the lipid and TAG biosynthesis

513.2 BIODIESEL

pathways in plants often results in moderate effects on seed oil content (Durrett et al., 2008;Thelen andOhlrogge, 2002). It is likely that expression of a single gene is not sufficient to drivemetabolic flux toward TAG accumulation.

In agreement with increased de novo biosynthesis of TAG during nitrogen deprivation,acyl-ACP thioesterase (also referred to as FAT1), a protein that takes part in fatty acid exportfrom the chloroplast to the ER, where TAG assembly occurs, is up-regulated (Miller et al.,2010). Moreover, mRNAs of phosphatidic acid phosphatases (PAPs) and phospholipid:diacylglycerolacyltransferase 1 (PDAT1), proteins that participate in biosynthesis of TAG,are also increased after nitrogen deprivation (Boyle et al., 2012; Miller et al., 2010). To char-acterize cellular functions of these isolated genes during lipid biosynthesis, reverse geneticshas been employed. Consistent with the functional role of PDAT1 during TAG biosynthesisfrom other organisms, Chlamydomonas pdat1 mutants accumulate less TAG than the parentalstrain (Boyle et al., 2012).

Nitrogen starvation has been shown to result in reorganization of the structure and break-down of the intracellular membrane systems (Fan et al., 2011; Martin et al., 1976; Moelleringand Benning, 2010). It is therefore reasonable to hypothesize that membrane lipids serve asbuilding blocks for TAG biosynthesis under nitrogen-deprived conditions. It will beimportant to understand how membrane lipids are recycled for TAG biosynthesis whenintracellular nitrogen is limited. In fact, PDAT1 (an acyl-CoA independent enzyme) may con-tribute to TAG synthesis by transferring an acyl group from phospholipid membranes toDAG during nitrogen deprivation (Oelkers et al., 2002). A greater understanding of theinterplay of membrane catabolism and its interaction with physiological adaptations willbe essential to devise a better strategy for metabolic engineering.

Recent studies have shown that fatty acid assembly in microalgae probably occurs in boththe ER and the chloroplast (Fan et al., 2011; Goodson et al., 2011). This indicates that two setsof acyltransferases are required to facilitate TAG synthesis in the ER or chloroplast. Howthese different sets of acyltransferases are coordinated and regulated during TAG biosynthe-sis and where they are located remain to be clarified.

In addition to enzymes known to take part in fatty acid and lipid biosynthesis, muchattention has been paid to identifying and isolating essential regulatory components thatact in the early signaling cascade to control lipid biosynthesis (Work et al., 2012). Transcrip-tional factors such as APETALA2 and Ethylene-Responsive Element Binding Protein(AP2-EREBP) and those belonging to the basic/Helix-Loop-Helix (bHLH) families have beenfound to be up-regulated under nitrogen-deficient conditions (Miller et al., 2010). Addition-ally, a SQUAMOSA promoter-binding protein domain transcriptional factor has been impli-cated in lipid biosynthesis under nitrogen-deficient conditions (Boyle et al., 2012). It will be ofgreat interest to test whether these transcriptional factors serve as molecular switches forbiosynthesis of fatty acid and storage lipids. Following the same logic, one of the AP2 domaintranscriptional regulators, WRINKLED 1 (WRI1), has been discovered to play an importantrole in the control of biosynthesis of storage lipids in plants (Bourgis et al., 2011; Cernac andBenning, 2004; Shen et al., 2010). Ectopic expression of WRI1 results in accumulation of TAGin developing seeds in Arabidopsis and maize (Cernac and Benning, 2004; Shen et al., 2010).In an important finding, comparative transcriptome analysis of C. reinhardtii, Phaeodactylumtricornutum, and Thalassiosira pseudonana has identified common regulatory genes whoseexpression is up-regulated under nitrogen-depleted conditions. Overexpressing one of the

52 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

genes has been shown to trigger lipid production in Chlamydomonas (Yohn et al., 2011).Ectopic expression or inducible expression of identified positive regulators of lipid biosyn-thesis and TAG assembly proteins will be preferable approaches to increase oil productionin microalgae.

As microalgae undergo metabolic changes under stress conditions, neutral lipids are con-centrated in a specialized structure called lipid droplets or lipid bodies. The globular lipid bodiesare filled with neutral lipids and enclosed by a single-layer lipid membrane (Goodson et al.,2011; Moellering and Benning, 2010). To decipher biochemical and cellular mechanismsunderlying the biogenesis, maintenance, and degradation of microalgal oil bodies, proteomicanalyses have been carried out to isolate and identify novel and conserved proteins directlyassociated with lipid bodies (Moellering and Benning, 2010; Nguyen et al., 2011). Consistentwith what was expected, proteins involved in lipid metabolism and catabolism wereassociated with lipid bodies (Moellering and Benning, 2010; Nguyen et al., 2011). These in-cluded acyl-coA synthetases, lipoxygenase, BTA1 (the enzyme required for the synthesisof membrane lipid diacylglyceryl-N-trimethylhomoserine), GPAT, LPAAT, PDAT, lyso-phosphatidyl lipid acyltransferases, and lipases. Other proteins potentially involved in lipidtrafficking such as small Rab-related GTPase, RAS GTPase, and subunits of the coat proteincomplex and its putative regulator, ARF1a, were also isolated (Moellering and Benning,2010). It is important to note that orthologs of these proteins have also been identified inthe lipid droplet proteome in animal species (Bartz et al., 2007; Cermelli et al., 2006). Isolationof these proteins in lipid bodies from different organisms suggests that mechanisms respon-sible for lipid biogenesis, assembly, and organization are evolutionally conserved. In additionto the proteins commonly found in the lipid body proteome, most of the isolated proteins arenovel. One of these proteins is named themajor lipid droplet protein (MLDP). MLDP is specificto the green algal lineage. Functional study of MLDP indicates that it is required to modulatelipid droplet size (Moellering and Benning, 2010). Further understanding of the biochemicaland molecular functions of these proteins will be important in devising an innovative planto increase oil body accumulation.

3.3 BIOHYDROGEN

The ability of microalgae to produce hydrogen was first reported by Gaffron and Rubinin 1942 (Gaffron and Rubin, 1942). However, the observed emission of hydrogen wastransient and the amount was very minimal. In the late 1990s, Melis and co-workers demon-strated that sulfur deprivation changes cellular metabolism and allows algal culture to switchfrom aerobic photosynthetic growth to an anaerobic physiology state. Switching to anaerobiccondition allows microalgal cultures to generate significant amounts of hydrogen for anextended period of time (Melis et al., 2000). This major breakthrough makes sustainablehydrogen production in a microalgal system a possibility. Over the years, extensive studieshave been done to understand the physiology and metabolic adaptation resulting fromsulfur depletion for better manipulation of biohydrogen production. Here our currentunderstanding of hydrogen production in microalgae is highlighted and possiblemetabolic engineering/biotechnology strategies for improving hydrogen production arediscussed.

533.3 BIOHYDROGEN

In green microalgae, production of hydrogen is catalyzed by [FeFe]-hydrogenases. [FeFe]-hydrogenases catalyze the reversible reduction of protons toH2 (Equation 3.1). Electrons usedfor the reduction reaction are derived from photosynthetic reductants. [FeFe]-hydrogenaseshave been isolated and identified frommicroalgae such as C. reinhardtii (Forestier et al., 2001;Happe and Kaminski, 2002), Scenedesmus obliquus (Florin et al., 2001), and Chlorella fusca(Winkler et al., 2002). Biochemical evidence supports the presence of hydrogenase activityin a variety of Chlorophycophyta (Brand et al., 1989). Because the molecular and physio-logical mechanisms of hydrogen production are better studied in the model organism C.reinhardtii, the prospects and implications of hydrogen production in microalgae will bebased on the recent genetic and biochemical studies in C. reinhardtii.

2Hþ þ 2e� !hydrogenaseH2 ð3:1Þ

Two highly similar [FeFe]-hydrogenases are present in the genome of C. reinhardtii and are

named HydA1 and HydA2. Both [FeFe]-hydrogenases are activated by anaerobic conditionsinduced by purging with neutral gas or by sulfur deprivation of the cultures (Forestier et al.,2003). Gene expression and mRNA stability as well as the enzymatic activities of HydA1 andHydA2 are extremely sensitive to oxygen. For this reason, hydrogen production rapidlystops as soon as cells begin oxygenic photosynthesis. Therefore, establishing anoxic cultureconditions is crucial for induction of hydrogen in algal cultures. In addition to creatinganaerobic conditions, electrons required to reduce Hþ ions are derived from photosystemII (PSII)-dependent activity (Antal et al., 2003; Antal et al., 2009) and PSII-independentplastoquinone (PQ) reduction pathways, as shown in Figure 3.2 (Chochois et al., 2009;Hemschemeier et al., 2008).

Through the PSII-dependent pathway, electrons required for hydrogen production aregenerated from light-dependent water-splitting reactions (photolysis). The electrons arepassed through the photosynthetic electron transport chain to reduce ferredoxin (FDX).Reduced FDX then serves as an electron donor in the reduction of twoHþ ions toH2 by hydro-genases. A chloroplast-located pyruvate:ferredoxin oxidoreductase has been proposed toregenerate reduced FDX, which is subsequently used for H2 production under anaerobicconditions (Atteia et al., 2006; Mus et al., 2007).

In addition to the light-dependent PSII pathway, electrons required for H2 production canbe derived from oxidation of endogenous reserves. It has been inferred that starch meta-bolism is important for H2 evolution. Studies from Chochois et al. (2009) suggest that starchcatabolism provides reductants to the PQ pool and the derived electrons are then passed tohydrogenases via a PSI-dependent electron transport chain. The reductants derived fromstarch catabolism are most likely to be NAD(P)H, which is further oxidized by athylakoid-associated type II NADH dehydrogenase (Desplats et al., 2009; Jans et al., 2008).The contribution of starch catabolism to H2 production has been verified by studies of mu-tants that failed to degrade starch (Chochois et al., 2010). These mutants have shown eitherslow or low hydrogen production by the PSII-independent pathway.

Asmentioned earlier, sulfur limitation is by far themost commonway to achieve extendedH2 production. Sulfur deprivation enhances H2 production by adjusting photosynthetic ca-pacity. The major effect of sulfur starvation is rapid inactivation of the PSII reaction center(Melis et al., 2000; Zhang and Melis, 2002) due to down-regulation of the de novo biosynthesisof the D1 protein (Wykoff et al., 1998). The decline in PSII activity reduces the amount of

PSII PSI

2H2O O2 + 4H+

PQ e-

e-

e-

PCe- e-

FNR

FDX

Chloroplast stroma

Thylakoidmembrane

ThylakoidLumen

ADP + Pi ATP

NADP+ NADPH

e-

HYD

2H+

H2

cyclic e- flow

NDA2

NADH NAD+

Starch reserves

pyruateglycolysis

photolysis

e-

H+

H+

PSII independentpathway

PSII dependentpathway

SIR

FTR

GOGAT

e-

e-e-

ATP synthaseCytb6f

e-

FIGURE 3.2 Electron transport pathways contributing to or competing with hydrogen production. The pathwaysthat contribute to hydrogen production are denoted as solid arrows. The pathways that potentially compete with thehydrogenase for electrons are marked as arrows with dashed lines. PQ, plastoquinone; HYD, [FeFe]-hydrogenase;Cytb6f, cytochrome b6f complex; FDX, ferredoxin; NDA2, type II NAD(P)H dehydrogenase; FNR, ferredoxin-NADPþ

reductase; SIR, sulfite reductase; FTR, ferredoxin-thioredoxin reductase; GOGAT, ferredoxin-dependent glutamatesynthase.

54 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

photosynthetic oxygen evolution. As a result, the rate of photosynthetic oxygen productionfalls below the rate of respiration and results in anaerobic conditions that enable the inductionof hydrogenase activity and H2 production.

In addition to establishing anaerobic culture conditions, removing sulfur stimulatesaccumulation of starch at an early stage (Tsygankov et al., 2002; Zhang and Melis, 2002).The mechanism of how sulfur deprivation transiently boosts starch accumulation remainsunclear. The accumulated starch reserves are subsequently catabolized via glycolysis to pro-duce reducing equivalents and to feed electrons into the PSII-independent hydrogenphotoproduction pathway (Hemschemeier et al., 2008; Kosourov et al., 2003). In addition,NAD(P)H produced from starch breakdown is oxidized via the mitochondrial respiratorychain, which in turn keeps oxygen levels sufficiently low to maintain the expression andactivity of hydrogenases (Melis et al., 2007). Therefore, sulfur starvation not only inducesand maintains anaerobic culture conditions under light, it also helps accumulate stored en-ergy from photosynthesis to drive PSII-independent hydrogen photoproduction. In essence,metabolic and physiological adaptation of microalgae by sulfur deprivation provides idealconditions for production of H2.

Although sulfur deprivation improves H2 production yield, it is still far from beingeconomically practical. In the batch system, sulfur deprivation results in production of1.2 mmol H2 per mol of chlorophyll per second for up to about 120 hours (Melis et al., 2000).

553.3 BIOHYDROGEN

The absence of sulfur eventually leads to lethal damage of cellular functions (Melis et al., 2000;Zhang et al., 2002). Various genetic andmetabolic engineering approaches have been utilizedto optimize hydrogen production under S-deprivation conditions. These strategies are aimedat (Antal et al., 2003) reducing the rate of oxygen evolution by decreasing PSII efficiency,(Antal et al., 2009) enhancing auxiliary electron transport pathways directing toward[FeFe]-hydrogenases, and (Atteia et al., 2006) improving [FeFe]-hydrogenases by decreasingoxygen sensitivity.

One of the major consequences of sulfur starvation is a decline of photosynthesis activitythat eventually leads to anaerobic culture conditions. DecreasingO2-evolution rates by reduc-ing photosynthesis ability can in theory achieve anaerobiosis. Indeed,mutants defective in D1protein (the major component of the PSII reaction center) display reduced PSII activity andsubstantially increased H2 production (Faraloni and Torzillo, 2010; Torzillo et al., 2009).In addition, knocking down a chloroplast sulfate permease using genetic engineering triggersa sulfur starvation response in sulfur-replete media. The resulting cells display reduced ratesof light-dependent oxygen evolution, low steady-state levels of the photosystem II D1 protein,establishment of anaerobiosis, and activation of [FeFe]-hydrogenases (Chen et al., 2005). Suchmicroalgal strains with low rates of photosynthesis/respiration are potentially useful for anintegrated and sustainable biohydrogen system (Melis and Melnicki, 2006). Following thesame logic, mutants mimicking similar metabolic responses that lead to anoxic conditionswill be good candidates to improve H2 production yield without losing cell viability fromprolonged S starvation.

Molecular engineering to redirect electrons to [FeFe]-hydrogenases offers another way toincrease hydrogen production. FDXs are important electron donors for [FeFe]-hydrogenases.However, FDXs also function to distribute high-energy electrons for CO2 fixation, nitrite andsulfite reduction, glutamate biosynthesis, cyclic electron flow, and reduction of thioredoxins(Figure 2). There are six [Fe2S2] ferredoxins present in Chlamydomonas (Winkler et al., 2010).Among them, PetF is thought to be the major ferredoxin involved in delivering electrons toHydA1 for H2 production in vivo (Jacobs et al., 2009). Recently, amino acid residues involvedin the direct interaction between FDX and hydrogenase have been identified (Chang et al.,2007; Long et al., 2008; Winkler et al., 2009). Enhancing electron transfer by manipulatingbinding affinity and kinetics between FDX and hydrogenase may provide a novel way toimprove hydrogen production efficiency.

Another way to direct electron flow toward [FeFe]-hydrogenases is to reduce the compet-itive electron sink (Figure 3.2). Cyclic electron flow (CEF) is important for optimal photosyn-thetic activity. Instead of passing electrons from reduced FDX to generate NADPH followingthe photosynthetic linear electron transport chain, CEF generates cyclic electron flow aroundPSI by passing electrons from reduced FDX to PQ through NADPH or directly to the Cytb6fcomplex to generate ATP (Rochaix, 2011). The balance between the linear photosynthesis elec-tron transport chain and CEF is required to modulate the ratio of ATP/NADPH to meet thecellular energy demands. Disrupting electron flow to CEF can in principle divert electronflow toward hydrogenase. Indeed, blocking CEF by antimycin A has demonstrated a twofoldimprovement in H2 production (Antal et al., 2009). A search for mutants with defective CEFhas been carried out to isolate enhancers of H2 production. Through fluorescence videoimaging, mutants unable to switch between linear and cyclic electron transport have beenisolated (Kruse et al., 1999). As expected, the proton gradient regulation like 1 (pgrl1) mutant

56 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

with impaired CEF shows improvedH2 production under both sulfur-repleted and -depletedconditions (Tolleter et al., 2011). To further channel electrons to hydrogenases via the light-independent PQ pathway, Baltz et al. overexpressed NDA2, a PQ-reducing type II NAD(P)Hdehydrogenase (Jans et al., 2008), in a pgrl1mutant background to enhance electron flow fromthe PQ pool (Desplats et al., 2009; Jans et al., 2008). This approach allows the pgrl1 mutant toincrease H2 by twofold (Baltz et al., Chlamydomonas meeting 2012). Similarly, the moc1 mu-tant that is defective in CEF and shows modified respiratory metabolism displays �5.4 timesincreased H2 production over the WT (Kruse et al., 2005; Schonfeld et al., 2004). Introductionof a hexose uptake protein from Chlorella kessleri allows Chlamydomonas to utilize externalglucose and leads to a further increase in hydrogen production (Doebbe et al., 2007). Theseexperiments provide proof-of-principle evidence that molecular andmetabolic engineering ispotentially useful in improving H2 production in microalgae.

To develop a culture system for continuous H2 production, an inducible chloroplast geneexpression system has been proposed (Surzycki et al., 2007). In this case, the PSII reactioncenter D2 protein (encoded by psbD) was engineered to be controlled in a copper-sensitiveand hypoxia-inductive post-transcriptional manner. Nac2, a gene required for D2 proteinexpression, has been engineered to be controlled by a cytochromeC6 (Cyc6) promoter. Thepromoter activity of Cyc6 is inhibited by copper and activated by hypoxia. Adding the copperleads to loss of PSII activity and subsequently anaerobic conditions and H2 production inlight. As cell cultures go anaerobic, repression of psbD is gradually relieved because theCyc6 promoter is activated by a low concentration of oxygen. Resumed expression of Nac2allows accumulation of D2 protein and subsequent photosynthetic growth of cell culture.As a result, manipulation of copper and subsequent creation of anaerobic conditions allowperiodic and reversible H2 production. This continuous culture system is potentially advan-tageous because it could operate without the penalty of nutrient starvation and reduce thecost associated with changes of growth media (Surzycki et al., 2007). However, the designof a bioreactor supporting this concept remains to be developed.

Despite the current progress, we are far from understanding the full picture of cellular andmetabolic changes and fermentative adaptations during anoxia. Optimization of H2 produc-tion requires an understanding of the combinatorial interplay between cellular physiologyandmetabolic pathways. The advantages ofChlamydomonas genetics allow large-scale geneticscreening for mutants that potentially affect or enhance H2 production (Ruhle et al., 2008;Schonfeld et al., 2004; Tolleter et al., 2011). Despite the differences in screening strategies,mutants with elevated hydrogen production have been isolated (Kruse et al., 2005; Tolleteret al., 2011). Because the mutants were tagged, disrupted genes could be readily isolatedand identified and their biological relevance to biohydrogen production could be easilyconfirmed. It is expected that a saturated mutant screen will yield more insights into molec-ularmechanisms underlyingmetabolic adaptation for hydrogen emission and pave an inroadfor future metabolic engineering.

Because [FeFe]-hydrogenases are irreversibly inactivated byO2 and their expression is alsonegatively regulated by O2, developing oxygen-tolerant [FeFe]-hydrogenases whose regu-lation is O2 independent would in theory circumvent limitations imposed by O2 sensitivity.Even though sequential mutagenesis followed by selection has been used to isolate oxygen-tolerant [FeFe]-hydrogenases (Flynn et al., 2002), the introduction of random mutations intothe genomemay result in reduced fitness that complicates further analysis. Other approaches,

573.3 BIOHYDROGEN

such as DNA shuffling, may be useful to generate oxygen-tolerant [FeFe]-hydrogenases(Nagy et al., 2007). Molecular designs of a [FeFe]-hydrogenases based on simulations ofpotential gas diffusion pathways have also been proposed (Ghirardi et al., 2007). Based on elec-tron paramagnetic resonance spectroscopy (Kamp et al., 2008) and X-ray absorptionspectroscopy analyses (Stripp et al., 2009a; Stripp et al., 2009b), the structural characterizationof the active site of the H-cluster explains the mechanism of oxygen sensitivity of [FeFe]-hydrogenases. The accumulated knowledge will be very useful for molecular engineering ofO2-tolerant [FeFe]-hydrogenases. Modification of [FeFe]-hydrogenases may not be sufficientto driveH2 production. It is known that thematuration of [FeFe]-hydrogenases requiresHydEFandHydG proteins whose regulation is also negatively regulated byO2.Without the help fromHydEF and HydG proteins during the maturation process, the engineered oxygen-tolerant[FeFe]-hydrogenasesmaynot suffice toproducehydrogen in thepresenceofO2.Awell-roundedstrategy, such as increasing the half-life of the activity of the [FeFe]-hydrogenases under aerobicconditions, could provide an alternative option (Ghirardi et al., 2007).

In addition to metabolic engineering, other strategies have been tested to optimize hydro-gen photoproduction. Unfortunately, manipulation of culture conditions such as adding lowlevels of sulfate (Kosourov et al., 2002), altering extracellular pH (Kosourov et al., 2003),adjusting light intensity (Laurinavichene et al., 2004), optimizing medium composition(Jo et al., 2006; Ma et al., 2011), or altering growth conditions (Kosourov et al., 2007) producesonly marginal improvements on H2 yield. Immobilization of cells on a solid surface has beendemonstrated to achieve better oxygen tolerance, light energy conversion efficiency, andduration of H2 photoproduction, and to therefore maximize H2 yield under sulfur-limitedconditions (Burgess et al., 2011; Hahn et al., 2007; Kosourov and Seibert, 2009; Laurinavicheneet al., 2006). To scale up production and extend the duration of H2 photoproduction, a two-stage chemostat bioreactor system that physically separates the photosynthetic growth phase(limited-sulfate conditions) and anaerobic H2 production phase (sulfur-deprived conditions)was designed (Fedorov et al., 2005). Based on the pilot experiment, H2 production can lastfor up to 4,000 hours (more than five months). It will be of great interest to assess H2 yieldof genetically optimized H2 producers (described previously) grown in such a two-stagechemostat bioreactor.

Recently, an integrated culture system combining photo-fermentation-based (carried outby photosynthetic green algae or cyanobacteria) and dark-fermentation-based (carriedout by anaerobic purple bacteria) H2 production units has been proposed (Melis andMelnicki, 2006). Co-cultivation of photosynthetic green algae and cyanobacteria allows utili-zation of a wide-range spectrum of solar irradiance and therefore improves solar energyutilization. In addition, the organic carbon and nitrogen generated from photo-fermentationcan be utilized to grow anaerobic bacteria for H2 production in a separate anaerobic bio-reactor. In return, the effluents from dark fermentation of an anaerobic bioreactor can berecycled to provide nutrients for photosynthetic algae and cyanobacteria in a photobiore-actor for H2 production. The idea is to create a self-sustainable system that integratesmetabolic systems of different microorganisms to optimize energy efficiency during H2

production processes (Eroglu and Melis, 2011; Ghirardi et al., 2009; Melis and Melnicki,2006). In addition to combining different biological systems, integration of biology andengineering in such a proposal will be very important to further advance the economics ofmicroalgae-based energy.

58 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

3.4 OTHER STRATEGIES

3.4.1 Optimization of Light Conversion Efficiency (LHCB)

Optimization of light conversion efficiency (LCE) is another way to make microalgae-basedbiofuels cost-effective. LCE is defined by Ghirardi et al. (2009) as the “fraction of the energycontent of the incident solar spectrum that is converted into chemical energy by the organ-ism.” It has been known that sunlight intensities are much higher than those requiredto saturate photosynthesis. To avoid overexcitation of the photosystem, plants and greenmicroorganisms deal with excess light by dissipating heat and emitting fluorescence. As aconsequence, the realistic LCE converts solar energy to biomass is much lower than thetheoretical calculation (Dismukes et al., 2008; Melis, 2009; Wijffels and Barbosa, 2010).

Another energy issue dealing with light efficiency is uneven distribution of light in a high-density cultivation system. For cells directly exposed to sunlight, up to 80% of the absorbedphotons could be wasted due to dissipation of excitation by nonphotochemical quenchingand photoinhibition of photosynthesis (Melis, 1999;Melis et al., 1999). On the other hand, cellsunderneath the culture are shaded from sunlight and have reduced photosynthesis rates.

To improve solar illumination distribution of the microalgal culture, mutants with reducedlight-harvesting chlorophyll antenna sizes that would allow for efficient utilization of lightenergy, and therefore would increase productivity, have been proposed. The rationale of thisapproach is to minimize light absorption by cells on the surface and to permit greater sunlightpenetrance into the deeper layers of the culture. This concept was experimentally validated byisolation and characterization of truncated light-harvesting chlorophyll antenna size (tla)mutants (Lee et al., 2002; Polle et al., 2000; Polle et al., 2003). Reduction of photosystemchlorophyll antenna size in tla mutants has been demonstrated to improve solar energyconversion efficiency and productivity. The notion has also been verified independently byanRNAi approach.Reductionof the light harvest complex I(LHCI) andLHCII antenna complexsystem by knocking down light harvest complex B major proteins results in improved photoncapture efficiency, enhanced growth rate, and reduced photoinhibition (Mussgnug et al., 2007).

In summary, accumulated experimental evidence indicates optimization of light-captureefficiency by genetic engineering can be very useful to improve culture productivity.Designs integrating growth optimization and fuel production will be important to makingmicroalgae-based fuel cost effective.

3.4.2 Recycling and Recovery of Co-products

To enhance the economics of microalgae-based biofuels, utilization of every ingredient ofthe raw biomass is important (Georgianna andMayfield, 2012; Sheehan et al., 1998). Whereasthe majority of fuels derived frommicroalgae have been focused on storage oils, the extractedoil accounts for only 37.9% of the energy and 27.4% of the initial fixed carbon (Lardon et al.,2009). The remaining carbon is stored in the leftover oil cakes composed of abundant proteinsand carbohydrates. Hence, recycling these nutrient elements may help increase biomassmargins of microalgae-based fuels (Lardon et al., 2009). Recycling algal waste by anaerobicdigestion has been proposed to support the microalgae production process (Ras et al., 2011;Zamalloa et al., 2012).

593.5 CHALLENGES AND PERSPECTIVES

Several innovativemetabolic engineering strategies have been proposed recently to reducethe energy debt and increase the margins of microalgae-based fuels. One of the approaches isto establish an integrated system that takes advantage of the amenable genetic modificationcapability of the Escherichia coli (E. coli) system. Although microalgae can grow photosynthet-ically to accumulate biomass for biodiesel purposes, the leftover paste can be utilized foralcohol-fuel production by feeding it into an engineered bacterial system. Huo et al. accom-plished this by genetically engineering an E. coli strain that is capable of converting the back-bone and side chains of amino acids in pretreated biomass into two-, four- and five-carbonalcohol fuels, ammonia, and other chemicals (Huo et al., 2011). In a small-scale experiment,the authors successfully converted hydrolyzed microalgal protein biomass into alcohol fuels.This demonstration supports the potential of using microalgal biomass as a feedstock forprotein-based biorefinaries.

3.5 CHALLENGES AND PERSPECTIVES

Even though the optimistic outlook on microalgae-based biofuels has driven microalgalresearch forward, we are still far from understanding the molecular networks underlyingthe complex metabolic flexibility and physiological adaptations to environmental cues ofphotosynthetic microalgae. Elucidation of molecular mechanisms of favorable traits suchas stress-induced oil accumulation and anaerobic fermentation capability is of fundamentalimportance to the basic biology and of practical importance to algal biotechnology. The recentefforts in sequencing algal genome sequences have facilitated isolation of genes involved inlipid biosynthesis, photosynthesis, anaerobic adaptation, and stress regulation. The utiliza-tion of reverse genetics techniques has allowed functional characterization of some of theisolated genes. Furthermore, integrated omics approaches have started to reveal novelinsights into the gene regulatory networks and cellular responses associated with metabolicfeatures for fuel production. The accumulated knowledge has generated testable hypothesesand provided strategies to increase biomass and improve fuel production. However, the mo-lecular toolbox required for reliable genetic manipulation of microalgae remains limited toonly a few species (e.g., C. reinhardtii, Volvox carteri, Nannochloropsis sp., and the diatomPhaeodactylum tricornutum) (Kilian et al., 2011; Leon and Fernandez, 2007; Schiedlmeieret al., 1994; Schroda, 2006; Siaut et al., 2007). For other species, genetic transformations havebeen documented sporadically but have not been robustly applied to routine genetic modi-fications. Lack of a reliable toolkit makes hypothesis-driven functional studies and practicalmanipulation in oleaginous species impossible. Development of custom-made moleculartoolkits for the chosen oleaginous algal species will be essential for metabolic engineering.Because genomic sequencing projects of various microalgae are in progress, the developmentof toolkits will accelerate in the coming years and shape the future of microalgalbiotechnology.

The recent advances in developing innovative technologies are aimed at improving theeconomics of microalgae-based biofuels. However, the practical application of the currenttechnology is still in its infancy, and most of the work has only been demonstrated at thelaboratory scale level. For instance, the proposed metabolic engineering strategies to improvebiodiesel production are designed to increase oil content at the per-cell level. Crucial to

60 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

overall yield relies on oil content at the per-culture basis. It is not clear whether small-scaleexperimental concepts can be directly translated into large-scale industrial setups. If not, whatfactors need to be considered and modified to allow laboratory oil producers to scale up toindustrial-level production? Until now, accurate assessment of energy balance and carbonreduction potential based on industrial-scale data spanning continuous seasons remains lim-ited. It is therefore difficult to assess the overall yield, energy balance, carbon mitigation, andenvironmental impacts of the yet-to-be-refined technology. Moreover, other interferencefactors such as parasite contamination, temperature fluctuation, weather influence, and lightpenetration that can potentially affect the productivity of the energy crop also need to be con-sidered during such an assessment. To make microalgae-based fuels a realistic industrialcommodity, multidisciplinary principles need to be integrated into current research strategiesto establish production platforms. In particular, integration of engineering and biology,followed by life-cycle-based long-term feedback evaluation/adjustment analyses of produc-tion pipelines, will be crucial to establishing solutions and optimizing protocols for energyproduction from microalgae.

Currently, the algal products (mostly food supplements and cosmetics products) on themarket cost approximately two orders of magnitude more than the current cost for biodieselproduction derived from oleaginous crops (Wijffels and Barbosa, 2010; Wijffels et al., 2010).Therefore, the practicality of producing microalgae-based fuels using the current technologyis still questionable (Chisti, 2008; Reijnders, 2008). Before the microalgae–to–fuels technologyis in place, incorporating the existing high-valued commodities into fuel production pipelinesmay provide a sustainable business model for microalgal biotechnology.

Acknowledgment

I am grateful to Dr. Lu-Shiun Her for his valuable comments on and suggestions regarding this chapter.

References

Antal, T.K., Krendeleva, T.E., Laurinavichene, T.V., Makarova, V.V., Ghirardi, M.L., et al., 2003. The dependence ofalgal H2 production on Photosystem II and O2 consumption activities in sulfur-deprived Chlamydomonasreinhardtii cells. Biochim. Biophys. Acta 1607, 153–160.

Antal, T.K., Volgusheva, A.A., Kukarskih, G.P., Krendeleva, T.E., Rubin, A.B., 2009. Relationships between H2

photoproduction and different electron transport pathways in sulfur-deprived Chlamydomonas reinhardtii.International Journal of Hydrogen Energy 34, 9087–9094.

Atteia, A., van Lis, R., Gelius-Dietrich, G., Adrait, A., Garin, J., et al., 2006. Pyruvate formate-lyase and a novel route ofeukaryotic ATP synthesis in Chlamydomonas mitochondria. J. Biol. Chem. 281, 9909–9918.

Bartz, R., Zehmer, J.K., Zhu, M., Chen, Y., Serrero, G., et al., 2007. Dynamic activity of lipid droplets: protein phos-phorylation and GTP-mediated protein translocation. J. Proteome. Res. 6, 3256–3265.

Bourgis, F., Kilaru, A., Cao, X., Ngando-Ebongue, G.F., Drira, N., et al., 2011. Comparative transcriptome and metab-olite analysis of oil palm and date palmmesocarp that differ dramatically in carbon partitioning. Proc. Natl. Acad.Sci. U. S. A. 108, 12527–12532.

Boyle, N.R., Page, M.D., Liu, B., Blaby, I.K., Casero, D., et al., 2012. Three acyltransferases and nitrogen-responsiveregulator are implicated in nitrogen starvation-induced triacylglycerol accumulation in Chlamydomonas. J. Biol.Chem. 287, 15811–15825.

Brand, J.J., Wright, J.N., Lien, S., 1989. Hydrogen production by eukaryotic algae. Biotechnol. Bioeng. 33, 1482–1488.Burgess, S.J., Tamburic, B., Zemichael, F., Hellgardt, K., Nixon, P.J., 2011. Solar-driven hydrogen production in green

algae. Adv. Appl. Microbiol. 75, 71–110.

613.5 CHALLENGES AND PERSPECTIVES

Cermelli, S., Guo, Y., Gross, S.P., Welte, M.A., 2006. The lipid-droplet proteome reveals that droplets are a protein-storage depot. Curr. Biol. 16, 1783–1795.

Cernac, A., Benning, C., 2004.WRINKLED1 encodes an AP2/EREB domain protein involved in the control of storagecompound biosynthesis in Arabidopsis. Plant J. 40, 575–585.

Chang, C.H., King, P.W., Ghirardi, M.L., Kim, K., 2007. Atomic resolution modeling of the ferredoxin:[FeFe] hydrog-enase complex from Chlamydomonas reinhardtii. Biophys. J. 93, 3034–3045.

Chen, H.C., Newton, A.J., Melis, A., 2005. Role of SulP, a nuclear-encoded chloroplast sulfate permease, in sulfatetransport and H2 evolution in Chlamydomonas reinhardtii. Photosynth. Res. 84, 289–296.

Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306.Chisti, Y., 2008. Biodiesel from microalgae beats bioethanol. Trends Biotechnol. 26, 126–131.Chochois, V., Dauvillee, D., Beyly, A., Tolleter, D., Cuine, S., et al., 2009. Hydrogen production in Chlamydomonas:

photosystem II-dependent and -independent pathways differ in their requirement for starch metabolism. PlantPhysiol. 151, 631–640.

Chochois, V., Constans, L., Dauvillee, D., Beyly, A., Soliveres,M., et al., 2010. Relationships between PSII-independenthydrogen bioproduction and starch metabolism as evidenced from isolation of starch catabolism mutants in thegreen alga Chlamydomonas reinhardtii. Int. J. Hydrogen Energ. 35, 10731–10740.

Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P., Del Borghi, M., 2009. Effect of temperature and nitrogen concen-tration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel produc-tion. Chemical Engineering and Processing 48, 1146–1151.

Dahlqvist, A., Stahl, U., Lenman, M., Banas, A., Lee, M., et al., 2000. Phospholipid:diacylglycerol acyltransferase: anenzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc. Natl.Acad. Sci. U. S. A. 97, 6487–6492.

Desplats, C., Mus, F., Cuine, S., Billon, E., Cournac, L., Peltier, G., 2009. Characterization of Nda2, a plastoquinone-reducing type II NAD(P)H dehydrogenase in chlamydomonas chloroplasts. J. Biol. Chem. 284, 4148–4157.

Dismukes, G.C., Carrieri, D., Bennette, N., Ananyev, G.M., Posewitz, M.C., 2008. Aquatic phototrophs: efficientalternatives to land-based crops for biofuels. Curr. Opin. Biotechnol. 19, 235–240.

Doebbe, A., Rupprecht, J., Beckmann, J., Mussgnug, J.H., Hallmann, A., et al., 2007. Functional integration of theHUP1 hexose symporter gene into the genome of C. reinhardtii: Impacts on biological H(2) production.J. Biotechnol. 131, 27–33.

Durrett, T.P., Benning, C., Ohlrogge, J., 2008. Plant triacylglycerols as feedstocks for the production of biofuels. Plant J.54, 593–607.

Eroglu, E., Melis, A., 2011. Photobiological hydrogen production: Recent advances and state of the art. Bioresour.Technol. 102, 8403–8413.

Fan, J., Andre, C., Xu, C., 2011. A chloroplast pathway for the de novo biosynthesis of triacylglycerol in Chlamy-domonas reinhardtii. FEBS Lett. 585, 1985–1991.

Faraloni, C., Torzillo, G., 2010. Phenotypic characterization and hydrogen producion in Chlamydomonas reinhardtiiQb-binding D1-protein mutants under sulfur starvation: changes in Chl fluorescence and pigment composition.J. Phycol. 46, 788–799.

Fedorov,A.S.,Kosourov,S.,Ghirardi,M.L., Seibert,M., 2005.ContinuoushydrogenphotoproductionbyChlamydomonasreinhardtii: using a novel two-stage, sulfate-limited chemostat system. Appl. Biochem. Biotechnol. 121–124, 403–412.

Florin, L., Tsokoglou, A., Happe, T., 2001. A novel type of iron hydrogenase in the green alga Scenedesmus obliquus islinked to the photosynthetic electron transport chain. J. Biol. Chem. 276, 6125–6132.

Flynn, T., Ghirardi, M.L., Seibert, M., 2002. Accumulation of O2-tolerant phenotypes in H2-producing strains ofChlamydomonas reinhardtii by sequential applications of chemical mutagenesis and selection. InternationalJournal of Hydrogen Energy 27, 1421–1430.

Forestier, M., Zhang, L., King, P., Plummer, S., Ahmann, D., et al., 2001. The cloning of two hydrogenase genes fromthe green alga Chlamydomonas reinhardtii. In Proceedings of the 12th International Congress on Photosynthesis(Melbourne, Australia: CSIRO Publishing).

Forestier, M., King, P., Zhang, L., Posewitz, M., Schwarzer, S., et al., 2003. Expression of two [Fe]-hydrogenases inChlamydomonas reinhardtii under anaerobic conditions. Eur. J. Biochem. 270, 2750–2758.

Gaffron, H., Rubin, J., 1942. Fermentative and Photochemical Production of Hydrogen in Algae. J. Gen. Physiol. 26,219–240.

Georgianna, D.R., Mayfield, S.P., 2012. Exploiting diversity and synthetic biology for the production of algal biofuels.Nature 488, 329–335.

62 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

Ghirardi, M.L., Posewitz, M.C., Maness, P.C., Dubini, A., Yu, J., Seibert, M., 2007. Hydrogenases and hydrogenphotoproduction in oxygenic photosynthetic organisms. Annu. Rev. Plant Biol. 58, 71–91.

Ghirardi, M.L., Dubini, A., Yu, J., Maness, P.C., 2009. Photobiological hydrogen-producing systems. Chem. Soc. Rev.38, 52–61.

Goodson, C., Roth, R.,Wang, Z.T., Goodenough, U., 2011. Structural Correlates of Cytoplasmic andChloroplast LipidBody Synthesis in Chlamydomonas reinhardtii and Stimulation of Lipid Body Production with Acetate Boost.Eukaryot. Cell 10, 1592–1606.

Hahn, J.J., Ghirardi, M.L., Jacoby, W.A., 2007. Immobilized algal cells used for hydrogen production. BiochemicalEngineering Journal 37, 75–79.

Happe, T., Kaminski, A., 2002. Differential regulation of the Fe-hydrogenase during anaerobic adaptation in the greenalga Chlamydomonas reinhardtii. Eur. J. Biochem. 269, 1022–1032.

Hemschemeier, A., Fouchard, S., Cournac, L., Peltier, G., Happe, T., 2008. Hydrogen production by Chlamydomonasreinhardtii: an elaborate interplay of electron sources and sinks. Planta 227, 397–407.

Hu, Q., Sommerfeld,M., Jarvis, E., Ghirardi,M., Posewitz,M., et al., 2008.Microalgal triacylglycerols as feedstocks forbiofuel production: perspectives and advances. Plant J. 54, 621–639.

Huo, Y.X., Cho, K.M., Rivera, J.G., Monte, E., Shen, C.R., et al., 2011. Conversion of proteins into biofuels by engineer-ing nitrogen flux. Nat. Biotechnol. 29, 346–351.

Jacobs, J., Pudollek, S., Hemschemeier, A., Happe, T., 2009. A novel, anaerobically induced ferredoxin inChlamydomonas reinhardtii. FEBS Lett. 583, 325–329.

Jans, F., Mignolet, E., Houyoux, P.A., Cardol, P., Ghysels, B., et al., 2008. A type II NAD(P)H dehydrogenasemediateslight-independent plastoquinone reduction in the chloroplast of Chlamydomonas. Proc. Natl. Acad. Sci. U. S. A.105, 20546–20551.

Jo, J.H., Lee, D.S., Park, J.M., 2006. Modeling and optimization of photosynthetic hydrogen gas production by greenalga Chlamydomonas reinhardtii in sulfur-deprived circumstance. Biotechnol. Prog. 22, 431–437.

Kamp, C., Silakov, A., Winkler, M., Reijerse, E.J., Lubitz, W., Happe, T., 2008. Isolation and first EPR characterizationof the [FeFe]-hydrogenases from green algae. Biochim. Biophys. Acta 1777, 410–416.

Kilian, O., Benemann, C.S., Niyogi, K.K., Vick, B., 2011. High-efficiency homologous recombination in the oil-producing alga Nannochloropsis sp. Proc. Natl. Acad. Sci. U. S. A.

Kosourov, S.N., Seibert,M., 2009.Hydrogenphotoproduction by nutrient-deprivedChlamydomonas reinhardtii cellsimmobilized within thin alginate films under aerobic and anaerobic conditions. Biotechnol. Bioeng. 102, 50–58.

Kosourov, S., Tsygankov, A., Seibert, M., Ghirardi, M.L., 2002. Sustained hydrogen photoproduction byChlamydomonas reinhardtii: Effects of culture parameters. Biotechnol. Bioeng. 78, 731–740.

Kosourov, S., Seibert, M., Ghirardi, M.L., 2003. Effects of extracellular pH on the metabolic pathways in sulfur-deprived, H2-producing Chlamydomonas reinhardtii cultures. Plant Cell Physiol. 44, 146–155.

Kosourov,S.,Patrusheva,E.,Ghirardi,M.L., Seibert,M.,Tsygankov,A., 2007.Acomparisonofhydrogenphotoproductionby sulfur-deprived Chlamydomonas reinhardtii under different growth conditions. J. Biotechnol. 128, 776–787.

Kruse, O., Nixon, P.J., Schmid, G.H., Mullinezux, C.W., 1999. Isolation of state transitionmutants of Chlamydomonasreinhardtii by fluorescence video imaging. Photosynth. Res. 61, 43–51.

Kruse, O., Rupprecht, J., Bader, K.P., Thomas-Hall, S., Schenk, P.M., et al., 2005. Improved photobiological H2

production in engineered green algal cells. J. Biol. Chem. 280, 34170–34177.La Russa,M., Bogen, C., Uhmeyer, A., Doebbe, A., Filippone, E., et al., 2012. Functional analysis of three type-2 DGAT

homologuegenes for triacylglycerol production in the greenmicroalgaChlamydomonas reinhardtii. J. Biotechnol .Lardon, L., Helias, A., Sialve, B., Steyer, J.P., Bernard, O., 2009. Life-cycle assessment of biodiesel production from

microalgae. Environ. Sci. Technol. 43, 6475–6481.Laurinavichene, T., Tolstygina, I., Tsygankov, A., 2004. The effect of light intensity on hydrogen production by sulfur-

deprived Chlamydomonas reinhardtii. J. Biotechnol. 114, 143–151.Laurinavichene, T.V., Fedorov, A.S., Ghirardi, M.L., Seibert, M., Tsygankov, A.A., 2006. Demonstration of sustained

hydrogen photoproduction by immobilized, sulfur-deprived Chlamydomonas reinhardtii cells. InternationalJournal of Hydrogen Energy 31, 659–667.

Lee, J.W., Mets, L., Greenbau, E., 2002. Improvement of photosynthetic CO2 fixation at high light intensity throughreduction of chlorophyll antenna size. Appl. Biochem. Biotechnol. 98–100, 37–48.

Lemaire, S.D., Guillon, B., Le Marechal, P., Keryer, E., Miginiac-Maslow, M., Decottignies, P., 2004. New thioredoxintargets in the unicellular photosynthetic eukaryote Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. U. S. A. 101,7475–7480.

633.5 CHALLENGES AND PERSPECTIVES

Leon, R., Fernandez, E., 2007. Nuclear transformation of eukaryotic microalgae: historical overview, achievementsand problems. Adv. Exp. Med. Biol. 616, 1–11.

Li, Y., Horsman, M., Wang, B., Wu, N., Lan, C.Q., 2008. Effects of nitrogen sources on cell growth and lipid accumu-lation of green alga Neochloris oleoabundans. Appl. Microbiol. Biotechnol. 81, 629–636.

Liu, Z.Y., Wang, G.C., Zhou, B.C., 2008. Effect of iron on growth and lipid accumulation in Chlorella vulgaris.Bioresour. Technol. 99, 4717–4722.

Long, H., Chang, C.H., King, P.W., Ghirardi, M.L., Kim, K., 2008. Brownian dynamics andmolecular dynamics studyof the association between hydrogenase and ferredoxin from Chlamydomonas reinhardtii. Biophys. J. 95,3753–3766.

Ma, W., Chen, M., Wang, L., Wei, L., Wang, Q., 2011. Treatment with NaHSO3 greatly enhances photobiological H2

production in the green alga Chlamydomonas reinhardtii. Bioresour. Technol. 102, 8635–8638.Martin, N.C., Chiang, K.S., Goodenough, U.W., 1976. Turnover of chloroplast and cytoplasmic ribosomes during ga-

metogenesis in Chlamydomonas reinhardi. Dev. Biol. 51, 190–201.Melis, A., 1999. Photosystem-II damage and repair cycle in chloroplasts: what modulates the rate of photodamage?

Trends Plant Sci. 4, 130–135.Melis, A., 2009. Solar energy conversion efficiencies in photosynthesis: Minimizing the chlorophyll antennae to

maximize efficiency. Plant Sci. 177, 272–280.Melis, A.,Melnicki,M.R., 2006. Integrated biological hydrogen production. International Journal of Hydrogen Energy

31, 1563–1573.Melis, A., Neidhardt, J., Benemann, J., 1999. Dunaliella salina (Chlorophyta) with small chlorophyll antenna sizes

exhibit higher photosynthetic productivities and photon use efficiencies than normally pigmented cells. J. Appl.Phycol. 10, 515–525.

Melis, A., Zhang, L., Forestier, M., Ghirardi, M.L., Seibert, M., 2000. Sustained photobiological hydrogen gas produc-tion upon reversible inactivation of oxygen evolution in the green algaChlamydomonas reinhardtii. Plant Physiol.122, 127–136.

Melis, A., Seibert, M., Ghirardi, M.L., 2007. Hydrogen fuel production by transgenicmicroalgae. Adv. Exp. Med. Biol.616, 110–121.

Merchant, S.S., Prochnik, S.E., Vallon, O., Harris, E.H., Karpowicz, S.J., et al., 2007. The Chlamydomonas genome re-veals the evolution of key animal and plant functions. Science 318, 245–250.

Miller,R.,Wu,G.,Deshpande,R.R.,Vieler,A.,Gartner,K., et al., 2010.Changes in transcript abundance inChlamydomonasreinhardtii following nitrogen deprivation predict diversion of metabolism. Plant Physiol. 154, 1737–1752.

Moellering, E.R., Benning, C., 2010. RNA interference silencing of a major lipid droplet protein affects lipid dropletsize in Chlamydomonas reinhardtii. Eukaryot. Cell 9, 97–106.

Msanne, J., Xu, D., Konda, A.R., Casas-Mollano, J.A., Awada, T., et al., 2012. Metabolic and gene expression changestriggered by nitrogen deprivation in the photoautotrophically grownmicroalgaeChlamydomonas reinhardtii andCoccomyxa sp. C-169. Phytochemistry.

Mus, F., Dubini, A., Seibert, M., Posewitz, M.C., Grossman, A.R., 2007. Anaerobic acclimation in Chlamydomonasreinhardtii: anoxic gene expression, hydrogenase induction, and metabolic pathways. J. Biol. Chem. 282,25475–25486.

Mussgnug, J.H., Thomas-Hall, S., Rupprecht, J., Foo, A., Klassen, V., et al., 2007. Engineering photosynthetic lightcapture: impacts on improved solar energy to biomass conversion. Plant Biotechnol. J. 5, 802–814.

Nagy, L.E., Meuser, J.E., Plummer, S., Seibert, M., Ghirardi, M.L., et al., 2007. Application of gene-shuffling for therapid generation of novel [FeFe]-hydrogenase libraries. Biotechnol. Lett. 29, 421–430.

Nguyen, H.M., Baudet, M., Cuine, S., Adriano, J.M., Barthe, D., et al., 2011. Proteomic profiling of oil bodies isolatedfrom the unicellular green microalga Chlamydomonas reinhardtii: with focus on proteins involved in lipid me-tabolism. Proteomics 11, 4266–4273.

Oelkers, P., Cromley, D., Padamsee, M., Billheimer, J.T., Sturley, S.L., 2002. The DGA1 gene determines a second tri-glyceride synthetic pathway in yeast. J. Biol. Chem. 277, 8877–8881.

Ohlrogge, J., Browse, J., 1995. Lipid biosynthesis. Plant Cell 7, 957–970.Polle, J.E., Benemann, J.R., Tanaka,A.,Melis, A., 2000. Photosynthetic apparatus organization and function in thewild

type and a chlorophyll b-less mutant of Chlamydomonas reinhardtii. Dependence on carbon source. Planta 211,335–344.

Polle, J.E., Kanakagiri, S.D., Melis, A., 2003. tla1, a DNA insertional transformant of the green alga Chlamydomonasreinhardtii with a truncated light-harvesting chlorophyll antenna size. Planta 217, 49–59.

64 3. METABOLIC ENGINEERING AND MOLECULAR BIOTECHNOLOGY OF MICROALGAE FOR FUEL PRODUCTION

Post-Beittenmiller, D., Roughan, G., Ohlrogge, J.B., 1992. Regulation of plant Fatty Acid biosynthesis: analysis of acyl-coenzymea andacyl-acyl carrier protein substrate pools in spinach andpea chloroplasts. Plant Physiol. 100, 923–930.

Ras, M., Lardon, L., Bruno, S., Bernet, N., Steyer, J.P., 2011. Experimental study on a coupled process of productionand anaerobic digestion of Chlorella vulgaris. Bioresour. Technol. 102, 200–206.

Reijnders, L., 2008. Do biofuels frommicroalgae beat biofuels from terrestrial plants? Trends Biotechnol. 26, 349–350;author reply 51–2.

Riekhof, W.R., Sears, B.B., Benning, C., 2005. Annotation of genes involved in glycerolipid biosynthesis inChlamydomonas reinhardtii: discovery of the betaine lipid synthase BTA1Cr. Eukaryot. Cell 4, 242–252.

Rochaix, J.D., 2011. Regulation of photosynthetic electron transport. Biochim. Biophys. Acta 1807, 375–383.Ruhle, T., Hemschemeier, A., Melis, A., Happe, T., 2008. A novel screening protocol for the isolation of hydrogen

producing Chlamydomonas reinhardtii strains. BMC Plant Biol. 8, 107.Schiedlmeier, B., Schmitt, R.,Muller,W., Kirk,M.M., Gruber, H., et al., 1994.Nuclear transformation of Volvox carteri.

Proc. Natl. Acad. Sci. U. S. A. 91, 5080–5084.Schonfeld, C., Wobbe, L., Borgstadt, R., Kienast, A., Nixon, P.J., Kruse, O., 2004. The nucleus-encoded protein MOC1

is essential for mitochondrial light acclimation in Chlamydomonas reinhardtii. J. Biol. Chem. 279, 50366–50374.Schroda, M., 2006. RNA silencing in Chlamydomonas: mechanisms and tools. Curr. Genet. 49, 69–84.Sheehan, J., Dunahay, T., Benemann, J., Roessler, P., 1998. A Look Back at the U.S. Department of Energy’s Aquatic

SpeciesProgram—Biodiesel from Algae, Close Out Report TP–580–24190. US Department of Energy’s Office ofFuels Development, Golden, CO.

Shen, B., Allen, W.B., Zheng, P., Li, C., Glassman, K., et al., 2010. Expression of ZmLEC1 and ZmWRI1 increases seedoil production in maize. Plant Physiol. 153, 980–987.

Siaut, M., Heijde, M., Mangogna, M., Montsant, A., Coesel, S., et al., 2007. Molecular toolbox for studying diatombiology in Phaeodactylum tricornutum. Gene 406, 23–35.

Stern, D.B., Harris, E.H. (Eds.), 2009. The Chlamydomonas Sourcebook, second ed. Organellar and MetabolicProcesses, vol. 2. Elsevier Ltd, p. 1071.

Stripp, S., Sanganas, O., Happe, T., Haumann, M., 2009a. The structure of the active site H-cluster of [FeFe] hydrog-enase from the green alga Chlamydomonas reinhardtii studied by X-ray absorption spectroscopy. Biochemistry48, 5042–5049.

Stripp, S.T., Goldet, G., Brandmayr, C., Sanganas, O., Vincent, K.A., et al., 2009b. How oxygen attacks [FeFe] hydrog-enases from photosynthetic organisms. Proc. Natl. Acad. Sci. U. S. A. 106, 17331–17336.

Surzycki, R., Cournac, L., Peltier, G., Rochaix, J.D., 2007. Potential for hydrogen productionwith inducible chloroplastg.ene expression in Chlamydomonas. Proc. Natl. Acad. Sci. U. S. A. 104, 17548–17553.

Thelen, J.J., Ohlrogge, J.B., 2002. Metabolic engineering of fatty acid biosynthesis in plants. Metab. Eng. 4, 12–21.Tolleter, D., Ghysels, B., Alric, J., Petroutsos, D., Tolstygina, I., et al., 2011. Control of Hydrogen Photoproduction by

the Proton Gradient Generated by Cyclic Electron Flow in Chlamydomonas reinhardtii. Plant Cell .Torzillo, G., Scoma, A., Faraloni, C., Ena, A., Johanningmeier, U., 2009. Increased hydrogen photoproduction by

means of a sulfur-deprived Chlamydomonas reinhardtii D1 protein mutant. International Journal of HydrogenEnergy 34, 4529–4536.

Tsygankov, A., Kosourov, S., Seibert, M., Ghirardi, M.L., 2002. Hydrogen photoproduction under continuousillumination by sulfur-deprived, synchronous Chlamydomonas reinhardtii cultures. International Journal ofHydrogen Energy 27, 1239–1244.

White, S.W., Zheng, J., Zhang, Y.M., Rock, 2005. The structural biology of type II fatty acid biosynthesis. Annu. Rev.Biochem 74, 791–831.

Wijffels, R.H., Barbosa, M.J., 2010. An outlook on microalgal biofuels. Science 329, 796–799.Wijffels, R.H., Barbosa, M.J., Eppink, M.H.M., 2010. Microalgae for the production of bulk chemicals and biofuels.

Biofuel. Bioprod. Bior. 4, 287–295.Winkler, M., Heil, B., Heil, B., Happe, T., 2002. Isolation andmolecular characterization of the [Fe]-hydrogenase from

the unicellular green alga Chlorella fusca. Biochim. Biophys. Acta 1576, 330–334.Winkler, M., Kuhlgert, S., Hippler, M., Happe, T., 2009. Characterization of the key step for light-driven hydrogen

evolution in green algae. J. Biol. Chem. 284, 36620–36627.Winkler, M., Hemschemeier, A., Jacobs, J., Stripp, S., Happe, T., 2010. Multiple ferredoxin isoforms in

Chlamydomonas reinhardtii - their role under stress conditions and biotechnological implications. Eur. J. CellBiol. 89, 998–1004.

653.5 CHALLENGES AND PERSPECTIVES

Work, V.H., D’Adamo, S., Radakovits, R., Jinkerson, R.E., Posewitz, M.C., 2012. Improving photosynthesis andmetabolic networks for the competitive production of phototroph-derived biofuels. Curr. Opin. Biotechnol. 23,290–297.

Wykoff, D.D., Davies, J.P.,Melis, A., Grossman,A.R., 1998. The regulation of photosynthetic electron transport duringnutrient deprivation in Chlamydomonas reinhardtii. Plant Physiol. 117, 129–139.

Xu, H., Miao, X., Wu, Q., 2006. High quality biodiesel production from a microalga Chlorella protothecoides byheterotrophic growth in fermenters. J. Biotechnol. 126, 499–507.

Yohn, C., Mendez, M., Behnke, C., Brand, A., 2011. Stress-induced lipid trigger. USA patent. USPTO Application :20120322157.

Zamalloa, C., Boon, N., Verstraete, W., 2012. Anaerobic digestibility of Scenedesmus obliquus and Phaeodactylumtricornutum under mesophilic and thermophilic conditions. Appl. Energ. 92, 733–738.

Zhang, L., Melis, A., 2002. Probing green algal hydrogen production. Philos. Trans. R. Soc. Lond. B Biol. Sci. 357,1499–1507; discussion 507–11.

Zhang, L., Happe, T., Melis, A., 2002. Biochemical and morphological characterization of sulfur-deprived andH2-producing Chlamydomonas reinhardtii (green alga). Planta 214, 552–561.