Environ. Sci. Technol. 2010, 44, 2243–2254
Challenges in Developing Biohydrogen as a Sustainable Energy Source: Implications for a Research Agenda L A U R A B . B R E N T N E R , * ,† J O R D A N P E C C I A , † A N D J U L I E B . Z I M M E R M A N †,‡ Environmental Engineering Program and School of Forestry and Environmental Studies, Yale University, New Haven, Connecticut
Received October 7, 2009 . Revised manuscript received February 8, 2010
The U.S. Department of Energy’s Hydrogen Program aims to develop hydrogen as an energy carrier to decrease emissions of greenhouse gases and other air pollutants and reduce the use of fossil fuels. However, current hydrogen production technologies are not sustainable as they rely heavily on fossil fuels, either directly or indirectly through electricity generation. Production of hydrogen by microorganisms, biohydrogen, has potential as a renewable alternative to current technologies. The state-of-the-art for four different biohydrogen production mechanisms is reviewed, including biophotolysis, indirect biophotolysis, photofermentation, and dark fermentation. Future research challenges are outlined for bioreactor design, optimization of bioreactor conditions, and metabolic engineering. Development of biohydrogen technologies is still in the early stages, although some fermentation systems have demonstrated efficiencies reasonable for implementation. To enhance the likelihood of biohydrogen as a feasible system to meet future hydrogen demands sustainably, directed investment in a strategic research agenda will be necessary.
Introduction The goal of developing a hydrogen economy is often stated broadly in terms of the need to provide a more sustainable energy system in the face of global climate change, dwindling fossil fuel supplies, and a push to decrease dependence on foreign oil. Hydrogen is characterized as a “clean fuel”, as the only byproduct of its use as an energy carrier is water vapor, producing no toxic emissions and adding no carbon dioxide (CO2) to the atmosphere. The U.S. Department of Energy (DOE) currently operates a Hydrogen Program for the development of hydrogen technologies to establish the “long-term viability of hydrogen as an energy carrier for transportation and stationary power.” (1) The program is focused primarily on lowering hydrogen production and delivery costs from a variety of sources, including fossil, nuclear, and renewable sources (wind, solar, geothermal, and hydroelectric). Currently, production of hydrogen in the U.S. relies heavily on fossil fuels, with 95% of the 9 million tons produced annually coming from steam methane reforming of natural gas (2). Nearly all of the remaining hydrogen is produced as a byproduct of chemical processes such as chlor-alkalai production, with water electrolysis contributing a tiny fragment to the current U.S. hydrogen supply (3). Globally, 40% of hydrogen is produced from natural gas, 30% from * Corresponding author e-mail:
[email protected]. † Environmental Engineering Program. ‡ School of Forestry and Environmental Studies. 10.1021/es9030613
2010 American Chemical Society
Published on Web 03/11/2010
crude oil, 18% from coal, and 4% from water electrolysis (4). Biological hydrogen, biohydrogen, production may provide a renewable, more sustainable alternative but has yet to reach a scale large enough for consideration in replacing a significant portion of the hydrogen supply. The Hydrogen Program’s aim for biohydrogen production is to verify the feasibility and competitiveness of hydrogen generation by microorganisms with current hydrogen production schemes by 2020 (1). Biohydrogen has the potential to considerably reduce costs and environmental impact as it can be produced with sunlight and minimal nutrients or organic waste effluents. Hydrogen (H2) producing microorganisms can be rapidly grown in bioreactors with relatively small energy and environmental footprints, making biohydrogen a renewable and low impact technology. Significant advances have been made over the past two decades in biohydrogen production: identifying capable microorganisms, engineering microorganisms to improve H2 production efficiency, and optimizing systems to maximize growth and hydrogen production potential. However, this biotechnology has yet to be utilized on a scale significant enough to be considered along with technologies currently in use for hydrogen production. If biohydrogen is to be considered a feasible alternative to conventional hydrogen production methods, a strategic research agenda needs to be identified. Figure 1 shows a breakdown of the budget for the DOE’s Hydrogen Program, by agencies within the DOE and among research sectors of the office of Energy Efficiency and Renewable Energy (EERE). Eighteen percent of the 2008 EERE hydrogen budget was directed at developing hydrogen production and delivery pathways, and only 11% of that budget was invested in researching biological production pathways. Given this level of investment by the only program aimed at launching the U.S. hydrogen economy, the hurdle is set high for establishing biohydrogen as a feasible and competitive sustainable technology. Hydrogen is a clean energy carrier, but current production technologies are not sustainable as they are largely dependent on carbon-based, nonrenewable resources (steam reformation of natural gas, petroleum refining, and coal gasification) (5). The limited supply of fossil fuels, predicted to peak for oil in 2023 and for natural gas and coal by 2050 (6), necessitate the adoption of alternative technologies for energy production, particularly H2 as an energy carrier. Biohydrogen production may be considered as a CO2 offset because it utilizes carbon sources that are already present in our environment. Other renewable hydrogen production options include electrolysis of water, possibly using wind power, photoelectrochemical, and thermochemical biomass conversion (gasification or pyrolysis) (7). Each of these approaches has associated challenges for sustainability including high electricity costs and VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2243
FIGURE 1. Budget breakdown for the Hydrogen Fuel Initiative for the U.S. DOE, Energy Efficiency and Renewable Energy (EERE) sector, and Hydrogen Production program within the EERE for the fiscal year 2008. The total Hydrogen Fuel Initiative budget for the U.S. DOE for 2008 was $281 million. precious metal catalysts. Biohydrogen represents a potential for a renewable and cost-effective option for hydrogen production. This review focuses on the current state of the biotechnology and the challenges that need to be addressed over the next ten years to verify feasibility of biohydrogen production as a competitive process in the hydrogen economy.
acids produced during dark fermentation (8-10). Table 2 presents a comparison of the production rates of systems utilizing the different mechanisms, in units of milliliters H2 per liter hour as a base comparison for bioreactor efficiency. Substrate conversion efficiency (hydrogen yield) (eq 1) is based on the fraction of moles of H2 of total theoretical moles available in the electron donor:
Biological Hydrogen Production
substrate conversion efficiency (%) ) moles H2 produced × 100% theoretical moles H2 per mole substrate
There are four basic mechanisms for biohydrogen production: biophotolysis, indirect biophotolysis, photofermentation, and fermentation. Table 1 provides a summary of these mechanisms including organisms, reactions, and key enzymes as well as pros and cons. Biophotolysis is associated with plant-type photosynthesis as it utilizes light energy to split water for hydrogen formation, generally performed by green algae under anaerobic conditions. Indirect biophotolysis typically involves cyanobacteria that make use of carbohydrate energy stores from photosynthesis to generate hydrogen from water. Photofermentation is a carried out by purple nonsulfur bacteria (PNSB) that utilize light energy to transform organic acids into H2 and CO2. Finally, fermentation is a dark process in which anaerobic bacteria break down carbohydrates to produce H2, among other gaseous byproducts (namely CO2). These mechanisms may also be used in combination to expand the technological possibilities. For example, photofermentation can produce hydrogen by utilizing light energy to further break down reduced organic 2244
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
(1)
Substrate conversion efficiency is not applicable to photoautotrophic mechanisms where water is the electron donor. For light-utilizing processes, photochemical efficiency (eq 2) is calculated as Photochemical efficiency (%) ) H2 mole production rate × H2 energy content per mole incident light energy (2)
Biophotolysis (Green Algae) Mechanisms. Biophotolysis, or photoautotrophic biohydrogen production, is the production of hydrogen from water using energy from absorbed light. The advantage of biophotolysis is that the electron donor for the production of
VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2245
purple nonsulfur bacteria
anaerobic bacteria
fermentation
cyanobacteria
indirect biophotolysis
photofermentation
green algae, cyanobacteria
organisms
biophotolysis
mechanism
2CH3COOH + 4H2 + 2CO2
C6H12O6 + 2H2O f
(y/2 + 2x - 2)H2 + xCO2
CxHyOz + (2x - z)H2O 98
light energy
2NH3 + H2 + 16 ADP + 16 Pi
N2 + 8H+ + 8e- + 16 ATP f
2NH3 + H2 + 16 ADP + 16 Pi
N2 + 8H+ + 8e- + 16 ATP f
12H2 + 6CO2
C6H12O6 + H2O 98
light energy
C6H12O6 + 6 O2
light energy
12H2O + 6CO2 98
light energy
2H2O 98 2H2 + O2
reaction
dehydrogenases, (fumarate) reductases, hydrogenases
nitrogenase
nitrogenase
high H2 production, utilizes waste streams, mixed-culture friendly
utilizes energy from sunlight to convert small organic acids or waste organic compounds to H2 and CO2 with no byproducts
heterocysts separate H2 production from O2-evolution within organisms
H2 produced from sunlight and water
[FeFe]-hydrogenase
[NiFe]-hydrogenase
pros
key enzymes
TABLE 1. Overview of Biohydrogen Mechanisms Including Organisms, Reactions, and Key Enzymes As Well As Pros and Cons
many byproducts, intensive biogas separation to retrieve H2, incomplete substrate utilization/low yields
bioreactor design challenges to maximize utilization of sunlight
O2 sensitivity, bioreactor design challenges to maximize utilization of sunlight, low H2 production efficiency bioreactor design challenges to maximize utilization of sunlight
cons
2246
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
organism/strain
Federov et Chlamydomonas al 2005 (31) reinhardtii Jo et al Chlamydomonas 2006 (23) reinhardtii Kim et al Chlamydomonas 2006 (24) reinhardtii Kruse et al Chlamydomonas 2005 (17) reinhardtii Stm6 Laurinavichene Chlamydomonas et al 2006 reinhardtii (32) Torzillo et al Chlamydomonas 2009 (33) reinhardtii D1 mutant Tsygankov Chlamydomonas et al 2006 reinhardtii (25) Melis et al Chlamydomonas 2000 (19) reinhardtii Antal and Gloeocapsa alpicola Linblad 2005 (37) Linblad et al Anabaena AMC414 2002 (39) Sveshnikov Anabaena variabilis et al 1997 PK84 (41) Borodin et Anabaena variabilis al 2000 (40) PK84 Troshina et Gloeocapsa alpicola al 2002 (38) CALU 743 Oh et al Rhodopseudomonas 2004 (29) palustris P4 Barbosa et Rhodopseudomonas sp. al 2001 (47) Barbosa et Rhodopseudomonas sp. al 2001 (47) Chen and Rhodopseudomonas Chang palustris WP3-6 2006 (48) Chen et al Rhogdopseudomonas 2008 (49) palustris WP3-5 Franchi et al Rhodobacter 2004 (50) sphaeroides SMV089 He et al Rhodobacter capsulatis 2005 (52) IR3
ref
biophotolysis biophotolysis biophotolysis biophotolysis biophotolysis biophotolysis biophotolysis biophotolysis indirect biophotolysis indirect biophotolysis indirect biophotolysis indirect biophotolysis photofermentation photofermentation photofermentation photofermentation photofermentation photofermentation photofermentation
water water water water water water water water/glycogen water water water water/glycogen fermentor effluent acetate acetate acetate acetate fermentor effluent lactate
mechanism biophotolysis
e- donor water
TABLE 2. Production Rates of Biohydrogen Systems
20.6
64.6
32.4
43.8
10.2
25
1.66
11.8
4.4
8.97
13.8
0.12
1.75
1.71
1.81
11.7
4.0
1.94
2.24
0.58
specific H2 production rate (mL H2 L-1 h-1)
84.8%
30.0%
61.0%
91.0%
50.4%
72.8%
70.0%
substrate conversion efficiency
88.10%
2.31%
6.20%
0.90%
0.14%
0.04%
1.33%
photochemical efficiency
672 h 285 h
80 µg Chl ml-1 12 µg Chl mL-1
360 h
5.51 g dcw L-1
93%
408 h
6.13 g dcw L-1 88.10%
0.9 g dcw L-1
240 h
200 h
0.4 g dcw L-1
0.56 g dcw L-1
200 h
0.6 g dcw L-1
8h
12 h
168 h
200 h
0.472 g dcw L-1
3.4-4.6 µg Chl mL-1
2.12 µg Chl mL-1
7 µg Chl mL-1
NR
6.0 × 106 cells mL-1
120 h
300 h
26 µg Chl L-1
18-22 µg Chl mL-1
116 h
96 h
4000 h
duration H2
0.96 g dcw L-1
6.0 × 106 cells mL-1
11.2 µg Chl mL-1
culture density
0.17 g dcw L-1
50%
87%
99.50%
92%
H2 fraction of biogas
VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2247
a
sucrose glucose
acidogenic sludge
acidogenic sludge/ Rhodopseudomonas capsulata cattle dung and sludge/ Rhodobacter sphaeroides SHC Clostridium pastegurianum/ Rhodopseudomonas palus sucrose
sucrose
formate
sucrose
anaerobic sludge
Escherichia coli SR13
glucose
mixed culture
fermentation
sucrose
fermentation/ photofermentation
fermentation/ photofermentation
fermentation/ photofermentation
fermentation
fermentation
fermentation
fermentation
fermentation
photofermentation
acetate,propianate
sucrose
photofermentation
mechanism
malate
e- donor
Rhodobacter sphaeroides KD131 Rhodopseudomonas capsulata anaerobic sludge (Clostridium, Sporolactobacillus) sewage sludge
organism/strain
8
25.9 (overall)
360 (dark), 22.7 (photo)
9.08 (photo)
9.3 × 106
3.0 × 10
505
192
56
541
23.2
82.5
specific H2 production rate (mL H2 L-1 h-1)
42.5%
27.6%
38.0%
16.8%
100.0%
9.5%
17.5%
14.5%
16.3%
46.7%
146%a
substrate conversion efficiency
2.39%
4.63%
photochemical efficiency
A substrate efficiency greater than 100% are based on the theoretical hydrogen yield of malate although additional subsrates were present.
Chen et al 2008 (8)
Tao et al 2007 (10)
Chen et al 2001 (59) Fang and Liu 2002 (63) Kyazze et al 2005 (60) Yoshida et al 2005 (64) Lee et al 2006 (61) Shi and Yu 2006 (9)
Kim et al 2006a (51) Shi and Yu 2006 (9) Fang et al 2001 (54)
ref
TABLE 2. Continued
88.40%
65%
42.70%
55.20%
64%
63%
85%
H2 fraction of biogas
8.21gdcwL-1
NR
0.40gdcwL-1
40 gVSSL-1
93.6gdcwL
10 d
382 h
50 d
20 d
5.72gVSSL-1 -1
21 d
1.0 gVSSL-1
1.2 gVSSL-1
20 gVSSL-1
90 d
72 h
1.78gdcwL-1 0.64g dwtL-1
duration H2
culture density
H2 is water, requiring no addition of organic substrates. Sunlight, CO2, and nutrients are the basic inputs to grow the green algae or cyanobacteria that carry out biophotolysis via the enzyme hydrogenase. Light energy is absorbed by photosystems I and II of microalgae, exciting the reaction centers that transfer energy through an electron transport chain, ultimately reducing ferredoxin, which provides electrons to the enzyme hydrogenase. It is thought that hydrogenases exist to provide an outlet for excess electrons when the carbon fixation component of the photosynthetic chain is compromised under special conditions (anaerobic, very lowPH2,lowlight)(11).Greenalgaepossess[FeFe]-hydrogenases, considered to be the more catalytically efficient of the hydrogenases (12-17). Cyanobacteria employ a wide variety of hydrogenases, mostly [NiFe]-hydrogenases, that may function as either uptake or bidirectional hydrogenases (18). Uptake hydrogenases counter H2 production by catalyzing H2 oxidation, bidirectional hydrogenases can catalyze both reduction and oxidation reactions of hydrogen. In biophotolysis, 1 mol of O2 is released for every 2 mol of H2 produced. O2 acts as a transcriptional repressor, an inhibitor of assembly, and an irreversible inhibitor of catalytic activity for [FeFe]-hydrogenases (12). Thus, sustained H2 production by biophotolysis requires the removal of O2 to produce anoxic conditions. Melis et al. (19) introduced a two-stage approach where green algae (Chlamydomonas reinhardtii) are grown under normal conditions and then deprived of sulfur to establish anaerobiosis. In sulfur-deficient environments, the repair cycle of photosystem II proteins is inhibited, restricting photosynthesis such that mitochondrial respiration can consume O2 at competitive rates in the light (20). This allows H2 production to continue uninhibited and produces a higher purity biogas. Technological Challenges. Three main issues to be addressed to improve biophotolysis systems include: (1) low photochemical efficiencies, (2) sensitivity of hydrogenases to O2, and (3) competition for reductant from ferredoxin between hydrogenases and other cellular functions. A photochemical efficiency of 10% is generally targeted, (7, 21, 22) but current efficiencies are far from reaching this target due to limitations in light penetration within photobioreactors and the transfer of light energy within cells. Hydrogenase sensitivity to O2 places strain on the duration of the H2 production phase. Photobioreactor design has been used to address 1 and 2, and genetic engineering has been used to address all three issues. Designs for photobioreactors are primarily based on the two-stage, sulfur-deprivation induced anaerobic system for C. reinhardtii described above. Photobioreactor design is aimed at optimizing cell growth in the first stage and cell maintenance in the second stage for light capture and length of H2 production phase. The first sustained hydrogen production by green algae using the two-stage system, by Melis et al. (19), produced H2 at a rate of 1.75 mL L-1 h-1 over a production phase of approximately 120 h. Optimizing nutrient concentration, CO2 addition and light intensity has provided minor improvements to H2 production rates (23-29). For cell growth, light can be inhibitory at both low and high intensities. Similarly, finding the optimal sulfur concentration that restricts photosystem II enough to limit O2 production but provides enough sulfur for minimal biosynthesis of other essential cell proteins has been a challenge for sustaining H2 production in the second stage (30). Response surface methodology (RSM) is a statistical tool used to analyze the influence of several independent variables on a dependent variable and is used to optimize the physiochemical factors within a bioreactor that influence H2 production. H2 production rates increased in sulfurdeprived C. reinhardtii by up to 1.55 times using RSM to optimize nutrients, temperature, pH, and light intensity (23). 2248
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
Bioreactor design innovations have also improved the biophotolysis process. A chemostat system considerably prolonged the H2 production phase for more than 166 days, albeit at a lower production rate of 0.58 mL H2 L-1 h-1 (31). Immobilization of sulfur-deprived algal cells in fiber glass matrices allowed for significant enhancement in both H2 production rates and length of production phase, with up to 11.7 mL H2 L-1 h-1 and 4 weeks duration (32). This is one of the highest reported H2 production rates for C. reinhardtii. Protein and genetic engineering efforts have further advanced H2 production rates and lengthened H2 production periods. Sulfur limitation limits competition for electrons by reducing rubisco, as well as limiting the sulfate reduction pathway (29), but competition for electron transport can be further limited. Directed evolution in C. reinhardtii by Kruse et al. (17) produced strain Stm6, a mutant with impaired cyclic electron transport in PSI. Stm6 mutants had larger starch reserves, lower dissolved oxygen concentrations, and greater electron flow to hydrogenase, producing a longer H2 production phase and greater H2 content biogas. Mutations of the PSII cofactor binding protein D1 may increase its electron donor capacity and thus have the potential to enhance H2 production in sulfur-deprived C. reinhardtii. Directed amino acid substitutions carried out by Torzillo et al. (33) created a C. reinhardtii D1 double mutant (two amino acid substitutions) characterized by a lower chlorophyll content, higher photosynthetic capacity, and a higher respiration rate that produced hydrogen at an elevated rate for a longer period of time compared with the wild-type. Other C. reinhardtii strains have been developed that downregulate (truncate) light harvesting complexes to improve photosynthetic quantum yield in algal bioreactors under high solar irradiance levels but have yet to be tested for their effect on H2 production (21, 34). The greater sensitivity of [FeFe]-hydrogenases to O2 makes it difficult to isolate them and limits accessibility to characterize the enzyme’s structure. Many engineering efforts are therefore focused on the more accessible, less O2sensitive, cyanobacterial [NiFe]-hydrogenases. A membranebound, soluble hydrogenase has been characterized in the bacterium Ralstonia eutropha which is relatively O2-tolerant, although it operates at activities 2 orders of magnitude lower than the O2-sensitive hydrogenases (15). Investigations have revealed that O2-tolerance in hydrogenases evolved through one of two structural advantages: the presence of an extra CN ligand at the active site or a narrow gas channel to the active site of the hydrogenase (15, 35). Researchers also strive to improve the catalytic efficiency of [NiFe]-hydrogenases. One approach is the modification of intramolecular electron transfer pathways in bidirectional [NiFe]-hydrogenase, to alter the redox potential at the active site for preferential catalysis in one direction (15).
Indirect Biophotolysis Mechanisms. Indirect biophotolysis separates the photosynthetic processes from hydrogen production, either temporally or spatially. Indirect biophotolysis avoids problems associated with enzymatic O2-sensitivity by disjoining the evolution of O2 from the production of H2. Generally, indirect biophotolysis is a mechanism employed by cyanobacteria. While the two-stage, sulfur-limiting C. reinhardtii system separates aerobic conditions from anaerobic conditions, it is still considered direct biophotolysis because the photosynthetic operations of the cells are still functioning and delivering electrons to the hydrogenases during anaerobiosis (36) (even though some electrons are derived from starch in these systems 26, 28). In indirect biophotolysis, cells utilize starch reserves from previous photosynthetic activities to provide the energy needed to produce H2 through fermentation. Indirect biophotolysis shares many of the advantages
of direct biophotolysis because water is still the electron donor and inorganic carbon the carbon source in this process (37). Filamentous cyanobacteria, such as those in the genus Anabaena, spatially separate the two processes by forming heterocysts. Heterocysts are specialized cells for nitrogen fixation that are found among chains of photosynthetic cells. They are fed by neighboring photosynthetic cells, so they do not need their own photosynthetic machinery, which would impede the function of the O2-sensitive nitrogenase enzymes that catalyze nitrogen fixation. Nitrogenases also catalyze the formation of H2 as an obligatory product of nitrogen fixation; according to eq 3 (nitrogenase isoenzymes vary on how many hydrogen ions are paired with the fixation):
N2 + 8H+ + 8e- + 16 ATP f 2NH3 + H2 + 16 ADP + 16 Pi (3) Indirect biophotolysis has also been developed in cyanobacteria, such as Gleocapsia alpicola or Synechtocystis sp., by temporal separation of photosynthesis and H2 production stages. Cells are grown photoautotrophically to accumulate stores of glycogen that is then fermented under dark incubation to drive production of H2. In addition to nitrogenase-catalyzed H2 production, cyanobacteria are equipped for hydrogen cycling with uptake hydrogenases and occasionally bidirectional hydrogenases. It is proposed that NAD(P)H, produced during the “dark” fermentation of glycogen, supplies the energy to a bidirectional hydrogenase that is induced under anaerobic conditions (38). By taking advantage of H2-producing pathways and engineering cells to impair H2 uptake, some relatively high production rates have been achieved. Technological Challenges. The sequencing and characterization of genes for the small and large subunits of uptake hydrogenase (hupS and hupL) for multiple cyanobacteria has allowed for genetic manipulations of these genes to improve H2 production (39). The heterocyst-forming Anabaena variabilis mutant PK84 is both uptake hydrogenase deficient and has an impaired reversible hydrogenase. Both Borodin et al. (40) and Sveshnikov et al. (41) saw enhanced H2 production in PK84 over wild-type strains, with even greater enhancement after limiting N2 in the headspace, reaching production rates of 4.4 and 8.97 mL H2 L-1 h-1, respectively. Sveshnikov et al. (41) reported that H2 production was not increased as much under N2-limiting conditions in the wild-type strain or in a mutant only deficient in uptake hydrogenase and that nitrogenase activity did not increase in any of the strains. This suggests that the productivity of PK84 is likely due to improvements in the efficiency of the bidirectional hydrogenase as well as the loss of the uptake hydrogenase. Another Anabaena uptake hydrogenase deficient mutant, AMC414, has increased H2 production in a photobioreactor with limited N2 (replaced air with argon gas) (39). In an airlift photobioreactor described by Linblad et al. (39), under simulated outdoor conditions with argon gas flowing through it, Anabaena AMC414 boasts one of the highest production rates among direct or indirect biophotolysis systems at 13.8 mL H2 L-1 h-1. The increase in H2 production was, as in PK84, not correlated with nitrogenase activity. AMC414 was also capable of producing H2 using air in the photobioreactor, while the wild-type strain was incapable of H2-production when exposed to air. However, the photochemical efficiency of the system is quite low at 0.042%, exposing one of the disadvantages of indirect biophotolysis systems. In temporally separated indirect biophotolysis systems, Gleocapsia alpicola CALU 743 have been shown to produce H2 at rates competitive with other biophotolysis systems at 11.8 mL H2 L-1 h-1 under dark, anaerobic conditions (38). This rate of H2 production was only sustained for 8 h.
Glycogen stores in the cells are finite and limit the length of time that energy is available to the cells and hydrogenase activity can continue. RSM for media composition has been used to increase glycogen stores up to 44-fold in cells in the optimized media over cells in a complete media, contributing to an increase in hydrogen production of 148-fold (42). In order to maintain H2 production, cells need to be continuously cycled between photosynthetic aerobic and dark fermentative stages. Engineering cells to increase glycogen storage could potentially increase the length of fermentative H2 production cycles. While indirect biophotolysis is able to mitigate some of the O2 sensitivity issues of direct biophotolysis, they still share many of the same challenges. Designs for efficient photobioreactors are needed to make the process cost-effective. Research efforts should be directed at more innovative photobioreactor designs to immobilize or better stabilize cells and operate on a continuous flow to allow for scale-up to industrial operations. Metabolic engineering efforts to increase efficiency of [NiFe]-hydrogenases, eliminate oxidation of H2 by uptake and bidirectional hydrogenases, improve glycogen accumulation, and reduce competition for electron flow may lead to competitive production rates in indirect biophotolysis systems. H2 production by indirect biophotolysis relies on glycogen stores in cells that are shared with other cellular processes requiring energy, thus metabolic engineering to increase glycogen available for hydrogen reduction would improve overall efficiency. Photochemical efficiency could also be improved by developing mutants with truncated light-harvesting complexes as mentioned above for algae.
Photofermentation (Purple Nonsulfur Bacteria) Mechanisms. Photofermentation, or photoheterotrophic hydrogen production, is a mechanism found among a diverse group of photosynthetic bacteria and well-characterized in PNSB. This mechanism is primarily mediated by the enzyme nitrogenase. Similarly to cyanobacteria, PNSB may also have bidirectional and uptake hydrogenases that contribute to hydrogen cycling within cells. PNSB have an advantage over cyanobacteria and algae because they exist solely under anaerobic conditions and do not have to separate their growth phase from their H2 generating activities. Photohetertrophs make use of energy from sunlight to oxidize organic compounds and generate the electron potential needed to drive H2 production. By utilizing energy from the sun to drive thermodynamically unfavorable reactions, PNSB can potentially divert 100% of electrons from an organic substrate to H2 production (43). This is a considerable advantage over fermentative bacteria, which can only divert a theoretical 1/3 of electrons (in practice about 15%) from highcarbohydrate waste streams for H2 production (44). Photoheterotrophs typically utilize the smaller organic acids that are often produced but not metabolized, during dark fermentation. Thus, waste streams from photofermentation contain fewer byproducts as the organic compounds are fully reduced to form H2 and CO2. Nitrogenases are expressed in bacteria growing on poor nitrogen sources, under ammonia deprivation, when cells must fix nitrogen to meet the needs of growing cells. Nitrogenases continue to function in the absence of nitrogen gas, directing the flow of electrons to the reduction of hydrogen instead of fixing nitrogen. Under such conditions, H2 production is accelerated, producing 4 H2 molecules for every 8 electrons and 16 ATP instead of the 1 hydrogen molecule produced during nitrogen fixation. Nitrogenase will continue to operate in this manner, as its activity is not subject to feedback inhibition by the production of H2 (43). For a more in depth review of photofermentation processes by PNSB, see the work of Harwood, 2008 (43). VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2249
It is also worth mentioning that some PNSB are capable of producing hydrogen via a water-gas shift reaction, where carbon monoxide from the atmosphere reacts with water, producing CO2 and H2: CO(g) + H2O(l) f CO2(g) + H2(g)
(4)
The bacterium Rubrivivax gelatinosus is capable of utilizing CO as a sole carbon source while generating near stoichiometric quantities of H2 (45, 46). This biohydrogen production pathway is thermodynamically favorable (4°GRxn ) -20 kJ mol-1), but the drawbacks would be the low purity of biogas (roughly 50%) and the difficulty in finding reliable feedstocks of CO. Technological Challenges. Much work has been done to optimize nutrient concentrations, electron-donating substrate, and light intensity for H2 production by various strains of PNSB. Barbosa et al. (47) demonstrated the balance involved in optimizing such conditions for Rhodopseudomonas sp., achieving photochemical efficiencies of 6.2% and 0.9% at production rates of 10.2 and 25 mL H2 L-1 h-1, respectively. The substrate conversion efficiency was much greater when light was provided in abundance. The photobioreactor developed by Chen and Chang (2006) for Rhodopseudomonas palustris WP3-6 is a promising balance of optimizing parameters. By stabilizing cells and introducing internal illuminating optical fibers to the photobioreactor, Chen and Chang (48) were able to achieve a 91% acetate conversion efficiency, a production rate of 43.8 mL H2 L-1 h-1 and a photochemical efficiency of 2.32%. Chen et al. (49) also demonstrated success with solar-excited fiber optics for internal illumination to save on energy, yielding production rates of 32.4 mL H2 L-1 h-1. As in cyanobacteria, impairment of uptake hydrogenase enzymes is a fundamental mutation for improving the efficiency of production. Rhodobacter sphaeroides SMV089 is an uptake hydrogenase deficient mutant with a reported production rate of 64.6 mL H2 L-1 h-1 (50). Within PNSB cells, the synthesis of polyhydroxybutyrate (PHB) or polyhydroxyalkanoate (PHA) requires multiple enzymes that compete for electrons with nitrogenase (43). The double knockout mutant Rhodobacter sphaeroides KD131, isolated by Kim et al. (51), is both uptake hydrogenase and PHB synthase impaired and has demonstrated a production rate of 82.5 mL H2 L-1 h-1. Some strains are developed specifically for their ability to utilize certain substrates efficiently. He et al. (52) have identified Rhodobacter capsulatis IR3 as the strain preferential for degrading lactate, a common fermentor effluent component, with a conversion efficiency of 84.8%. The strain produced a biogas of up to 93% H2, suggesting the potential for photofermentation systems to be connected directly fuel cells. By utilizing fermentation waste streams, photofermentation systems can improve the overall efficiency by degrading fermentation byproducts and produce a higher purity biogas with environmental, economic and energy benefits. Shi and Yu (9) describe the efficiency of Rhodopseudomonas capsulata in producing H2 from a synthetic fermentor effluent containing primarily propionate and acetate as organic substrates, with 47% substrate efficiency and 4.62% photochemical efficiency. Investigations by Oh et al. (53) focus on utilizing a single organism, Rhodopseudomonas palustris P4 that can perform both dark and light fermentation to increase substrate conversion efficiency on glucose. The production rate of this system is low, at 1.66 mL H2 L-1 h-1, but the combined fermentations in a single strain did improve H2 yield on glucose over dark fermentation and presents a novel approach for the combined systems. 2250
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
Dark Fermentation (Anaerobic Bacteria) Mechanisms. In dark fermentation, microorganisms anaerobically break down carbohydrate-rich substrates into organic acids and alcohols, releasing H2 and CO2 in the process. The substrate for dark fermentation can often be supplied by several wastewater streams, performing the dual functions of treatment and energy production. A variety of microorganisms can be used, and the consortium that develops in such a reactor will affect the distribution of byproducts, including the makeup of biogas. Typically, fermentors are seeded with a mixture of undefined microorganisms and the strains selected for are based upon the feedstock delivered to the bioreactor and the reactor conditions (residence time, temperature, etc.). Hydrogen-producing sludges are often dominated by Clostridium species (54-57). Members of this genus develop heat resistant spores, and sludge samples are heat-treated to remove methanogens or other bacteria that interfere with biohydrogen production by utilizing the H2 being produced. This is one advantage of dark fermentation systems, as communities of microorganisms are generally more stable and adaptable to changes in environment and feedstock than pure cultures (58), making them better suited for continuous operations and waste stream applications. Pure cultures are more ideally suited for specific tasks and metabolic engineering. Fermentative biohydrogen production is characterized by high production rates but low substrate efficiencies. The low hydrogen yield stems from the fact that carbohydrates are not fully metabolized, leading to incomplete conversion to H2 and CO2 and the generation of byproducts, such as organic acids, in the waste stream. The biogas produced from dark fermentation contains much lower fractions of H2 than are achievable in biophotolysis or photofermentation systems, so a separation step is needed to recover H2. The redox potential needed for electron carriers for hydrogen reduction, ferredoxin and NADH, can only be maintained when the partial pressure of H2 is less than 0.3 atm and 6 × 10-4 atm, respectively (44). Thus, continuous flushing is needed to maintain H2 production. Utilization of waste streams presents challenges for managing infiltration of competing microorganisms from nonsterile feedstocks by pretreatment or maintaining specific environmental conditions to favor H2producing consortia. In addition, identifying consortia or specific strains to metabolize particular feedstocks is a challenge. Technological Challenges. Several different bioreactor configurations have been designed to improve biohydrogen fermentations through advancements in biomass concentration and substrate conversion efficiency. Batch reactors are often used in laboratory experiments; however, they are not practical for industrial applications that would require larger, long-term production cycles. Continuously stirred tank reactor (CSTR) systems allow for steady state operation for longer ranges of time, from 20 to 90 days (54, 59, 60). Chen et al. (59) operated a simple CSTR with production rates comparable to some of the better biophotolysis or photofermentation systems, at 56 mL H2 L-1 h-1 at a biomass concentration of 1.2 g volatile suspended solids (VSS) L-1. Bioreactors that assist in the formation of granules may help stabilize cells and increase biomass concentration for improved biohydrogen production. With the formation of a granular sludge in a CSTR, Fang et al. (54) was able to achieve a 20 g VSS L-1 biomass concentration and a production rate of 541 mL H2 L-1 h-1. Further improvement in biomass concentration and production rates were obtained by Lee et al. (61) using a carrier-induced granular sludge bed (CIGSB) bioreactor with agitation, reaching 40 g VSS L-1 and 9300 mL H2 L-1 h-1. Other improvements can be made by optimizing bioreactor conditions, such as stirring speeds, pH, or substrate
concentration. The 40 g VSS L-1 obtained by Lee et al. (61) was achieved by adding agitation to a CIGSB system that was attaining concentrations up to 26 g VSS L-1 (62). Fang and Liu (63) investigated the effect of pH on substrate conversion in a CSTR with only 1.0 g VSS L-1 and were able to achieve a relatively high substrate conversion of 17.5% for glucose conversion to H2. This substrate conversion efficiency is slightly greater than the 16.33% of the system by Fang et al. (54) or the 17% of the system by Lee et al. (61), even though these systems had a much greater biomass concentrations (See Table 2). Kyazze et al. (60) evaluated the influence of substrate concentration on conversion efficiency in a CSTR, with the aim of lowering energy costs by reducing energy required to heat reactor feedstocks. A production rate of 505 mL H2 L-1 h-1 was maintained at a loading rate of 40 g sucrose L-1, in a reactor sparged with nitrogen. While this is a reasonable production rate, the hydrogen yield in this system was only 9.5%, indicating a high organic loading rate in the waste stream. Strategies for genetic engineering to enhance fermentative biohydrogen production are similar to other systems: inactivation of uptake hydrogenases to reduce oxidation of hydrogen, increased activation of H2-evolving pathways, and inactivation of competing pathways. Escherichia coli is a popular species to work with because of its well characterized genome and metabolism. In E. coli, electrons from sugar metabolism are directed to formate for proton reduction. Yoshida et al. (64) found a 2.8-fold increase in hydrogen production in a strain of E. coli genetically modified to overexpress a formate hydrogen lyase (FHL). The FHL complex breaks down formate (CHOO-) into CO2 and H2, a common step in the metabolic breakdown of sugars (glucose breaks down into pyruvate, which is split into acetyl-CoA and formate). Using formate as a substrate allows for complete conversion when overexpressing this complex (64). Yoshida et al. (64) reported an unprecedented 300 000 mL H2 L-1 h-1, 2 orders of magnitude higher than any other system that has been described, by creating a high-cell-density reactor fed formate. Repressing the expression of lactate dehydrogenase, ldhA, or fumarate reductase, frd, reduces competition for electrons from pyruvate in the glucose metabolic pathway and has led to improvements in hydrogen production rates and substrate conversion. Maeda et al. (65, 66) successfully increased both substrate conversion efficiency (by up to 50%) and hydrogen production rates (up to 141-fold increase over wild-type) by combining overexpression of FHL complex genes with the inactivation of several hydrogenase uptake genes and redirecting metabolic pathways by deleting ldhA and frd. Efforts to create recombinant E. coli by adding efficient hydrogenases have also led to improved hydrogen production, with production rates of up to 1.4 L H2 L-1 h-1 (67). Hallenbeck and Ghosh (36) recently wrote a review of the state of technology for fermentative biohydrogen production that thoroughly covers the current efforts for metabolic engineering to improve hydrogen yields. Matching feedstocks to cultures can present a challenge for fermentative hydrogen production. E. coli strains tend to have limited substrate range and expanding the metabolic capabilities is necessary for utilizing waste streams. Penfold and Macaskie (68) have demonstrated a simple solution to this with the insertion of a plasmid to provide the necessary gene for sucrose metabolism in an H2-producing E. coli strain. Cellulosic digestion is of interest for the utilization of biomass waste from forestry, agriculture, and municipal waste as feedstocks for biohydrogen production. Certain strains of Clostridia are capable of metabolizing cellulosic substrates via the secretion of an enzyme complex to the outside of the cell called a “cellulosome”, which binds to cellulose and catalyzes hydrolysis (69). The main challenge for developing a cellulosic fermentation pathway for hydrogen synthesis is
the difficulty in regulating all of the factors that determine the endproducts of the system. This involves a delicate balance of reactor conditions, microbial cultures, and biogas removal to ensure continued production of H2. There is potential to recover ethanol or H2 from direct cellulosic fermentation, without the extensive pretreatment currently used in cellulosic ethanol production. Levin et al. (70) provide a useful extensive review of the challenges for biohydrogen production by direct lignocelluloses fermentation. Dual Systems. To make up for the shortcomings of individual technologies, one approach is to combine technologies. The most common combination is a two-stage dark and photofermentation system. A dark anaerobic reactor produces a mix of H2 and CO2 as it breaks down complex sugars, potentially from a waste stream, leading to the production of small organic acids. The small organic acids produced in dark fermentation are then fed to a photofermentation bioreactor for further breakdown into H2 and CO2, to increase the overall hydrogen yield and produce a waste stream with fewer organic byproducts. It has been established that fermentation effluent can be used for photofermentative hydrogen production (8-10, 50, 53) and two-stage systems have been shown to reduce chemical oxygen demand (COD) load in fermentor effluent by up to 90% (8). Reported substrate conversion efficiencies are 1.5-4.5-fold more efficient in twostage systems than fermentation systems using sugars as substrate. Others have attempted a dual system that combines the two mechanisms by coculture. These systems have limited success because of the difficulty of reaching optimal conditions for the different organisms as well as balancing the reaction rates to achieve a stable consortium (71). Melis and Melnicki (72) ran trials of a dual biophotolysis and photofermentation system, combining Rhodospirillum rubrum and Chlamydomonas reinhardti. This dual system theoretically utilizes a fuller spectrum of solar irradiation, because PNSB typically utilize light from the near infared side of the spectrum while algae utilize the visible spectrum. R. rubrum and C. reinhardtii were successfully cocultivated, but the hydrogen productivity of this system was not given. Another possibility for process design is the use of a single strain to perform both light- and dark-fermentation. Rhodopseudomonas palustris P4 is able to produce H2 by dark fermentation of glucose, and cells transferred to the light are capable of H2 production and consumption of acetate, one of the main byproducts of glucose fermentation (53).
Future Directions for Biohydrogen Research On the basis of the production rates given in Table 2, dark fermentation is the leading mechanism for biohydrogen production. Production rates in Table 2 have been converted to units of milliliters H2 per liter hour in order to compare the size of bioreactor systems required to produce a given quantity of hydrogen. Dark fermentation systems are described with 2-4 orders of magnitude higher hydrogen production rates than photofermentation and biophotolysis systems but at very low substrate conversion efficiency and low H2 enrichment of biogas. Photofermentation systems have the next highest production rates and outperform dark fermentation in substrate conversion efficiency and purity of biogas. Biophotolysis and indirect biophotolysis are able to produce gas of high H2 purity (up to 99.5% (33)) but do not produce hydrogen at competitive rates to photofermentation and fermentation. The overall substrate conversion efficiencies for dual dark and photofermentation systems shows promise and the production rates for dual systems could theoretically be at least as productive as the most productive dark fermentation system. A recent technoeconomic analysis of biohydrogen production, undertaken as part of the EERE Hydrogen Production VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2251
TABLE 3. Future Research Directions for Biohydrogen bioreactor engineering biophotolysis
indirect biophotolysis
environmental optimization
• increase light penetration
• sulfur concentration
• immobilize cells
• light intensity
• continuous system for ongoing production • increase light penetration
• N2 concentration in air space
• stabilize cells
• light intensity
• continuous system for ongoing production • continuous air flow
photofermentation
dark fermentation
• increase light penetration
• nitrogen source
• stabilize cells
• light intensity
• continuous system for ongoing production
• optimal substrate
• maximize cell concentration
• optimal substrate/waste stream • substrate concentration
• granular formation • design for scale-up
• conditions to favor H2-producing microorganisms
metabolic engineering • reduce competition with hydrogenase for electrons • restrict light harvesting complex for improved quantum yield, increased light penetration (truncated antenna) • improve hydrogenase tolerance to O2 • impair uptake hydrogenases, restrict bidirectional hydrogenases • improved efficiency of hydrogenase, favor reduction of hydrogen in bidirectional hydrogenases • increase glycogen storage for nonfilamentous (temporally separated) systems • restrict light harvesting complex for improved quantum yield, increased light penetration (truncated antenna) • impair uptake hydrogenases, restrict bidirectional hydrogenases • impair PHA- or PHB-synthase (reduce competition for electrons with nitrogenase) • improved efficiency of hydrogenase, favor reduction of protons in bidirectional hydrogenases • improve substrate utilization (yield) • direct electrons from pyruvate to formate • reduce competition for electrons from pyruvate by competing pathways in glucose metabolism
• gas separation technology to purify H2
program in 2009 (73), provides a different comparison of these biological production pathways for producing 10 000 kg H2 d-1. Biophotolysis chemostat systems, with 2% photochemical efficiency and a production rate of 124 mL H2 L-1 h-1, were predicted to produce H2 at a cost of $8.15 kg-1. Increasing photochemical efficiency to 9.2% could reduce costs to $2.99 kg H2-1. A similar chemostat system for photofermentation by PNSB, at 3.5% photochemical efficiency and a production rate of 244 mL H2 L-1 h-1, would generate H2 at $10.36 kg-1. The higher production price is due to higher feedstock costs for PNSB, as well as increased capital costs to separate H2 (using current standard technology) based on an assumption of 95% H2 in biogas. Biohydrogen production by fermentation of lignocelluloses feedstocks was also evaluated (73) (including pretreatment of lignocelluloses), operating at about 940 mL H2 L-1 h-1 to have a price of $4.33 kg H2-1. Again, the price is driven up by the cost of feedstocks, in this case assumed to be corn stover (due to costs associated with delivery), and gas separation. On the basis of the costs estimated by James et al (73), biological sources could be competitive with steam methane reforming, based on 2003 cost estimates of $5 gge-1 ($4.55 kg H2-1) for this standard technology. It should be noted, however, that the estimates in the study by James et al. (73) 2252
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
are based on ambitious assumptions of enhanced productivity, using production rates for biophotolysis systems 2 orders of magnitude greater than those currently reported (see Table 2). Such production rates and photochemical efficiencies would come from genetic engineering to truncate light-harvesting complexes and improve hydrogenase O2tolerance. Production costs for fermentation and photofermentation could be reduced in identifying waste streams and colocating hydrogen production with industries generating this waste. Potential substrates for fermentative hydrogen production include energy crops (sugar beet, grasses, including lignocellulose fractions), solid waste (food waste, organic fraction of municipal solid waste), and industrial wastewaters (food industries, pulp and paper industry) (74). Research is needed to determine what pretreatment or bioreactor conditions are needed such that nonsterile waste streams will not contaminate the system and disrupt H2 production. Managing gas streams to maintain redox potentials and separate H2 inexpensively will be fundamental for application of the technology (44). Table 3 summarizes the current research challenges for bioreactor design and metabolic engineering for each of the four biohydrogen mechanisms that will need to be pursued in order to provide a sustainable alternative technology for hydrogen production.
Current biohydrogen technology is not ready for industrial scale production of hydrogen. A decentralized strategy may be more appropriate for hydrogen to reduce costs associated with transport and storage of the elusive, small moleculesized gas. A major recommendation for the U.S. DOE Hydrogen Initiative from the National Academy of Engineering is to explore decentralized systems for hydrogen energy (75). Applications could include on-site production for hydrogen fueling stations for fuel-cell vehicles or on site fermentors with direct links to fuel cells for home-based energy. Levin et al. (76) calculated that a bioreactor of approximately 1000 L, with a hydrogen production rate of approximately 2.95 L L-1 h-1, connected to a proton exchange membrane fuel cell (PEMFC) of 5.0 KW (50% efficiency, 95% utilization) could be used to provide sufficient energy to meet an average residential electricity load. The next phase of biohydrogen research needs to include pilot scale demonstration projects to explore the opportunities for industrial scale production, as described by James et al. (73), as well as localized production and energy generation.
Appendix A Abbreviations U.S. DOE EERE PNSB PS I, II RSM PHB CSTR CIGSB VSS FHL
United States Department of Energy Energy Efficiency and Renewable Energy purple nonsulfur bacteria photosystem I, II response surface methodology polyhydroxybutyrate continuously stirred tank reactor carrier-induced granular sludge bed volatile suspended solids formate hydrogen lyase
Literature Cited (1) U.S. Department of Energy Hydrogen Program FY2008 Progress Report; U.S. Department of Energy: Washington, DC, 2008. (2) U.S. Department of Energy Energy Efficiency and Renewable Energy Fuel Cell Technologies Program Hydrogen Production Natural gas reforming. http://www1.eere.energy.gov/ hydrogenandfuelcells/production/natural_gas.html (accessed February 2010). (3) Roadmap on manufacturing R&D for the hydrogen economy; U.S. Department of Energy Interagency Working Group: Washington, DC, 2005. (4) Das, D. Advances in biohydrogen production processes: an approach towards commercialization. Int. J. Hydrogen Energy, 2009, 34, 7349-7357. (5) Hankamer, B.; Lehr, F.; Rupprecht, J.; Mussgnug, J. H.; Posten, C.; Kruse, O. Photosynthetic biomass and H2 production by green algae: from bioengineering to bioreactor scale-up. Physiol. Plantarum 2007, 131, 10–21. (6) Mason, J. E. World energy analysis: H2 now or later. Energy Policy 2007, 35, 1315–1329. (7) Turner, J.; Sverdrup, G.; Mann, M. K.; Maness, P.-C.; Kroposki, B.; Ghirardi, M. L.; Evans, R. J.; Blake, D. Renewable hydrogen production. Int. J. Hydrogen Energy 2008, 32, 379–407. (8) Chen, C.-Y.; Yang, M.-H.; Yeh, K.-L.; Liu, C.-H.; Chang, J.-S. Biohydrogen production using sequential two-stage dark and photo fermentation processes. Int. J. Hydrogen Energy 2008, 33, 4755–4762. (9) Shi, X.-Y.; Yu, H.-Q. Continuous production of hydrogen from mixed volatile fatty acids with Rhodopseudomonas capsulata. Int. J. Hydrogen Energy 2006, 31, 1641–1647. (10) Tao, Y.; Chen, Y.; Wu, Y.; He, Y.; Zhou, Z. High hydrogen yield from a two-step proces of dark- and photo-fermentation of sucrose. Int. J. Hydrogen Energy 2007, 32, 200–206. (11) Akkerman, I.; Janssen, M.; Rocha, J. M. S.; Reith, J. H.; Wijffels, R. H., Photobiological hydrogen production: Photochemical efficiencey and bioreactor design. In Biomethane and Biohydrogen: Status and perspectives of biological methane and hydrogen production; Reith, J. H., Wijffels, R. H., Barten, H., Eds.; Dutch Biological Hydrogen Foundation: Petten, The Netherlands, 2003; pp 124-145.
(12) Seibert, M.; King, P. W.; Posewitz, M. C.; Melis, A.; Ghirardi, M. L., Photosynthetic water-splitting for hydrogen production. In Bioenergy; Wall, J. D., Harwood, C. S., Demain, A., Eds.; ASM Press: Washington, DC, 2008; pp 273-291. (13) Esper, B.; Badura, A.; Rogner, M. Photosynthesis as a power supply for (bio-)hydrogen production. Trends Plant Sci. 2006, 11 (11), 543–549. (14) Ghirardi, M. L.; Posewitz, M. C.; Maness, P.-C.; Dubini, A.; Yu, J.; Seibert, M. Hydrogenases and hydrogen production in oxygenic photosynthetic organisms. Annu. Rev. Plant Biol. 2007, 58, 71–91. (15) Rousset, M.; Cournac, L. Towards hydrogenase engineering for hydrogen production. In Bioenergy; Wall, J. D., Harwood, C. S., Demain, A., Eds.; ASM Press: Washtington, DC, 2008; pp 249258. (16) Frey, M. Hydrogenases: Hydrogen-activating enzymes. Chembiochem 2002, 3, 153–160. (17) Kruse, O.; Rupprecht, J.; Bader, K.-P.; Thomas-Hall, S.; Schenk, P. M.; Finazzi, G.; Hankamer, B. Improved photobiological H2 production in engineered green algal cells. J. Biol. Chem. 2005, 280 (40), 34170–34177. (18) Tamagnini, P.; Axelsson, R.; Lindberg, P.; Oxelfelt, F.; Wunschiers, R.; Lindblad, P. Hydrogenases and hydrogen metabolism of cyanobacteria. Microbiol. Molec. Biol. Rev. 2002, 66 (1), 1–20. (19) Melis, A.; Zhang, L.; Forestier, M.; Ghirardi, M. L.; Seibert, M. Sustained photobiological hydrogen gas production upon reversible inactivtion of oxygen evolution in the green alga Chlamydomonas reinhardtii. Plant Physiol. 2000, 122, 127–135. (20) Wykoff, D. D.; Davies, J. P.; Melis, A.; Grossman, A. R. The regulation of photosynthetic electron transport during nutrient deprivtion in Chlamydomonas reinhardtii. Plant Physiol. 1998, 117, 129–139. (21) Mussgnug, J. H.; Thomas-Hall, S.; Rupprecht, J.; Foo, A. Engineering photosynthetic light capture: impacts on improved solar energy to biomass conversion. Plant Biotechnol. J. 2007, 5, 802–814. (22) Melis, A. Maximizing light utilization efficiency and hydrogen production in microalgal cultures; U.S. Department of Energy: Washington, DC, 2008; pp 187-190. (23) Jo, J. H.; Lee, D. S.; Park, J. M. Modeling and optimization of photosynthetic hydrogen gas production by green alga Chlamydomonas reinhardtii in sulfur-deprived circumstance. Biotechnol. Prog. 2006, 22, 431–437. (24) Kim, J. P.; Kang, C. D.; Park, Y. H.; Kim, M. S.; Sim, S. J. Enhanced hydrogen production by controlling light intensity in sulfurdeprived Chlamydomonas reinhardtii culture. Int. J. Hydrogen Energy 2006, 31, 1585–1590. (25) Tsygankov, A. A.; Kosourov, S. N.; Tolstygina, I. V.; Ghirardi, M. L.; Seibert, M. Hydrogen production by sulfur-deprived Chlamydomonas reinhardtii under photoautotrophic conditions. Int. J. Hydrogen Energy 2006, 31, 1574–1584. (26) Kosourov, S.; Seibert, M.; Ghiardi, M. L. Effects of extracellular pH on the metabolic pathways in sulfur-deprived H2-producing Chlamydomonas reinhardtii cultures. Plant Cell Physiol. 2003, 44 (2), 146–155. (27) Kosourov, S.; Tsygankov, A.; Seibert, M.; Ghirardi, M. L. Sustained hydrogen photoproduction by Chlamydomonas reinhardtii: effects of culture parameters. Biotechnol. Bioeng. 2002, 78 (7), 731–740. (28) Laurinavichene, T.; Tolstygina, I. V.; Tsygankov, A. The effect of light intensity on hydrogen production by sulfur-deprived Chlamydomonas reinhardtii. J. Biotechnol. 2004, 114, 143–151. (29) Zhang, L.; Melis, A. Probing green algal hydrogen production. Philosophical Trans. R. Soc. London Biol. Sci. 2002, 357, 1499– 1509. (30) Laurinavichene, T.; Kosourov, S.; Ghirardi, M. L.; Seibert, M.; Tsygankov, A. Prolongation of H2 photoproduction by immobilized sulfur-limited Chlamydomonas reinhardtii cultures. J. Biotechnol. 2008, 134, 275–277. (31) Fedorov, A. S.; Kosourov, S.; Ghiardi, M. L.; Seibert, M. Continuous hydrogen photoproduction by Chlamydomonas reinhardtii. Appl. Biochem. Biotechnol. 2005, 121-124, 403– 412. (32) Laurinavichene, T.; Fedorov, A. S.; Ghiardi, M. L.; Seibert, M.; Tsygankov, A. Demonstration of sustained hydrogen photoproduction by immobilized, sulfur-deprived Chlamydomonas reinhardtii cells. Int. J. Hydrogen Energy 2006, 31, 659–667. (33) Torzillo, G.; Scoma, A.; Faraloni, C.; Ena, A.; Johanningmeier, U. Increased hydrogen photoproduction by means of a sulfurdeprived Chlamydomonas reinhardtii D1 protein mutant. Int. J. Hydrogen Energy 2009, 34, 4529–4536. VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2253
(34) Polle, J. E. W.; Kanakagiri, S.-D.; Melis, A. tla1, a DNA insertional transformant of the green alga Chlamydomonas reinhardtii with a truncated light-harvesting chlorophyll antenna size. Planta 2003, 217, 49–59. (35) Cohen, J.; Kim, K.; Posewitz, M. C.; Ghiardi, M. L.; Schulten, K.; Seibert, M.; King, P. W. Molecular dynamics and experimental investigation of H2 and O2 diffusion in [Fe]-hydrogenase. Biochem. Soc. Trans. 2005, 33 (1), 80–83. (36) Hallenbeck, P. C.; Ghosh, D. Advances in fermentative biohydrogen production: the way forward. Trends Biotechnol. 2009, 27 (5), 287–297. (37) Antal, T. K.; Krendeleva, T. E.; Laurinavichene, T.; Makarova, V.; Ghiardi, M. L.; Rubin, A. B.; Tsygankov, A.; Seibert, M. The dependence of algal H2 production on Photosystem II and O2 consumption activities in sulfur-deprived Chlamydomonas reinhardtii cells. Biochim. Biophys. Acta 2003, 1607, 153–160. (38) Troshina, O.; Serebryakova, L.; Sheremetieva, M.; Linblad, P. Production of H2 by the unicellular cyanobacterium Gleocapsa alpicola CALU 743 during fermentation. Int. J. Hydrogen Energy 2002, 27, 1283–1289. (39) Linblad, P.; Christensson, K.; Lindberg, P.; Fedorov, A. S.; Pinto, F.; Tsygankov, A. Photoproduction of H2 by wildtype Anabaena PCC 7120 and a hydrogen uptake deficient mutant: from laboratory experiments to outdoor culture. Int. J. Hydrogen Energy 2002, 27, 1271–1281. (40) Borodin, V. B.; Tsygankov, A.; Rao, K. K.; Hall, D. O. Hydrogen production by Anabaena variabilis PK84 under simulated outdoor conditions. Biotechnol. Bioeng. 2000, 69 (5), 478–485. (41) Sveshnikov, D. A.; Sveshnikova, N. V.; Rao, K. K.; Hall, D. O. Hydrogen metabolism of mutant forms of Anabaena variabilis in continuous cultures and under nutritional stress. FEMS Microbiol. Lett. 1997, 147, 297–301. (42) Burrows, E. H.; Chaplen, F. W. R.; Ely, R. L. Optimization of media nutrient composition for increased photofermentative hydrogen production by Synechocystis sp. PCC 6803. Int. J. Hydrogen Energy 2008, 33, 6092–6099. (43) Harwood, C. S., Nitrogenase-catlyzed hydrogen production by purple non-sulfur photosynthetic bacteria. In Bioenergy; Wall, J. D., Harwood, C. S., Demain, A., Eds.; ASM Press: Washington, D.C., 2008. (44) Angenent, L. T.; Karim, K.; Al-Dahhan, M. H.; Wrenn, B. A.; Domiguez-Espinosa, R. Production of bioenergy and biochemicals from industrial and agricultural wastewater. Trends Biotechnol. 2004, 22, 477–485. (45) Maness, P.-C.; Weaver, P. F. Hydrogen production from a carbonmonoxide oxidation pathway in Rubrivivax gelatinosus. Int. J. Hydrogen Energy 2002, 27, 1407–1411. (46) Maness, P.-C.; Huang, J.; Smolinski, S.; Tek, V.; Vanzin, G. Energy generation from the CO oxidation-hydrogen production pathway in Rubrivivax gelatinosus. Appl. Environ. Microbiol. 2005, 71 (6), 2870–2874. (47) Barbosa, M. J.; Rocha, J. M. S.; Tramper, J.; Wijffels, R. H. Acetate as a carbon source for hydrogen production by photosynthetic bacteria. J. Biotechnol. 2001, 85, 25–33. (48) Chen, C.-Y.; Chang, J.-S. Enhancing phototropic hydrogen production by solid-carrier assisted fermentation and internal optical-fiber illumination. Process Biochem. 2006, 41, 2041– 2049. (49) Chen, C.-Y.; Saratale, G. D.; Lee, C.-M.; Chen, P.-C.; Chang, J.-S. Phototrophic hydrogen production in photobioreactors coupled with solar-energy-excited optical fibers. Int. J. Hydrogen Energy 2008, 33, 6886–6895. (50) Franchi, E.; Tosi, C.; Scolla, G.; Penna, G. D.; Rodriguez, F.; Pedroni, P. M. Metabolically engineered Rhodobacter sphaeroides RV strains for improved biohydrogen photoproduction combined with disposal of food wastes. Mar. Biotechnol. 2004, 6, 552–565. (51) Kim, M. S.; Baek, J.-S.; Lee, J. K. Comparison of H2 accumulation by Rhodobacter sphaeroides KD131 and its uptake hydrogenase and PHB synthase deficient mutant. Int. J. Hydrogen Energy 2006, 31 (1), 121–127. (52) He, D.; Bultel, Y.; Magnin, J.-P.; Roux, C.; Willison, J. C. Hydrogen photosynthesis by Rhodobacter capsulatus and its coupling to a PEM fuel cell. J. Power Sources 2005, 141, 19–23. (53) Oh, Y.-K.; Seol, E.-H.; Kim, M.-S.; Park, S. Photoproduction of hydrogen from acetate by a chemoheterotrophic bacterium Rhodopseudomonas palustris P4. Int. J. Hydrogen Energy 2004, 29, 1115–1121.
2254
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
(54) Fang, H. H. P.; Liu, H.; Zhang, T. Characterization of a hydrogenproducing granular sludge. Biotechnol. Bioeng. 2001, 78 (1), 44–52. (55) Fang, H. H. P.; Zhang, T.; Li, C. Characterization of Fehydrogenase genes diversity and hydrogen-producing population in acidophilic sludge. J. Biotechnol. 2006, 126, 357–364. (56) Fang, H. H. P.; Zhang, T.; Liu, H. Microbial diversity of a mesophilic hydrogen-producing sludge. Appl. Microbiol. Biotechnol. 2002, 58, 112–118. (57) Wang, C.-C.; Chang, C. W.; Chu, C. P.; Lee, D. J.; Chang, B.-V.; Liao, C. S. Producing hydrogen from wastewater sludge by Clostridium bifermentans. J. Biotechnol. 2003, 102, 83–92. (58) Kleerebezem, R.; van Loosdrecht, M. Mixed culture biotechnology for bioenergy production. Curr. Opin. Biotechnol. 2007, 18, 207–212. (59) Chen, C.-C.; Lin, C.-Y.; Chang, J.-S. Kinetics of hydrogen production with continuous anaerobic cultures utilizing sucrose as the limiting substrate. Appl. Microb. Biotechnol. 2001, 57, 56–64. (60) Kyazze, G.; Martinez-Perez, N.; Dinsdale, R.; Premier, G. C.; Hawkes, F. R. Influence of substrate concentration on the stability and yield of continuous biohydrogen production. Biotechnol. Bioeng. 2006, 93 (5), 971–979. (61) Lee, K.-S.; Lo, Y.-C.; Lin, P.-J.; Chang, J.-S. Improving biohydrogen production in a carrier-induced granular sludge bed by altering physical configuration and agitation pattern of the bioreactor. Int. J. Hydrogen Energy 2006, 31, 1648–1657. (62) Lee, K.-S.; Wu, J.-F.; Lo, Y.; Lo, Y.-C.; Lin, P.-J.; Chang, J.-S. Anaerobic hydrogen production with an efficient carrierinduced granular sludge bed bioreactor. Biotechnol. Bioeng. 2004, 87 (5), 648–657. (63) Fang, H. H. P.; Liu, H. Effect of pH on hydrogen production from glucose by a mixed culture. Bioresour. Technol. 2002, 82, 87–93. (64) Yoshida, A.; Nishimura, T.; Kawaguchi, H.; Inui, M.; Yukawa, H. Enhanced hydrogen production from formic acid by formate hydrogen lyase-overexpressing Escherichia coli strains. Appl. Environ. Microbiol. 2005, 71 (11), 6762–6768. (65) Maeda, T.; Sanchez-Torres, V.; Wood, T. K. Enhanced hydrogen production from glucose by metabolically engineering Escherichia coli. Appl. Environ. Microbiol. 2007, 77, 879–890. (66) Maeda, T.; Sanchez-Torres, V.; Wood, T. K. Metabolic engineering to enhance bacterial hydrogen production. Microb. Biotechnol. 2007, 1 (1), 30–39. (67) Chittibabu, G.; Nath, K.; Das, D. Feasibility studies on the fermentative hydrogen production by recombinant Escherichia coli BL-21. Process Biochem. 2006, 41 (3), 682–688. (68) Penfold, D. W.; Macaskie, L. E. Production of H2 from sucrose by Escherichia coli strains carrying the pUR400 plasmid, which encodes invertase activity. Biotechnol. Lett. 2004, 26, 1879–1883. (69) Lamed, R.; Setter, E.; Bayer, E. Characterization of a cellulosebinding, cellulose-containing complex in Clostridium thermocellum. J. Bacteriol. 1983, 156, 828–836. (70) Levin, D. B.; Carere, C. R.; Cicek, N.; Sparling, R. Challenges for biohydrogen production via direct lignocellulose fermentation. Int. J. Hydrogen Energy 2009, 34, 7390–7403. (71) Redwood, M. D.; Paterson-Beedle, M.; Macaskie, L. E. Integrating dark and light bio-hydrogen production strategies: towards the hydrogen economy. Rev. Environ. Sci. Biotechnol. 2009, 8, 149– 185. (72) Melis, A.; Melnicki, M. R. Integrated biological hydrogen production. Int. J. Hydrogen Energy 2006, 31, 1563–1573. (73) James, B. D.; Baum, G. N.; Perez, J.; Baum, K. N. Technoeconomic boundary analysis of biological pathways to hydrogen production; US Department of Energy: September 2009. (74) Bartacek, J.; Zabranska, J.; Lens, P. N. L. Developments and constraints in fermentative hydrogen production. Biofuels, Bioprod. Biorefining 2007, 1, 201–214. (75) National Academy of Engineering Committee on Alternatives and Strategies for Future Hydrogen Production and Use, The hydrogen economy: opportunities, costs, barriers and R&D needs, 9780309091633; National Academies Press: Washington, DC, 2004. (76) Levin, D. B.; Pitt, L.; Love, M. Biohydrogen production: prospects and limitations to practical application. Int. J. Hydrogen Energy 2004, 29, 173–185.
ES9030613
