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Continuous microbial production of hydrogen gas
0360-3199/82/0806234)6 $03.00/0 Pergamon Press Ltd. © 1982 International Association for Hydrogen Energy.
Int. J. Hydrogen Energy, Voi. 7, No. 8, pp. 623~528, 1982. Printed in Great Britain.
C O N T I N U O U S MICROBIAL P R O D U C T I O N OF H Y D R O G E N GAS J. D. BROSSEAUand J. E. ZAJIC Saskatchewan Research Council, Saskatoon, Saskatchewan, Canada and Department of Biochemical Engineering, University of Western Ontario, London, Ontario. Canada
(Received for publication 11 November 1981) Abstract--Techniques utilized to study microbial hydrogen gas production involved intermittent flow digesters, chemostats (continuous culture), immobilized cells and immobilized enzymes. Overall, the chemostat appears to have the greatest potential to maximize hydrogen gas productivity. Immobilized systems tended to result in lower hydrogen-producing activities which may have been due to diffusion limitation. Organic compounds, such as carbohydrates, stimulate the greatest amount of H2. waste treatment facilities the reported composition of gas evolved ranges from 65 to 90% (v/v) methane, 5 to 35% (v/v) carbon dioxide, 0 to 10% (v/v) hydrogen and small amounts of nitrogen [25]. Similar results were obtained with manure [26] and in rumen gas production where hydrogen gas usually constitutes only 0.05% (v/v) of the total rumen gas production [21] or 3 x 10 -4 atm. [27]. Hydrogen is synthesized first and is then replaced by methane in these environments [27, 28]. Studies have shown that pure cultures of methanogenic bacteria use carbon dioxide as an electron acceptor and molecular hydrogen as an electron donor according to equation (1) [10, 29].
INTRODUCTION There are basically four processes available for the production of hydrogen gas (H2) from non-fossil energy sources. These include water electrolysis, thermochemical and radiolytic processes [1]. Nuclear, either fission or fusion [2], solar [3] or geothermal [4] energies are potential alternative heat sources for the thermal production of hydrogen from water. Electrolytic hydrogen is presently produced competitively for industrial use, but only in areas where electricity is inexpensive and readily available. For the economic considerations involving the use of electrolytic hydrogen as a fuel source in the future, see refs. [5, 6].
4H2 + CO2-~ CH4 + 2H20.
RESULTS AND DISCUSSION
Biological production of hydrogen by mixed cultures A further process for the production of fuels is biologically based. An example is the microbial formation of hydrogen or methane under anaerobic conditions. The history of methane production is reviewed in refs. [7-10]. Engineering theory and design are treated by Andrews and Graef [11] and Lawrence [12]. Hydrogen gas production is reviewed in refs. [13-18]. A small group of bacteria is responsible for the formation of methane (methanogenesis). The methanogenic bacteria are readily found in anaerobic environments where organic matter is being decomposed. Such environments include swamps [8], lake sediments and rice paddies [19], oil strata [20], coal fields [19], digestive tract of animals, ruminants [21] and non-ruminants [22], cattle waste [23] and sewage sludge digestion [24]. A microbial food chain exists in these mixed-culture environments. Cellulose or other organic polymers are hydrolysed by extracellular enzymes to carbohydrates, which are in turn fermented to a variety of organic acids, alcohols, carbon dioxide and hydrogen. Acetate, methanol, formate or carbon dioxide and hydrogen are the preferred substrates for methanogenic bacteria. Traces of hydrogen gas are usually found along with carbon dioxide in methane fermentations. In anaerobic 623
(1)
Certain other carbohydrates or intermediates are first converted to carbon dioxide which is reduced to methane if hydrogen gas is present [30]. Evidence to support hydrogen utilization in methane synthesis is supported by three lines of evidence. First, Stephenson and Stickland [31], Hungate [27] and Bryant et al. [29] demonstrated an absolute dependence of methane formation upon the presence of hydrogen by the methanogenic organisms. Second, studies with specific inhibitors of methanogenesis [28, 32, 33], in anaerobic digestion demonstrated expected hydrogen gas accumulation in every instance. However, accumulation of hydrogen gas was never as great as expected. Third, the addition of glucose [34] led to high rates of hydrogen gas and carbon dioxide production within 24 h. The disappearance of hydrogen was followed by the concomitant appearance of methane.
Substrates associated with hydrogen evolution Hydrogen gas can be produced from sewage, garbage and agriculture wastes by microbial processes. The evolution of hydrogen gas by anaerobic digesting sewage sludge was studied by disrupting the interaction of the methanogenic and heterotrophic (acidogenic) bacteria involved [28]. Disruption was achieved by either inhibiting methanogenesis with carbon tetrachloride or by adding pulse loads of specific organic substrates.
J. D. BROSSEAU AND J. E. ZAJIC
624 II
o Total gas I0
• H2
T
A C02 • CH4
T._
E
2
o
i 0
I
Time o f t ~
l~ul~,
days
Fig. 1. Rates of gas evolution from a digester pulse loaded with glucose at 13.3 mg m1-1 [28]. The addition of organic substrates to intermittent flow digesters was for the purpose of increasing the specific growth rate of the heterotrophic bacteria to a greater extent than that of the methanogenic bacteria. Hydrogen gas evolved only after pulse loads of carbohydrates (cellulose, starch, glucose). Glucose induced more hydrogen than starch or cellulose. At the highest glucose loading levels (13.3 mg ml-t), the maximum rate of hydrogen production lagged behind the maximum rate of gas evolution (Fig. 1). The amount of hydrogen that accumulated increased with the difference in the growth rates of the hydrogen-forming acidogenic bacteria and the hydrogen-consuming methanogenic bacteria. The greatest amounts of hydrogen occurred after methanogenesis had failed.
Continuous hydrogen fermentation of glucose Continuous-flow microbiological culture techniques
(chemostat) have been applied to fermentations such as biological waste treatment processes for many years [11]. Monod [35] was primarily responsible for the initiation of the continuous culture technique on a laboratory scale. Continuous culture systems have certain advantages [36]. Productivity is one of them. The productivity of a continuous culture is usually higher than that for batch culture. One of the reasons is that the growth rate can be held constant. It also requires time and work to clean, refill, sterilize and re-establish a cell population under batch culture conditions. There are also difficulties to be considered. These include problems with maintenance, sterility and culture stability. A chemostat culture system was applied to the production of biomass and hydrogen gas by Otrobacter intermedius [37]. The growth medium used, based on previous batch results, was (% w/v): glucose, 0.77; (NH4)2SO4, 0.1; Na2S203"5HzO, 0.1; MgSO4, 0.05; NaCI, 0.2; KH2PO4, 2.0; KzHPO4, 0.5. By using chemostat cultures, biomass, substrate and gas balances can be readily determined for each dilution rate (D) and therefore for each specific growth rate (#). This provides an excellent means for kinetic and physiological studies, such as the evaluation of maintenance metabolism (m) of the micro-organism. This parameter is a measure of the energy expended in functions other than those required for cell growth [38]. The slower the growth rate the larger the proportion of the total substrate consumed for maintenance. This gives, as a result, less available substrate for biomass synthesis which, in turn, diminishes the substrate yield coefficient YxJs [39]. A series of experiments showed the relationship between dilution rate (D) and cell concentration (X), biomass productivity (Px), glucose utilized (S) and the rate of hydrogen production (PH0 (Fig. 2). Biomass productivity was calculated by the equation Px = X D (g 1. of reactor vol. h-l). The productivity curve (Fig. 2) declines very sharply at a dilution rate of 0.26 h-L The productivity was 0.2 g 1-~ h-L The calculation of Px in batch experiments
t.0-
08-
4 •
;.
o.6-
.~ .
24
~14o 30 ~o
~o1~ ~,o 16
04
20
;2
i
02--
0--
0
OI Dilution rote,
02
,o
0.3
h -I
Fig. 2. The relationship between biomass concentration (X), productivity (XD), glucose utilized (So - S) mmol 1.-1 and gas (H2, PHz) productivity (mmol 1.-] h -1) of Citrobacter intermedius in a 11. chemostat fermenter as a function of dilution rate. The pH was controlled at 6.0 ± 0.1, temperature at 37°C, and agitation at 1.3 m s-L
625
CONTINUOUS MICROBIAL PRODUCTION OF HE Table 1. Effect of the dilution, rate on biomass concentration, yield, H2 and biomass productivities by C. intermedius in continuous culture
D
X
Ys
% H2
Yr~rs
Yco21s
YHZX
Px
P~
PH2
Pco:
0.22 0.20 0.17 0.12 0.07
0.88 0.90 0.86 0.79 0.50
0.114 0.112 0.106 0.094 0.064
57 65 69 74 80
575 493 426 360 267
489 265 191 126 76
28 24 22 21 23
0.19 0.18 0.14 0.13 0.03
10 6.5 4.5 2.4 1.0
5.7 4,2 3.1 1.8 0.8
4.3 2.3 1.4 0.6 0.2
D = Dilution rate (h -~) =/~ (at steady state); X = biomass (g 1.-1); S = glucose (g 1. 1); Ys = yield of biomass (g) per (g) glucose; Ynvs = yield of H2 (retool) per tool of glucose = Pnvso; Yco~s = yield of CO2 (mmol) per mol of glucose = Pcovso; YH2/x = specific yield of H2 per g of biomass = PHxxo; Px = biomass productivity (XD, g l.-~h-l); Pa = gas productivity (mmol per 1. of reactor per h); PH2 = H2 productivity (mmol per 1. of reactor per h); Pco2 = CO2 productivity (mmol per 1. of reactor per h); SD = tool glucose utilized per 1. of reactor per h.
did not include the time required for cell harvesting and preparation for another run. The maximum overall batch productivity (Px) obtained was approximately 0.31 times that obtained in continuous culture at a dilution rate of 0.22 h -1 (Table 1, Fig. 2). The yields and productivities of gas (H2) and biomass produced (Fig. 2) indicate that the concentration of biomass plays an important role in hydrogen production (Table 1). Hydrogen gas yields and rates increased with increases in biomass and dilution rate. The maximum hydrogen yield, Yn~s, was 575 at D = 0.22 h -Z and the maximum productivity, PHi, was 5.7, compared with 2.5 in batch. The specific yield, Yn~x,was relatively constant at all dilution rates but reached a maximum of 28 at 0.22 h -1. The biomass productivity at the latter dilution rate was also maximum with a value of 0.19. The overall yield (Pn~o) of hydrogen was 22.5 mmol (45 rag) of hydrogen per 1. of feed. This corresponds to a 5.8% (w/w) yield per g of glucose. A comparison was made with batch and continuous cultures of C. intermedius on a similar medium. The results shown in Table 2 indicate that the batch fermentation yields were larger whereas the continuous culture productivities (D = 0.22 h -~) were greater. For example, the hydrogen yields (YH~s) were 50% larger with batch while the hydrogen productivity was 220% Table 2. Comparison between batch 1- and 14-1. fermentors and 1 1. chemostat fermenter Fementer (1.)
Yx/s
YHz/S
Ycozs
Px
PH21s
1~ 14b 1c
28.0 31.0 20.6
710 1,140 575
390 600 489
0.06* 0.07* 0.19
32* 57* 126
Yx/s = Biomass, g mol-~ of glucose; Ym/s = yield of H2 (raM) per tool of glucose; Yco~s = yield of CO2 (mM) per tool of glucose; Px = biomass productivity, g 1.-~ h-~; PH~s = H2 productivity, mM I-I2per mol of glucose per h. * Includes cultivation, harvesting and preparation time. b = Batch, c = chemostat.
greater with continuous culture. The main reason for the productivity increase over the batch fermentation appears to be the time required for harvesting and preparation of another run included in the bath productivity determinations. The novel introduction of a continuous culture system to microbial hydrogen production with C. intermedius has demonstrated the usefulness and widespread applicability of the technique to productivity improvement and process development. In general, although the maximum biomass and gas yields obtained were slightly lower as expected when compared with bath results, the maximum overall productivities were 220% higher. Similar increases in productivity reported herein can be extrapolated to other microbial hydrogen producing systems irrespective of the type of anaerobe or substrate utilized. When comparing processes for production formation it is important to consider productivities, final product concentrations as well as product recovery systems. If the productivity of a continuous culture process is greater than a batch process, it may not be economical unless final product concentrations are comparable [36]. In general, many fermentation processes have high recovery costs which can govern their economics rather than fermentation costs. In this study, the fermentation costs appeared to be relatively high especially since a microbial hydrogen production process has 'in-house' applicability reducing recovery costs to a minimum. One technique than can be applied to increase feed concentrations of substrate (glucose), output concentrations and productivities of hydrogen is that of cell recycle. A theoretical approach has indicated that product yields and productivities would be increased seven-fold (seven times) with the application of optimum cell recycle [37]. The involvement of a hydrogen producer with a greater capability than the anaerobe utilized in the present study may allow for a further two- to three-fold increase in hydrogen productivity. This increase along with the increase from cell recycle could clearly reduce the demonstrated cost of hydrogen for a better competitive outlook for microbial hydrogen
626
J. D. BROSSEAU AND J. E. ZAJIC
producing processes. With the escalating costs of fossil fuel and construction costs in the world today, the outlook for microbial production of hydrogen may indeed be economically competitive in the near future. There are apparently other anaerobes that can be obtained from stock culture collections that produce hydrogen in greater amounts from glucose. However, they usually require significant and costly media extracts containing amino acids or vitamins [40]. Other anaerobes can also produce greater quantities of hydrogen on synthetic medium containing cellulose but their rates of growth and gas production are too limiting to warrant any process development work [41].
Immobilized enzymes Some investigations on enzyme immobilization have centred on the chemical or photochemical reduction of ferredoxin by illuminated chloroplasts, followed by the hydrogenase-catalysed reaction between the reduced ferredoxin and protons to produce hydrogen. One scheme which was recently studied was the immobilization of chloroplasts and hydrogenase in separate reactors. Reduced ferredoxin would flow through the hydrogenase bed catalysing hydrogen production, and oxidized ferredoxin would circulate from the hydrogenase bed to the chloroplast bed where it would be reduced by photosynthetic interactions, and oxygen would be formed [42]. Hydrogenase and ferredoxin were obtained from Clostridium pasteurianum. Sodium dithionite was also used in place of the photo-illuminated chloroplast preparation to reduce oxidized ferredoxin. One main problem encountered was the presence of dissolved oxygen in the solution, reoxidizing the reduced ferredoxin before it reached the hydrogenase bed. A relatively rapid separation of the reduced ferredoxin and oxygen produced in the chloroplast reactor was suggested. A second major problem in this and other hydrogenase immobilizations is that activities obtained are low, 5% of free enzyme [42, 43]. On increasing positive charge density in the microenvironment by using polyethyleneimine-cellulose and increasing the volume of counter-anion by substituting phosphate for chloride in the buffer, great increases in stabilization on immobilizing compared with rates of air inactivation were observed using DEAE-cellulose and Tris-HCl buffer [44]. Hydrogenase is absorbed on polyethyleneimine--cellulose with 50% retention of total activity and an increase of half-life for air inactivation from 4 min to 30 h, an improvement of some 400-fold. Similar results are found by using poly(L-lysine)-Sepharose with 60% total activity retention. When, in addition, Tris-H3PO4 is substituted for Tris-HC1, the halflife lengthens almost 3000-fold (Fig. 3), as expected from salting-out experiments. A laboratory has shown [45] that platinum and palladium catalysts can replace hydrogenase for evolution of hydrogen from reduced methyl viologen produced chemically or photosynthetically by chloroplasts. Although there are no efficient synthetic systems for
~1~ I00L
i ,,,
0
2
I Time,
weeks
Fig. 3. Influence of counter-ion on air inactivation of clostridial hydrogenase absorbed on polyethyleneimine-cellulose. Curve (a): The anion exchanger was previously washed with TrisHCI buffer (pH 8) and air inactivation proceeded in 3 mM Tris-HC! buffer (pH 8); curve (b): the anion exchanger was previously washed with Tris--H3PO4 buffer (pH 8) and oxygen inactivation was observed in 3 mM Tris--H3PO4 buffer (pH 8) [44].
direct photolysis of water, this laboratory has developed a totally synthetic system for producing hydrogen from organic compounds in the light [45, 46]. Methyl viologen is reduced when irradiated near 440 nm in the presence of catalytic quantities of proflavin. Hydrogen is evolved from photo-reduced methyl viologen in the presence of hydrogenase [47] or platinum and palladium catalysts. The system is shown schematically in Fig. 4. This system would be practical for light-driven hydrogen production if a municipal or industrial waste product would serve as electron donor. Thiol compounds as electron donors are of particular interest since they are oxidized to disulphides in the photoreaction and the reaction could -OOCCH2\ / CH~COO~"N ~ -OOCcH~NCH2CH2N~. CHzCO0+ H2N , . EOTA Proflavin (IO0) + (I) C H ~ - ' N ~ N ~ - - CH~ Methyl viok~en (IO) hv I 450nm
NHz
I Reducedmethylviolooen I Hydrogenose I or catalysis
I "2,met",' v~,o~en I Fig. 4. Synthetic system for production of hydrogen from organic compounds in the light. EDTA is only one of many possible electron donors. The numbers in parentheses refer to the quantities of the different components required. Proflavin and methyl viologen are required in catalytic quantities. The yield of H2 is stoichiometric to EDTA [46].
CONTINUOUS MICROBIAL PRODUCTION OF H2
627
S-
I q~,O~,06' °
E .
lo
250
~
£ I
0
I
I0 Time,
20
:~ 0 30
h
Fig. 5. Progress curves for hydrogen evolution. Native cells (0.1 g wet) and immobilized cells (0.1 g wet) in 2 ml of 0.1 M phosphate buffer (pH 7.7) containing 0.25 M glucose were incubated at 37°C. O O, Native cells; 0----------0,immobilized cells [48]~ be made cyclic if coupled to a second reaction, chemical or biological, dark or light, which reduced disulphides back to thiols. Immobilized cells
The immobilization of C. butryicum in 10% polyacrylamide gel afforded the hydrogen producing system protection against inactivation by oxygen compared with non-immobilized (free) cells [48]. Maximum hydrogen evolution by free cells occurred within 4 h but lasted only 7 h (Fig. 5). The rate was slow with the immobilized system but evolved hydrogen for more than 20 days (Fig. 6). Protection against acidifying conditions was also demonstrated although the immobilization experiments of the whole cells were batch in nature and non-continuous. The hydrogen-producing system of immobilized whole cells was protected from the deleterious effects of oxygen observed with native cells, although the rate of hydrogen evolution of immobilized cells decreased with extended incubation periods. However, the rate of hydrogen evolution by immobilized cells increased again after being resuspended in a new media. The biochemical fuel cell contained cells of C. butyricum entrapped in polyacrylamide [49]. The gel-cell suspension was polymerized on a platinum black electrode so that one side was coated (0.1 cm thick) with the gel. The electrode was immersed in a glucose-phosphate buffer solution that was connected to a cathode chamber with a bridge. A carbon electrode served as the cathode. The anode potential of the cell was - 0 . 6 5 V against a saturated calomel electrode. A constant current of 1.1-1.2mA was obtained during 15 days of operation. The current was generated by the electrochemical oxidation of hydrogen and formic acid produced by the entrapped cells. The fuel cell required large amounts of cells to produce a relatively small amount of power: 2 4 W kg ~ dry immobilized cells.
~
I
5
I0
Time,
i
15
20
days
Fig. 6. Continuous hydrogen production. Native (0.2 g wet, O .... ©) and immobilized (0.2 g wet, 0------0) cells in 0.1 M phosphate buffer containing 0.25 M glucose were incubated at 37°C for 24 h [48].
CONCLUSION Almost all work on microbial hydrogen gas production has emphasized the biochemical mechanisms involved in such systems. The development of practical systems will require greater emphasis on the bioengineering aspects touched upon above. For example, the effect of increasing mass density with continuous culture systems for cell recycle is apparent. It would be advantageous to investigate stirred tank immobilized whole cells with eventual application to continuous cell systems. Thus, continuing cell techniques should be of expanding practical and academic significance in the future. REFERENCES 1. J. W. Michel, 166th A.C.S. natn. Meet. Div. Fuel Chem. 18, 1 (1973). 2. D. J. Rose, Science 184, 351 (1974). 3. M. Calvin, Science 184, 375 (1974). 4. G. R. Robson, Science 184, 371 (1974). 5. D. P. Gregory, D. Y. Ng and G. M. Long, in Electrochemistry of Cleaner Environments (L O. Blockris, ed.). Plenum Press, New York (1972). 6. D. P. Gregory and J. B. Pangborn, A. Rev. Energy 1,279 (1976). 7. M. J. Pine, in Advances in Chemistry (R. F. Gould, ed.) Series 105. American Chemical Sac., Washington, D.C. (1971). 8. R. S. Wolfe, Adv. Microbial. Physiol. 6, 107 (1971). 9. G. T. Taylor, Process Biochem. 10, 29 (1975). 10. J. G. Zeikus, Bact. Rev. 41,514 (1977). 11. G. G. Andrews and S. P. Graef, in Advances in Chemistry (R. F. Gould, ed.) Series 105. American Chemical Sac., Washington, D.C. (1971). 12. A. E. Lawrence, in Advances in Chemistry, (R. F. Gould, ed.) Series 105. American Chemical Sac., Washington, D.C. (1971). 13. G. E. Zobel, BullAm. Ass. Petrol. Geol. 31, 1709 (1947). 14. H. Gest, Bact. Rev. 18, 43 (1954). 15. C. T. Grey and H. Gest, Science 148, 186 (1965).
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16. L. E. Mortenson and J. Chen, in Microbial Iron Metabolism (J. B. Neilands, ed.). Academic Press, New York (1974). 17. J. A. Cole, Adv. Microbiol. Physiol. 14, 1 (1977). 18. J. E. Zajic, N. Kosaric and J. D. Brosseau, Adv. Biochem. Engng 9, 57 (1978). 19. T. Koyama, J. geophys. Res. 68, 3971 (1963). 20. F. A. Eksertsev, Geochemistry 18, 432 (1960). 21. R. E. Hungate (ed.), The Rumen and its Microbes, p. 273. Academic Press, New York (1966). 22. D. H. Calloway, Nature 212, 1238 (1966). 23. V. H. Varel, H. R. Isaacson and M. P. Bryant, Appl. envir. Microbiol. 33, 298 (1977). 24. R. M. Sykes and E. J. Kirsch, Devs ind. Microbiol. 11, 357 (1970). 25. W. D. Hatfield, Ind. Engng Chem. 20, 174 (1928). 26. M. Coppenet and G. Ducet, Ann. Agron. 18, 33 (1948). 27. R. E. Hungate, Arch. Mikrobiol. 59, 158 (1967). 28. R. M. Sykes, Ph.D. thesis, Purdue University, Lafayette, Indiana (1970). 29. M. D. Bryant, B. C. McBride and R. S. Wolfe, J. Bact. 95, 1118 (1968). 30. J. G. Zeikus, P. J. Weimer, D. R. Nelson and L. Daniels, Archs Microbiol. 104, 129 (1975). 31. M. Stephenson and L. H. Stickland, Biochem. J. 27, 1528 (1933). 32. E. A. Wolin, R. S. Wolfe and M. J. Wolin, J. Bact. 87, 933 (1964). 33. P. G. Thiel, Water Res. 3, 215 (1969).
34. I. Yamane and K. Sato, PI. Nutr. 10, 127 (1964). 35. J. Monod, Annls Inst. Pasteur, Paris 79, 390 (1950). 36. D. I. C. Wang, C. L. Cooney, A. L. Demain, D. Dunnill, A. E. Humphrey and M. D. Lilly, in Fermentation and Enzyme Technology. John Wiley, Toronto (1979). 37. J. D. Brosseau, Ph.D. thesis, University of Western Ontario, London, Ontario (1980). 38. S. J. Pirt, Proc. R. Soc. Ser. B163, 224 (1965). 39. S. J. Pirt, in Principles of Microbe and Cell Cultivation. Hoisted Press, John Wiley, New York (1975). 40. P. S. May, G. C. Blanchard and R. T. Foley. 18th A. Proc. Power Sources Conf., 19--21 May. Session on Fuel Cell Batteries (1964). 41. P. H. Weimer and J. G. Zeikus, Appl. environ. Microbiol. 33,289 (1977). 42. B. Z. Egan and C. D. Scott, Chem. Tech. 8,304 (1978). 43. D. A. Lappi, F. E. Stolzenbach, N. O. Kaplan and M. D. Kamen, Biochem. biophys. Res. Commun. 69,878 (1976). 44. M. Klibanov, N. O. Kaplan and D. Kamen, Proc. natn. Acad. Sci. U.S.A. 75, 3640 (1978). 45. A. I. Krasna, in Biological Solar Engery Conversion, p. 53. Academic Press, New York (1977). 46. A. I. Krasna, Photochem. Photobiol. 29, 267 (1979). 47. L. Yu and M. J. Wolin, J. Bact. 98, 51 (1969). 48. I. Karube, T. Matsunaga, S. Tsuri and S. Suzuki, Biochim. biophys. Acta 444, 338 (1976). 49. I. Karube, T. Matsunaga, S. Tsuri and S. Suzuki, Biotech. Bioengng 19, 1727 (1977).
Int. J. Hydrogen Energy, Voi. 7, No. 8, pp. 623~528, 1982. Printed in Great Britain.
C O N T I N U O U S MICROBIAL P R O D U C T I O N OF H Y D R O G E N GAS J. D. BROSSEAUand J. E. ZAJIC Saskatchewan Research Council, Saskatoon, Saskatchewan, Canada and Department of Biochemical Engineering, University of Western Ontario, London, Ontario. Canada
(Received for publication 11 November 1981) Abstract--Techniques utilized to study microbial hydrogen gas production involved intermittent flow digesters, chemostats (continuous culture), immobilized cells and immobilized enzymes. Overall, the chemostat appears to have the greatest potential to maximize hydrogen gas productivity. Immobilized systems tended to result in lower hydrogen-producing activities which may have been due to diffusion limitation. Organic compounds, such as carbohydrates, stimulate the greatest amount of H2. waste treatment facilities the reported composition of gas evolved ranges from 65 to 90% (v/v) methane, 5 to 35% (v/v) carbon dioxide, 0 to 10% (v/v) hydrogen and small amounts of nitrogen [25]. Similar results were obtained with manure [26] and in rumen gas production where hydrogen gas usually constitutes only 0.05% (v/v) of the total rumen gas production [21] or 3 x 10 -4 atm. [27]. Hydrogen is synthesized first and is then replaced by methane in these environments [27, 28]. Studies have shown that pure cultures of methanogenic bacteria use carbon dioxide as an electron acceptor and molecular hydrogen as an electron donor according to equation (1) [10, 29].
INTRODUCTION There are basically four processes available for the production of hydrogen gas (H2) from non-fossil energy sources. These include water electrolysis, thermochemical and radiolytic processes [1]. Nuclear, either fission or fusion [2], solar [3] or geothermal [4] energies are potential alternative heat sources for the thermal production of hydrogen from water. Electrolytic hydrogen is presently produced competitively for industrial use, but only in areas where electricity is inexpensive and readily available. For the economic considerations involving the use of electrolytic hydrogen as a fuel source in the future, see refs. [5, 6].
4H2 + CO2-~ CH4 + 2H20.
RESULTS AND DISCUSSION
Biological production of hydrogen by mixed cultures A further process for the production of fuels is biologically based. An example is the microbial formation of hydrogen or methane under anaerobic conditions. The history of methane production is reviewed in refs. [7-10]. Engineering theory and design are treated by Andrews and Graef [11] and Lawrence [12]. Hydrogen gas production is reviewed in refs. [13-18]. A small group of bacteria is responsible for the formation of methane (methanogenesis). The methanogenic bacteria are readily found in anaerobic environments where organic matter is being decomposed. Such environments include swamps [8], lake sediments and rice paddies [19], oil strata [20], coal fields [19], digestive tract of animals, ruminants [21] and non-ruminants [22], cattle waste [23] and sewage sludge digestion [24]. A microbial food chain exists in these mixed-culture environments. Cellulose or other organic polymers are hydrolysed by extracellular enzymes to carbohydrates, which are in turn fermented to a variety of organic acids, alcohols, carbon dioxide and hydrogen. Acetate, methanol, formate or carbon dioxide and hydrogen are the preferred substrates for methanogenic bacteria. Traces of hydrogen gas are usually found along with carbon dioxide in methane fermentations. In anaerobic 623
(1)
Certain other carbohydrates or intermediates are first converted to carbon dioxide which is reduced to methane if hydrogen gas is present [30]. Evidence to support hydrogen utilization in methane synthesis is supported by three lines of evidence. First, Stephenson and Stickland [31], Hungate [27] and Bryant et al. [29] demonstrated an absolute dependence of methane formation upon the presence of hydrogen by the methanogenic organisms. Second, studies with specific inhibitors of methanogenesis [28, 32, 33], in anaerobic digestion demonstrated expected hydrogen gas accumulation in every instance. However, accumulation of hydrogen gas was never as great as expected. Third, the addition of glucose [34] led to high rates of hydrogen gas and carbon dioxide production within 24 h. The disappearance of hydrogen was followed by the concomitant appearance of methane.
Substrates associated with hydrogen evolution Hydrogen gas can be produced from sewage, garbage and agriculture wastes by microbial processes. The evolution of hydrogen gas by anaerobic digesting sewage sludge was studied by disrupting the interaction of the methanogenic and heterotrophic (acidogenic) bacteria involved [28]. Disruption was achieved by either inhibiting methanogenesis with carbon tetrachloride or by adding pulse loads of specific organic substrates.
J. D. BROSSEAU AND J. E. ZAJIC
624 II
o Total gas I0
• H2
T
A C02 • CH4
T._
E
2
o
i 0
I
Time o f t ~
l~ul~,
days
Fig. 1. Rates of gas evolution from a digester pulse loaded with glucose at 13.3 mg m1-1 [28]. The addition of organic substrates to intermittent flow digesters was for the purpose of increasing the specific growth rate of the heterotrophic bacteria to a greater extent than that of the methanogenic bacteria. Hydrogen gas evolved only after pulse loads of carbohydrates (cellulose, starch, glucose). Glucose induced more hydrogen than starch or cellulose. At the highest glucose loading levels (13.3 mg ml-t), the maximum rate of hydrogen production lagged behind the maximum rate of gas evolution (Fig. 1). The amount of hydrogen that accumulated increased with the difference in the growth rates of the hydrogen-forming acidogenic bacteria and the hydrogen-consuming methanogenic bacteria. The greatest amounts of hydrogen occurred after methanogenesis had failed.
Continuous hydrogen fermentation of glucose Continuous-flow microbiological culture techniques
(chemostat) have been applied to fermentations such as biological waste treatment processes for many years [11]. Monod [35] was primarily responsible for the initiation of the continuous culture technique on a laboratory scale. Continuous culture systems have certain advantages [36]. Productivity is one of them. The productivity of a continuous culture is usually higher than that for batch culture. One of the reasons is that the growth rate can be held constant. It also requires time and work to clean, refill, sterilize and re-establish a cell population under batch culture conditions. There are also difficulties to be considered. These include problems with maintenance, sterility and culture stability. A chemostat culture system was applied to the production of biomass and hydrogen gas by Otrobacter intermedius [37]. The growth medium used, based on previous batch results, was (% w/v): glucose, 0.77; (NH4)2SO4, 0.1; Na2S203"5HzO, 0.1; MgSO4, 0.05; NaCI, 0.2; KH2PO4, 2.0; KzHPO4, 0.5. By using chemostat cultures, biomass, substrate and gas balances can be readily determined for each dilution rate (D) and therefore for each specific growth rate (#). This provides an excellent means for kinetic and physiological studies, such as the evaluation of maintenance metabolism (m) of the micro-organism. This parameter is a measure of the energy expended in functions other than those required for cell growth [38]. The slower the growth rate the larger the proportion of the total substrate consumed for maintenance. This gives, as a result, less available substrate for biomass synthesis which, in turn, diminishes the substrate yield coefficient YxJs [39]. A series of experiments showed the relationship between dilution rate (D) and cell concentration (X), biomass productivity (Px), glucose utilized (S) and the rate of hydrogen production (PH0 (Fig. 2). Biomass productivity was calculated by the equation Px = X D (g 1. of reactor vol. h-l). The productivity curve (Fig. 2) declines very sharply at a dilution rate of 0.26 h-L The productivity was 0.2 g 1-~ h-L The calculation of Px in batch experiments
t.0-
08-
4 •
;.
o.6-
.~ .
24
~14o 30 ~o
~o1~ ~,o 16
04
20
;2
i
02--
0--
0
OI Dilution rote,
02
,o
0.3
h -I
Fig. 2. The relationship between biomass concentration (X), productivity (XD), glucose utilized (So - S) mmol 1.-1 and gas (H2, PHz) productivity (mmol 1.-] h -1) of Citrobacter intermedius in a 11. chemostat fermenter as a function of dilution rate. The pH was controlled at 6.0 ± 0.1, temperature at 37°C, and agitation at 1.3 m s-L
625
CONTINUOUS MICROBIAL PRODUCTION OF HE Table 1. Effect of the dilution, rate on biomass concentration, yield, H2 and biomass productivities by C. intermedius in continuous culture
D
X
Ys
% H2
Yr~rs
Yco21s
YHZX
Px
P~
PH2
Pco:
0.22 0.20 0.17 0.12 0.07
0.88 0.90 0.86 0.79 0.50
0.114 0.112 0.106 0.094 0.064
57 65 69 74 80
575 493 426 360 267
489 265 191 126 76
28 24 22 21 23
0.19 0.18 0.14 0.13 0.03
10 6.5 4.5 2.4 1.0
5.7 4,2 3.1 1.8 0.8
4.3 2.3 1.4 0.6 0.2
D = Dilution rate (h -~) =/~ (at steady state); X = biomass (g 1.-1); S = glucose (g 1. 1); Ys = yield of biomass (g) per (g) glucose; Ynvs = yield of H2 (retool) per tool of glucose = Pnvso; Yco~s = yield of CO2 (mmol) per mol of glucose = Pcovso; YH2/x = specific yield of H2 per g of biomass = PHxxo; Px = biomass productivity (XD, g l.-~h-l); Pa = gas productivity (mmol per 1. of reactor per h); PH2 = H2 productivity (mmol per 1. of reactor per h); Pco2 = CO2 productivity (mmol per 1. of reactor per h); SD = tool glucose utilized per 1. of reactor per h.
did not include the time required for cell harvesting and preparation for another run. The maximum overall batch productivity (Px) obtained was approximately 0.31 times that obtained in continuous culture at a dilution rate of 0.22 h -1 (Table 1, Fig. 2). The yields and productivities of gas (H2) and biomass produced (Fig. 2) indicate that the concentration of biomass plays an important role in hydrogen production (Table 1). Hydrogen gas yields and rates increased with increases in biomass and dilution rate. The maximum hydrogen yield, Yn~s, was 575 at D = 0.22 h -Z and the maximum productivity, PHi, was 5.7, compared with 2.5 in batch. The specific yield, Yn~x,was relatively constant at all dilution rates but reached a maximum of 28 at 0.22 h -1. The biomass productivity at the latter dilution rate was also maximum with a value of 0.19. The overall yield (Pn~o) of hydrogen was 22.5 mmol (45 rag) of hydrogen per 1. of feed. This corresponds to a 5.8% (w/w) yield per g of glucose. A comparison was made with batch and continuous cultures of C. intermedius on a similar medium. The results shown in Table 2 indicate that the batch fermentation yields were larger whereas the continuous culture productivities (D = 0.22 h -~) were greater. For example, the hydrogen yields (YH~s) were 50% larger with batch while the hydrogen productivity was 220% Table 2. Comparison between batch 1- and 14-1. fermentors and 1 1. chemostat fermenter Fementer (1.)
Yx/s
YHz/S
Ycozs
Px
PH21s
1~ 14b 1c
28.0 31.0 20.6
710 1,140 575
390 600 489
0.06* 0.07* 0.19
32* 57* 126
Yx/s = Biomass, g mol-~ of glucose; Ym/s = yield of H2 (raM) per tool of glucose; Yco~s = yield of CO2 (mM) per tool of glucose; Px = biomass productivity, g 1.-~ h-~; PH~s = H2 productivity, mM I-I2per mol of glucose per h. * Includes cultivation, harvesting and preparation time. b = Batch, c = chemostat.
greater with continuous culture. The main reason for the productivity increase over the batch fermentation appears to be the time required for harvesting and preparation of another run included in the bath productivity determinations. The novel introduction of a continuous culture system to microbial hydrogen production with C. intermedius has demonstrated the usefulness and widespread applicability of the technique to productivity improvement and process development. In general, although the maximum biomass and gas yields obtained were slightly lower as expected when compared with bath results, the maximum overall productivities were 220% higher. Similar increases in productivity reported herein can be extrapolated to other microbial hydrogen producing systems irrespective of the type of anaerobe or substrate utilized. When comparing processes for production formation it is important to consider productivities, final product concentrations as well as product recovery systems. If the productivity of a continuous culture process is greater than a batch process, it may not be economical unless final product concentrations are comparable [36]. In general, many fermentation processes have high recovery costs which can govern their economics rather than fermentation costs. In this study, the fermentation costs appeared to be relatively high especially since a microbial hydrogen production process has 'in-house' applicability reducing recovery costs to a minimum. One technique than can be applied to increase feed concentrations of substrate (glucose), output concentrations and productivities of hydrogen is that of cell recycle. A theoretical approach has indicated that product yields and productivities would be increased seven-fold (seven times) with the application of optimum cell recycle [37]. The involvement of a hydrogen producer with a greater capability than the anaerobe utilized in the present study may allow for a further two- to three-fold increase in hydrogen productivity. This increase along with the increase from cell recycle could clearly reduce the demonstrated cost of hydrogen for a better competitive outlook for microbial hydrogen
626
J. D. BROSSEAU AND J. E. ZAJIC
producing processes. With the escalating costs of fossil fuel and construction costs in the world today, the outlook for microbial production of hydrogen may indeed be economically competitive in the near future. There are apparently other anaerobes that can be obtained from stock culture collections that produce hydrogen in greater amounts from glucose. However, they usually require significant and costly media extracts containing amino acids or vitamins [40]. Other anaerobes can also produce greater quantities of hydrogen on synthetic medium containing cellulose but their rates of growth and gas production are too limiting to warrant any process development work [41].
Immobilized enzymes Some investigations on enzyme immobilization have centred on the chemical or photochemical reduction of ferredoxin by illuminated chloroplasts, followed by the hydrogenase-catalysed reaction between the reduced ferredoxin and protons to produce hydrogen. One scheme which was recently studied was the immobilization of chloroplasts and hydrogenase in separate reactors. Reduced ferredoxin would flow through the hydrogenase bed catalysing hydrogen production, and oxidized ferredoxin would circulate from the hydrogenase bed to the chloroplast bed where it would be reduced by photosynthetic interactions, and oxygen would be formed [42]. Hydrogenase and ferredoxin were obtained from Clostridium pasteurianum. Sodium dithionite was also used in place of the photo-illuminated chloroplast preparation to reduce oxidized ferredoxin. One main problem encountered was the presence of dissolved oxygen in the solution, reoxidizing the reduced ferredoxin before it reached the hydrogenase bed. A relatively rapid separation of the reduced ferredoxin and oxygen produced in the chloroplast reactor was suggested. A second major problem in this and other hydrogenase immobilizations is that activities obtained are low, 5% of free enzyme [42, 43]. On increasing positive charge density in the microenvironment by using polyethyleneimine-cellulose and increasing the volume of counter-anion by substituting phosphate for chloride in the buffer, great increases in stabilization on immobilizing compared with rates of air inactivation were observed using DEAE-cellulose and Tris-HCl buffer [44]. Hydrogenase is absorbed on polyethyleneimine--cellulose with 50% retention of total activity and an increase of half-life for air inactivation from 4 min to 30 h, an improvement of some 400-fold. Similar results are found by using poly(L-lysine)-Sepharose with 60% total activity retention. When, in addition, Tris-H3PO4 is substituted for Tris-HC1, the halflife lengthens almost 3000-fold (Fig. 3), as expected from salting-out experiments. A laboratory has shown [45] that platinum and palladium catalysts can replace hydrogenase for evolution of hydrogen from reduced methyl viologen produced chemically or photosynthetically by chloroplasts. Although there are no efficient synthetic systems for
~1~ I00L
i ,,,
0
2
I Time,
weeks
Fig. 3. Influence of counter-ion on air inactivation of clostridial hydrogenase absorbed on polyethyleneimine-cellulose. Curve (a): The anion exchanger was previously washed with TrisHCI buffer (pH 8) and air inactivation proceeded in 3 mM Tris-HC! buffer (pH 8); curve (b): the anion exchanger was previously washed with Tris--H3PO4 buffer (pH 8) and oxygen inactivation was observed in 3 mM Tris--H3PO4 buffer (pH 8) [44].
direct photolysis of water, this laboratory has developed a totally synthetic system for producing hydrogen from organic compounds in the light [45, 46]. Methyl viologen is reduced when irradiated near 440 nm in the presence of catalytic quantities of proflavin. Hydrogen is evolved from photo-reduced methyl viologen in the presence of hydrogenase [47] or platinum and palladium catalysts. The system is shown schematically in Fig. 4. This system would be practical for light-driven hydrogen production if a municipal or industrial waste product would serve as electron donor. Thiol compounds as electron donors are of particular interest since they are oxidized to disulphides in the photoreaction and the reaction could -OOCCH2\ / CH~COO~"N ~ -OOCcH~NCH2CH2N~. CHzCO0+ H2N , . EOTA Proflavin (IO0) + (I) C H ~ - ' N ~ N ~ - - CH~ Methyl viok~en (IO) hv I 450nm
NHz
I Reducedmethylviolooen I Hydrogenose I or catalysis
I "2,met",' v~,o~en I Fig. 4. Synthetic system for production of hydrogen from organic compounds in the light. EDTA is only one of many possible electron donors. The numbers in parentheses refer to the quantities of the different components required. Proflavin and methyl viologen are required in catalytic quantities. The yield of H2 is stoichiometric to EDTA [46].
CONTINUOUS MICROBIAL PRODUCTION OF H2
627
S-
I q~,O~,06' °
E .
lo
250
~
£ I
0
I
I0 Time,
20
:~ 0 30
h
Fig. 5. Progress curves for hydrogen evolution. Native cells (0.1 g wet) and immobilized cells (0.1 g wet) in 2 ml of 0.1 M phosphate buffer (pH 7.7) containing 0.25 M glucose were incubated at 37°C. O O, Native cells; 0----------0,immobilized cells [48]~ be made cyclic if coupled to a second reaction, chemical or biological, dark or light, which reduced disulphides back to thiols. Immobilized cells
The immobilization of C. butryicum in 10% polyacrylamide gel afforded the hydrogen producing system protection against inactivation by oxygen compared with non-immobilized (free) cells [48]. Maximum hydrogen evolution by free cells occurred within 4 h but lasted only 7 h (Fig. 5). The rate was slow with the immobilized system but evolved hydrogen for more than 20 days (Fig. 6). Protection against acidifying conditions was also demonstrated although the immobilization experiments of the whole cells were batch in nature and non-continuous. The hydrogen-producing system of immobilized whole cells was protected from the deleterious effects of oxygen observed with native cells, although the rate of hydrogen evolution of immobilized cells decreased with extended incubation periods. However, the rate of hydrogen evolution by immobilized cells increased again after being resuspended in a new media. The biochemical fuel cell contained cells of C. butyricum entrapped in polyacrylamide [49]. The gel-cell suspension was polymerized on a platinum black electrode so that one side was coated (0.1 cm thick) with the gel. The electrode was immersed in a glucose-phosphate buffer solution that was connected to a cathode chamber with a bridge. A carbon electrode served as the cathode. The anode potential of the cell was - 0 . 6 5 V against a saturated calomel electrode. A constant current of 1.1-1.2mA was obtained during 15 days of operation. The current was generated by the electrochemical oxidation of hydrogen and formic acid produced by the entrapped cells. The fuel cell required large amounts of cells to produce a relatively small amount of power: 2 4 W kg ~ dry immobilized cells.
~
I
5
I0
Time,
i
15
20
days
Fig. 6. Continuous hydrogen production. Native (0.2 g wet, O .... ©) and immobilized (0.2 g wet, 0------0) cells in 0.1 M phosphate buffer containing 0.25 M glucose were incubated at 37°C for 24 h [48].
CONCLUSION Almost all work on microbial hydrogen gas production has emphasized the biochemical mechanisms involved in such systems. The development of practical systems will require greater emphasis on the bioengineering aspects touched upon above. For example, the effect of increasing mass density with continuous culture systems for cell recycle is apparent. It would be advantageous to investigate stirred tank immobilized whole cells with eventual application to continuous cell systems. Thus, continuing cell techniques should be of expanding practical and academic significance in the future. REFERENCES 1. J. W. Michel, 166th A.C.S. natn. Meet. Div. Fuel Chem. 18, 1 (1973). 2. D. J. Rose, Science 184, 351 (1974). 3. M. Calvin, Science 184, 375 (1974). 4. G. R. Robson, Science 184, 371 (1974). 5. D. P. Gregory, D. Y. Ng and G. M. Long, in Electrochemistry of Cleaner Environments (L O. Blockris, ed.). Plenum Press, New York (1972). 6. D. P. Gregory and J. B. Pangborn, A. Rev. Energy 1,279 (1976). 7. M. J. Pine, in Advances in Chemistry (R. F. Gould, ed.) Series 105. American Chemical Sac., Washington, D.C. (1971). 8. R. S. Wolfe, Adv. Microbial. Physiol. 6, 107 (1971). 9. G. T. Taylor, Process Biochem. 10, 29 (1975). 10. J. G. Zeikus, Bact. Rev. 41,514 (1977). 11. G. G. Andrews and S. P. Graef, in Advances in Chemistry (R. F. Gould, ed.) Series 105. American Chemical Sac., Washington, D.C. (1971). 12. A. E. Lawrence, in Advances in Chemistry, (R. F. Gould, ed.) Series 105. American Chemical Sac., Washington, D.C. (1971). 13. G. E. Zobel, BullAm. Ass. Petrol. Geol. 31, 1709 (1947). 14. H. Gest, Bact. Rev. 18, 43 (1954). 15. C. T. Grey and H. Gest, Science 148, 186 (1965).
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