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Production Hidrogen
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Iowa State UniversityDigital Repository @ Iowa State University
Retrospective Theses and Dissertations
2006
Biological hydrogen production by anaerobicfermentationWen-Hsing ChenIowa State University
Follow this and additional works at: http://lib.dr.iastate.edu/rtd
Part of the Environmental Engineering Commons
This Dissertation is brought to you for free and open access by Digital Repository @ Iowa State University. It has been accepted for inclusion inRetrospective Theses and Dissertations by an authorized administrator of Digital Repository @ Iowa State University. For more information, pleasecontact [email protected].
Recommended CitationChen, Wen-Hsing, "Biological hydrogen production by anaerobic fermentation " (2006). Retrospective Theses and Dissertations. Paper1863.
Biological hydrogen production by anaerobic fermentation
by
Wen-Hsing Chen
A thesis submitted to the graduate faculty
in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
Major: Civil Engineering (Environmental Engineering)
Program of Study Committee: Shihwu Sung (Major Professor)
Kenneth J. Koehler Say Kee Ong Timothy Ellis
Dennis Bazylinski
Iowa State University
Ames, Iowa
2006
Copyright © Wen-Hsing Chen, 2006. All rights reserved.
UMI Number: 3243819
32438192007
UMI MicroformCopyright
All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.
ProQuest Information and Learning Company 300 North Zeeb Road
P.O. Box 1346 Ann Arbor, MI 48106-1346
by ProQuest Information and Learning Company.
ii
TABLE OF CONTENTS
LIST OF FIGURES ........................................................................................................... iv
LIST OF TABLES............................................................................................................. vi
ABSTRACT...................................................................................................................... vii
CHAPTER 1. INTRODUCTION .......................................................................................1 1.1 Research Background................................................................................................1 1.2 Research Goals..........................................................................................................3 1.3 References .................................................................................................................5
CHAPTER 2. LITERATURE REVIEW ............................................................................8 2.1 Hydrogen Production ................................................................................................8 2.1.1 Water electrolysis...............................................................................................8 2.1.2 Thermal-chemical hydrogen production............................................................9 2.1.3 Biological hydrogen production ......................................................................10
2.2 Dark Fermentation ..................................................................................................11 2.2.1 Fundamentals of dark fermentation .................................................................11
2.3 Environmental Factors Affecting Hydrogen Production ........................................14 2.3.1 Inocula and start-up..........................................................................................14 2.3.2 Substrate...........................................................................................................15 2.3.3 pH.....................................................................................................................16 2.3.4 Hydraulic retention time ..................................................................................17 2.3.5 Product inhibition.............................................................................................18
2.4 Fluorescence in situ Hybridization..........................................................................19 2.4.1 Hybridization reaction kinetics ........................................................................20 2.4.2 Technical aspects of FISH ...............................................................................21 2.4.3 FISH application on wastewater treatment ......................................................23
2.5 References ...............................................................................................................25
CHAPTER 3. KINETIC STUDY OF BIOLOGICAL HYDROGEN PRODUCTION BY ANAEROBIC FERMENTATION ............................................................40
3.1 Abstract ...................................................................................................................40 3.2 Introduction .............................................................................................................41 3.3 Materials and Methods............................................................................................44 3.3.1 Seed microorganisms .......................................................................................44 3.3.2 Hydrogen production experiments...................................................................44 3.3.3 Analysis............................................................................................................45 3.3.4 Data analysis ....................................................................................................45
3.4 Results and Discussion............................................................................................46 3.4.1 Kinetic analysis of hydrogen production .........................................................46 3.4.2 Variations of biomass, pH and carbohydrate at different substrate
concentrations ..................................................................................................48 3.4.3 Growth kinetics of hydrogen-producing bacteria for three different substrates..........................................................................................................49
3.5 Conclusions .............................................................................................................51 3.6 References ...............................................................................................................52
iii
CHAPTER 4. BIOLOGICAL HYDROGEN PRODUCTION IN ANAEROBIC SEQUENCING BATCH REACTOR........................................................68
4.1 Abstract ...................................................................................................................68 4.2 Introduction .............................................................................................................69 4.3 Materials and Methods............................................................................................71 4.3.1 Seed sludge and substrate preparation .............................................................71 4.3.2 Reactor operation .............................................................................................72 4.3.3 Analysis............................................................................................................73
4.4 Results and Discussion............................................................................................74 4.4.1 Effect of HRT ..................................................................................................74 4.4.2 Effect of pH......................................................................................................77 4.4.3 Effect of substrate concentration .....................................................................81 4.4.4 Effect of cyclic duration...................................................................................85
4.5 Conclusions .............................................................................................................87 4.6 References ...............................................................................................................88
CHAPTER 5. DIAGNOSIS OF HYDROGEN FERMENTATION BY FLUORESCENCE IN SITU HYBRIDIZATION...................................103
5.1 Introduction ...........................................................................................................103 5.2 Materials and Methods..........................................................................................105 5.2.1 Reactor operation ...........................................................................................105 5.2.2 FISH...............................................................................................................105 5.2.3 Microscopy and quantificantion analysis.......................................................106
5.3 Results and Discussion..........................................................................................107 5.3.1 Hybridization performance from different 16S rDNA probes.......................107 5.3.2 Quantification of bacteria community ...........................................................109
5.4 Conclusions ...........................................................................................................110 5.5 References .............................................................................................................110
CHAPTER 6. CONCLUSIONS ......................................................................................120 6.1 Closing Statements................................................................................................120 6.2 Recommendations for Future Work......................................................................123
ACKNOWLEDGEMENTS.............................................................................................125
iv
LIST OF FIGURES
Figure 2.1 Catabolic pathway of Clostridia in hydrogen fermentation (Jones and Woods, 1989; Mitchell, 2001).....................................................................................39
Figure 3.1 Cumulative hydrogen production with time: (a) sucrose, (b) NFDM, and (c) food waste ......................................................................................................61
Figure 3.2 Hydrogen yield at different S/X ratios: (a) sucrose, (b) NFDM, and (c) food waste ..............................................................................................................62
Figure 3.3 Specific hydrogen production rate at different S/X ratios: (a) sucrose; (b) NFDM; and (c) food waste ............................................................................63
Figure 3.4 Metabolism of hydrogen-producing bacteria at different sucrose concentrations: (a) mixed liquor volatile suspended solid concentration, (b) final pH, (c) removed carbohydrate concentration, and (d) carbohydrate removal efficiency..........................................................................................64
Figure 3.5 Metabolism of hydrogen-producing bacteria at different NFDM concentrations: (a) mixed liquor volatile suspended solid concentration, (b) final pH, (c) removed carbohydrate concentration, and (d) carbohydrate removal efficiency..........................................................................................65
Figure 3.6 Metabolism of hydrogen-producing bacteria at different food waste concentrations: (a) mixed liquor volatile suspended solid concentration; (b) final pH; (c) removed carbohydrate concentration; and (d) carbohydrate removal efficiency..........................................................................................66
Figure 3.7 Effect of substrate concentration on the hydrogen production rate from different substrate...........................................................................................67
Figure 4.1 Performance of ASBR operated at HRT of 16 h with pH controlled of 6.7: (a) Gas production rate; (b) gas content; (c) acid concentration; and (d) solvent concentration .....................................................................................98
Figure 4.2 Performance of hydrogen fermentation at different pHs: (a) hydrogen yield; (b) hydrogen conversion efficiency; (c) hydrogenic activity; (d) carbohydrate removal efficiency..........................................................................................99
Figure 4.3 Variation of soluble microbial product concentration and MLVSS concentration at different pHs: (a) soluble microbial product concentration; (b) HBu/HAc ratio; and (c) MLVSS concentration .....................................100
Figure 4.4 (a) MLVSS and effluent VSS concentrations; (b) SRT; and (c) F/M ratio at different sucrose concentration ....................................................................101
Figure 4.5 Effect of batch feeding on substrate concentration at different cyclic duration in ASBR (Sung and Dague, 1995) ...............................................................102
v
Figure 5.1 Hydrogen-producing sludge hybridized with the EUB338 probe and stained with DAPI. Scale bars. 5 µm ......................................................................116
Figure 5.2 Clostridium pasteurianum hybridized with the ARC915 probe and stained with DAPI. Scale bars, 5 µm ......................................................................117
Figure 5.3 Escherichia coli hybridized with the ARC915 probe and stained with DAPI. Scale bars, 5 µm...........................................................................................118
Figure 5.4 (a) Hydrogen-producing sludge hybridized with the CLOST I probe labeled by Oregon-green, and stained with DAPI; (b) hydrogen-producing sludge hybridized with the CLOST I probe labeled by Texas-red, and stained with DAPI; and (c) Clostridium pasteurianum hybridized with the CLOST I probe labeled by Texas-red, and stained with DAPI. Scale bars, 5 µm................119
vi
LIST OF TABLES Table 1.1 Life time of fossil fuels with the current consumption rate (Lodhi, 1997)......7
Table 2.1 Summary of hydrogen production from thermo-chemical reactions (Sørensen, 2005) ..............................................................................................................37
Table 2.2 Comparison of biological hydrogen production processes (Nath and Das, 2004) ..............................................................................................................38
Table 3.1 Characteristics of NFDM and food waste......................................................57
Table 3.2 Components of the food waste (on wet weight basis) (Li et al., 2003) .........58
Table 3.3 Estimated parameters of Gompertz equation for hydrogen production.........59
Table 3.4 Summary of growth kinetics of hydrogen-producing bacteria ......................60
Table 4.1 Summary of experimental design ..................................................................94
Table 4.2 Results of hydrogen production at different HRT .........................................95
Table 4.3 Hydrogen production and soluble microbial products in ASBR at each influent sucrose concentration .......................................................................96
Table 4.4 Cyclic effect on hydrogen production and soluble microbial products in ASBR .............................................................................................................97
Table 5.1 16S rDNA oligonucleotide probes used for hybridization...........................114
Table 5.2 Summary of the percentage of EUB338 to DAPI at different operating conditions .....................................................................................................115
vii
ABSTRACT
Considering the energy security and the global environment, there is a pressing
need to develop non-polluting and renewable energy sources. Alternatively, hydrogen is
a clean energy carrier, producing water as its only by-product when it burns. Anaerobic
bioconversion of organic wastes to hydrogen gas is an attractive option that not only
stabilizes the waste/wastewater, but also generates a benign renewable energy carrier.
The purposes of this study were to determine the kinetics of hydrogen production using
different characteristics of substrates and to evaluate hydrogen production potential from
different operating conditions in continuous operation.
The growth kinetics of hydrogen-producing bacteria using three different
substrates including sucrose, non-fat dry milk (NFDM), and food waste were investigated
through a series of batch experiments. The results demonstrated that hydrogen
production potential and hydrogen production rate increased with an increasing substrate
concentration. The maximum hydrogen yields from sucrose, NFDM, and food waste
were 234, 119, and 101 mL/g COD, respectively. The low pH (pH < 4) inhibited
hydrogen production and resulted in lower carbohydrate fermentation at high substrate
concentrations. The Michaelis-Menten equation was employed to model the hydrogen
production rate at different substrate concentrations. The equation gave a good
approximation of the maximum hydrogen production rate and the half saturation constant
(KS) with correlation coefficient (R2) over 0.85. The values of half saturation constant
(KS) for sucrose, NFDM, and food waste were 1.4, 6.6, and 8.7 g COD/L, respectively.
Based on the Ks values, the substrate affinity of the enriched hydrogen-producing culture
viii
was found to depend on the carbohydrate content of the substrate. The substrate
containing high carbohydrates showed a lower KS value. The maximum hydrogen
production rate was governed by the complexity of carbohydrates in the substrate.
Biological hydrogen production from sucrose-rich substrate was investigated in an
anaerobic sequential batch reactor (ASBR). The goal of this study was to investigate the
effect of different hydraulic retention times (HRT) (8, 12, 16, 24, and 48 h), pHs (4.9, 5.5,
6.1, and 6.7), substrate concentrations (15, 25, and 35 g COD/L), and cyclic durations (4,
6, and 8 h) on biological hydrogen production. The maximum hydrogen yield of 2.53
mol H2/mol sucrose consumed and the maximum hydrogenic activity of 538 mL H2/g
VSS-d were obtained at HRT of 16h, pH 4.9, sucrose concentration of 25 g COD/L, and
feeding cycle of 4 h. Methane was detected in the biogas when solids retention time
(SRT) exceeded 100 h at pH of 6.7. Based on the low ethanol concentration of nearly
300 mg/L, the metabolic pathway shift to solvent fermentation was not observed at pH of
4.9. The ratios of butyrate (HBu) to acetate (HAc) decreased from 1.25 to 0.54 mol/mol,
when the sucrose concentration was increased from 15 to 35 g COD/L. This suggests that
the metabolic pathway of acetate fermentation was predominant at higher sucrose
concentrations. Hydrogen production was found to improve at a shorter feeding cycle of
4 h.
Fluorescent in situ hybridization (FISH) was applied for identifying and
quantifying the specific microbial populations in the study. Most bacteria successfully
identified by an EUB338 probe were counted and the percentages of 16S rDNA of
EUB338 to DAPI at different reactor operating conditions were determined. Due to the
false positive hybridization results, the ARC915 probe was found unsuitable for
ix
identifying cells belonging to the domain Archaea in this study. FISH results using the
probe CLOST I were not fully determined because of the difficulty of recognizing the
hybridized clostridia cluster I. Therefore, a correlation between hydrogen production and
the number of Clostridium belonging to clostridia cluster I was not determined.
1
CHAPTER 1. INTRODUCTION
1.1 Research Background
For more than a century, fossil fuels have been extensively used to satisfy the
needs of humans. Today, applications of fossil fuels in humans’ daily lives are very
diverse— fossil fuels furnish electricity, power transportation, provide materials for
clothes and construction, etc. However, there is apprehension that the world’s fossil fuels
are declining and soon will disappear. Based on current consumption rates, they will last
for some finite periods as shown in Table 1.1. As shown in the table, humans must face
the possibility of an oil shortage within 60 years. Even without an immediate risk of
exhausting fossil fuels, fossil fuels might be exhausted within a few centuries.
Another problem is that serious environmental consequences of the extensive use
of fossil fuels have already begun to surface. The excessive use of fossil fuels is one of
the primary causes of global warming and acid rain, which have started to affect the
earth’s climate, weather conditions, and vegetation and aquatic ecosystems (Hansen et al.,
1981). Some researchers believe that greenhouse gas emissions have already caused a
global warming of 0.5ºC to 1.5ºC. Meanwhile, the issue of ozone depletion in the
stratospheric zone is becoming critical, due to the breakage of ozone by N2OX gases
emitted by the combustion of fossil fuels. In addition to this environmental damage, the
economic loss by environmental damage worldwide has been evaluated (Barbir et al.,
1990), and was estimated at $2,360 billion per year or $460 per capita per year in 1990.
It is believed and likely that the cost will be higher sixteen years later—2006.
2
Considering today’s energy security and needs and the global environment, there
is a pressing need to develop non-polluting and renewable energy sources. Hydrogen is a
reasonable alternative energy carrier producing water as its only by-product when it burns.
Therefore, hydrogen is an excellent energy substitute for fossil fuels. The complete
concept of hydrogen production, application, and its scientometric analysis has been
studied since the middle 1980s. The concept comprises hydrogen production from non-
renewable and renewable energy sources, hydrogen transportation and storage, hydrogen
utilization, and hydrogen safety issues (Goltsov et al., 2006). As an aspect of the impact
of hydrogen on human society, some researchers are currently evaluating the future of a
hydrogen economy (Winter, 2005; Milciuvience, et al., 2006), and are beginning to
predict the development of hydrogen civilization and its culture (Ohta, 2006).
As mentioned previously, hydrogen can be produced from non-renewable (coal,
nuclear energy) and renewable energy sources (sun, hydro, wind, biomass, tides, and so
forth). One aspect that must be addressed is that the non-renewable energy sources will
be depleted completely at some point in the future. The process of hydrogen production
from non-renewable sources still has a chance to release environmentally unfriendly or
hazardous wastes. However, renewable energy sources are unlimited, and the process of
hydrogen production has little impact on the environment. Therefore, unquestionably, a
worldwide research goal is the use of hydrogen as a carrier of energy generated from
renewable energy sources (Da Silva et al., 2005; Sherif et al., 2005; Tsai, 2005).
Biological processes evolving hydrogen gas are categorized as a renewable
energy source. This has been the subject of basic and applied research for several
decades. In these biological processes, hydrogen production is carried out by
3
microorganisms; those that can split water into hydrogen and oxygen molecules or those
different that can ferment organic materials into hydrogen (Levin et al., 2004). Based on
the metabolic pathways performed by different groups’ microorganisms, biological
hydrogen production processes can be classified as: (1) biophotolysis of water using
algae and cyanobacteria, (2) photo fermentation of organic materials by photosynthetic
bacteria, and (3) dark fermentation of organic materials using fermentative bacteria
(Hallenbeck and Benemann, 2002). A novel hybrid system of combining dark and photo
fermentations has also been investigated to enhance hydrogen production (Das and
Veziroğlu, 2001).
Hydrogen can be produced from renewable raw materials such as organic wastes.
This is advantageous because there is a need to dispose of human-derived wastes in an
environmentally friendly manner. Some of these wastes are by-products/residuals of
food processing plants and agricultural entities. As an aspect of waste stabilization and
waste reuse, hydrogen production through anaerobic fermentation couples waste
reduction/treatment with recovery of renewable bioenergy. The goal of the proposed
project is to develop an anaerobic fermentation process that generates hydrogen gas from
organic wastes. A process like this may significantly enhance economic viability either
by utilizing hydrogen as a fuel source or as a raw material for industries that consume
hydrogen.
1.2 Research Goals
Hydrogen production through dark fermentation has many advantages compared
to other biological hydrogen production methods because of its ability to continuously
4
produce hydrogen from renewable sources, such as carbohydrate-rich wastes, without an
input of an external energy. Therefore, the proposed research focuses on hydrogen
production through dark fermentation. The main goal of the proposed study is to
understand growth kinetics of hydrogen-producing bacteria through a series of batch
assays, and to explore the potential of hydrogen production in a continuously operating
bioreactor. Fluorescence in situ hybridization (FISH0 technique will be applied to link
specific microbial types with hydrogen production in a continuous bioreactor. The
specific objectives of this study are:
(1) To investigate the effects of different substrates (sucrose, nonfat dry milk (NFDM)
and food waste) on hydrogen production potential with naturally occurring
inocula, i.e., anaerobic digested sludge, and to study the kinetics of hydrogen
production with these substrates, using modified Gompertz and Michaelis-Menten
equations.
(2) To determine the optimal conditions needed to operate an anaerobic sequencing
batch reactor (ASBR) for hydrogen production. Hydraulic retention time (HRT),
pH, substrate concentration, and cyclic durations of ASBR are the primary
parameters optimized for maximizing hydrogen production.
(3) To apply the molecular technique, e.g., FISH to identify and quantify the
hydrogen producing population in the ASBR. The change in microbial population
diversity in the reactor will be continuously monitored, using specific gene probes
with unique sequences.
5
1.3 References
Barbir, F., Veziroglu, T.N., Plass, H.J., Jr., 1990. Environmental damage due to fossil
fuels use. Int. J. Hydrogen Energy 15, 739-749.
Das, D., Veziroğlu, T.N., 2001. Hydrogen production by biological processes: a survey of
literature. Int. J. Hydrogen Energy 26, 13-28.
Da Silva, E.P., Neto, A.J.M., Ferreira, P.F.P., Camargo, J.C., Apolinario, F.R., Pinto,
C.S., 2005. Analysis of hydrogen production from combined photovoltaics, wind
energy and secondary hydroelectricity supply in Brazil. Solar Energy 78, 670-677.
Goltsov, V.A., Veziroglu, T.N., Goltsova, L.F., 2006. Hydrogen civilization of the
future – a new conception of the IAHE. Int. J. Hydrogen Energy 31, 153-159.
Hallenbeck, P.C., Benemann, J.R., 2002. Biological Hydrogen Production; Fundamentals
and Limiting Processes. Int. J. Hydrogen Energy 27, 1185-1193.
Hansen, J., Johnson, D., Lacis, A., Lebedeff, S., Lee, P., Rind, D., Russell, G., 1981.
Climate impact of increasing atmospheric carbon dioxide. Science 213(4511),
957-966.
Levin, D.B., Pitt, L., Love, M., 2004. Biohydrogen production: prospects and limitations
to practical application. Int. J. Hydrogen Energy 29, 173-185.
Lodhi, M.A.K., 1997. Photovoltaics and hydrogen: future energy options. Energy
Convers. 38, 1881-1893.
Milciuviene, S., Milcius, D., Praneviciene, B., 2006. Towards hydrogen economy in
Lithuania. Int. J. Hydrogen Energy 31, 861-866.
6
Ohta, T., 2006. Some thoughts about the hydrogen civilization and the culture
development. Int. J. Hydrogen Energy 31, 161-166.
Sherif, S.A., Barbir, F., Veziroglu, T.N., 2005. Wind energy and the hydrogen economy.
Solar Energy 78, 647-660.
Tsai, W.T., 2005. Current status and development policies on renewable energy
technology research in Taiwan. Renewable and Sustainable Energy Reviews 9,
237-253.
Winter, C.J., 2005. Into the hydrogen energy economy – milestones. Int. J. Hydrogen
Energy 30, 681-685.
7
Table 1.1 Life time for fossil fuels (Lodhi, 1997)
Fossil types Annual rate of consumption Period to last
Oil 2 х 109 barrels 60 yrs
Natural gas 2 x 1012 m3 120 yrs
Coal 3 x 109 metric tons Several centuries
8
CHAPTER 2. LITERATURE REVIEW
2.1 Hydrogen Production
Hydrogen is the most abundant element on earth. It can be found in a many
sources—water, hydrocarbon fuels, inorganic substances, etc. Based on diverse sources
of hydrogen, different technologies have been developed to produce hydrogen gas: (1)
water electrolysis, (2) thermo-chemical hydrogen production, and (3) biological hydrogen
production.
2.1.1 Water electrolysis
Water electrolysis has been a well-known method to uses electric energy to
convert water into molecular hydrogen oxygen gases (Busby, 2005). Since water
electrolysis needs intensive electric energy input, the cost of water electrolysis depends
on the cost of electricity derived from varieties sources. It is common to generate
electricity by combusting fossil fuels (oil, coal, and natural gas) in power plants. In the
U.S., over 50% of the electricity is generated from coal-fired power plants. However,
such combustion is accompanied with an emission of air pollutants, including the
greenhouse gases CO2 and N2O. Other approaches for electrolysis are electricity from
either nuclear power or solar/wind power. Yet, some problems from these sources
include dealing with the production of nuclear waste from the nuclear plants and the
upfront capital costs and maintenance expenses for solar/wind power plants. In sum, the
electrolysis of water to generate hydrogen gas is only cost-effective in some areas where
9
cheap electricity is available or in some applications that demand pure molecular
hydrogen.
2.1.2 Thermo-chemical hydrogen production
Current technologies using thermo-chemical reaction to generate hydrogen gas
include steam reforming, partial oxidation/autothermal reforming, and gasification of
coal and woody biomass (Sørensen, 2005). Steam reforming of natural gas is the process
by which methane and water are heated to a temperature of between 700-1,100 °C and
exposed to a pressure between 3-25 bar with a catalyst. The products are a mixture of
hydrogen and carbon monoxide (Song and Guo, 2006). The mixture of hydrogen and
carbon monoxide is called synthesis gas.
In the process of partial oxidation, methane and oxygen react at high temperatures
and pressure without a catalyst to produce hydrogen and carbon monoxide. A
combination of partial oxidation and steam reforming is an autothermal reforming
process. An autothermal reforming process has been demonstrated to pose higher energy
efficiency than the partial oxidation process (Docter and Lamm, 1999). In addition, it
provides advantages of smaller reaction units and is a more cost-effective for synthesis
gas production (Song and Guo, 2006).
Gasification is the oldest known method for hydrogen production. Gasification
starts with the conversion of coal and woody biomass in the presence of steam and
oxygen at high temperatures into a mixture of hydrogen and carbon monoxide (Stiegel
and Ramezan, 2006). Table 2.1 summaries the main stochiomatric equations of hydrogen
production from thermo-chemical reactions (Sørensen, 2005).
10
2.1.3 Biological hydrogen production
It is well-known there exists three microbial groups that have the potential for
hydrogen production in a biochemical reaction. The first group consists of the
photosynthetic green algae and cyanobacteria. These organisms are autotrophs and
directly split water to molecular hydrogen and oxygen in the presence of light
(Hallenbeck and Benemann, 2002). This biological reaction of splitting water into
hydrogen and oxygen is classified as two categories: (1) direct photolysis by green algae
and (2) indirect photolysis by cyanobacteria (Levin et al., 2004). Since this reaction
requires only water and sunlight, and generates oxygen, it is an attractive option from the
perspective of environmental protection. However, there are still some limitations for the
photolysis. The drawback encountered using green algae is the inhibition by the presence
of oxygen accompanied with the production of hydrogen during direct photolysis (Nath
and Das, 2004). In addition, the cyanobacteria examined so far shows lower
photochemical efficiency, due to the complicated reaction systems needed to overcome
the large Gibb’s free energy (+237 kJ/mol hydrogen) requirements (Miyake, 1998).
The second and third groups of bacteria are heterotrophs that use organic
substrates for hydrogen production. These heterotrophic microorganisms produce
hydrogen under anaerobic conditions, either in the presence or absence of light.
Accordingly, this process is classified as photo fermentation or dark fermentation.
Photosynthetic purple non-sulfur bacteria carry out photo fermentation using organic
acids as a carbon source while light is supplied as an energy source (Levin et al., 2004).
Therefore, thermodynamically, hydrogen production through photo fermentation is not
favorable without light as a source of energy.
11
On the other hand, anaerobic non-photosynthetic fermentative bacteria use
carbohydrate-rich substrates as sources of carbon and energy driving dark fermentation
(Hawkes et al., 2002). Research reports that hydrogen production can be achieved from
renewable feedstock, such as biomass-derived sugars, organic wastes, etc. (Nandi and
Sengupta, 1998; Lay et al., 1999). From an environmental engineering viewpoint, dark
fermentation is of great interest— not only for stabilizing human-derived organic
wastes—but producing a clean and sustainable energy carrier. The comparison of the
aforementioned biological hydrogen production is listed in Table 2.2.
2.2 Dark Fermentation
Dark fermentation is a catabolic process in which bacteria convert sugars and
proteins to carboxylic acids, hydrogen gas, carbon dioxide and organic solvents. This
biological chemical reaction inhibited by the presence of oxygen and thus is only carried
out under anaerobic conditions (Levin et al., 2004).
2.2.1 Fundamentals of dark fermentation
In dark fermentation, different groups of bacteria are known to be responsible for
hydrogen production such as Enterobacter, Clostridium, and Bacillus. Fang et al. (2002)
reported that in a mixed culture study where hydrogen was produced, about 70% of the
population was of the genus Clostridium and 14% belonged to genus Bacillus. A study
conducted in our laboratory also showed that hydrogen production was correlated to the
presence of Clostridium species in the bioreactor (Duangmanee et al., 2002).
12
Clostridium is an anaerobic spore former. In response to hostile conditions, such
as the presence of oxygen, heat, low or high pH, alcohol, toxic compounds, etc.,
Clostridium species are changed from vegetative cells to endospores, which is a stress-
resistant state with greatly reduced metabolic activity. Due to endospore production, the
members of genus Clostridium can be isolated from mixed cultures by heating the
cultures to 70ºC for 10 min to kill vegetative Clostridium cells and nonspore-forming
organisms (Bergey’s manual, 1984). In addition to the selection process of Clostridium
by heat treatment, some species require heat activation for endospore prior to germination
(Mead, 1992).
According to its catabolism, Clostridium can be classified as two groups—
saccharolytic and proteolytic—of fermenting bacteria. Saccharolytic clostridia ferment
carbohydrates, consisting of simple sugars, disaccharides, oligosaccharides, and cellulose;
whereas, proteolytic clostridium hydrolyzes protein and ferments amino acids (Ljungdahl
et al., 1989). However, most proteolytic clostridium can also ferment carbohydrates, and,
hence, carbohydrates are very common substrates for the genus of Clostridium.
With regard to dark fermentation, saccharolytic Clostridium species metabolize
simple sugars using the Embden-Meyerhof-Parnas (EMP) glycolytic pathway, where
glucose is converted to pyruvate, an intermediate of hexose metabolism (Schroder et al.,
1994). Pyruvate is then oxidized by the enzyme pyruvate-ferredoxin oxidoreductase to
yield acetyl CoA, carbon dioxide, and reduced ferredoxin (Hallenbeck and Benemann,
2002). The re-oxidation of reduced ferredoxin is catalyzed by the enzyme hydrogenase
and generates hydrogen gas. The fermentation pathway is shown in Fig. 2.1.
13
Using glucose as a model substrate, hydrogen production is accompanied with
either acetate formation (Eq. 2.1) or butyrate formation (Eq. 2.2) (Miyake, 1998). In
acetate fermentation, 4 ATP molecules are produced; whereas, 3 ATP molecules are
produced in butyrate fermentation. Thus, for the microorganisms, it seems that the
acetate fermentation is energetically more favorable than the butyrate fermentation.
However, based on Gibb’s free energy change, butyrate fermentation is the more
dominant reaction, where it only yields 3.3 ATP molecules and the maximum hydrogen
production of 2.5 moles H2/mole glucose stoichiometrically (Eq. 2.3) (Wood, 1961).
Possible explanations why clostridia use the butyrate fermentation pathway during
hydrogen production are 1) the formation of one equivalent butyrate leads to less
acidification of microorganisms’ environment than the two equivalents of acetate and 2)
generation of a higher amount of butyrate may deplete excess reducing equivalents
(Ljungdahl et al., 1989).
C6H12O6 + 2H2O → 2 CH3COOH + 2CO2 + 4H2 ∆G°-184 kJ (2.1)
C6H12O6 → CH3CH2CH2COOH + 2CO2 +2H2 ∆G°-257 kJ (2.2)
C6H12O6 + 0.5 H2O → 0.75 CH3CH2CH2COOH + 0.5 CH3COOH + 2 CO2 + 2.5 H2 (2.3)
As evident from Fig. 2.1, saccharolytic Clostridium species can convert glucose
not just into hydrogen and organic acids, but solvents as well. In a batch culture,
Clostridium sp. produced hydrogen and organic acids at an exponential growth phase;
whereas, the metabolism shifted to solvent production during the stationary growth phase
(Afschar et al., 1986; Brosseau et al., 1986). Clostridium acetobutylicum, a species know
to produce hydrogen and organic acids, is favored to produce organic solvents at pHs
below 5 (Gottwald and Gottschalk, 1985). In addition, the metabolic pathway of
14
Clostridium pasteurianum showed an abrupt shift from hydrogen and acid production to
solvent production under iron and phosphate limitations, high substrate concentrations
(125 g glucose/L), and the appearance of carbon monoxide (an inhibitor of hydrogenase)
(Dabrock et al., 1992).
2.3 Environmental Factors Affecting Hydrogen Production
In dark fermentation, there are several factors can heavily impact the performance
of biological hydrogen production. They mainly include inocula, substrate, pH, hydraulic
retention time (HRT), and product inhibition.
2.3.1 Inocula and start-up
To carry out biological hydrogen production, abundance of Clostridium species
could be extracted from soil, anaerobic digested sludge, compost, etc. (Van Ginkel et al.,
2001; Lay et al., 2003; Khanal et al., 2005). Since Clostridium can tolerate heat ,which
its morphology and physiology shift to endospore, the heat shock method is a common
treatment to select spores forming Clostridium, while killing nonspore forming bacteria,
such as methanogens, sulfate reducing bacteria, etc. Boiling anaerobic digested sludge
has been broadly used to differentiate Clostridium from other microbial populations in
hydrogen fermentation studies (Lay, 2000; Duangmanee et al., 2002). Meanwhile,
baking compost or soil is the other heat shock method, while the inoculum is obtained
from the solid phase (Fan and Chen, 2004; Khanal et al., 2004; Lay et al., 2005).
In addition to heat shock treatment, acid and base treatments have been developed
as alternative processes for the selection of Clostridium. Research reported that adjusting
15
pH of inocula to 3 or 10 efficiently selected Clostridium and inhibited methanogens
(Chen et al., 2002). Meanwhile, a successful hydrogen production was observed from
different types of pre-acidified inocula including activated sludge, anaerobic digested
sludge, refuse compost, watermelon soil, kiwi soil, and lake sediment (Kawagoshi et al.,
2005). Another study adopted acid-pretreated anaerobic digested sludge for investigating
the effect of sludge immobilization by ethylene-vinyl acetate copolymer to achieve the
goal of preventing biomass washout at a low hydraulic retention time (Wu et al., 2005).
2.3.2 Substrate
Research investigating hydrogen fermentation has been conducted using various
types of substrates. Several studies on the biological hydrogen production from simple
sugars, e.g., glucose, and sucrose have been reported (Majizat et al., 1997; Mizuno et al.,
2000; Van Ginkel et al., 2001; Duangmanee et al., 2002; Lin and Jo, 2003; Khanal et al.,
2004). Some investigators also studied the hydrogen production potential of complex
substrates, e.g., food and food processing wastes (Shin et al., 2004; Van Ginkel et al.,
2005; Wu and Lin, 2004), cellulose containing waste (Okamoto et al., 2000), municipal
solid wastes (Ueno et al., 1995), activated sludge (Wang et al., 2003), etc.
In addition to testing the availability of hydrogen production from different
feedstocks, Lay et al. (2003) compared hydrogen production from carbohydrate-rich,
protein-rich, and fat-rich organic solid wastes. Their batch study results indicated that
hydrogen producing microbes could evolve much more hydrogen from carbohydrate-rich
organic waste. The same conclusion was also obtained from the study of converting bean
curd manufacturing waste (protein-rich waste), rice and wheat bran (carbohydrate-rich
16
waste) into hydrogen, where rice and wheat bran were more favorable for hydrogen
fermentation (Noike and Mizuno, 2000). On the other hand, many studies have already
concentrated on studying the utilization of carbohydrate-rich organic wastes, e.g., rice
winery wastewater (Yu et al., 2002; Yu et al., 2003), and starch-manufacturing
wastewater (Yokoi et al., 2002; Hussy et al., 2003) to carry out hydrogen fermentation.
2.3.3 pH
As mentioned in Section 2.2.1, research reported that pH could affect the
metabolic pathway of dark fermentation in a pure culture. Hence, there is a need to
investigate the effect of pH on hydrogen production in mixed cultures. Many studies
determined the optimal pH from various types of substrates. Batch studies indicated that
the optimum initial pH for hydrogen production using sucrose as a limiting substrate
ranged from 5.5 to 5.7 (Van Ginkel et al., 2001; Khanal et al., 2004; Wang et al, 2005).
Zhang et al. (2003) reported that the optimal initial pH for converting starch to hydrogen
was found at 6.0 under thermal conditions. In addition, a study showed that an initial pH
6.0 was favorable for hydrogen production from cheese whey (Ferchichi et al., 2005).
Fang et al. (2006) found that a better performance of hydrogen production from rice
slurry was obtained at an initial pH of 4.5. Based on these studies, it can be concluded
that an initial pH at slightly acidophilic conditions helps to enhance hydrogen production.
Even though there have been many investigations of the initial pH on hydrogen
production in batch studies, the optimal pH determined from continuous operation is still
limited. In a continuous operation, a pH of 5.5 was found to be the optimum for
hydrogen production from glucose (Fang and Liu, 2002). Lay (2000) optimized
17
hydrogen production by controlling the pH at 5.2 in a starch-synthetic wastewater. In
addition, Lay and his coworkers determined the optimum pH of 5.8, based on the
statistical contour plot analysis in a complete mixed bioreactor converting beer
processing wastes into hydrogen (Lay et al., 2005).
2.3.4 Hydraulic retention time (HRT)
A kinetic study of hydrogen production using sucrose as a limiting substrate
showed the maximum specific growth rate of 0.172 h-1 for hydrogen producers, which
allowed them to retain a continuous stirred tank reactor (CSTR), operating at a short HRT
(Chen et al., 2001). An investigation of the effects of HRT on hydrogen production
indicated that the maximum hydrogen yield of 1.76 mol H2/mol glucose was obtained
from a CSTR operated at HRT of 6 h (Lin and Chang, 1999). In addition to the hydraulic
effect on CSTR, hydrogen fermentation could be carried out at further shorter HRT in the
high rate bioreactor, which can maintain the biomass with an unlimited sludge retention
time. In a three-phase fluidized-bed bioreactor, HRT could be reached as short as 2 h to
accomplish the best hydrogen yield of 2.67 mol H2/mol sucrose (Wu et al., 2003). On the
other hand, a maximal hydrogen yield of 3.03 mol H2/mol sucrose was found at a HRT of
0.5 h in a carried-induced granular sludge bed bioreactor (Lee at al., 2004).
Since varying HRT altered the organic loading rate simultaneously, there is a
concern with the ambiguity between the effect of HRT and the organic loading rate on
hydrogen production. Therefore, a study, examining the influence of HRT and substrate
concentration on continuous hydrogen production by the granular acidogenic sludge at a
18
constant organic loading rate, reported that the maximum yield occurred at HRT of 13.7 h
with a sucrose concentration of 14.3 g/L (Liu and Fang, 2002).
At short HRT, hydrogen consumers, primarily methanogens, could essentially be
washed-out or depleted. Control of HRT could be another strategy to limit the hydrogen
consumers without a pretreatment of seed sludge. Lin and Jo (2003) operated a hydrogen
producing anaerobic sequencing batch reactor (ASBR) with a gradually reducing HRT
until 8 h to completely inhibit methane production.
2.3.5 Product inhibition
During hydrogen fermentation, acetate and butyrate productions are always
accompanied with hydrogen production. However, these products could result in the
feedback of product inhibition to the microbes’ activities. Therefore, product inhibition
is always the critical factor leading to a worse performance scenario in biological
hydrogen reactions. In an early study, Heyndrickx and his coworkers (1987) reported no
significant difference of hydrogen production was found when adding acetic acid up to
18.0 g/L. However, the addition of butyric acid higher than 17.6 g/L began to inhibit the
activity of Clostridium butyricum. In addition, van den Heuvel et al. (1988) agreed with
these results that only butyric acid up to 17.6 g/L inhibited the mixed culture acidogenic
bacteria growth, but acetic acid did not inhibit bacteria growth. An investigation reported
that the IC50 values, which the butyric concentration cause 50% inhibition at bioactivity
of hydrogen producing bacteria, were estimated as 19.39 and 20.78 g/L with respect to
hydrogen production rate and hydrogen yield (Zheng and Yu, 2005).
19
Hydrogen partial pressure in the liquid phase is another factor that might interfere
with biological hydrogen production. Lamed et al. (1988) studied the effect of stirring on
hydrogen production from cellulose and cellobiose. They found a three-fold hydrogen
content with less hydrogen production in the unstirred culture broth when compared to
that in the stirred culture. Accordingly, Lay (2000) demonstrated that increasing the
agitation speed from 100 to 700 rpm in a lab-scale completed mixed reactor could double
the daily hydrogen production rate from starch. The other approach reducing the
hydrogen content showed that the process of nitrogen gas sparging could help enhance
hydrogen yield from 0.85 mol H2/mol glucose to 1.43 mol H2/mol glucose (Mizuno et al.,
2000).
2.4 Fluorescence in situ Hybridization (FISH)
In recent years, culture-independent molecular techniques have been successfully
applied to study the microbial community structure. In particular, fluorescence in situ
hybridization (FISH) technique with 16S rRNA-targeted oligonucleotide probes has been
broadly used to evaluate the phylogenetic identity, morphology, and to quantify and
verify the presence of microorganisms in the environment (Olsen et al., 1986; Amann et
al., 1995). It is a unique and powerful technique, due to its convenient and precise
targeting to specific microbial groups or species. Compared with other hybridization
techniques, there is no need to process RNA or DNA extraction. Therefore, carrying out
FISH saves more time than other hybridization techniques.
20
2.4.1 Hybridization reaction kinetics
Hybridization is a dynamic reaction where denatured target sequences and
complementary single stranded DNA or RNA probes anneal, forming stable double
stranded hybrid molecules (Swiger and Tucker, 1996). Successful hybridization is
performed at just below the melting temperature (Tm) of probes and their targets. The
Tm is the temperature at which half the DNA is present in its single stranded (denatured)
form. It has been well understood that the Tm of any given strand duplex depends on its
base composition and sodium concentration (Schildkraut and Lifson, 1965). The
relationship among these parameters to predict the Tm is determined by:
Tm (ºC) = 16.6 log M + 0.41 (%G+C) + 81.5 (2.4)
where M is the sodium concentration and % G+C is the base composition. However, at
high sodium concentrations (above 0.4 M), sodium concentration has a minor effect on
the Tm. Therefore, the equation can be modified as:
Tm (ºC) = 0.41 (%G+C) + 81.5 (2.5)
In practice, formamide is added to the hybridization solution to reduce the Tm of
probes and targets, so that hybridization can be carried out at lower temperatures. Within
the range of experimental conditions, the Tm is decreased by about 0.7ºC for each
percentage of formamide present in the hybridization solution (Chan et al., 1990).
However, a disadvantage of adding formamide to the hybridization solution may require
a longer hybridization time.
In addition, Swiger and Tucker also (1996) concluded that hybridization can be
optimized by pH, probe length, and probe concentration. Typically, pH of the
hybridization solution is maintained at 6.5 to 7.5. A higher pH can be used to achieve
21
more stringent hybridization conditions, where the binding between the probe and the
target sequence is more specific. Regarding the effect of the probe on hybridization, the
hybridization rate is proportional to the square root of the probe fragment length, while
the higher probe concentration contributes to the higher hybridization rate (Wetmur, 1975;
Wetmur and Davidson, 1968; Britten and Davidson, 1985). On the other hand, the probe
length and the probe concentration affect the rate of formation of the initial short
specifically based-paired region (nucleation reaction).
2.4.2 Technical aspects of FISH
The methodology of FISH technique involves the development of oligonucleotide
probes to permeate microbial cells (Hugenholtz et al., 2002). The probes can be designed
to target microorganisms from species to domain. The probes enter the cells to only
hybridize with their complementary target sequence in the ribosomes. The common
length of an oligonucleotide probe is between 15 and 30-base pair (bp). Probes are
typically labeled with a fluorochrome at the 5’end through an aminolinker during
synthesis (Moter at al., 1998). Therefore, the cells retaining the probes can be observed
under either epifluorescence microscope or confocal laser scanning microscope.
Several crucial steps contribute to FISH work, which include: (1) fixing the
specimen, (2) hybridizing the specific target sequences by the respective probes, (3)
removing unbound probes by a washing step, and (4) mounting and visualizing (Moter
and Göbel, 2000). Fixation of living cells simply means an immediate stop of life
processes taking place within and around the cells. Fixation must be carried out prior to
hybridization in order to achieving good permeabilization for probe penetration,
22
sustaining the maximum level of the target RNA, and maintaining the cell’s integrity. In
general, chemicals used for fixation can be categorized into two groups. For Gram-
negative bacteria, paraformaldehyde of 3 to 4% (v/v) is sufficient for fixation. For Gram-
positive bacteria, either ethanol of 50% (v/v) or ethnol/formalin of 9 to 1 (v/v) is
recommended to use (Jurtshuk et al., 1992; Brown-Howland et al., 1992).
As mentioned previously about hybridization kinetics, the maximum
hybridization reaction with the minimum degree of potential nonspecific targeting with
other sequences can be achieved by adjusting the sodium concentration and the
formamide concentration in the step of hybridization. In addition, hybridization time
must be determined experimentally to optimize hybridization. After hybridization, a
post-hybridization wash is required. This helps remove unbound probes and nonspecific
binding, but does not dissociate the perfectly matched hybrid molecules. Such stringency
of the wash can be regulated by adjusting the sodium concentration in the washing
solution (Lathe, 1985).
Most fluorochromes labeled on the probes will quench rapidly with UV light
excitation. Therefore, prior to making visualization under an epifluorescence microscope,
one must alleviate the quenching problem by mounting antifading agents on the
specimens (Johnson and Nogueira Araujo, 1981). The function of antifading agents is to
extend the intensity of the photon emissions from fluorochromes. Antifading agents are
now commercially available.
For visualization, an epifluorescence microscope, equipped with a lamp and
narrow-band-pass filters, is used to tool the observation. The fluorochrome excited by
the lamp emits the fluorescent light, while the special filter allows the specific
23
wavelength of fluorescent light transmission to obtain the unique signal. Therefore, the
cells containing the hybrid molecules can be observed under the microscope.
2.4.3 FISH application on wastewater treatment
The biological treatment process has been developed for nearly a century. The
first activated-sludge system was built to remove organic matter in Manchester (Ardern
and Lockett, 1914). Thereafter, the outline of the role of microbial consortia in the
biological treatment process was approximately developed regarding the stabilization of
organic matters, but the entire microbial community in the treatment process is still not
completely understood. Nevertheless, a progressive improvement in molecular
biotechnology during the past two decades helps to deeply inspect every part of microbial
populations and their functions in the treatment process.
FISH has been applied to quantify and identify microbial populations in
biological nutrient removal process. Pijuan et al. (2003) confirmed that the
polyphosphate accumulating organisms (PAOs), Accumulibacter, made up about 55% of
the total biomass in a lab-scale enhanced biological phosphorus removal (EBPR) reactor;
whereas, Wong et al. (2005) found that Rhodocyclus-related PAOs of 4 to 18% of
EUBmix-stained cells were the predominant population in full-scale enhanced biological
phosphorous removal plants. Meanwhile, other researches also proved the presence of
Rhodocyclus-related PAOs in biological phosphorous removal process (Onuki et al., 2002
and Zilles et al., 2002).
In addition to removing phosphorous, researchers also investigated the microbial
populations working on nitrification in the biological nitrogen removal process. The
24
study of nitrification in a full-scale activated sludge plant explored Nitrosospira genus,
ammonia oxidizing bacteria (AOB), and Nitrobacter genus, nitrite oxidizing bacteria
(NOB), which were predominant in the activated sludge system (Coskuner and Curtis,
2002). However, a controversy result showed that Nitrosomonas (AOB) and Nitrospira
(NOB) were more dominant in a fixed biofilm reactor (Kim et al., 2004). With regard to
nitrification efficiency, Hall et al. (2003) attempted to establish a correlation between
Nitrospira quantitative data and nitrate production rate determined in batch tests. A
further investigation of two continuous systems indicated that the proportions of AOB in
a combined activated sludge-rotating biological contactor process (AS-RBC) was 2.6
times that in A2O system, where the ratio was close to the ammonia oxidization rate of
2.9 times (You et al., 2003).
Recently, FISH was used to diagnose the sludge bulking problem in an activated
sludge system. Gaval et al. (2002) carried out FISH to explore the effects of oxygen
deficiencies and loading shocks on filamentous bacteria in activated sludge. Additionally,
Gaval and Pernelle (2003) further studied the impact of the repetition of oxygen
deficiencies on the filamentous bacteria. Since the competition between filaments and
floc formers has been well described using kinetic selection, a study combining the
substrate uptake test and quantitative FISH re-evaluated the differences in kinetic growth
of bulking and non-bulking activated sludge (Lou and de los Reyes III, 2005). To be
more specific, the ratios of the filaments Eikelboom Type 021N to activate sludge in
terms of concentration and in terms of fluorescent signal were determined for
constructing their own relationship (Guan et al., 2003). By applying FISH technique to
identify the filamentous bacteria, environmental engineers designed new oligonucleotide
25
probes for detecting the filamentous Nostocoida limicola population (Liu and Seviour,
2001).
In anaerobic digestion studies, FISH has been used for purposes, e.g., the
determination of the ratio of Bacteria and Archaea (Tay et al., 2001), the effects of
micro-aeration on the phylogenetic diversity in a thermophilic anaerobic digester (Tang
et al., 2004), hydrolytic activity of alpha-amylase in anaerobic digested sludge (Higuchi
et al., 2005), and so forth. Furthermore, many studies have elaborated on the structure of
the microbial community using FISH. Kuang et al. (2002) studied the influence of co-
substrates on methanogenic activity and their structure in anaerobic digesters treating
oleate. Shigematsu et al. (2003) analyzed the structure of acetate-degrading
methanogenic consortia at different HRTs. O’Sullivan et al. (2005) investigated an
enriched cellulose degrading bacterial community from an anaerobic batch reactor. A
comparable study was performed by Song et al. (2005), but he and his coworkers worked
on both cellulolytic microbial community and methanogenic populations. In sum, FISH
technique has been broadly used in anaerobic digestion, and it is believed that it may
become an essential tool to evaluate the performance of anaerobic digesters.
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37
Table 2.1 Summary of hydrogen production from thermo-chemical reactions (Sørensen,
2005)
Thermo-chemical reaction Stochiomatric equation
Steam reforming CH4 + H2O → CO + 3H2 ∆Hº = -252.3 kJ / mol
Partial oxidation CH4 + 1/2O2 → CO + 2H2 ∆Hº = -35.7 kJ / mol
Autothermal reforming CH4 + 3/2O2 → CO + 2H2 ∆Hº = -519.3 kJ / mol
Gasification and woody biomass conversion
C + H2O → CO + H2 ∆Hº = -138.7 kJ / mol
Table 2.2 Comparison of biological hydrogen production processes (Nath and Das, 2004)
Biological process
Type of microorganisms
Advantages
Disadvantages
Direct biophotolysis
Green algae
Produce H
2 directly from
water and sunlight
Require high intensity of light
Solar conversion energy
increased by tenfolds as
compared to trees, crops
O2 can be dangerous for the
system
Indirect biophotolysis
Cyanobacteria
Produce H
2 from water
Lower photochem
ical
efficiency
Has the ability to fix N
2 from
atmosphere
Uptake hydrogenase enzymes
are to be removed to stop
degradation of H2
About 30% O
2 present in gas
mixture
O2 has an inhibitory effect on
nitrogenase
Photo fermentation
Photosynthetic bacteria
A wide spectral light energy
can be used by these bacteria
Light conversion efficiency is
very low, only 1 to 5 %
Use different waste materials
like distillery effluents, waste
etc
O2 is a strong inhibitor of
hydrogenase
Dark fermentation
Fermentative bacteria
Produce H
2 without light
Relatively lower achievable
yields of H2
A variety of carbon sources
CO2 in gas mixture has to be
separated
Produce valuable metabolites
Anaerobic process with no O
2
limitation problem
38
39
Fig. 2.1 Catabolic pathway of Clostridia in hydrogen fermentation (Jones and Woods,
1989; Mitchell, 2001)
Glucose
Pyruvate
2 NAD+
2 NADH
2 ADP
2 ATP
Lactate
NADH NAD+
Fd Ox
Fd Red
Fd Ox
Fd Red
NAD+
NADH
NADH
NAD+
H2
COA
2CO2
Acetyl-COA Acetate
ADP ATP
Ethanol
NADH NAD+
COA COA
Acetoacetyl-COA
COA
Acetoacetate Acetone
CO2
Isopropanol
Butyryl-COA
ADP ATP
COA
Butyrate Butanol
NADH NAD+
COA
2 NADH
2 NAD+
40
CHAPTER 3. KINETIC STUDY OF BIOLOGICAL
HYDROGEN PRODUCTION BY ANAEROBIC
FERMENTATION
A paper published to International Journal of Hydrogen Energy
Wen-Hsing Chen, Shen-Yi Chen, Samir Kumar Khanal and Shihwu Sung
3.1 Abstract
The growth kinetics of hydrogen-producing bacteria using three different
substrates, namely sucrose, non-fat dry milk (NFDM), and food waste were investigated
in dark fermentation through a series of batch experiments. The results showed that
hydrogen production potential and hydrogen production rate increased with an increasing
substrate concentration. The maximum hydrogen yields from sucrose, NFDM, and food
waste were 234 mL/g COD, 119 mL/g COD, and 101 mL/g COD, respectively. The low
pH (pH < 4) inhibited hydrogen production and resulted in lower carbohydrate
fermentation at high substrate concentration. Michaelis-Menten equation was employed
to model the hydrogen production rate at different substrate concentrations. The equation
gave a good approximation of the maximum hydrogen production rate and the half
saturation constant (Ks) with correlation coefficient (R2) over 0.85. The Ks values of
sucrose, NFDM, and food waste were 1.4 g COD/L, 6.6 g COD/L, and 8.7 g COD/L,
respectively. Based on Ks values, the substrate affinity of the enriched hydrogen
41
producing culture was found to depend on carbohydrate content of the substrate. The
substrate containing high carbohydrate showed a lower Ks value. The maximum
hydrogen production rate was governed by the complexity of carbohydrates in the
substrate.
Keywords: Anaerobic fermentation; Food waste ; Carbohydrate; Hydrogen production;
Michaelis-Menten.
3.2 Introduction
There has been a renewed research interest on biological hydrogen production
lately. This was mainly attributed to growing global environmental concerns due to
increasing use of fossil-derived fuels and energy insecurity due to political instability in
major oil exporting countries. As a sustainable and clean energy source with minimal or
zero use of hydrocarbons, hydrogen is a promising alternative to fossil fuel. Hydrogen
can be generated by thermochemical, electrochemical or microbial fermentation
processes. However, thermochemical process needs hydrocarbon feedstocks, which
mostly comes from fossil fuels, where as electrochemical process requires supply of
electricity. Hydrogen production through microbial fermentation of renewable
feedstocks, such as biomass-derived sugars, organic wastes, and carbohydrate-rich
wastewater does not require input of external energy.
There are three microbial groups that have been studied to produce hydrogen.
The first group consists of the cyanobacteria which are autotrophs and directly
decompose water to hydrogen and oxygen in the presence of light energy by
42
photosynthesis (Hallenbeck and Benemann, 2002). Since this reaction requires only
water and sunlight and generates oxygen, it is attractive from the viewpoint of
environmental protection. However, the cyanobacteria examined so far showed rather
low rates of hydrogen production due to the complicated reaction pathway that needed to
overcome a large Gibb’s free energy (+237 kJ/mol hydrogen) requirement. Other
drawbacks are the requirement of a carrier gas to collect the evolved gas from the culture
and the difficulty of reactor design to maintain and allow sun light penetration into a
highly turbid bioreactor. Ready separation of oxygen and hydrogen is another important
issue yet to be resolved.
The second and third groups of bacteria are heterotrophs which use organic
substrates for hydrogen production. The heterotrophic microorganisms produce
hydrogen under anaerobic condition either in presence or absence of light energy.
Accordingly, the process is classified as photo fermentation or dark fermentation.
Hydrogen production through photo fermentation is carried out by photosynthetic purple
non-sulfur bacteria whereas hydrogen production through dark fermentation is carried out
by fermentative bacteria, primarily clostridia. Thermodynamically, hydrogen production
through photo fermentation is also not favorable unless light energy is supplied.
Additionally, light conversion efficiency, photoinhibition at high solar light intensities,
and design of efficient photobioreactors are other limitations of light fermentation
(Wakayama and Miyake, 2001).
Hydrogen production through dark fermentation has advantages over the other
processes because of its ability to continuously produce hydrogen from a numbers of
renewable feedstocks without an input of external energy. From environmental
43
engineering stands point, this group of bacteria is of great interest as they not only
stabilize the human derived organic wastes, but also produce a clean and renewable
energy source. In dark fermentation, different groups of bacteria were known to be the
responsible for hydrogen production such as Enterobacter, Clostridium and Bacillus.
Fang et al. (2002) reported that about 70% of population was of genus Clostridium and
14% belonged to Bacillus species in a mixed culture study. Our previous study also
showed that the hydrogen production was directly correlated to Clostridium population in
the bioreactor (Duangmanee et al., 2002).
Promising results on hydrogen production were obtained using different
substrates. In early studies, researches have explored the hydrogen production potential
of simple synthetic substrates in batch cultures (Van Ginkel et al., 2001; Khanal et al.,
2004) and from continuous operation (Lin and Chang, 1999; Duangmanee et al., 2002;
Khanal et al., 2005). The potentials of hydrogen production from complex substrates, e.g.
municipal solid wastes, cellulose containing wastes, starch-manufacturing wastewater,
and activated sludge were also reported by several investigators (Ueno et al., 1995;
Okamoto et al., 2000; Yokoi et al., 2002; Wang et al., 2003). However, the kinetic study
of hydrogen production from different characteristics of substrates in dark fermentation
has rarely been reported. Therefore, the goals of this study were two folds: (a) to study
the kinetics of biohydrogen production of different substrates (sucrose, non-fat dry milk
(NFDM) and food waste) using modified Gompertz and Michaelis-Menten equations;
and (b) to investigate the effects of these substrates on the hydrogen production potential
by enriched culture of hydrogen producers.
44
3.3 Materials and Methods
3.3.1 Seed microorganisms
The seed sludge for hydrogen production experiments was collected from a local
anaerobic digester. The anaerobically digested sludge was then filtrated through a 20-
mesh sieve, and was stored at 4˚C before inoculation.
3.3.2 Hydrogen production experiments
The hydrogen production experiments were conducted in a series of 250-mL
serum bottles with 30 mL of seed sludge (concentration of 2.8 to 3.0 g/L), 1 mL of
nutrient solution, and 5 ml of 0.72 M KHCO3. The nutrient solution composed of
NH4HCO3 (160 g/L); KH2PO4 (80 g/L); FeCl2·4H2O (70.5 g/L); NaCl (0.4 g/L);
MgSO4·7H2O (4 g/L); CaCl2·2H2O (0.4 g/L); MnSO4·7H2O (0.6 g/L); and
Na2MoO4·2H2O (0.4 g/L). Different concentrations of substrates (e.g. sucrose, NFDM
and food waste) were placed into the serum bottles. The characteristics of NFDM and
food waste used in this study are shown in Table 3.1. The food waste consisted of
produce, deli and wax-coated cardboard. The representative components of food waste
are shown in Table 3.2. The components of food waste and their percentage (wet weight
basis) were selected based on waste generation pattern of local grocery stores. Initially,
the pH in each serum bottle was adjusted to 5.5 ± 0.1 using 0.5 N potassium hydroxide or
hydrochloric acid. The serum bottles were purged with nitrogen gas, sealed with butyl
rubber stoppers, and then incubated in a shaker at 180 rpm and 36±1oC. During the test,
biogas samples were collected routinely and analyzed for hydrogen and methane contents.
Mixed liquor samples from each serum bottle were drawn at the end of the test, and
45
analyzed for chemical oxygen demand (COD), volatile suspended solids (VSS), pH, and
residual carbohydrate.
3.3.3 Analysis
The biogas production was measured regularly by plunger displacement method
(Owen et al., 1979). The hydrogen and methane contents in biogas were analyzed by a
gas chromatograph (Gow-Mac series 350) equipped with a thermal conductivity detector
and two columns. The biogas hydrogen content was measured using a 2.4 m x 6 mm
stainless column packed with Porapak Q (80/100 mesh). Nitrogen was used as a carrier
gas at a flow rate of 30 mL/min. The temperatures for the injection port, the column and
the detector were set at 100, 50 and 100 °C, respectively. Methane gas was determined
with a 2.4 m x 6 mm stainless column packed with Porapak T (80/100 mesh) and the
temperatures for the detector, the injection port, and the column were set at 200, 160, and
70 ºC, respectively. Helium was used as a carrier gas at a flow rate of 35 mL/min. COD
and VSS of mixed liquor were measured according to Standard Methods (1995).
Residual carbohydrate in the mixed liquor was determined following the method
described in Dubois et al. (1956).
3.3.4 Data analysis
In this study, cumulative hydrogen production curves with respect to time were
obtained first from the hydrogen production experiments; then the modified Gompertz
equation was applied to determine the hydrogen production potential (H), hydrogen
production rate (R), and lag phase (λ) (Lay, 2001; Van Ginkel et al., 2001).
46
( )
+⋅
⋅= 1expexp λ-tH
eR- HH(t) (3.1)
Where, H(t) is cumulative hydrogen production (mL) at time t; λ is time of lag-phase (h);
H is hydrogen production potential (mL); R is hydrogen production rate (mL/h); and e is
exp(1) , i.e. 2.71828. These parameters in Eq. (3.1) were estimated by minimizing the
sum square of errors (SSE) between experimental data and estimation from the models.
This estimation was carried out by using the ‘Solver’ function in Microsoft Excel 2002.
The significance of the estimated parameters was tested by analysis of variance
(ANOVA).
3.4 Results and Discussion
3.4.1 Kinetic analysis of hydrogen production
The cumulative hydrogen production curves obtained from the experiments with
different substrates (sucrose, NFDM, and food waste) are presented in Fig. 3.1. During
the experiment, no methane was detected in the biogas. The substrate concentration was
found to affect the hydrogen production significantly. Additionally, substrate inhibition
was not observed for these organic substrates. The kinetic parameters estimated based on
Eq (1) are listed in Table 3.3. Hydrogen production was well correlated to the modified
Gompertz equation (R2 > 0.98). Moreover, the results of analysis of variance (ANOVA)
suggest that the estimated parameters (H, R and λ) at different concentrations for the
same substrate were statistically significant different (p < 0.001) at a confidence interval
47
(CI) of 99.9% (Wackerly et al., 2002). Hydrogen yield and specific hydrogen production
rate were calculated from H and added substrate and from R and biomass concentration,
respectively. As shown in Table 3.3, the maximum hydrogen production potentials were
at the substrate concentrations of 9.0, 64.0 and 32.3 g COD/L for sucrose, NFDM, and
food waste, respectively. In addition, the maximum hydrogen production rates were
achieved at the substrate concentrations of 4.5, 64.0 and 32.3 g COD/L for sucrose,
NFDM, and food waste, respectively. The maximum specific hydrogen production rates
also occurred at the same substrate concentrations. However, the hydrogen yields from
NFDM and food waste were much lower than that from sucrose, and decreased at a
higher NFDM and food waste concentration. This might be attributed to lower
carbohydrate contents in NFDM and food waste than that in sucrose (Table 3.1).
The efficiency of biohydrogen production is highly related to the optimal control
of substrate to biomass (S/X) ratio. This ratio significantly affects the metabolic and
kinetic characteristics of microorganisms (Lui, 1996). Fig. 3.2 and Fig. 3.3 show the
variations of hydrogen yield and specific hydrogen production rate at various S/X ratios
for different substrates. As evident from Fig. 3.2(a), it was apparent that the highest
hydrogen yield of 234 mL/g COD from sucrose was achieved at an S/X ratio of 7.3 g
COD/g VSS. For food waste, the hydrogen yield reached a peak value of 101 mL/g COD
at an S/X ratio of 7.8 g COD/g VSS (Fig. 3.2(c)). However, for NFDM, the maximum
hydrogen yield ranged from 114 to 119 mL/g COD at S/X ratios below 14.7 g COD/g
VSS; then decreased with an increasing S/X ratio (Fig. 3.2(b)). Sucrose and food waste
presented a different kinetic behavior compared to NFDM.
48
As shown in Fig. 3.3, the specific hydrogen production rate remained between
120 to 140 mL/g VSS-h at S/X ratios over 3.6 g COD/g VSS from sucrose (Fig. 3.3(a)).
However, for NFDM and food waste, the specific hydrogen production rate increased
slowly to the maximum value with an increasing S/X ratio (Fig. 3(b) and Fig. 3.3(c)).
For pure carbohydrate or carbohydrate-rich substrate, e.g. sucrose, the maximum specific
hydrogen production rate was accomplished at a lower S/X ratio.
3.4.2 Variations of biomass, pH and carbohydrate at different substrate
concentrations
The biomass concentration, pH, removed carbohydrate concentration, and
carbohydrate removal efficiency at different substrate concentrations after the cessation
of hydrogen production, are presented in Fig. 3.4, Fig. 3.5, and Fig. 3.6. It was apparent
from the figures that the final biomass level and removed carbohydrate concentration
increased with an increasing sucrose concentration (Fig. 3.4(a) and Fig. 3.4(c)). However,
a lower final pH was observed at a higher sucrose concentration (Fig. 3.4(b)). It reflects
that the removed carbohydrate was used by hydrogen-producing bacteria for their growth,
and organic acid production. This finding was in close agreement with that of
Heyndrickx et al. (1987).
The final pH remained about 4 at an initial sucrose concentration greater than 7 g
COD/L. Meanwhile the carbohydrate removal efficiency decreased to about 75% and
continued to decline at higher sucrose concentration (Fig. 3.4(d)). Roychowdhury et al.
(1988) reported that hydrogen production by Clostridium sp. was inhibited in the pH
range of 4 to 5. Thus, carbohydrates at high sucrose concentrations in this study could
49
not be metabolized further at low pH. The results also explain that hydrogen production
potential (Table 3.3) did not increase with the increase in sucrose concentration, but
stayed roughly plateau at high sucrose concentration.
Similar results were also observed for NFDM and food waste. The final pH of
less than 4 was observed at higher substrate concentrations from NFDM and food waste
in comparison to sucrose. No apparent change in final pH was observed at NFDM
concentration greater than 16.0 g COD/L, and on the other hand the carbohydrate
removal efficiency declined gradually at high NFDM concentration (Fig. 3.5). Table 3
clearly shows the increase in hydrogen production potential at high NFDM concentration
was limited. For food waste, the final pH and the removed carbohydrate concentrations
were not significantly different at food waste concentration higher than 9.5 g COD/L (Fig.
3.6). Based on removed carbohydrate and the final biomass levels, it was apparent that
hydrogen-producing bacteria quickly converted soluble part of food wastes into hydrogen;
but not the particulate fraction due to rate limiting hydrolysis step. Different initial food
waste concentrations were employed for the determination of growth kinetics. Hence, the
final biomass concentrations were mostly contributed by the particulate fraction of food
wastes.
3.4.3 Growth kinetics of hydrogen-producing bacteria for three different substrates
Fig. 3.7 presents the growth kinetics of hydrogen-producing bacteria for three
different substrates. These results showed the dependence of hydrogen production rate
(R) on substrate concentration based on Michaelis-Menten equation:
50
SK
SRR
S
max
+= (3.2)
Where, R is hydrogen production rate (mL/h), KS is the half saturation constant, and is the
substrate concentration that yields Rmax/2, Rmax is the maximum hydrogen production rate
(mL/h) and S is the substrate concentration. Least square method was used to determine
Rmax and KS using Michaelis-Menten equation similar to that obtained using Eq. (3.1).
The Rmax and KS values obtained from Eq. (3.2) are given in Table 3.4. The
correlation coefficients (R2) obtained by the nonlinear regression analysis of Eq. (3.2)
were greater than 0.85 for all substrates. The effect of substrate concentration on
hydrogen production was well described by the Michaelis-Menten equation. In
Michaelis-Menten equation, KS represents the substrate affinity of the microorganisms.
The KS values for sucrose, NFDM, and food waste were 1.4 g COD/L, 6.6 g COD/L, and
8.7 g COD/L, respectively, and the substrate affinity of the hydrogen-producing bacteria
decreased in the order: sucrose > NFDM > food waste. Lay et al. (2003) concluded that
hydrogen-producing bacteria were more effective in hydrogen production from
carbohydrate-rich substrate.
The maximum hydrogen production rate decreased in the order: food waste >
NFDM > sucrose. The maximum hydrogen production rate of sucrose, NFDM, and food
waste were 13.9, 25.6, and 29.9 mL/h, respectively. It is believed that simple sugar
(monosaccharide) contributed to the most part of carbohydrate in the food waste
(Velterop and Vos, 2001; Barry et al., 2004). The hydrogen production rate from a
simple sugar was higher than from a disaccharide, and the hydrogen production from
51
lactose was reported to be higher than that from sucrose (Woodward et al., 2000), which
further supplements our data.
For comparison, the growth kinetics of biological hydrogen production from
several studies along with this research is summarized in Table 3.4. The high maximum
specific growth rates (µmax) were reported in these studies, which clearly suggest that
continuous hydrogen-producing bioreactor could be operated at a shorter hydraulic
retention time (HRT). Studies have also demonstrated that reactors running at low HRTs
presented a better performance in terms of hydrogen production. At HRT of 4 h, the
highest hydrogen production of nearly 7 L/day was achieved in the continuous stirred
tank reactor (Majizat et al., 1997). Chang et al. (2002) reported an optimal hydrogen
production rate of 0.42 L/h/L in a fixed-bed reactor operating at an HRT of 2 h. Owing
to short operating HRT in the hydrogen-producing bioreactors, methane gas was not
detected; even though the seeding inocula was not pre-treated to inactivate methanogens
(Lin and Jo, 2003). In addition, the KS values for biological hydrogen fermentation were
much higher than that of traditional anaerobic digestion. It further suggested that the
operation of hydrogen bioreactor requires a high influent substrate concentration or high
organic loading rate.
3.5 Conclusion
Anaerobic bioconversion of organic wastes to hydrogen gas is an attractive option
that not only stabilizes the waste/wastewater but also generates benign renewable energy.
Three different substrates were selected in this study to investigate the substrate affinity
of mixed microbial culture for biological hydrogen production. In general, the hydrogen
52
production rate increased with increasing substrate concentration. The substrate affinity
significantly affected the hydrogen yield. The hydrogen yield from sucrose was found to
be much higher than that from NFDM and food waste. It suggested that higher hydrogen
yield from sucrose was attributed to higher carbohydrate content. However, the substrate
could not be completely metabolized by hydrogen-producing bacteria at a higher
concentration due to low pH condition, which inhibited hydrogen-producing bacteria.
The effects of substrate concentration on the biohydrogen production were well described
by Michaelis-Menten equation.
3.6 References
APHA, 1995. Standard methods for the examination of water and wastewater. 19th edn,
American Public Health Association, Washington, D.C.
Barry, G.H., Castle, W.S., Davies, F.S., 2004. Rootstocks and plant water relations affect
sugar accumulation of citrus fruit via osmotic adjustment. J. Am. Soc. Hort. Sci.
129, 881-889.
Brosseau, J.D., Zajic, J.E., 1982. Continuous microbial production of hydrogen gas. Int. J.
Hydrogen Energy 7, 623-628.
Chang, J.S., Lee, K.S., Lin, P.J., 2002. Biohydrogen production with fixed-bed
bioreactors. Int J Hydrogen Energy 27, 1167-1174.
Chen, C.C., Lin, C.Y., Chang, J.S., 2001. Kinetics of hydrogen production with
continuous anaerobic cultures utilizing sucrose as the limiting substrate. Appl.
Microbiol. Biotechnol. 57, 56-64.
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57
Table 3.1 Characteristics of NFDM and food waste
NFDM Food Waste
COD, g/g NFDM 1.00 TS, g/L 91.3
TOC, g/g NFDM 0.21 VS, g/L 79.6
TKN, % as N 5.4 COD, g/L 142.3
Total phosphate, % 2.2 Soluble COD, g/L 62.1
Lactose, g/100g NFDM 51.0 Carbohydrates, g/L 58.0
Protein, g/100g NFDM >36.0 Proteins, g/L 19.8
Fat, g/100g NFDM <1.0 Fat, g/L 11.2
Ash, g/100g NFDM 8.2 TKN, g/L as N 1.0
Total phosphate, g/L 0.4
58
Table 3.2 Components of the food waste (on wet weight basis) (Li et al., 2003)
Potato 17.6% Orange 23.5%
Tomato 8.1% Apple 6.0%
Lettuce 4.3% Banana 5.4%
Cabbage 1.4% Watermelon 2.7%
Beans 3.7% Grape 2.7%
Pepper 1.4%
Fruit
(43%)
Strawberry 2.7%
Vegetable
(38%)
Carrot 1.5% Bread 8.0%
Chicken 2.6% Corn 5.0%
Beef 1.7% Egg 0.5%
Meat
(5%)
Pork 0.7%
Other
(14%)
Cheese 0.5%
Table 3.3 Estimated param
eters of Gompertz equation for hydrogen production
Substrate
Concentration
(g COD/L)
H
(mL)
R
(mL/h)
λ
(h)
Hydrogen yield
(mL/g COD)
Specific hydrogen
production rate
(mL/g VSS·h)
R2
0.3
2
0.2
6
45
3
0.9972
0.6
9
1.3
8
112
15
0.9974
1.1
25
5.1
9
146
55
0.9960
2.2
68
11.3
10
202
122
0.9986
4.5
157
13.6
11
234
147
0.9986
9.0
244
11.4
12
182
124
0.9994
Sucrose
17.9
240
10.6
12
89
115
0.9998
2.0
35
4.3
6
117
53
0.9937
4.0
71
8.9
7
119
108
0.9983
8.0
136
14.6
7
114
179
0.9976
16.0
224
19.9
6
93
243
0.9990
32.0
268
21.1
4
56
258
0.9986
64.0
292
24.3
4
30
298
0.9985
NFDM
96.0
248
22.0
3
17
269
0.9972
1.1
9
3.4
4
52
39
0.9882
2.3
30
6.7
4
86
77
0.9975
4.6
69
12.1
4
101
139
0.9991
9.5
77
14.4
4
54
165
0.9991
16.1
110
17.5
3
45
201
0.9988
Food waste
32.3
178
25.0
3
37
286
0.9981
59
Table 3.4 Summary of growth kinetics of hydrogen-producing bacteria
Test type
Culture
Test
temperature
(ºC)
Test
substrate
µmax
(h-1)
Rmax
(mL·h
-1)
KS
(g COD/L)
YX/S
(g biomass/mole
substrate)
Reference
Continuous
Citro
bact
er
inte
rmed
ius
37
Glucose
0.220
121.4
NA
20.6
Brosseau and
Zajic (1982)
Continuous
Mixed
35
Sucrose
0.172
NA
0.068
34.2
Chen et al.
(2001)
Continuous
Mixed
37
Glucose
0.333
NA
NA
45.0
Horiuchi et al.
(2002)
Batch
Ente
robact
er
cloaca
e 37
Glucose
0.568
NA
3.914
15.1
Kumar et al.
(2000)
Batch
Cald
icel
lulo
sir-
upto
r
sacc
haro
lyticu
s
70
Sucrose
0.130
67.2
0.801
NA
Van Niel et al.
(2003)
This study
Batch
Mixed
35
Sucrose
ND
13.9
1.446
ND
Batch
Mixed
35
NFDM
ND
25.6
6.616
ND
Batch
Mixed
35
Food
waste
ND
29.9
8.692
ND
NA: Not available
ND: Not determined
60
61
200
150
100
50
020151050
250
200
150
100
50
06050403020100
400
300
200
100
02520151050
0 g COD/L
0.3 g COD/L
0.6 g COD/L
1.1 g COD/L
2.2 g COD/L
4.5 g COD/L
9.0 g COD/L
17.9 g COD/L
0 g COD/L
2.0 g COD/L
4.0 g COD/L
8.0 g COD/L
16.0 g COD/L
32.0 g COD/L
64.0 g COD/L
96.0 g COD/L
0 g COD/L
1.1 g COD/L
2.3 g COD/L
4.6 g COD/L
9.5 g COD/L
16.1 g COD/L
32.3 g COD/L
Time (hours)
Cumulative H
2 production (mL)
(a)
(b)
(c)
Fig. 3.1 Cumulative hydrogen production with time: (a) sucrose, (b) NFDM, and (c)
food waste
62
120
80
40
06050403020100
120
80
40
0150100500
250
200
150
100
50
035302520151050
Substrate to microorganism ratio (g COD/g VSS)
Hydrogen yield (mL/g COD)
(a)
(c)
(b)
Fig. 3.2 Hydrogen yield at different S/X ratios: (a) sucrose, (b) NFDM, and (c) food
waste
63
300
200
100
06050403020100
300
200
100
0150100500
160
120
80
40
035302520151050
Substrate to microorganism ratio (g COD/g VSS)
Specific hydrogen production rate (mL/g VSS-h)
(a)
(c)
(b)
Fig. 3.3 Specific hydrogen production rate at different S/X ratios: (a) sucrose; (b)
NFDM; and (c) food waste
64
100
80
60
40
20
020151050
7
6
5
4
3
2
8000
6000
4000
2000
0
2500
2000
1500
1000
500
0
VSS concentration
(mg/L)
Removed carbohydrate
concentration
(m
g/L as glucose)
Carbohydrate removal
efficiency (%)
Sucrose concentration (g COD/L)
Final pH
(a)
(d)
(c)
(b)
Fig. 3.4 Metabolism of hydrogen-producing bacteria at different sucrose concentrations:
(a) mixed liquor volatile suspended solid concentration, (b) final pH, (c)
removed carbohydrate concentration, and (d) carbohydrate removal efficiency
65
7
6
5
4
3
2
20
15
10
5
0
x103
12
8
4
0
x103
100
80
60
40
20
0120100806040200
VSS concentration
(mg/L)
Final pH
Removed carbohydrate
concentration
(m
g/L as glucose)
Carbohydrate removal
efficiency (%)
NFDM concentration (g COD/L)
(a)
(d)
(c)
(b)
Fig. 3.5 Metabolism of hydrogen-producing bacteria at different NFDM concentrations:
(a) mixed liquor volatile suspended solid concentration, (b) final pH, (c)
removed carbohydrate concentration, and (d) carbohydrate removal efficiency
66
100
80
60
40
20
035302520151050
6000
4000
2000
0
12
8
4
0
x103
7
6
5
4
3
2
VSS concentration
(mg/L)
Final pH
Removed carbohydrate
concentration
(mg/L as glucose)
Carbohydrate removal
efficiency (%)
Food waste concentration (g COD/L)
(a)
(b)
(c)
(d)
Fig. 3.6 Metabolism of hydrogen-producing bacteria at different food waste
concentrations: (a) mixed liquor volatile suspended solid concentration; (b) final
pH; (c) removed carbohydrate concentration; and (d) carbohydrate removal
efficiency
67
30
20
10
0403020100
16
12
8
4
020151050
30
20
10
0100806040200
Sucrose concentration (g COD/L)
Food waste concentration (g COD/L)
NFDM concentration (g COD/L)
H2 production rate (mL/h)
R2=0.8576
R2=0.9759
R2=0.9801
Fig. 3.7 Effect of substrate concentration on the hydrogen production rate from different
substrate
68
CHAPTER 4. BIOLOGICAL HYDROGEN PRODUCTION
IN ANAEROBIC SEQUENCING BATCH REACTOR
A paper to be submitted to Water Research Journal
Wen-Hsing Chen, Shihwu Sung
4.1 Abstract
Biological hydrogen production from sucrose-rich substrate was investigated in
an anaerobic sequential batch reactor (ASBR). The goal of this study was to investigate
the effect of different hydraulic retention times (HRTs) (8, 12, 16, 24, and 48 h), pHs (4.9,
5.5, 6.1, and 6.7), substrate concentrations (15, 25, and 35 g chemical oxygen demand
(COD/L)), and cyclic durations (4, 6, and 8 h) on biological hydrogen production. The
maximum hydrogen yield of 2.53 mol H2/mol sucrose consumed and the maximum
hydrogenic activity of 538 mL H2/g VSS-d were obtained at HRT of 16h, pH 4.9, sucrose
concentration of 25 g COD/L, and feeding cycle of 4 h. Methane was detected in the
biogas when solids retention time (SRT) exceeded 100 h at pH of 6.7. Based on the low
ethanol concentration of nearly 300 mg/L, the metabolic pathway shifting to solvent
fermentation was not observed at pH of 4.9. The ratios of butyrate (HBu) to acetate
(HAc) decreased from 1.25 to 0.54 mol/mol when the sucrose concentration was
increased from 15 to 35 g COD/L. This apparently suggests that the metabolic pathway
69
of acetate fermentation was predominant at higher sucrose concentration. The hydrogen
production was found to improve at shorter feeding cycle of 4 h.
Keywords: Biological hydrogen production; pH; HRT; Substrate concentration;
Anaerobic sequencing batch reactor
4.2 Introduction
There has been a renewed research interest on biological hydrogen production
lately. This was mainly attributed to growing global environmental concerns and energy
insecurity. As a sustainable and clean energy source with minimal or zero use of
hydrocarbons, hydrogen is a promising alternative to petroleum fuels. Hydrogen can be
generated by thermochemical, electrochemical or microbial fermentation processes.
However, thermochemical process needs hydrocarbon feedstock, which mostly comes
from fossil fuels, where as electrochemical process requires supply of electricity.
Hydrogen production through microbial fermentation of renewable feedstock, such as
biomass-derived sugars, organic wastes, and carbohydrate-rich wastewater does not
require input of external energy.
Promising results on biological hydrogen production were obtained using
different substrates. In early studies, researchers have explored the hydrogen production
potential of simple synthetic substrates in batch experiments (Van Ginkel et al., 2001;
Khanal et al., 2004) and in continuous operation (Lin and Chang, 1999; Duangmanee et
al., 2002a; Khanal et al., 2005). The potential of hydrogen production from complex
substrates, e.g., municipal solid wastes (Ueno et al., 1995), cellulose containing wastes
70
(Okamoto et al., 2000), starch-manufacturing wastewater (Yokoi et al., 2002), and
activated sludge (Wang et al., 2003) was reported by several investigators. In these
studies, successful hydrogen production was obtained by suppressing the growth of
hydrogen consumers, primarily, methanogens. The pre-treatment of seed inoculum was
one of the approaches to suppress hydrogen consumers. Lay et al. (2003) studied the
heat-treatment option to obtain an enriched culture of hydrogen producers from
anaerobically digested sludge. Chen et al. (2002) employed acid pre-treatment of seed
sludge for selection of hydrogen producers.
High maximum specific growth rates (µmax) of 0.17 to 0.57 h-1 were reported for
hydrogen producers (Brosseau and Zajic, 1982; Kumar et al., 2000; Chen et al., 2001;
Horiuchi et al., 2002). This suggests that completely mixed hydrogen-producing
bioreactor could be operated at a much shorter hydraulic retention time (HRT) in
comparison to conventional anaerobic digester. Earlier studies also demonstrated that
reactors running at low HRTs yielded a better performance in terms of hydrogen
production (Lin and Chang, 1999; Gavala et al., 2006). Owing to short HRT in the
hydrogen-producing bioreactor, methane gas was not detected; even though the seed
inocula were not pre-treated to inactivate methanogens (Lin and Jo, 2003). In addition, a
kinetic analysis investigating the substrate affinity indicated that pure carbohydrate or
carbohydrate-rich substrate was ideal for hydrogen production (Chen et al., 2006). The
study also concluded that the half saturation constant KS values for biological hydrogen
fermentation were much higher than that of a traditional anaerobic digestion. Moreover,
due to low substrate affinity of hydrogen-producing bacteria, the operation of hydrogen
bioreactor requires a high influent substrate concentration or high organic loading rate.
71
With respect to bioreactor configuration, researchers examined high rate
anaerobic bioreactors that maintain long solids retention time (SRT) at short HRT, for
better hydrogen yield. In a three-phase fluidized bed reactor, hydrogen yield of 2.67 mol
H2/mol sucrose was obtained at HRT as short as 2 h (Wu et al., 2003). A maximal
hydrogen yield of 3.03 mol H2/mol sucrose was found at HRT of 0.5 h in a carrier-
induced granular sludge bed bioreactor (Lee et al., 2004).
As indicated earlier, hydrogen fermentation experiment was conducted in the
bioreactors operated under continuous feeding mode. Biohydrogen production from
bioreactors operated under batch-mode feeding, particularly, an anaerobic sequencing
batch reactor (ASBR) has not been well examined. ASBR is a typically operated under
batch-mode feeding with four different cycles; feed, react, settling and decant (Sung and
Dague, 1995). In ASBR, the loss of biomass is low irrespective of HRT, since the
biomass is allowed to settle down before the supernatant is withdrawn. Fresh substrate is
fed to the bioreactor in each feeding cycle followed by react cycle. To maximize
hydrogen yield in ASBR, the operating conditions need to be optimized. Based on this
premise, the goal of this study was to evaluate the hydrogen production potential at
different HRT, pH, substrate concentration, and cyclic duration in ASBR.
4.3 Materials and Methods
4.3.1 Seed sludge and substrate preparation
The seed sludge for hydrogen production experiments was collected from a local
anaerobic digester, located in Ames, Iowa, USA. Anaerobic digested sludge was
pretreated before seeding the bioreactor as described elsewhere (Chen et al., 2002).
72
Sucrose was used as the limiting substrate to mimic the carbohydrate-rich wastewater.
Every liter of sucrose solution was supplemented with 5.85 g of NH4HCO3, and 6.7 ml of
trace mineral stock solution composed of KH2PO4 (150 g/L); MgSO4·7H2O (10 g/L);
NaCl (1 g/L); Na2MoO4·2H2O (1g/L); CaCl2·2H2O (1 g/L); MnSO4·7H2O (1.5 g/L);
FeCl2 (0.28 g/L); CoCl2·6H2O (0.24 g/L); NiCl2·6H2O (0.12 g/L) and ZnCl2 (0.06 g/L).
The concentrations of NH4HCO3 and KH2PO4 in the synthetic wastewater were then
adjusted for obtaining chemical oxygen demand (COD) to nitrogen to phosphor ratio of
nearly 100:5:1.
4.3.2 Reactor operation
The ASBR had an active volume of 3.0 L and operated under a mesophilic
condition (35±1ºC). Mixing was achieved by using a mechanical mixer rotating at 300
rpm during the react phase. It took about 90 min for biomass to settle in each cycle prior
to the decant phase. An on-line pH controller was used to maintain a constant pre-set pH
by adding either 2 N hydrochloric acid or 2 N of the mixture of sodium hydroxide and
potassium hydroxide. According to the principle of ASBR, complete mixing condition
occurs during reaction phase (Sung and Dague, 1995). Therefore, the pH controller was
set only to function in react phase.
The seed sludge was kept under completely mixed condition at 100 rpm with
sucrose concentration of 6.25 g COD/L for one day during the start-up. The pH was not
controlled in this stage. The bioreactor was run under ASBR mode after the start-up.
ASBR was operated at different HRT, pH, sucrose concentration, and cyclic regime to
determine the optimum condition for maximum hydrogen yield. The experimental design
73
is summarized in Table 4.1.
4.3.3 Analysis
Soluble chemical oxygen demand (SCOD), effluent volatile suspended solids
(VSS), and mixed liquor volatile suspended solids (MLVSS) were measured following
the procedures listed in Standard Methods (1998). Residual carbohydrate in the mixed
liquor was determined following the method described in Frølund et al. (1996).
Oxidation reduction potential (ORP) was measured by an on-line pH/ORP controller
(Consort R305). The hydrogen and methane contents in biogas were analyzed by a gas
chromatograph (Gow-Mac series 350) equipped with a thermal conductivity detector and
two Porapak columns. The hydrogen content was measured using a 2.4 m x 6 mm
stainless column packed with Porapak Q (80/100 mesh). Nitrogen was used as a carrier
gas at a flow rate of 30 mL/min. The temperatures for the injection port, the column and
the detector were set at 100, 50 and 100 °C, respectively. Methane gas was determined
with a 2.4 m x 6 mm stainless column packed with Porapak T (80/100 mesh) and the
temperatures for the detector, the injection port, and the column were set at 200, 160, and
70 ºC, respectively. Helium was used as a carrier gas at a flow rate of 35 mL/min.
Solvents and individual volatile fatty acids (VFAs) were analyzed using another gas
chromatograph (HP5730A) equipped with a flame ionization detector (FID). The column
used was a 6' by 2 mm packed glass – carbopack BAW 80/120, carbowax 20m. The
operational temperatures of the injection port, the oven and detector were maintained at
250, 100, and 250°C, respectively. Helium was used as the carrier gas at a flow rate of
20 mL/min.
74
4.4 Results and Discussion
4.4.1 Effect of HRT
The ASBR was operated at a stepwise decrease in HRTs from 48, 24, 16, 12, to 8
h. The effect of HRT on hydrogen production is summarized in Table 4.2. As apparent
from the table, hydrogen yield was found to increase with decrease in HRT. However,
with further decrease in HRT below 12 h, the hydrogen yield decreased significantly.
The maximum hydrogen yield of 0.75 mole H2/mole sucrose was obtained at an HRT of
24 h. Hydrogenic activity also increased with decrease of HRT, and the maximum
hydrogenic activity of 106 mL H2/g VSS/d was achieved at HRT of 16 h. The lowest
ORP of approximately -400 mV was observed at HRT of 16 h. The ORP, however,
increased to about -300 mV, when the HRT was reduced to 12 h, and it continued to
increase to -220 mV with reduction in HRT of 8 h. Tanisho (2001) summarized that the
optimum for forming molecular hydrogen from proton is at ORP of -414 mV. The ORP
variation showed a significant impact on hydrogen content. At the higher ORP, the
hydrogen content was lower. This finding was in close agreement with that of Hussy’s et
al. (2005), who reported that the metabolism of hydrogen fermentation is associated with
ORP. This result further explains why hydrogen yield and hydrogenic activity were
lower at HRT shorter than 16 h. In addition, methane gas was detected in the biogas
throughout this study. However, at HRT of 8 h, no methane gas was detected in the
biogas. Thus, maintaining an optimal reducing environment is essentially important to
enhance the hydrogen-producing anaerobes and maximize the overall hydrogen yield.
Based on the earlier results, the maximum hydrogen production was achieved by
gradually reducing HRT to 16 h. Therefore, in this phase of the study, we repeated the
75
same experiment with directly operating the reactor at HRT of 16 h to confirm whether
the maximum hydrogen production could be obtained. The performance of ASBR on
hydrogen production is shown in Fig. 4.1. As evident from Fig. 4.1 (a), hydrogen
production rate reached its maximum and remained fairly constant at approximately 2.0
L/L/d. The maximum hydrogen production rate was sustained for 6 days and then it
gradually reduced to zero. The biogas production rate remained fairly constant at about
3.5 L/L/d. As shown in Fig. 4.1 (b), the hydrogen content followed the same trend as that
of hydrogen production rate. The hydrogen content in the biogas dropped to nearly zero;
while the carbon dioxide content increased to about 85% at the end of 19 days of
operation. Interestingly, the methane gas also began to appear in the biogas after day 15,
and reached as high as 20% on day 19. The VFA and solvent concentrations are
presented in Fig. 4.1 (c) and 4.1 (d), respectively. As evident from the figures, butyrate
and ethanol were the major VFA and solvent, respectively of hydrogen fermentation.
The maximum concentrations of butyrate and ethanol were approximately 7,600 and 290
mg/L, respectively. The butyrate and ethanol productions showed a similar trend with
that of hydrogen production. Based on acid and solvent production data, it becomes
apparent that hydrogen production followed butyrate fermentation pathway. This finding
was in close agreement with several studies (Chang and Lin, 2004; Shin and Youn, 2005;
Kyazze et al., 2006).
Many researches reported no methane production during hydrogen fermentation
at neutral pH, when the seed sludge was acid-, heat-, or base-pretreated (Chen et al., 2001;
Lin and Cheng, 2006; Lin et al., 2006). Without heat treatment of seed sludge to
eliminate hydrogen consumers, methane gas was detected in a completely mix reactor
76
operated at HRT of as short as 10 h under slightly acidic condition (pH 5.5) (Kraemer
and Bagley, 2005). Thus, thermal or chemical treatment of seed sludge is essential to
effectively suppress hydrogen consumers. However, the results of the previous studies
contradict the finding of this study, which showed that methane gas could still be
produced.
There could be various possible reasons for the contradictory results among these
studies. The thermal/chemical treatment and short HRT are two key methods of
obtaining enriched culture of hydrogen-producing bacteria. The thermal/chemical
treatment eliminates hydrogen consumers during the start-up (Lay, 2000; Chen et al.,
2002). The high specific growth rate of hydrogen-producing bacteria allows them to
grow faster than their washout rate in a completely mixed reactor operating at a short
HRT (Chen et al., 2006). Thus, the short HRT facilitates the washout of the hydrogen
consumers, especially methanogens from the hydrogen-producing bioreactor. The lack of
any of the two selection processes could increase the possibility of methane production.
Solids retention time (SRT) is another issue which may influence methane
production in a hydrogen-producing reactor. Based on the early results, successful
hydrogen production was achieved at a short HRT without sludge recirculation.
Accordingly, SRT as short as HRT could also select hydrogen producers without
methane gas generation. However, in ASBR system, only the suspended sludge in the
supernatant is discharged during the decant phase. Thus, SRT is always greater than
HRT in a properly operated ASBR system. The longer SRT in an ASBR system may be
responsible for potential survival of hydrogen consumers under neutral pH condition.
77
Based on earlier discussion, SRT of an ASBR system can be given by the following
expression.
θθXe
Xc = (4.1)
Where, θc is SRT, θ is HRT, X is mixed liquor VSS concentration, and Xe is effluent
VSS concentration. According to Eq. (4.1), the ABSR was operated at SRT of
approximately 100 h.
To prevent methane production at long SRT, the well acclimated hydrogen-
producing sludge is suggested as seed inoculum instead of thermal/chemical pretreated
anaerobic sludge. Earlier studies reported no methane production from high rate
bioreactors inoculated with well acclimated hydrogen-producing sludge (Wu et al., 2003;
Lee et al., 2004; Lee et al., 2006). The well acclimated hydrogen-producing sludge was
developed in a CSTR. The acclimated hydrogen-producing sludge has already been
subjected to thermal/chemical treatment and hydraulic selection. Based on this premise,
seeding an ASBR with the well acclimated hydrogen-producing sludge could minimize
the growth of methanogens.
4.4.2 Effect of pH
The reactor was operated at a HRT of 16 h with sucrose concentration of 25 g
COD/L. The pHs was varied from 4.9 to 6.7 to determine the optimum condition. With
different phases of feed, react, settle and decant in each cycle, the pH was controlled only
in the react phase due to the complete mix condition in the stage. Fig. 4.2 (a) shows the
maximum hydrogen yield of approximately 2.5 mol H2/mol sucrose consumed at pH of
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4.9 and then it decreased with increase in pH. Figs. 4.2 (b) and (c) indicate the maximum
hydrogen conversion efficiency and hydrogenic activity of roughly 1.4 mol H2/mol
sucrose fed and 540 mL H2/g VSS/d, respectively at pH of 4.9. Similar to hydrogen yield,
hydrogen conversion efficiency and hydrogenic activity were inversely correlated to pH.
The correlation coefficients were all greater than 0.6. Based on these results, it can be
concluded that hydrogen-producing bacteria were more active at pH of 4.9 in ASBR. In
addition, the results of carbohydrate removal efficiency as shown in Fig. 4.2 (d)
explained the discrepancy between hydrogen yield and hydrogen conversion efficiency at
pH of 4.9. However, there was no significant difference at pHs ranging from 5.5 to 6.7.
It was due to lower carbohydrate removal efficiency of approximately 56% at pH of 4.9;
but it reached plateau at nearly 100% at pHs ranging from 5.5 to 6.7.
Fig. 4.3 shows the variation of soluble microbial product and MLVSS
concentrations at different pHs. As shown in Fig. 4.3 (a), acetate concentration of about
2,000 mg/L at pHs of 4.9 and 6.7 was relatively low in comparison to that of at pHs of
5.5 to 6.1 (about 4,000 mg/L). Butyrate concentration of over 6,000 mg/L was observed
at pH range of 5.5 to 6.7. It, however, decreased to around 2,300 mg/L at pH of 4.9. The
results also show that the butyrate concentration was still higher than the acetate
concentration at pH ranging from 4.9 to pH 6.7. In addition, the appearance of
propionate demonstrated that propionate fermentation was carried out by hydrogen-
producing-bacteria or by other microbial populations (Hwang et al., 2004; Hallenbeck,
2005). With regard to the sum of acetate, butyrate and propionate concentrations, the
total VFA of approximately 6,000 mg/L as acetic acid at pH 4.9 was half of that at pH
between 5.5 and 6.7. The apparently suggests that less carbohydrate was consumed by
79
hydrogen-producing bacteria which lead to less total VFA production as shown in Fig.
4.2 (d). The ratio of butyrate to acetate (HBu/HAc) increased with increase pH as shown
in Fig. 4.3 (b). The HBu/HAc ratio was greater than 1.0 mol/mol at pH of 5.5 to 6.7, but
it decreased to less than 0.8 mol/mol at pH of 4.9. Thus, butyrate fermentation at pH
equal or greater than 5.5 appeared to be a dominant metabolic pathway in dark
fermentation.
Solvent concentrations were low at the near-detection limit in the study. Ethanol
was the prevailing species among solvents (Fig. 4.3 (a)). Apparently, no obvious effect
on ethanol concentration was observed at pH of 4.9 to 6.7. The variation of MLVSS
concentration at different pHs is presented in Fig. 4.3 (c). As seen in the figure, MLVSS
concentration as high as 15,000 mg/L was observed at near neutral pH condition, it
however, reduced to approximately 6,000 mg/L at pH of 4.9. MLVSS concentration was
significantly affected by pH and showed an increasing trend with the pH. The higher
MLVSS concentration obtained under neutral pH condition is in close agreement with the
pure culture study investigating hydrogen production from Clostridium butyricum CGS5
(Chen et al., 2005).
Earlier studies from batch experiments reported that the optimum initial pH for
hydrogen production from sucrose was ranged from 5.5 to 5.7 (Van Ginkel et al., 2001;
Khanal et al., 2004; Wang et al, 2005), and from starch was reported at around 6.0
(Zhang et al., 2003). In addition, one study showed that the initial pH of 6.0 was also
favorable for hydrogen production from cheese whey (Ferchichi et al., 2005). The initial
pH of 6.5 was optimal for hydrogen fermentation from 5-carbon xylose (Lin et al., 2006).
With regard to continuous operation, the pH of 5.5 was found to be optimum for
80
hydrogen production from glucose (Fang and Liu, 2002). Lay (2000) reported an
optimum pH of 5.2 for maximum hydrogen production from synthetic starch wastewater.
Based on statistical contour plot analysis, Lay and his coworkers determined an optimum
pH of 5.8 for hydrogen production in a complete-mixed bioreactor fermenting beer
processing wastes (Lay et al., 2005). From these studies, it can be concluded that the
slightly acidic pH of 5.5 to 6.0 helps to enhance the hydrogen production. However, our
finding, which the pH of 4.9 was the optimum for hydrogen-producing bacteria in ASBR,
was different from these reports. Our study found that higher hydrogen yield was
obtained at pH of 4.9 in ASBR. Based on the trend of hydrogen production at pH
ranging from 4.9 to 6.7 in the study, it could be recommended that hydrogen fermentation
is possible to achieve at even lower controlled pH without deterioration in the system
performance. Fang’s et al. (2006) investigation, which the better performance of
hydrogen production from rice slurry was at pH of 4.5, supported the possibility of our
assumption. In addition, the advantage of operating ASBR at pH as low as 4.9 is
probably more advantageous in terms of growth of hydrogen consumers even at long
SRT.
Previous research discussed the metabolic pathway shift from acid and hydrogen
fermentation to solvent fermentation at pH below 5.0 (Gottwald and Gottschalk, 1985;
Dabrock et al., 1992). Lay (2000), however, found that solvent fermentation prevailed at
pH lower than 4.1. Based on soluble microbial product data as revealed in Fig. 4.3 (a),
the metabolic pathway shift was not observed at pH of 4.9 in the study. In addition, it has
been well known that the hydrogen production was directly correlated to Clostridium
population in the bioreactor (Duangmanee et al., 2002). In response to hostile conditions,
81
such as oxygen, heat, acid, base, etc., the physiology of Clostridium may change from
vegetative to an endospore. It is also well understood that solvent fermentation was
associated with the early steps in sporulation (Rogers and Gottschalk, 1993). Therefore,
the neglected amount of solvent production in the study elaborated that the Clostridium
population was mostly as vegetative cell.
4.4.3 Effect of substrate concentration
The effect of sucrose concentration (from 15 to 35 g COD/L) on hydrogen
fermentation was investigated in the study. The ASBR was operated at a HRT of 16 h
with pH controlled at 4.9. The performance of hydrogen-producing bioreactor is
presented in Table 4.3. As evident from the table, carbohydrate removal efficiency
decreased with the increase of sucrose concentration. Carbohydrate removal efficiency at
sucrose level of 15 g COD/L decreased from nearly 100% to less than 40% at 35 g
COD/L. The removed carbohydrates with respect to the influent sucrose of 15, 25, and
35 g COD/L were approximately 13.0, 12.5, and 11.9 g/L as glucose, respectively. The
utilization of carbohydrate by hydrogen-producing bacteria was not obvious among the
three different sucrose concentrations. This difference is also reflected by the slight
variation in hydrogen yields of 1.86 to 2.53 mol H2/mol sucrose consumed at sucrose
concentrations ranging from 25 and 35 g COD/L. However, it is evident that hydrogen
conversion efficiency decreased with the increase of sucrose concentration. Due to the
high carbohydrate removal efficiency, hydrogen conversion efficiency of 1.81 mol
H2/mol sucrose fed at sucrose concentration of 15 g COD/L was close to its hydrogen
yield. With respect to hydrogen production rate, the peak hydrogenic activity of nearly
82
540 mL H2/g VSS/d was achieved at sucrose concentration of 25 g COD/L. Based on the
results of hydrogen yield, hydrogen conversion efficiency, and hydrogenic activity, the
optimal sucrose concentration was 25 g COD/L. Thereafter, hydrogen fermentation at
sucrose concentration of 25 g COD/L was repeated to confirm the results. As revealed in
Table 4.3, the results between the original experiment and the repeated test were very
close.
In addition to hydrogen production, the level of soluble microbial product
associated with the production of hydrogen was also affected by sucrose concentration.
As evident from the table, the maximum acetate concentration ranging from 1,680 to
2,020 mg/L was appeared at sucrose concentration between 25 to 35 g COD/L. The
maximum butyrate concentration of approximately 2,600 mg/L was generated at sucrose
concentration of 15 g COD/L, and then it decreased with the increase of sucrose
concentration. Based on butyrate and acetate concentrations, HBu/HAc ratios were 1.25,
0.79, and 0.54 mol/mol at sucrose concentration of 15, 25, and 35 g COD/L, respectively.
It suggests that the metabolic pathway of acetate fermentation would be more prominent
than butyrate fermentation at higher sucrose concentration. The solvents including
ethanol, 1-propanol, butanol, acetone and 2-propanol were also measured in the study.
Ethanol was only the solvent species produced above the detection limit, and its
background concentration between 166 and 281 mg/L was not affected by sucrose
concentration.
Fig. 4.4 presents MLVSS and effluent VSS concentrations, SRT, and food to
microorganism (F/M) ratio at each sucrose concentration. As seen in Fig. 4.4 (a),
MLVSS concentrations were stabilized between 5,650 and 6,350 mg/L at sucrose
83
concentration of 15 to 35 g COD/L. The lowest effluent VSS concentration of 630 mg/L
was obtained at sucrose concentration of 15 g COD/L, and then it increased
proportionally with sucrose concentration. Based on Eq. (4.1), SRT was calculated at
different sucrose concentration and is presented in Fig. 4.4 (b). In contrast to effluent
VSS concentration, SRT was inversely proportional to sucrose concentration. SRT as
long as 145 h was achieved at sucrose concentration of 15 g COD/L, and then it
decreased to 50 h at 35 g COD/L. Meanwhile, the results of SRT and hydrogen yield
illustrated that longer SRT would not benefit in terms of hydrogen yield. As indicated in
Fig. 4.4 (c), F/M ratio was also determined at different sucrose concentration and
MLVSS concentration. Since MLVSS concentration reached plateau in the designed
range of sucrose concentration, F/M ratio was directly proportional to sucrose
concentration. The F/M ratios were 3.64, 6.52, and 8.53 g COD/g VSS/d at sucrose
concentrations of 15, 25, and 35 g COD/L, respectively. The previous kinetic study
reported that high F/M ratio is required for hydrogen fermentation (Chen et al., 2006). In
the study, it was found that high F/M ratio was accompanied with low carbohydrate
removal efficiency. It reflected that at higher F/M ratio more carbohydrate residual
remained in the reactor. The higher amount of carbohydrate residual in ASBR might
agitate the suspension of hydrogen-producing bacteria due to internal gassing which
leaded to the poor VSS settling during settle phase.
It is well known that hydrogen production is accompanied by either acetate (Eq.
4.2) or butyrate formation (Eq. 4.3) (Miyake, 1998). In acetate fermentation, 4 moles of
hydrogen are produced from 1 mole of glucose, whereas 2 moles of hydrogen are
produced in butyrate fermentation. It suggests that the higher yield of hydrogen
84
production is obtained through acetate fermentation instead of butyrate fermentation. In
addition to the sole fermentation pathway of hydrogen production, it is commonly carried
out by the combination of acetate fermentation and butyrate fermentation, where 2 moles
of hydrogen is generated (Eq. 4.4) (Hallenbeck, 2005). The stoichiometrical ratio of HBu
to HAc ratio is 1.5 mol/mol. Researchers attempted to adopt HBu/HAc ratio as an
indicator to evaluate the performance of hydrogen production (Chen, et al., 2002; Kim et
al., 2006). However, this ratio does not fully elaborate the complex biochemical reaction
during the fermentation. The explanation was possibly due to the existence of
homoacetogens, mainly homoacetogenic clostridia, growing heterotrophically by
converting simple sugar to acetic acid without evolving hydrogen (Eq. 4.5) (Ljungdahl et
al., 1989). Meanwhile, homoacetogenic clostridia are able to grow autotrophically with
acetate production on a mixture of hydrogen and carbon dioxide as the only energy and
carbon source (Eq. 4.6). Accordingly, it makes the metabolic pathway of hydrogen
fermentation more complicated, and fails to examine the hydrogen yield based on
HBu/HAc ratio.
C6H12O6 + 2H2O → 2 CH3COOH + 2CO2 + 4H2 ∆G°-184 kJ (4.2)
C6H12O6 → CH3CH2CH2COOH + 2CO2 +2H2 ∆G°-257 kJ (4.3)
C6H12O6 + 0.5 H2O → 0.75 CH3CH2CH2COOH + 0.5 CH3COOH + 2 CO2 + 2.5 H2 (4.4)
C6H12O6 → 3 CH3COOH (4.5)
2CO2 + 4H2 → CH3COOH + 2 H2O (4.6)
85
4.4.4 Effect of cyclic duration
Hydrogen fermentation in ASBR was examined based on the cycle durations of 4,
6, and 8 h corresponding to 6, 4, and 3 cycles/d, respectively. The reactor was operated
at a 16-h HRT and an optimal pH of 4.9 determined previously. Influent sucrose
concentration was kept constant at 15 g COD/L. The performance of hydrogen
fermentation at different cyclic regime is summarized in Table 4.4. As evident from the
table, hydrogen content of approximately 37% was achieved at 4-h duration, and then
decrease linearly with increase in duration. This linear trend was followed by hydrogen
yield and hydrogen conversion efficiency as well. The maximum hydrogen yield of 1.86
mol H2/mol sucrose consumed and the maximum hydrogen conversion efficiency of 1.81
mol H2/mol sucrose fed were achieved at duration of 4 h. Since carbohydrate removal
efficiency achieved close to 100%, hydrogen yield and hydrogen conversion efficiency
were statistically equal throughout the cyclic durations. Hydrogenic activity was also
found to decrease with increase of cyclic duration. Hydrogenic activity of 388 mL H2/g
VSS/d at duration of 4 h was as twice as higher of that at duration of 8 h.
The cyclic regime also affected MLVSS concentration, effluent VSS
concentration, SRT, F/M ratio, and HBu/HAc ratio as summarized in Table 4.4. As
revealed in the table, the difference in MLVSS concentration between 4-h and 8-h cyclic
durations was of about 2,000 mg/L. The effluent VSS concentration ranging from 600 to
750 mg/L and did not vary much among cyclic durations, based on Eq. (4.1). The SRT
was, however, depended on the variation of MLVSS concentration. The results of
hydrogen yield and SRT reflected that the lower hydrogen yield could be obtained at
longer SRT in hydrogen fermentation. As same as the performance of hydrogen
86
fermentation, F/M ratio was decreased with the increase of cyclic duration. The
maximum hydrogen yield appeared at F/M ratio of 3.64 g COD/g VSS/d with 4-h cyclic
duration. With regard to HBu/HAc ratio, the results indicates that no significant variation
on HBu/HAc ratios at cyclic durations between 4 h and 6 h. However, due to the level of
acetate (data not shown), HBu/HAc ratio was dramatically decreased to 0.49 mol/mol at
8-h cyclic duration.
As discussed earlier, the cyclic regime enabled to impact the performance of
hydrogen production in ASBR. The longer cyclic duration could reduce hydrogen yield
and hydrogenic activity. This might be explained by the microbial population shift,
which facilitates the growth of other non-hydrogen-producing bacteria (Duangmanee et
al., 2002b) in the reactor. Lay et al. (2005) demonstrated that not only Clostridium
population, but also the other microbial population was able to prevail in the hydrogen-
producing bioreactor. There is still a need to explain the shift of microbial population.
Fig. 4.5 illustrates the variation of substrate concentration at different cyclic duration in
ASBR (Sung and Dague, 1995). The corresponding cycles to the cyclic durations of 4, 6,
and 8 h are 6, 4, and 3 cycles/d, respectively. As shown in the figure, substrate
concentration at different cyclic regime is high immediately after feeding and declines
until the next cycle. Nevertheless, the initial quantity of substrate fed into the reactor at
different cyclic duration was diverse. Although the organic loading rate on daily basis
was maintained at 22.5 g COD/L/d at each cyclic regime, organic loading rates on cyclic
basis were varied based on the cyclic regime. The organic loading rates on cyclic basis
were 3.75, 5.63, and 7.50 g COD/L/cycle at cyclic durations of 4, 6, and 8 h, respectively.
According to substrate affinity described in Monod kinetics, the microbial population can
87
be selected by substrate concentration. Therefore, this change might give an opportunity
to develop other microbial populations in the reactor. In addition, the speculation of
microbial population shift could also be supported based on the results of HBu/HAc
ratios. The lower HBu/HAc ratio at longer cyclic duration was possibly attributed to the
existence of homoacetogens which are able to convert simple sugar to acetic acid without
evolving hydrogen (Ljungdahl et al., 1989). Due to the blooming of homoacetogens,
acetate became the main byproduct instead of hydrogen and butyrate.
4.5 Conclusions
Hydrogen production in ASBR was optimized by adjusting HRT, pH, substrate
concentration, and cyclic regime. At very short HRT, ORP increased to a level hostile to
the survival of hydrogen-producing bacteria in ASBR. Since long SRT could be
achieved in ASBR, due to its capability of maintaining a high level of biomass, it could
enhance the potential of developing hydrogen consumers such as methanogens. A pH of
4.9 significantly impacted the growth of hydrogen-producing bacteria. However, a
higher hydrogen yield, hydrogenic activity, and hydrogen conversion efficiency were
achieved at pH 4.9. Different to early reports where a pH of 5.0 was a critical point for
hydrogen fermentation, acid and hydrogen fermentation shifting to solvent fermentation
did not appear at pH 4.9. Moreover, to prevent the growth of hydrogen consumers at a
long SRT, it was recommended to operate ASBR at a pH of 4.9. Substrate concentration
could affect the metabolic pathway, where acetate fermentation was more prevalent than
butyrate fermentation at higher sucrose concentrations. Lower sucrose concentrations
lead to higher hydrogen conversion efficiency, but lowered the hydrogen yield. In
88
contrast, reverse results were obtained at higher sucrose concentrations. The
performance of hydrogen production evaluated based on hydrogen conversion efficiency
or hydrogen yield could significantly contribute to the different costs of operating a full-
scale hydrogen-producing reactor. Cyclic duration in ASBR obviously influenced
hydrogen production, which a short cyclic duration or a high cycle frequency helped
enhance hydrogen production. In this study, the cyclic duration of 4 h was recommended
for running ASBR.
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Yokoi, H., Maki, R., Hirose, J., Hayashi, S., 2002. Microbial production of hydrogen
from starch-manufacturing wastes. Biomass Bioenergy 22, 389-395.
Zhang, T., Liu, H., Fang, H.H.P., 2003. Biohydrogen production from starch in
wastewater under thermophilic condition. J. Environ. Management 69, 149-156.
94
Table 4.1 Summary of experimental design
Testing variables Reactor operating conditions
HRT 8, 12, 16, 24, and
48 h
pH
Sucrose concentration
Feed/decant
6.7±0.05
25 g COD/L
6 cycles/day
pH 4.9, 5.5, 6.1, and
6.7±0.05
HRT
Sucrose concentration
Feed/decant
16 h
25 g COD/L
6 cycles/day
Sucrose
concentration
15, 25, and 35 g
COD/L
pH
HRT
Feed/decant
4.9±0.05
16 h
6 cycles/day
Cyclic duration 4, 6, and 8 h/cycle
pH
HRT
Sucrose concentration
4.9±0.05
16 h
15 g COD/L
Table 4.2 Results of hydrogen production at different HRTs
HRT
(h)
ORP
(-mV)
H2
(%)
CH
4
(%)
Hydrogen yield
(mol/mol sucrose consumed)
Hydrogenic activity
(mL H
2/g VSS/d)
8
220±19
1.1±0.3
0.2±0.1
0.01±0.00
3±1
12
301±40
3.6±3.0
2.2±0.9
0.10±0.08
7±5
16
395±8
18.0±2.0
2.9±0.9
0.67±0.13
106±20
24
334±14
18.8±5.2
7.9±1.5
0.75±0.12
73±8
48
344±8
12.8±4.4
2.1±0.4
0.17±0.02
8±1
All data reported as Mean ± SD, n ≥ 3
95
Table 4.3 Hydrogen production and soluble microbial products in ASBR at each influent sucrose concentration
Soluble microbial products (m
g/L)
Sucrose
concentration
(g COD/L)
Carbohydrate
removal
efficiency
(%)
H2
(%)
H2 yield
a H2 conversion
efficiency
b
Hydrogenic
activityc
HAcd
HPre
HBuf
Ethanol
15
97.2±3.1
37.4±1.4
1.86±0.12
1.81±0.13
431±92
1417±105
387±26
2601±233
197±21
25
56.1±6.0
40.9±1.4
2.53±0.28
1.41±0.15
538±65
2017±457
615±34
2254±264
281±60
35
38.2±6.7
36.1±3.8
2.36±0.35
0.91±0.22
443±93
1687±504
617±120
1271±286
166±40
25 (repeated)
63.8±2.4
36.7±0.5
2.14±0.18
1.36±0.12
551±25
1736±254
503±15
1731±26
201±35
All data reported as Mean ± SD, n ≥ 3
a Hydrogen yield, mol/mol sucrose consumed
b Hydrogen conversion efficiency, mol/mol sucrose fed
c Hydrogenic activity, mL H
2/g VSS/d
d Acetate
e Propionate
f Butyrate
96
Table 4.4 Cyclic effect on hydrogen production and soluble microbial products in ASBR
Cyclic
duration
(h)
Carbohydrate
removal
efficiency
(%)
H2
(%)
H2 yield
a H2 conversion
efficiency
b
Hydrogenic
activityc
MLVSS
(mg/L)
Effluent
VSS
(mg/L)
SRT
(h)
F/M
d
(g COD/L/d)
HBu/HAce
(mol/mol)
4
97.2±3.1
37.4±1.4
1.86±0.12
1.81±0.13
388±39
6223±549
630±51
145±34
3.64±0.34
1.25±0.10
6
98.7±0.8
32.6±1.5
1.52±0.22
1.50±0.21
217±14
8273±293
742±155
148±34
2.72±0.09
1.33±0.19
8
99.3±0.4
28.7±1.0
1.10±0.09
1.09±0.09
170±18
8510±522
672±65
204±24
2.65±0.15
0.49±0.08
All data reported as Mean ± SD, n ≥ 3
a Hydrogen yield, mol/mol sucrose consumed
b Hydrogen conversion efficiency, mol/mol sucrose fed
c Hydrogenic activity, mL H
2/g VSS/d
d Food to microorganism ratio
e Butyrate to acetate ratio
97
98
8
6
4
2
0
H2
Biogas
100
80
60
40
20
0
H2 (%)
CH4 (%)
CO2 (%)
8000
6000
4000
2000
0
Acetate
Butyrate
Propionate
500
400
300
200
100
0
20151050
Ethanol
Butanol
1-Propanol
Gas production rate
(L/L/d)
Gas content
(%
)Acid concentration
(m
g/L)
Solvent concentration
(mg/L)
Time (days)
(a)
(d)
(c)
(b)
Fig. 4.1 Performance of ASBR operated at HRT of 16 h with pH controlled of 6.7: (a)
Gas production rate; (b) gas content; (c) acid concentration; and (d) solvent
concentration
99
100
80
60
40
20
0
7.06.56.05.55.04.5
700
600
500
400
300
200
100
0
2.0
1.5
1.0
0.5
0.0
3.0
2.5
2.0
1.5
1.0
0.5
0.0 H
2 yield
(mol H2/mol sucrose consumed)
H2 conversion efficiency
(m
ol H2/mol sucrose fed)
Hydrogenic activity
(m
L H
2/gVSS/d)
Carbohydrate removal
efficiency
(%
)
pH
R2=0.7171
R2=0.8127
R2=0.6242
(a)
(b)
(c)
(d)
Fig. 4.2 Performance of hydrogen fermentation at different pHs: (a) hydrogen yield; (b)
hydrogen conversion efficiency; (c) hydrogenic activity; (d) carbohydrate
removal efficiency
100
14
12
10
8
6
4
2
0
x103
HAc
HPr
HBu
Ethanol
Total VFA
3.0
2.5
2.0
1.5
1.0
0.5
0.0
15
10
5
0
x103
7.06.56.05.55.04.5
Soluble m
icrobial product
(mg/L)
HBu to HAc ratio
(mol HBu/mol HAc)
MLVSS
(mg/L)
pH
R2=0.7848
R2=0.8818
(a)
(c)
(b)
Fig. 4.3 Variation of soluble microbial product concentration and MLVSS concentration
at different pHs: (a) soluble microbial product concentration; (b) HBu/HAc
ratio; and (c) MLVSS concentration
101
8000
6000
4000
2000
0
4000
3000
2000
1000
0
12
10
8
6
4
2
0
40353025201510
200
150
100
50
0
Sucrose concentration (g COD/L)
F/M ratio
(g COD/g VSS/d)
SRT
(h)
MLVSS concentration
(mg/L)
Effluent VSS concentration
(m
g/L)
R2=0.9968
R2=0.9079
R2=0.9897
(c)
(b)
(a)
Fig. 4.4 (a) MLVSS and effluent VSS concentrations; (b) SRT; and (c) F/M ratio at
different sucrose concentration
102
Fig. 4.5 Effect of batch feeding on substrate concentration at different cyclic duration in
ASBR (Sung and Dague, 1995)
Cyclic duration (h)
Substrate concentration
(g COD/L)
103
CHAPTER 5. DIAGNOSIS OF HYDROGEN
FERMENTATION BY FLUORESCENCE IN SITU
HYBRIDIZATION
5.1 Introduction
Considering the energy security and the global environment, there is a pressing
need to develop non-polluting and renewable energy sources. Alternatively, hydrogen is
a clean energy carrier, producing water as its only by-product when it burns. Therefore,
hydrogen is a potential energy substitute for fossil fuels. Promising results on biological
hydrogen fermentation, using simple synthetic substrates, were obtained from completely
mixed reactors (Lin and Chang, 1999; Duangmanee et al., 2002; Khanal et al., 2005).
The investigation of hydrogen production, using real complex wastes, was also reported
by several researchers, e.g., rice winery wastewater (Yu et al., 2002; Yu et al., 2003),
sugar beets (Hussy et al., 2005), and beer processing waste (Lay et al., 2005). However,
among these studies, it should be also noted that microbial community was the key
element to complete the metabolic pathway of hydrogen fermentation. Therefore, there is
a need to perform the microbial analysis for biological hydrogen fermentation.
The biological treatment process has been developed for nearly a century. The
first activated-sludge system was built to remove organic matter in Manchester (Ardern
and Lockett, 1914). Thereafter, the outline of the role of microbial consortia in the
biological treatment process was approximately developed regarding the stabilization of
organic matters, but the entire microbial community in the treatment process is still not
104
completely understood. Nevertheless, a progressive improvement in molecular
biotechnology during the past two decades helps to deeply inspect every part of microbial
populations and their functions in the treatment process. Fluorescent in situ hybridization
(FISH) has been applied successfully to quantify and identify microbial populations in
biological wastewater treatment, such as biological nutrient removal (Coskuner and
Curtis, 2002; Pijuan et al., 2003), sludge bulking problems (Gaval et al., 2002; Lou and
de los Reyes III, 2005), etc.
In anaerobic digestion, the application of FISH has been achieved, based on
different purposes, e.g., the determination of the ratio of Bacteria and Archaea (Tay et al.,
2001), the effects of micro-aeration on the phylogenetic diversity in a thermophilic
anaerobic digester (Tang et al., 2004), hydrolytic activity of alpha-amylase in anaerobic
digested sludge (Higuchi, et al., 2005), and so forth. Furthermore, many studies have
elaborated the structure of the microbial community by using the FISH technique. Kuang
et al. (2002) studied the influence of co-substrates on methanogenic activity and their
structure in anaerobic digesters treating oleates. Shigematsu et al. (2003) analyzed the
structure of acetate-degrading methanogenic consortia at different HRTs. O’Sullivan et
al. (2005) investigated enriched cellulose degrading bacterial communities from an
anaerobic batch reactor. A comparable study was performed by Song et al. (2005), but he
and his coworkers worked on both cellulolytic microbial communities and methanogenic
populations.
With these successful efforts and a demand for microbial population analysis, the
FISH technique was used to identify and quantify the specific microbial populations in
the hydrogen fermentation study. The microbial cells collected were based on different
105
operating conditions of the bioreactor and will be quantified for assisting to diagnose the
performance of hydrogen production.
5.2 Materials and Methods
5.2.1 Reactor operation
An ASBR was seeded with sludge from a local anaerobic digester, located in
Ames, Iowa, USA. The reactor was subject to 3 to 6 feed/decant cycles. Sucrose
concentration of 15 to 35 g/L was prepared as substrate. Every liter of sucrose solution
was supplemented with 5.85 g/L NH4HCO3 and 6.7 mL of trace mineral solution
composed of KH2PO4 (150 g/L); MgSO4·7H2O (10 g/L); NaCl (1 g/L); Na2MoO4·2H2O
(1g/L); CaCl2·2H2O (1 g/L); MnSO4·7H2O (1.5 g/L); FeCl2 (0.28 g/L); CoCl2·6H2O (0.24
g/L); NiCl2·6H2O (0.12 g/L) and ZnCl2 (0.06 g/L). The ASBR had an active volume of
3.0 L and operated under a mesophilic condition (35±1ºC). Mixing was achieved by
using a mechanical mixer rotating at 300 rpm in the react phase. The biomass took 90
min in each cycle to settle prior to the decant phase. An on-line pH controller was used
to maintain a constant pH from 4.9 to 6.1 by adding 2 N hydrochloric acid or 2 N of the
mixture of sodium hydroxide and potassium hydroxide. Mixing is only carried out in the
react phase in an ASBR (Sung and Dague, 1995). Therefore, the pH controller was set to
function in the react phase.
5.2.2 FISH
FISH was performed as described by Weber et al. (2001) with some modifications.
Periodically, samples were taken from the ASBR. The samples were fixed in 100%
106
ethanol for 2 h at 4°C, and then pelleted by centrifugation (10,000xg, 3 min), followed by
preservation in a 1:1 mixture of ethanol and PBS (43.3 mM NaCl, 3.3 mM
Na2HPO4·7H2O). Three µL of fixed samples were spotted on the wells of Teflon-coated
slides. The spotted samples were air dried and subsequently dehydrated in 50, 80, and
96% ethanol (2 min each). Hybridization was performed in a chamber at a constant
temperature of 46ºC for 1.5 h. During the hybridization step, sixteen µL of the mixture
of 16S rDNA oligonucleotide probes and hybridization buffer composed of NaCl (0.9M),
Tris-HCl buffer (20mM, pH 7.2), SDS (0.01%), and formamide (probe-dependent
concentration) were added to the wells. The probes’ specificity and their conditions were
given in Table 5.1. After hybridization, a washing step was conducted by rinsing the
wells with a preheated washing buffer, consisting of NaCl (probe-dependent
concentration), Tris-HCl buffer (20mM, pH 8.0), and SDS (0.01%), to remove the excess
DNA probe from the cells. The washing step was carried out at 48ºC for 20 min.
Finally, the whole cells were counterstained with 20 µL of 4’,6-diamidino-2-
phenylindole (DAPI; 1 µg/mL) for 30 min at room temperature on the slides. Prior to
optical examination, the wells were mounted with antifading agent Citifluor AF1.
5.2.3 Microscopy and quantification analysis
The hybridization and DAPI results were observed by an epifluorescent
microscope (Zeiss, Axioplan 2) equipped with a CCD camera (Zeiss, AxioCam MRc). To
differentiate hybridized cells from the total number of cells, the hybridized cells labeled
by fluorochromes of Oregon-green or Texas-red would be illuminated at the excitation
wavelengths of 495 and 583 nm, respectively, while the total number of cells stained by
107
DAPI would be visible at the excitation wavelength of 345 nm. Thereafter, the images
taken from the camera were imported to Meta Imaging Series 6.1 for cell counting. The
quantification of cells hybridized with the specific probe relative to the total number of
cells would be completed.
5.3 Results and Discussion
5.3.1 Hybridization performance from a different 16S rDNA probe
The EUB338 probe was successfully hybridized with bacterial cells and the
hybridization results are as shown in Fig. 5.1. The figure indicates that only the active
bacteria observed from a fluorescent microscope would be hybridized with an EUB338
probe. On the other hand, the total microbial cells were determined by DAPI staining.
To examine the existence of methane producers, ARC915 probe was chose to identify the
Archaea population. However, there was a doubt on the performance of hybridization
with the ARC915 probe, due to the false positive results as indicated in Fig. 5.2. As
evident from this figure, it was found that Clostridium pasteurianum, a negative control,
was able to hybridize with the ARC915 probe. The conflict result was against the
principle of classification between the domain bacteria and the domain Archaea.
Thereafter, an experiment using Escherichia coli as a negative control was carried out to
confirm the specificity of the ARC915 probe. Fig. 5.3 illustrates the performance of
hybridization between the ARC915 probe and the Escherichia coli. As revealed in this
figure, Escherichia coli could not hybridize with the ARC915 probe and remained visible
under staining by DAPI. This ensured that the ARC915 probe was still not unique to the
domain bacteria. Therefore, due to the potential of combining the ARC915 probe to
108
Clostridium pasteurianum‘s ribosomal RNA, ARC915 probe was determined unsuitable
to recognize the Archaea population in this case. In short, there was still an unknown of
why the hybridization would occur between the ARC915 probe and Clostridium
pasteurianum‘s ribosomal RNA.
According to the ribosomal database project II (RDP II), the clostridia cluster I is
the main division, composing nearly 50% of genus Clostridium. Hydrogen-producing
Clostridium (Clostridium acetobutylicum, Clostridium pasteurianum, Clostridium
beijerinckii, etc.) mostly belongs to cluster I. Hence, the CLOST I probe was used to
hybridize with the clostridia cluster I in this study. Fig. 5.4 indicates the results of
microbe cells hybridized with the CLOST I probe labeled with Oregon-green or Texas-
red. As illustrated in Fig. 5.4(a), several individual cells became illuminated and could
be recognized as the clostridia cluster I. However, it would be difficult to distinguish the
hybridization results from those green clusters. The autofluorescence could occur in
those green clusters. Therefore, it could not be confirmed that hybridization was
absolutely performed in those green clusters. In addition, the hybridization results in Fig.
5.4 (b) show that the background was too strong to observe the microbial cells after
imaging from the CCD camera. As revealed in this figure, the microbial cells visible
from the DAPI stained were encompassed by the background color under CLOST I
hybridization. The weak fluorescent signal from the microbial cells still could be
detected under a fluorescent microscope, but could not be pictured by the CCD camera.
In contrast, the strong background was not viewed from the positive control as illustrated
in Fig. 5.4(c). A possible explanation was that some end products in the mixture culture
might disturb the in situ hybridization and, therefore, might contribute to the interference.
109
Nevertheless, speculation could question why the strong background did not occur while
the EUB338 and ARC915 probes were used to hybridize with microbe cells. In sum,
there is a need for an intensive study to improve the hybridization by the CLOST I probe
or use a different probe to target clostridia cluster I.
5.3.2 Quantification of the bacterial community
Due to the inability of determining clostridia cluster I, the correlation between the
performance of hydrogen production and the quantity of Clostridium could not be
established. Nevertheless, even without the hybridization results of the clostridia cluster
I, the microbial cells bound by the EUB338 probe could still be counted from the Meta
Imaging Series 6.1. During this stage, the performance of microbial cell counting could
be evaluated. The percentages of 16S rDNA of EUB338 to DAPI at different operating
conditions are shown in Table 5.2.
As evident from the table, the standard deviations were all over 10% of their
mean. Two main causes could lead to such large standard deviations. The sludge sample
usually appears as flocs, not individual microbial cell. Therefore, prior to hybridization,
the sludge sample had to be ground to disperse flocs in the fixation step. However, the
observation from a fluorescent microscope still found a significant amount of flocs
remained. This amount could contribute the high standard deviation during cell counting.
In addition to the interference of the floc’s appearance, the image transferring the
3-dimensional of the microbial cells to a 2-dimensional photo could mislead the
microbial cell count. Due to a visual limitation, the microbial cells overlapped on the
110
others and could not be viewed nor counted. Indirectly, those microbial cells stacked
from the others could not be counted from the software.
5.4 Conclusions
FISH was attempted as the application for the quantification of the specific
microbial populations in this study. Most bacteria successfully identified by the EUB338
probe were counted and the percentages of 16S rDNA of EUB338 to DAPI at different
reactor operating conditions were determined. False positive results hybridized by the
ARC915 probe suggested that the ARC915 probe was inappropriate to use for
recognizing domain Archaea in this study. With regard to the hybridization with the
CLOST I probe, the background issue and the microbial cell clumps hindered observation
of the hybridization performance. Therefore, the microbial cells hybridized by the
CLOST I probe could not be fully determined. Accordingly, the number of the cells of
clostridia cluster I from the microbial cell counting could not be obtained. The affiliation
between hydrogen production and the quantity of clostridia cluster I could not be
established. This study applied FISH to identify and quantify the specific microbial
populations, however, the results still remained inconclusive.
5.5 Reference
Ardern, E., Lockett, W.T., 1914. Experiments of the oxidation of sewage without the aid
of filters. J. Soc. Chem. Ind. 33, 523-539.
111
Coskuner, G., Curtis, T.P., 2002. In situ characterization of nitrifiers in an activated
sludge plant: detection of Nitrobacter Spp. J. Appl. Microbiol. 93, 431-437.
Duangmanee, T., Chyi, Y., Sung, S., 2002. Biohydrogen production in mixed culture
anaerobic fermentation. In: Proc. 14th World Hydrogen Energy Conf., Québec,
Canada.
Gaval, G., Duchène, P., Pernelle, J.J., 2002. Filamentous bacterial population dominance
in activated sludges subject to stresses. Water Sci. Technol. 46, 49-53.
Higuchi, Y., Ohashi, A., Imachi, H., Harada, H., 2005. Hydrolytic activity of alpha-
amylase in anaerobic digested sludge. Water Sci. Technol. 52, 259-266.
Hussy, I., Hawkes, F.R., Dinsdale, R., Hawkes, D.L., 2005. Continuous fermentative
hydrogen production from sucrose and sugarbeet. Int. J. Hydrogen Energy 30,
471-483.
Khanal, S.K., Chen, W.H., Li, L., Sung, S., 2005. Biohydrogen production in continuous
flow reactor using mixed microbial culture. Water Environ. Res. 78 (2), 110-117.
Kuang, Y., Lepesteur, M., Pullammanappallil, P., Ho, G.E., 2002. Influence of co-
substrates on structure of microbial aggregates in long-chain fatty acid0fed
anaerobic digesters. Lett. Appl. Microbiol. 35, 190-194.
Lay, J.J., Tsai, C.J., Huang, C.C., Chang, J.J., Chou, C.H., Fan, K.S., Chang, J.I., Hsu,
P.C., 2005. Influences of pH and hydraulic retention time on anaerobes
converting beer processing wastes into hydrogen. Water Sci. Technol. 52, 123-
129.
Lin, C.Y., Chang, R.C., 1999. Hydrogen production during the anaerobic acidogenic
conversion of glucose. J. Chem. Technol. Biotechnol. 74, 498-500.
112
Liu, W.T., Chan, O.C., Fang, H.H.P., 2002. Characterization of microbial community in
granular sludge treating brewery wastewater. Water Res. 36, 1767-1775.
Lou, I., de los Reyes III, F.L., 2005. Substrate uptake tests and quantitative FISH show
differences in kinetic growth of bulking and non-bulking activated sludge.
Biotech. Bioeng. 92, 729-739.
O’Sullivan, C.A., Burrel, P.C., Clarke, W.P., Blackall, L.L., 2005. Structure of a
cellulose degrading bacterial community during anaerobic digestion. Biotech.
Bioeng. 92, 871-878.
Pijuan, M., Saunders, A.M., Guisasola, A., Baeza, J.A., Casas, C., Blackall, L.L., 2003.
Enhanced biological phosphorus removal in a sequencing batch reactor using
propionate as the sole carbon source. Biotech. Bioeng. 85, 56-67.
Shigematsu, T., Tang, Y.Q., Kawaguchi, H., Ninomiya, K., Kijima, J., Kobayashi, T.,
Morimura, S., Kida, K., 2003. Effect of dilution rate on structure of a mesophilic
acetate-degrading methanogenic community during continuous cultivation. J.
Biosci Bioeng. 96, 547-558.
Song, H, Clarke, W. P., Blackall, L.L., 2005. Concurrent microscopic observations and
activity measurements of cellulose hydrolyzing and methanogenic populations
during the batch anaerobic digestion of crystalline cellulose. Biotech. Bioeng. 91,
369-378.
Sung, S., Dague, R.R., 1995. Laboratory studies on the anaerobic sequencing batch
reactor. Water Environ. Res. 67, 294-301.
113
Tang, Y., Shigematsu, T., Ikbal, Morimura, S., Kida, K., 2004. The effects of micro-
aeration on the phylogenetic diversity of microorganisms in a thermophilic
anaerobic municipal solid-waste digester. Water Res. 38, 2537-2550.
Tay, T.L.S., Ivanov, V., Kim, I S., Feng, L., Tay, J.H., 2001. Quantification of ratios of
Bacteria and Archaea in methanogenic microbial community by fluorescence in
situ hybridization and fluorescence spectrometry. World J. Mirco. Biotech. 17,
583-589.
Weber, S., Stubner, S., Conrad, R., 2001. Bacterial populations colonizing and degrading
rice straw in anoxic paddy soil. Appl. Environ. Microbiol. 67, 1318-1327.
Yu, H., Zhu, Z., Hu, W., Zhang, H., 2002. Hydrogen production from rice winery
wastewater in an upflow anaerobic reactor by using mixed anaerobic cultures. Int.
J. Hydrogen Energy 27, 1359-1365.
Yu, H., Hu, Z., Hong, T., 2003. Hydrogen production from rice winery wastewater by
using a continuously-stirred reactor. J. Chem. Eng. Japan 36, 1147-1151.
Table 5.1 16S rDNA oligonucleotide probes used for hybridization
Probe Nam
e Specificity
Sequence of probe (5’-3’)
FA
(%)a
NaCl
(mM)b
Fluorc
Reference
EUB338
Bact
eria
GCTGCCTCCCGTAGGAGT
20
0.17
Texas-red
Liu et al., 2002
ARC915
Arc
haea
GTGCTCCCCCGCCAATTCCT
20
0.17
Texas-red
Liu et al., 2002
CLOST I
clostridia cluster I
and II
TTCTTCCTAATCTCTACGCA
30
0.1
Oregon-green/Texas-red
Weber et al., 2001
a FA, formam
ide concentration
b NaCl concentration in washing buffer
c Fluor, fluorochrome
114
115
Table 5.2 Summary of the percentage of EUB338 to DAPI at different operating
conditions
Operation conditions EUB338/DAPI (%)a
4.9 83 ± 10
5.5 75 ± 13 pH
6.1 37 ± 14
15 78 ± 8
25 83 ± 10 Sucrose concentrations
(g COD/L) 35 76 ± 15
4 78 ± 8
6 67 ± 11 Cyclic durations
(h) 8 68 ± 12
a Mean ± SD (n ≥ 35)
116
Fig. 5.1 Hydrogen-producing sludge hybridized with the EUB338 probe and stained
with DAPI. Scale bars. 5 µm
EUB338 DAPI
117
Fig. 5.2 Clostridium pasteurianum hybridized with the ARC915 probe and stained with
DAPI. Scale bars, 5 µm
ARC915 DAPI
118
Fig. 5.3 Escherichia coli hybridized with the ARC915 probe and stained with DAPI.
Scale bars, 5 µm
ARC915 DAPI
119
Fig. 5.4 (a) Hydrogen-producing sludge hybridized with the CLOST I probe labeled by
Oregon-green, and stained with DAPI; (b) hydrogen-producing sludge
hybridized with the CLOST I probe labeled by Texas-red, and stained with
DAPI; and (c) Clostridium pasteurianum hybridized with the CLOST I probe
labeled by Texas-red, and stained with DAPI. Scale bars, 5 µm
CLOST I
(a)
DAPI
CLOST I
(b)
DAPI
CLOST I
(c)
DAPI
120
CHAPTER 6. CONCLUSIONS
6.1 Closing Statements
Anaerobic bioconversion of organic wastes to hydrogen gas is an attractive option
that not only stabilizes the waste/wastewater, but also generates a benign renewable
energy carrier. To fully establish the knowledge of hydrogen fermentation, this study
was begun to determine the growth kinetics of hydrogen production in dark fermentation
from different characteristics of substrates. Based on the kinetic study, a suitable waste
was selected as limiting substrates for further investigation in continuous operation. The
following study was attempted to obtain the maximum hydrogen production by fine-
tuning the operation conditions of ASBR. In addition, since the performance of hydrogen
fermentation was related to the appearance of microbial populations, microbial analysis
was carried out during the study.
The study of growth kinetics was performed by a series of batch experiments.
Three different substrates (sucrose, NFDM, and food waste) were selected in this study to
investigate the substrate affinity of mixed microbial culture for biological hydrogen
production. The effects of substrate concentration on the biohydrogen production were
well described by Michaelis-Menten equation. In general, the hydrogen production rate
increased with increasing substrate concentration in the exponential stage, but it
plateaued at high substrate concentrations. The substrate affinity significantly affected
the hydrogen yield. The KS values of sucrose, NFDM, and food waste were 1.4, 6.6, and
8.7 g COD/L, respectively. Nevertheless, the maximum hydrogen yields from sucrose,
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NFDM, and food waste were 234, 119, and 101 mL/g COD, respectively. Hydrogen
yield from sucrose was found to be much higher than the yield from NFDM and food
waste. This suggested that higher hydrogen yield from sucrose was attributed to higher
carbohydrate content. It also recommended that, due to high biochemical reaction rate of
hydrogen evolution, the substrate contained a high fraction of particulates that would not
appropriate for hydrogen fermentation. Without pH control, the substrate could not be
completely metabolized by hydrogen-producing bacteria at a higher concentration, owing
to a pH lower than 4, which inhibited hydrogen-producing bacteria.
In the kinetic study, it found that soluble carbohydrate-rich waste was the ideal
carbon and energy sources for hydrogen fermentation. Therefore, sucrose was chosen to
mimic the real carbohydrate-rich waste for the continuous operation in the following
study. Hydrogen production in ASBR was optimized by adjusting HRT, pH, substrate
concentration, and cyclic regime. In the study, the maximum hydrogen yield of 2.53 mol
H2/mol sucrose consumed and the maximum hydrogenic activity of 538 mL H2/g VSS/d
were obtained at HRT of 16h, pH 4.9, sucrose concentration of 25 g COD/L, and with 4-h
cyclic duration.
At very short HRT, ORP increased to a level hostile to the survival of hydrogen-
producing bacteria in ASBR. Since long SRT could be achieved in ASBR, due to its
capability of maintaining a high level of biomass, it could enhance the potential of
developing hydrogen consumers such as methanogens. A pH of 4.9 significantly
impacted the growth of hydrogen-producing bacteria. However, a higher hydrogen yield,
hydrogenic activity, and hydrogen conversion efficiency were achieved at pH 4.9.
Different to early reports where a pH of 5.0 was a critical point for hydrogen
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fermentation, acid and hydrogen fermentation shifting to solvent fermentation did not
appear at pH 4.9. Moreover, to prevent the growth of hydrogen consumers at a long SRT,
it was recommended to operate ASBR at a pH of 4.9. Substrate concentration could
affect the metabolic pathway, where acetate fermentation was more prevalent than
butyrate fermentation at higher sucrose concentrations. Lower sucrose concentrations
lead to higher hydrogen conversion efficiency, but lowered the hydrogen yield. In
contrast, reverse results were obtained at higher sucrose concentrations. The
performance of hydrogen production evaluated based on hydrogen conversion efficiency
or hydrogen yield could significantly contribute to the different costs of operating a full-
scale hydrogen-producing reactor. Cyclic duration in ASBR obviously influenced
hydrogen production, which a short cyclic duration or a high cycle frequency helped
enhance hydrogen production. In this study, the cyclic duration of 4 h was recommended
for running ASBR.
FISH was applied to the quantification of the specific microbial populations in
this study. Most bacteria successfully identified by the EUB338 probe were counted and
the percentages of 16S rDNA of EUB338 to DAPI at different reactor operating
conditions were determined. False positive results hybridized by the ARC915 probe
suggested that the ARC915 probe was inappropriate to use for recognizing domain
Archaea in the study. With regard to the hybridization with the CLOST I probe, the
background issue and the microbial cell clumps hindered observations of the
hybridization performance. Therefore, the microbial cells hybridized by the CLOST I
probe could not be fully determined. Accordingly, the number of clostridia cluster I from
the microbial cell counting could not be obtained. The correlation between hydrogen
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production and the quantity of clostridia cluster I could not be established. The study
applied FISH to identify and quantify the specific microbial populations, which still
remained inconclusive.
6.2 Recommendations for Future Work
The survey of biological hydrogen production from this study gives a direction for
future investigations, where some prospects may be considered as potential research.
They include:
1. Use real soluble organic waste.
Our study demonstrated that soluble carbohydrate-rich wastes are the eligible
candidates for hydrogen fermentation. Therefore, regardless of the synthetic
wastes, there is a need to examine hydrogen fermentation from real organic
wastes such as molasses, sugar cane, rice winery waste water, etc.
2. Develop a two-stage hydrogen-producing reactor—dark fermentation followed by
photo fermentation.
Although many studies, including our research, attempted to maximize hydrogen
yields by using diverse reactor configurations, e.g., fine-tuning operating
conditions, etc., due to the limitation of the stochiometric principle, there still was
no significant breakthrough from dark fermentation. Nevertheless, combining
two metabolic pathways of dark fermentation and photo fermentation may
enhance the entire hydrogen yield to the level of providing sufficient amounts for
commercialization. The concept can be conceived in a two-stage hydrogen-
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producing reactor, where the first reactor—working with dark fermentation—is
followed by the second reactor, executing photo fermentation.
3. Adopt suitable molecular biology tools for microbial analysis.
Our study showed difficulties in using the FISH technique to identify the
microbial population, particularly the genus Clostridia. The problems might be
the interference of byproducts, nonspecific staining, wrong DNA probe, and
inappropriate hybridization conditions. Therefore, there may be a need to apply
different molecular biology tools. Terminal restriction fragment length
polymorphism (T-RFLP) or real-time polymerase chain reaction (real-time PCR)
can be used as alternative tools to examine the microbial populations. The
mechanism of T-RFLP is in comparison with the specific 16s rDNA fragments
cut by the restriction enzymes with those known microorganisms to determine the
microbial population. Real-time PCR is used to simultaneously quantify and
amplify a specific given DNA for determining the presence of the specific
microbial population and its quantity.
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ACKNOWLEDGEMENTS
There are several people to whom I owe a great thankfulness for this work. With
sincere appreciation, I would like to acknowledge my major professor, Dr. Shihwu Sung,
for his unlimited support and his guidance of this study. He invested me plenty of time
and effort to make me become what a Ph.D. should be. He critiqued my study and
challenged my thought. Except for the disciplines, he encouraged me when I faced the
frustration. He inspired me as a friend. I never met a professor like him who intended to
enlighten me seeing the world and appreciating the life.
Special thanks to Dr. Samir Kumar Khanal’s invaluable guidance. He also
challenged my research and encouraged me to achieve higher goals. As a non-native
English speaker, he improved my writing skill in the two years working with him.
I would like to thank Dr. Dennis Bazylinski and Tim Williams’ assistance on
microbial pure culture development, and thank Dr. Kenneth Koehler’s instruction on
statistical analysis. I feel grateful to my committee members, Dr. Say Kee Ong and Dr.
Timothy Ellis, for their comments on this dissertation.
Dr. Robert Doyle in Ultrahigh Resolution Centre provided me free facilities
including chemicals, microbial cell counting software, etc. Dr. Harry Horner and his
group in Bessey Microscopy Facility offered me the use of fluorescence miscroscope.
Without several laboratory facilities, I could not complete this work.
I am also appreciative of all other individuals who offered a great help in the
biotech group of Civil, Construction, and Environmental Engineering department.
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Finally, I would like to thanks my family for believing in me. With their strong support, I
can accomplish one of the important goals in my life.