Fermentative hydrogen production by the newly isolated Enterobacter asburiae SNU-1

International Journal of Hydrogen Energy 32 (2007) 192 – 199 www.elsevier.com/locate/ijhydene

Fermentative hydrogen production by the newly isolated Enterobacter asburiae SNU-1 Jong-Hwan Shin a , Jong Hyun Yoon a , Eun Kyoung Ahn a , Mi-Sun Kim b , Sang Jun Sim c , Tai Hyun Park a,∗ a School of Chemical and Biological Engineering, Seoul National University, Seoul 151-744, Republic of Korea b Biomass Research Team, Korea Institute of Energy Research, Daejeon 305-343, Republic of Korea c Department of Chemical Engineering, Sungkyunkwan University, Suwon 440-746, Republic of Korea

Available online 26 September 2006

Abstract A new fermentative hydrogen-producing bacterium was isolated from a domestic landfill and identified as Enterobacter asburiae using 16S rRNA gene sequencing and DNA–DNA hybridization methods. The isolated bacterium, designated as Enterobacter asburiae SNU-1, is a new species that has never been examined as a potential hydrogen-producing bacterium. This study examined the hydrogen-producing ability of Enterobacter asburiae SNU-1. During fermentation, the hydrogen was mainly produced in the stationary phase. The hydrogen yield based on the formate consumption was 0.43 mol hydrogen/mol formate. This strain was able to produce hydrogen over a wide range of pH (4–7.5), with the optimum pH being pH 7. The level of hydrogen production was also affected by the initial glucose concentration, and the optimum value was found to be 25 g glucose/l. The maximum and overall hydrogen productivities were 398 and 174 ml/l/hr, respectively, at pH 7 with an initial glucose concentration of 25 g/l. This strain could produce hydrogen from glucose and many other carbon sources such as fructose, sucrose, and sorbitol. 䉷 2006 International Association for Hydrogen Energy. Published by Elsevier Ltd. All rights reserved. Keywords: Enterobacter asburiae SNU-1; Hydrogen production; Fermentation; Formate decomposition

1. Introduction Biological processes for the production of hydrogen are more environment-friendly and less energy intensive than thermochemical and electrochemical processes [1–3]. Biohydrogen can be produced by photosynthetic microorganisms [4] or non-photosynthetic fermentative bacteria. Non-photosynthetic fermentative bacteria produce hydrogen without light under anaerobic conditions, which makes the bioreactor design much simpler than for photosynthetic microorganisms. These fermentative bacteria include Enterobacter [5–9], Bacillus [10], Rhodopseudomonas [11], and Clostridium [12] species. Some of these are facultative anaerobes and others are obligate anaerobes. Some Enterobacter species (facultative anaerobe) can degrade soluble starch [13], while some Clostridium species

∗ Corresponding author. Tel.: +82 2 880 8020; fax: +82 2 875 9348.

E-mail address: [email protected] (T.H. Park).

(obligate anaerobe) can degrade insoluble starch and cellulose without any pretreatment. The hydrogen production yield per unit carbon source is higher with obligate anaerobes than with facultative anaerobes. Despite this many studies on hydrogen production have used facultative anaerobes. This is in part due to the difficulty in maintaining the strict anaerobic conditions needed with obligate bacteria [14]. Obligate anaerobes are extremely sensitive to trace amounts of dissolved oxygen, resulting in the need for expensive reducing agents to be added to the culture medium. However, facultative anaerobes are less sensitive to dissolved oxygen, and the activity of the enzyme involved in hydrogen production recovers rapidly from oxygen damage when depleted in the culture medium. Compared with obligate anaerobes, the rates of cell growth and hydrogen production are higher, and the total operation time for the hydrogen production is lower. Various facultative anaerobes have been examined for their potential in hydrogen production. Enterobacter cloacae

0360-3199/$ - see front matter 䉷 2006 International Association for Hydrogen Energy. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.ijhydene.2006.08.013

J.-H. Shin et al. / International Journal of Hydrogen Energy 32 (2007) 192 – 199

IIT-BT 08 was reported to show a high maximum volumetric hydrogen production rate of 447 ml H2 /l/h [15]. Other facultative anaerobic Enterobacter aerogenes strains such as HU-101, AY-2, and HO-39 have been also investigated [6,8]. As part of an ongoing study aimed at isolating new microorganism with a higher capability of hydrogen production than previously reported ones, a new fermentative bacterium, Enterobacter asburiae, was isolated from a landfill. It was found that this strain has unique properties in hydrogen production, which has not been observed in other hydrogen-producing strains. In this study, the hydrogen production performance by this strain was examined under various culture conditions in order to determine the optimal conditions for the maximal hydrogen production. 2. Materials and methods 2.1. Isolation of the bacterial strain The hydrogen-producing bacterial strain was isolated from a domestic landfill. The strain was isolated using a method described elsewhere [16]. Soil samples were collected from several domestic landfill areas and incubated in 50 ml of an LB medium for several hours. After incubation, 0.5 ml of the supernatant was transferred to 50 ml of minimal medium consisting of 50 mM L-homophenylalanine (L-HPA) as the sole nitrogen source, 50 mM potassium phosphate buffer (pH 7.2), 100 mM glycerol, 1 g/l MgSO4 ·7H2 O, 0.2 mM CaCl2 , 0.1 mg/l ZnCl2 , 0.1 mg/l MnSO4 · 4H2 O, 0.02 mg/l H3 BO3 , 0.1 mg/l CuSO4 · 5H2 O, 0.05 mg/l CoCl2 , 0.1 mg/l NiSO4 · 6H2 O, 2.0 mg/l NaMoO4 , and 4.0 mg/l FeSO4 · 7H2 O. After cultivation at 37 ◦ C for 2 days, 0.5 ml of the culture broth was diluted in 50 ml of the same minimal medium (100-fold dilution). These procedures were repeated three times in order to simulate the enrichment culture. After the enrichment steps, the culture broth was spread out on an agar plate with the same composition as the minimal media. Each colony on the plate was cultivated in the same type of minimal or LB media. The method used to isolate the strain was originally used for the screening of microorganisms possessing an amino acid transferase [15]. The microorganisms screened by this method also showed extremely high dehydrogenase activity, which is a malate dehydrogenase-like enzyme (personal communication). Microorganisms possessing a high dehydrogenase activity that is involved in NADH production are expected to be good hydrogen producers. Among the 30–40 colonies obtained after the enrichment steps, several isolates were further examined for hydrogen production. The strain with the highest level of hydrogen production in the evolved gas among the isolates was selected. This strain was stored in a 25% glycerol solution at −70 ◦ C for further use. 2.2. Biochemical test and fatty acid analysis An API 20E Kit (BioMerieux, St. Louis, MO, USA) was used for the biochemical tests. The fatty acids were analyzed to determine the taxonomy of the isolate. The fatty acid methyl

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esters were prepared from the biomass grown on triptic soy agar (TSA) at 30 ◦ C for 1 day and were identified using a Microbial Identification System (Microbial ID, Inc., USA) [17]. 2.3. 16S rDNA sequencing and phylogenetic analysis 2.3.1. DNA extraction The chromosomal DNA was isolated using a slight modification of the method reported by Pitcher et al. [18]. Small amounts of biomass from the glucose–yeast extract agar plates were mixed and gently homogenized in 100 l of T.E. buffer (pH 8) supplemented with 50 mg/ml lysozyme (Sigma, Ltd., Poole, Dorset, UK). The resulting solutions were incubated at 37 ◦ C overnight. The aqueous layers were separated by centrifugation and extracted with chloroform isoamyl alcohol (25:1, vol/vol). After the RNase treatment, the DNA was extracted with phenol and chloroform. 2.3.2. Determination of 16S rDNA sequence A large fragment of the 16S rRNA gene was amplified by PCR using the universal primers 27F (5 -AGA GTT TGA TCM TGG CTC AG-3 ) and 1522R (5 -AAG GAG GTG WTC CAR CC-3 ) [19]. The PCR products were purified using a Wizard PCR Preps DNA Purification System (Promega, USA) according to the manufacturer’s instructions and sequenced using a BigDyeTM Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, USA) and a model 3100 automatic sequencer (Applied Biosystems, USA). The closest known relatives of the new isolates were determined by performing a sequence database search. The sequences of the closely related strains were retrieved from GenBank and the Ribosomal Database Project (RDP) libraries. The nucleotide (NT) sequence similarities were calculated using the PHYDIT program [20]. 2.3.3. Phylogenetic analysis Phylogenetic analysis was performed using the neighborjoining methods [21]. Evolutionary distance matrices were generated using the method reported by Jukes and Cantor [22]. The resultant neighbor-joining tree topology was evaluated using bootstrap analyses based on 1000 samples [23]. Alignment and phylogenetic analyses were carried out using the PHYDIT program (available at http://chunlab.snu.ac.kr/jphydit/). 2.4. Strain identification by DNA hybridization DNA–DNA hybridization was carried out based on a membrane filter technique using a DIG High Prime DNA Labeling and Detection Starter Kit II (Roche Molecular Biochemicals, Germany). The genomic DNA (300 ng) was denatured using the alkaline method, and immobilized on a nylon membrane (Hybond-N+, Amersham, UK) by applying a low vacuum. The DNA preparations (2000 ng) were labeled using the DIG High Prime DNA Labeling and Detection Starter Kit II according to the manufacturer’s protocol. The membranes were then prehybridized in a hybridization solution at 42 ◦ C for 30 min. The

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actual prehybridization was carried out in a hybridization solution containing the labeled DNA (25 ng/ml) at 65 ◦ C for 16 h. After hybridization, the membranes were washed twice in a primary washing solution (2 × SSC and 0.1% SDS) and twice in a secondary solution (0.5×SSC and 0.1% SDS) at 65 ◦ C. The detection reagents were added to the membranes for 5 min at room temperature, and the membranes were exposed to autoradiography film (Hyperfilm-ECL, Amersham Biosciences, UK) for 15 min. The signal intensities were determined using the TINA 2.0 program. The signal produced by the self-hybridization of the probe with the homologous target DNA was considered to be 100%, and the percentage similarity was determined for the duplicate samples. 2.5. Bacterial cell culture The cells were cultured at 37 ◦ C in a 1 l fermentor (working volume 400 ml) under anaerobic conditions and stirred at an impellor speed of 220 rpm. The seed was cultured in a 120 ml serum bottle (working volume 20 ml) at 37 ◦ C and agitated in a shaking incubator at 220 rpm. The serum bottle was purged with nitrogen gas to produce the required anaerobic conditions and sealed with a 12 mm-thick butyl rubber septum and an aluminum cap. After seed cultivation, the inoculum was transferred anaerobically into the fermentor at the late exponential phase. After inoculation, the fermentor was purged with nitrogen gas for 20 min to produce the anaerobic conditions. The PYG medium containing 10 g/l glucose, 20 g/l peptone, 3 g/l yeast extract, 0.5 g/l MgSO4 · 7H2 O, and 2 g/l KH2 PO4 was used for fermentation. The pH was set to various levels in the fermentor using 2 N HCl and 2 N NaOH. The concentrations of all components in medium except for glucose were fixed in all experiments. The cell concentration was determined by measuring the absorbance at 680 nm using a spectrophotometer (Genesys, Spectronic Instruments, USA). One unit of absorbance was equal to 0.42 g dry cell/l. 2.6. Analysis of liquid and gas phases The glucose concentration was analyzed using an enzymatic method (Glucose HK Assay Kit, Sigma Chemical Co., St. Louis, USA). The organic acids and alcohols in the culture broth were analyzed by high-performance liquid chromatography (HPLC) (HP1100, Agilent Technologies, USA), which was equipped with an Aminex HPX-87H packed column (⭋300 × 7.8 mm; Bio-Rad, USA). Prior to injection, the liquid samples were purified through a 0.2 m disposable filter. A refractive index detector (RID) was used to quantify the organic acids and alcohols. A 0.01 N H2 SO4 solution at 0.6 ml/min was used as the mobile phase and the column temperature was kept at 40 ◦ C. The injection volume of the samples was 10 l. The H2 concentration in the gas phase was analyzed by gas chromatography (HP 5890 Series II, Hewlett-Packard, USA), which was equipped with a thermal conductivity detector and a Carboxen-1000 column (Supelco, USA) [24]. Argon (Ar) was used as the carrier gas at 58 ml/min. The oven, injector, and detector were maintained at 30, 120, and 120 ◦ C, respectively.

3. Results and discussion 3.1. Phenotypic characterization of the hydrogen-producing isolate The hydrogen-producing bacterium was isolated from several domestic landfill areas. Biochemical test carried out to identify the isolate (data not shown) indicated that the strain belonged to the genus Enterobacter and had a 90.9% similarity to Enterobacter cloacae. Table 1 shows the fatty acid composition of the isolate. The result was compared with the TSBA40 (aerobic bacteria database) and CLIN40 databases (clinical bacteria database), which indicated an 84.2% similarity with the Escherichia coli GC subgroup C on the TSBA40 database and a 62.7% similarity with Escherichia hermannii on the CLIN40 database. 16S rRNA gene full sequencing was carried out to identify the isolated strain, and the 1377 bp sequence was determined. Based on the similarity of the 16S rRNA gene, the isolate was found to be analogous to Enterobacter asburiae JCM 6051T (99.56%), Pantoea agglomerans JCM 1236T (99.42%), and Enterobacter cancerogenus LMG 2693T (99.27%). A phylogenetic tree was constructed (Fig. 1). Because the species of the isolate could not be determined by the 16S rDNA analysis, DNA–DNA hybridization was performed for more accurate identification using a DIG High Prime DNA Labeling and Detection Starter Kit II (Roche Molecular Biochemicals, Germany). The isolate had the highest similarity to Enterobacter asburiae JCM 6051T (76.7%). Two strains are considered to be the same species if the similarity is higher than 70%. Therefore, the isolate was determined to belong to Enterobacter asburiae Table 1 Fatty acid composition of the isolated strain (analyzed using Hewlet-Packard model 6890A gas chromatography and MIDI Aerobe method, Chem Station ver. 4.02) Fatty acid

Composition (%)

Unknown 10.928 C12:0 C13:0 C14:0 C13:0 3OH/15:1 i I/H C15:0 C14:0 2OH C16:1 ISO I/14:0 3OH C16:1 7c/15 iso 2OH C16:0 C17:1 8c C17:0 CYCLO C17:0 C18:1 7c

0.72 2.95 0.57 6.88 0.62 2.93 0.85 7.91 21.04 24.09 0.62 5.74 1.98 21.53

Identification result (similarity)

TSBA40a Escherichia coli GC subgroup C (0.842)

CLIN40a

Klebsiella planticola Enterbacter cloacae Escherichia hermannii Enterobacter cloacae

TSBA40: aerobic bacteria, CLIN40: clinical bacteria. a Comparison databases.

(0.835) (0.713) (0.627) (0.602)

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195

Plesiomonas shigelloides ATCC 14029T (X74688) Enterobacter pyrinus KCTC 2520T (AJ010486) 85

Pantoea ananatis LMG 2665T (Z96081)

97

Pantoea stewartii subsp. indologenes LMG 2632T (Y13251)

99

56

Pantoea stewartii subsp. stewartii ATCC 8199T (U80208) Enterobacter gergoviae JCM 1234T (AB004748) Enterobacter hormaechei CIP 103441T (AJ508302) Enterobacter kobei CIP 105566 (AJ508301) Enterobacter aerogenes JCM 1235T (AB004750)

56

Enterobacter nimipressuralis LMG 10245T (Z96077)

50 61

Enterobacter amnigenus JCM 1237T (AB004749) Enterobacter intermedius JCM 1238T (AB004747) Enterobacter cowanii CIP 107300T (AJ508303)

56

Enterobacter cloacae ATCC 13047T (AJ251469) 99

Enterobacter dissolvens LMG 2683T (Z96079)

Pantoea agglomerans JCM 1236T (AB004691) 51

Enterobacter asburiae SNU-1 Enterobacter asburiae JCM 6051T (AB004744) 71

Enterobacter cancerogenus LMG 2693T (Z96078) Enterobacter sakazakii JCM 1233T (AB004746)

1%

Fig. 1. Neighbor-joining tree showing the phylogenetic position of the isolated strain, Enterobacter asburiae SNU-1, based on the 16S rDNA sequences of Enterobacteriae. The numbers at the nodes indicate the levels of bootstrap support based on the neighbor-joining analysis of 1000 resampled data sets: only the values  50% are given. The scale bar indicates 0.01 nucleotide substitutions per nucleotide position (1%).

and was designated Enterobacter asburiae SNU-1. The major fermentative bacteria used for hydrogen production includes several Enterobacter and Clostridium species. Enterobacter aerogenes and Enterobacter cloacae are known to produce hydrogen from carbohydrates. The Enterobacter species isolated in this study is a new species that has not been examined for hydrogen production. 3.2. Batch fermentation profiles of the isolated strain Fig. 2 shows the fermentation profiles of the isolated hydrogen-producing bacterial strain, Enterobacter asburiae SNU-1. Hydrogen production began when cell growth entered the late exponential phase, and the rate of hydrogen production reached a maximum in the early stationary phase (Fig. 2a, b). Most of the hydrogen was produced in the stationary phase. Hydrogen production in the stationary phase might be due to the decomposition of formate that accumulates during the exponential phase.

Fig. 2c shows the pH and metabolite profiles during the anaerobic culture when the pH was not controlled. The pH decreased from 7.1 to 5.7 and reached a minimum after 7 h. The pH then increased from 5.7 to 6.6. The initial decrease in pH was attributed to the accumulation of organic acids [5–9,12]. The time to reach the minimum pH matched the time to reach the maximum formate concentration. The concentrations of the other acidic compounds were relatively low. This suggests that the decrease in pH before 7 h was mainly due to the accumulation of formate in the fermentation broth, and the increase in pH after 7 h was caused by the consumption of formate. At this point, the cells almost reached the stationary phase because the glucose was almost completely consumed. The cells then need to use another carbon source, which was formate in this case. At this point, formate consumption began, which resulted in an increase in pH and hydrogen production. This hydrogen production is believed to occur through formate decomposition, which works at high formate concentrations via formate hydrogen lyase (FHL) [25].

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Cell concentration (g/l)

10 1 8 6 4 Cell Glucose

2

0.1

0 0

2

4

6

8

10

12

14

16

18

Culture time (hr)

350

8.0

300

7.5

250 7.0 pH

200 6.5

150 6.0

100 50

5.5

0 75

5.0 60 Metabolite concetration (mM)

H2 productivity (ml/l/h)

H2 production/unit volume (ml/l)

(a)

60

45

30

15

formate ethanol butanediol acetate lactate succinate butylate

50 40 30 20 10 0

0 0

(b)

Residual glucose concentration (g/l)

12

2

4

6

8

10

12

14

16

0

18

Culture time (hr)

2

4

6

(c)

8

10

12

14

16

18

Culture time (hr)

Fig. 2. Fermentation profiles of Enterobacter asburiae SNU-1. The initial glucose concentration was 10 g/l, and the pH was not controlled. (a) The profiles of cell growth and glucose consumption, (b) the profiles of H2 production per unit volume and the H2 productivity, and (c) the profiles of pH and metabolites.

The hydrogen yield based on the formate consumption was 0.43 mol hydrogen/mol formate (Fig. 3). The glucose concentration was zero in the stationary phase and the concentrations of other metabolites were maintained at a constant level, as shown in Fig. 2c. Therefore, the level of formate production would be negligible in the stationary phase. However, between 7 and 8 h, the yield was 1.43 mol hydrogen/mol formate, which was the deceleration phase. It is believed that a yield > 1.0 mol hydrogen/mol formate, which is the theoretical maximum, is caused by the production of formate. The calculation of the yield was based on the assumption that no formate would be produced. However, formate might be still produced during this period. This formate production was not consid-

ered in this calculation, which resulted in an overestimation of the yield. Biohydrogen from Enterobacter species is mainly produced through two different ways. One is via formate decomposition, which evolves hydrogen by FHL [1]: HCOOH → H2 + CO2 . The other one is the NADH pathway, which evolves hydrogen by hydrogenase through the reoxidation of NADH that is produced via the glycolysis [26]: NADH + H+ → NAD+ + H2 .

J.-H. Shin et al. / International Journal of Hydrogen Energy 32 (2007) 192 – 199 14

Hydrogen concentration (mM)

12 10 8

0.43 (mol H2/mol formate)

6 4

1.4

2

(mol H2/mol formate)

0 0.0

2.5

5.0

7.5 10.0 12.5 15.0 Formate concentration (mM)

17.5

197

The local maximum at pH 5.5 might be caused by a different effect of pH on the cell growth and the hydrogen production per unit cell. The pH 7 was favorable to the cell growth while pH 5.5 was favorable to the specific hydrogen production. The level of hydrogen production per unit volume without pH control was similar to that with the pH set to 6 or 6.5. This is because the pH varied in this range during the period of hydrogen production, as shown in Fig. 2c. Although the level of hydrogen production had a local maximum at pH 5.5, it has a global maximum at pH 7. When the pH was set to pH 7, the level of hydrogen production/unit volume and the maximum hydrogen productivity were 665 ml/l and 175 ml/l/h, respectively.

20.0

3.4. Effect of initial glucose concentration

Fig. 3. Hydrogen production yield based on the level of formate consumption.

3.3. Effect of pH on hydrogen production As fermentation progresses, various organic acids accumulate resulting in a significant decrease in pH in the culture medium. This decrease in pH caused a significant decrease in hydrogen production. Table 2 shows the effect of pH on the growth of Enterobacter asburiae SNU-1 and the level of hydrogen production. The results show that Enterobacter asburiae SNU-1 can produce hydrogen at pH 4–7.5, whereas, strictly anaerobic Clostridium butyricum CGS5, which was isolated from anaerobic sewage sludge, did not produce any hydrogen at pH 5 [12]. However, pH control is important for the efficient production of hydrogen by Enterobacter asburiae SNU-1. The level of cell growth increased with increasing pH from 4 and reached a maximum at pH 7, as shown in Table 2. However, the hydrogen production behavior was different from that of cell growth. A local maximum for hydrogen production was obtained at pH 5.5, i.e. the level of hydrogen production was higher at pH 5.5 than either pH 5 or 6. Although the reason for this is unclear, this phenomenon was repeatedly observed.

The initial glucose concentration usually plays an important role in hydrogen production [12,13]. The effect of the initial glucose concentration on hydrogen production was examined during anaerobic cultivation at pH 7. Both the final cell concentration and the level of hydrogen production increased with increasing initial glucose concentration up to 25 g/l, as shown in Fig. 4. The final level of hydrogen production per unit working volume and the overall hydrogen productivity reached a maximum, when the initial glucose concentration was 25 g/l. This means that the optimal glucose concentration for hydrogen production is 25 g/l. The maximum and overall hydrogen productivity at an initial glucose concentration of 25 g/l was 398 and 174 ml/l/hr, respectively, as shown in Fig. 4. The final level of hydrogen production per unit working volume decreased when the initial glucose concentration was 50 g/l. In this case, the glucose was not completely consumed even when cells entered the stationary phase. This was attributed to the depletion of other nutrients before the glucose had been completely consumed. However, the residual glucose was still consumed during the stationary phase, and hydrogen production continued for up to 46 h (data not shown).

Table 2 Effect of pH on cell growth and hydrogen production pH

NCc 4.0 5.0 5.5 6.0 6.5 7.0 7.5

Maximum cell concentration (g/l)

1.15 0.69 1.14 1.29 1.41 1.67 1.75 1.07

Cell yield (g cell/g glucose)

0.115 0.093 0.097 0.125 0.131 0.144 0.169 0.107

H2 yielda (mol H2 /mol glucose)

0.26 0.08 0.32 0.41 0.31 0.30 0.54 0.06

H2 productivity (ml/l/hr)

H2 production/unit volume

Maximum

Overallb

(ml/l)

(mM)

58 63 129 129 96 126 175 24

34 37 66 76 55 61 94 11

325 74 396 515 384 364 665 74

14.5 3.3 17.7 23.0 17.2 16.2 29.7 3.3

Initial glucose concentration = 10 g/l. a H yield = (total cumulative H produced)/(total glucose consumed). 2 2 b Overall H productivity = (total cumulative H produced)/(total H production time)/(working volume). 2 2 2 c NC: not controlled.

J.-H. Shin et al. / International Journal of Hydrogen Energy 32 (2007) 192 – 199

2500

3.0

2000 1500 1000 500 0

200

160 2.5 120

2.0 Cell H2 productivity Residual glucose H2 production

1.5 1.0

80

40 0.5 0.0

0 0

10

20 30 40 Initial glucose concentration (g/l)

50

30 25 20 15 10 5

Residual glucose concentration (g/l)

3.5

Overall H2 productivity (ml/l/hr)

3000

Maximum cell concentration (g/l)

H2 production/unit volume (ml/l)

198

0

60

Fig. 4. Effect of the initial glucose concentration on cell growth and hydrogen production. The pH was controlled at 7. The overall H2 productivity = (total cumulative H2 produced)/(total H2 production time)/(working volume).

as soluble starch, and alcohols such as sorbitol and glycerol. However, cellulose was unsuitable as a carbon source for hydrogen production without a pretreatment.

1800 1600

H2 production/unit volume (ml/l)

1400

4. Conclusions

1200 1000 800 600 400 200 0 glucose fructose

sucrose

lactose cellulose

starch

sorbitol glycerol

Carbon source

Fig. 5. Hydrogen production of Enterobacter asburiae SNU-1 from various carbon sources. The concentration of all the carbon sources was 10 g/l. The cells were cultured in a 120 ml serum bottle (working volume 80 ml) at 37 ◦ C under anaerobic conditions without pH control.

3.5. Hydrogen production of Enterobacter asburiae SNU-1 from various carbon sources Fermentative bacteria use various carbon sources for hydrogen production. Therefore, the level of hydrogen production of Enterobacter asburiae SNU-1 from various carbon sources was examined. The concentration of each carbon source was fixed at 10 g/l. Fig. 5 shows that Enterobacter asburiae SNU1 can produce hydrogen from various carbon sources such as monosaccharides (glucose, fructose), disaccharides (sucrose, lactose), polysaccharides (starch), and alcohols (sorbitol, glycerol). However, the amount produced from lactose, starch, and glycerol was quite low. The highest level of hydrogen production was obtained from sucrose but a similar amount of hydrogen was produced from glucose and fructose. Enterobacter asburiae SNU-1 was also able to utilize polysaccharides such

A hydrogen-producing bacterial strain was isolated from a domestic landfill, and identified as a new species, Enterobacter asburiae SNU-1. The level of fermentative hydrogen production by this strain was influenced by the medium pH and the initial glucose concentration. The optimum initial glucose concentration and pH were 25 g/l and 7, respectively, which resulted in a respective maximum and overall hydrogen productivity of 398 and 174 ml/l/hr, respectively. A direct comparison of the hydrogen production potential between various strains is difficult because the level of hydrogen production is reported in different ways including the volumetric hydrogen production rate and the specific hydrogen production rate, etc. Recently, the highest maximum volumetric rate of hydrogen production per unit working volume from C. butyricum CGS5 was reported to be 209 ml H2 /l/h from 20 g COD sucrose/l (=17.8 g/l) [12]. In this study, Enterobacter asburiae SNU-1 had a higher hydrogen production potential than that reported for C. butyricum CGS5. This strain has unique properties in hydrogen production. Hydrogen production by microorganisms usually shows growth-associated behavior. However, this strain produced hydrogen in the stationary phase. Hydrogen production in the stationary phase is considered to be due to the result of decomposition of the formate that accumulated during the exponential phase. This formate is decomposed by the FHL complex, which is more active at high formate concentrations. This non-growth-associated hydrogen production is suitable for applications in fuel cells. Although formic acid can be used directly as a fuel source in fuel cells, the hydrogen-fueled fuel cell is more efficient. In this aspect, the harvested cells of this strain can be used as a bioreformer, which convert the formic

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