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Evaluation of the potential hydrogen production by diazotrophic Burkholderia species
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Evaluation of the potential hydrogen production by diazotrophic Burkholderia species Horacio Terrazas-Hoyos a, Enrique Portugal-Marı´n b, Enrique Sa´nchez-Salinas a, Ma. Laura Ortiz-Herna´ndez a,* a
Laboratorio de Investigaciones Ambientales, Centro de Investigacio´n en Biotecnologı´a, Universidad Auto´noma del Estado de Morelos, Av. Universidad 1001, Col. Chamilpa, C.P. 62210 Cuernavaca, Mor, Mexico b Laboratorio de Geotermia, Instituto de Investigaciones Ele´ctricas, Reforma 113, Col. Palmira, C.P. 62490 Cuernavaca, Mor, Mexico
article info
abstract
Article history:
Hydrogen (H2) is considered one of the most promising fuels for sustainable energy.
Received 14 September 2013
Because nitrogenase produces H2 as a normal by-product, we tested the N2-fixing bacterial
Received in revised form
strains Burkholderia unamae and Burkholderia tropica to determine their H2 production ca-
16 November 2013
pacities. To maximize H2 production, several culture conditions were tested and optimized,
Accepted 6 December 2013
including atmospheric conditions, carbon sources and chemical compounds such as
Available online 4 January 2014
enzyme cofactors and sugar cane molasses. The results showed that both strains were capable of H2 production. The culture medium with the highest H2 yield was composed of
Keywords:
1% v/v molasses enriched with Na2MoO4 (0.2 g/L), FeSO4 (0.2 g/L) and cysteine (0.02 g/L)
Biohydrogen
under a partial vacuum (air 20% v/v) without Ar final atmosphere. Under these conditions,
Diazotrophic
the highest H2 production rate obtained was 24.64 mmol H2/L for B. unamae. The present
Burkholderia
study contributes an optimization process for H2 production in N2-fixing Burkholderia
Molasses
species. We propose further research and development to improve H2 production rates in order to make biohydrogen a tangible reality. Copyright ª 2013, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved.
1.
Introduction
World energy is almost completely dependent on fossil fuels. However, the intensive use of these fuels generates severe negative impacts on the global environment, contributing to global climate change, environmental degradation, health problems, and the rapid depletion of fossil resources [1,2]. Hydrogen (H2) is considered one of the most promising fuels for sustainable energy, mainly because it has an energy yield of 122 kJ/g, which is 2.75 times greater than that of hydrocarbon fuels [1,2]. Its use has a minimal environmental
impact because its utilization via combustion or fuel cells produces pure water [3]. At present, most H2 produced globally is produced through steam methane reforming of natural gas, which is a non-renewable process [4]. Thus, the biological production of H2 (biohydrogen) is potentially regarded as one of the most promising alternatives and an exciting new area of technology development that offers the potential production of usable H2 from a variety of renewable sources [2,5,6]. Biological systems provide a wide range of approaches to for generating H2, including direct bio-photolysis by green algae, indirect bio-photolysis by cyanobacteria, photo-
* Corresponding author. Tel.: þ52 777 329 7057; fax: þ52 777 329 7030. E-mail address: [email protected] (Ma.L. Ortiz-Herna´ndez). 0360-3199/$ e see front matter Copyright ª 2013, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.ijhydene.2013.12.049
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fermentation by anaerobic photosynthetic bacteria and dark fermentation by anaerobic fermentative bacteria [1,7]. All biological processes for H2 generation depend on enzymes such as hydrogenase and nitrogenase [8]. Although dark fermentation and photo-fermentation are considered the most promising processes with the highest yields reported [1,3,9], no biological process for H2 production has been found to be completely viable for various reasons. The profitability of solar-powered biohydrogen production processes is dependent on the light to H2 conversion efficiency. In fermentative and anoxygenic processes, the limiting step in terms of efficiency is the conversion of light to sugars [10]. Due to the above, in this report, we explore the manipulation of the biological nitrogen fixation (BNF) reaction as an alternative approach for H2 generation. BNF is performed by a variety of microorganisms and is a key process in which dinitrogen is reduced to form ammonia, a molecule used by living systems for the synthesis of many organic compounds [11]. N2 is an essential element for all living organisms. Although N2 gas is abundant in the atmosphere, it cannot be readily used by most organisms. The nitrogenase complex is the enzyme responsible for BNF in all nitrogen (N2)-fixing prokaryotes [8]. Nitrogenase is a metalloprotein containing iron/sulfur (Fe/S) clusters. It is composed of both an Fe-binding subunit and a molybdenumiron (MoeFe)-binding subunit to which the Fe subunit transfers electrons that are used to reduce substrates [11,12]. The Fe protein, also called component II or dinitrogenase reductase, transfers one electron at a time from a [4Fe4S] cluster to the MoeFe protein, otherwise termed ‘component I’ or ‘dinitrogenase’ [11]. Cycles of association and dissociation between these two proteins, coupled to the hydrolysis of MgATP, drive this enzymatic reaction forward [8]. Nitrogenase catalyzes the reduction of dinitrogen to ammonia, using electrons from low-potential reductants (ferredoxina or flavodoxin), according to the following reaction [11e13]:
N2 þ 8Hþ þ 8e þ 16ATP / 2NH3 þ H2 þ 16ADP þ 16Pi As seen in the reaction above, H2 is an obligate by-product of the nitrogenase reaction, reducing protons (Hþ) to H2 [14]. The key point of this study lies in the fact that this reaction is susceptible to manipulation, given that in the absence of N2, the total electron flux is directed to protons, producing only H2. Thus, H2 production by nitrogenase is not dependent upon the reduction of N2; in an argon (Ar) atmosphere, nitrogenase produces only H2 [15]. The reaction is irreversible and can provide continuous production of H2 even under an atmosphere of 100% H2 [13] according to the following reaction:
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H2 is evolved per N2 reduced, H2/N2 is more than 1.0 even under 50 atm of N2 [16]. Hence, another major unresolved problem is the ratio of H2 evolved per N2 reduced [12]. For all of these reasons, we believe that the nitrogenase BNF reaction should be further studied and manipulated in order to determine its maximal potential for H2 generation. To that end, in the present study, we tested N2-fixing bacteria from Burkholderia genus in different culture conditions to evaluate their H2 production capacity [17,18]. More than 60 bacterial species presently classified as Burkholderia are widely distributed in the natural environment [19]. In recent years, this genus has emerged as an important plant-associated bacterial species [20,21]. Until recently, among Burkholderia species, only Burkholderia vietnamiensis was recognized as a N2-fixing organism [22,19]. In 2002, novel N2-fixing Burkholderia species, including Burkholderia unamae and Burkholderia tropica, were isolated from the rhizosphere and inner tissues of field-grown sugar cane varieties [23,24]. Further studies have revealed that these diazotrophic species exhibit some plant growth-promoting activities, as well as potential for biocontrol and bioremediation processes [25]. This study aimed to determine whether the N2-fixing Burkholderia species B. unamae and B. tropica (which are phylogenetically distant from the pathogenic Burkholderia cepacia complex) are capable of producing H2 in sufficient amounts to consider using them a viable process. Several culture conditions were tested and optimized, including atmospheric conditions and the influence of different carbon sources and chemical compounds such as enzyme cofactors and sugar cane molasses in order to improve the H2 yield [26]. To make biological H2 production an economically viable process, the substrates used need to be abundant, low-cost resources that require minimal pretreatment [3]. Pure carbohydrate sources are expensive raw materials for real scale H2 production, which can only be viable when based on renewable and low-cost sources [27]. Thus, the emphasis has shifted from laboratory based pure substrates to cheap and easily available natural substrate and waste materials [9] such agroindustrial waste that could contain substrates and optimal nutrients for biohydrogen production. In the last few years, there has been an increasing interest in working with sugar cane molasses due to the possibility of using it as a costefficient carbon and energy source to cultivate microorganisms [28,29]. Sugar cane molasses is a by-product of the sugar cane industry. It is a viscous liquid rich in noncrystallized carbohydrates (sucrose, glucose and fructose). Currently this by-product is mainly used for animal feed, fermentation to ethanol and yeast production [30].
2.
Materials and methods
2.1.
N2-fixing Burkholderia strains used
2Hþ þ 2e þ 4ATP / H2 þ 4ADP Although it is well known that FeMo-co is the active center of nitrogenase [11,12], the exact sites for N2 and Hþ binding and reduction, the physiological substrates of nitrogenase, are still unknown. Additionally, the overall stoichiometric equation is still ambiguous. Although in the accepted equation one
Two Burkholderia strains were used in this study, B. unamae and B. tropica. The strains were previously isolated by Castro et al. [21] from two sugar cane crops in Mexico (in the states of Oaxaca and Morelos). Previous analyses of the N2-fixing abilities of these strains were based on acetylene reduction activity (ARA) and the presence of nifH genes [21].
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2.2.
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Initial culture medium
The previously described N2-free semisolid BAz medium (composition in g/L: azelaic acid, 5.0; K2HPO4, 0.4; KH2PO4, 0.4; MgSO4$7H2O, 0.2; CaCl2, 0.02; Na2MoO4$H2O, 0.002; FeCl3, 0.01; and agar, 2.3) was used as an enrichment culture for N2-fixing Burkholderia species. Additionally, a BAc medium (in wt/v; 0.2% azelaic acid, 0.02% L-citrulline, 0.04% K2HPO4, 0.04% KH2PO4, and 0.02%, MgSO4$7H2O) was used as a selective medium to maintain the viability of pure cultures [20].
2.3.
pH optimization for cell growth
In order to determine the optimal pH values for growth of the diazotrophic strains tested, cell growth was tested at pH 5.0, 5.5, 6.0, 6.5, 7.0 and 7.5 based on the original soil pH where the bacterial strains were isolated. pH was adjusted using potassium hydroxide (KOH), and the N2-free BAz medium was used. Bacterial growth was determined by measuring the OD610 every 12 h over a total incubation period of 96 h.
2.4.
H2 measurement
A gas chromatograph equipped with a thermal conductivity detector (GC-TCD) (Agilent Technologies 7890A) and a megabore column (0.53 mm 30 m) was used for H2 detection in cultivated samples. After initial screening experiments, analytical conditions for the GC-TCD were determined as follows: temperatures of the injection port, oven and detector were 150 C, 40 C and 150 C, respectively. N2 was used as carrier gas with a flow rate of 3 mL/min. Calibration curves were constructed with a standard gas mixture (PRAXAIR: 30% N2; 30% CH4; 25% H2; 5% He; 5% Ar and 5% CO mol/mol) before the injection of the microbiological samples. The mathematical method for the H2 yield calculation was implemented according to the proposal by Juantorena et al. [31,32]. Statistical analysis consisted of analysis of
variance (ANOVA) and Tukey’s test (a ¼ 0.05) using the statistical package SAS 9.1 [33] as reported by Burrows et al. [4].
2.5. Bioreactors used for cell cultivation and H2 measurement assays Gas sampling flaks (Fig. 1a) with a total volume of 310 mL equipped with Teflon valves [34] were used as bioreactors throughout all H2 measurement assays, providing microenvironments isolated from the external environment. The bioreactors used had the following characteristics: fully sealed, resistant to sudden temperature changes and easy to use, handle and transport. The bioreactors had a connector valve of 9 mm in thickness for proper coupling to the injection system for subsequent gas quantification. These bioreactors attached to the GC-TCD for gas detection are shown in Fig. 1b.
2.6.
We subdivided this step based on the culture condition(s) evaluated. Each stage is comprised of various H2 measurement assays described below. Based on previous results (data not shown), the incubation times of all H2 measurement assays were divided into two different phases [31]. The first phase (aerobic phase) occurred prior to atmospheric manipulation, exclusively promoting biomass production, and was set at two days. The second phase (microaerobic phase) refers to the incubation of the strains after the atmospheric manipulation (vacuum and Ar injection) and was set at three days. Bioreactors containing 200 mL of the appropriate medium were inoculated at an OD610 of 0.5 with an aliquot of 2 mL of a 24 h culture of each selected strain [35] in BAz medium. Each assay was carried out in triplicate. Based on previous results, all H2 measurement assays were executed at a pH of 6.0 and a temperature of 35 C. It is worth mentioning that during the first set of results, a bacterial consortium consisting of a mixed culture of the two Burkholderia species was evaluated on its H2 production capacity along with the same strains tested individually. However, contrary to the expected, the H2 production rates did not show a significant increase (data not shown).
2.6.1.
Fig. 1 e Gas sampling flasks used as bioreactors: (a) unattached bioreactor and its length, (b) bioreactor attached to the GC-TCD for gas detection.
Optimization of culture conditions for H2 production
Atmospheric conditions
Oxygen (O2) concentration plays a crucial role in obligate and facultative aerobic N2-fixing bacteria like Burkholderia sp. because they have to tolerate a phenomena previously described as the “O2 paradox”. In these organisms, a minimal O2 concentration is necessary to support aerobic respiration and ATP synthesis. However, when O2 is present at a concentration that is too high, the nitrogenase can be irreversibly damaged [36,37]. Therefore, it was necessary to work at microaerophilic conditions. After each bioreactor was inoculated, a vacuum was generated for a total time of 4 min, with the culture stirred for 3 s every minute to eliminate the O2. Based on the results (data not shown) of testing a wide range of different argon (Ar) concentrations (25, 50, 60, 70, 80 and 90% (v/v)), Ar flux was set to 3 0.1 mL/s at different times [31,32]. The highest H2 quantifiable detections were obtained under an 80% Ar
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atmosphere. Based on these results, an atmospheric condition of 80% v/v Ar was used for further testing.
2.6.2.
Carbon sources
The carbon sources selected and their concentration were based on previous reports in the literature using the same diazotrophic species [20,24]. Pure carbon sources tested were as follows: succinic acid, mannitol, fructose, sucrose and glucose. The culture medium was composed of the mineral salts present in the BAz medium, substituting the azelaic acid with the individual carbon source to be tested at a concentration of 0.5%. Based on prior results, an 80% v/v Ar atmosphere was used. Following the first set of results, on a separate experiment, the two carbon sources with the highest H2 yield (sucrose and succinic acid) were combined at 0.5% wt/ v each (final carbon source-concentration of 1.0% wt/v) and supplemented to the mineral salts medium (BAz) and tested for H2 production.
2.6.3.
Chemical compounds as enzyme cofactors
Because nitrogenase is a metalloprotein, different chemical compounds were tested [38] to evaluate their influence as stimulators of nitrogenase activity and subsequent H2 production [39]. The following chemical compounds were selected: ferrous sulfate (FeSO4), 0.2 g/L; vanadium oxide (V2O5), 0.02 g/L and the amino acid cysteine, 0.02 g/L. The original Na2MoO4 concentration in BAz was also modified from 0.002 g/L to 0.2 g/L. The carbon sources in the medium were sucrose and succinic acid. Based on the first set of results, the chemical compounds with the highest H2 yield were combined and tested.
2.6.4.
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production, including the cost of using Ar. Given that Ar is used to promote anaerobic conditions, it was inferred that there should be no significant difference between using it and making a vacuum for the final anaerobic atmospheric concentration.
3.
Results
3.1.
pH optimization for cell growth
The results show that the optimal pH for cell growth for both B. unamae and B. tropica is close to 6.0 (Fig. 2), where highest absorbance values were obtained. In addition, for both strains, the lowest absorbance values were obtained at a pH of 5.0. Based on these results, all further H2 measurement assays were executed at a pH of 6.0.
3.2.
H2 yield with different carbon sources
After evaluating the five different carbon sources separately, the results obtained (Fig. 3) indicate that the individual carbon source with the highest H2 production was sucrose, with rates of 0.34 mmol H2/L for B. unamae and 0.49 mmol H2/L for B. tropica. After sucrose, succinic acid showed the highest H2 production rates, with 0.17 mmol H2/L for B. unamae and 0.12 mmol H2/L for B. tropica. The carbon source with the lowest H2 yield was mannitol.
Molasses as a culture medium
Sugar cane molasses from a local factory located in Zacatepec, Morelos, Mexico, was used. In laboratory conditions, we determined its moisture percentage at 46% and pH at 6.4. The composition of the original sugar cane molasses was not determined. It is well known that the composition highly varies depending on cane varieties, soil composition and climate. However, different studies in Mexico using sugar cane molasses show an average carbon source composition of: sucrose, w68%; glucose, w13% and fructose, w18% [30]. In this report, we evaluated three different treatments using molasses as a culture medium at a constant concentration of 1% v/v (based on previous tests where different concentrations were evaluated; data not shown). The first treatment consisted of using only molasses as the culture medium. For the second treatment, the pure molasses medium was supplemented with Na2MoO4 (0.2 g/L), FeSO4 (0.2 g/ L) and cysteine (0.02 g/L) to stimulate nitrogenase activity. The third treatment composition was identical to the second with the addition of the original mineral salts in BAz.
2.6.5.
Variation of atmospheric condition: Ar suppression
This final stage consisted of testing the two H2 measurement assays with the highest H2 yield under a different atmospheric condition, eliminating the Ar injection and under a partial vacuum (air 20% v/v). This variation was assessed due to the economic aspects of maintaining a cost-effective process for biohydrogen
Fig. 2 e Growth of bacterial strains tested at different pH values. a) B. unamae, b) B. tropica.
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tested, three (Na2MoO4, FeSO4 and cysteine) resulted in an increase in H2 production rates compared to previous assays. The Na2MoO4 (0.2 g/L) showed the highest increase in H2 production, with a rate of 4.02 mmol H2/L for B. unamae and 3.56 mmol H2/L for B. tropica. Treatment with V2O5 showed no significant increase in H2 production rates in any of the two strains tested. However, the combination of the three chemical compounds (Na2MoO4, FeSO4 and cysteine) exhibited a surprisingly high H2 production rate, significantly superior to all previous culture conditions tested. For B. unamae, H2 production was 17.61 mmol H2/L and for B. tropica, 12.35 mmol of H2/L.
Fig. 3 e H2 production rates of B. unamae and B. tropica under the influence of different carbon sources. Error bars indicate standard errors from the mean for at least three independent experiments.
Furthermore, the combination of sucrose and succinic acid (individual carbon sources with the highest H2 yield), exhibited significantly higher H2 production rates of 1.22 mmol H2/L for B. unamae and 1.03 mmol H2/L for B. tropica. This significant increase is partly due to the final carbon source-percentage increase (from 0.5% when tested individually to 1.0% wt/v when combined), although the final H2 production rates exceeded the rates expected when combining the two carbon sources.
3.3. H2 yield with the addition of different chemical compounds as enzyme cofactors The results obtained at this stage represent a critical point for the purposes of this study, revealing the positive impact of most of the chemical compounds tested. The results (Fig. 4) indicate that out of the four different chemical compounds
3.4.
H2 yield with molasses as a culture medium
After evaluating the influence of the three different treatments using molasses, the results obtained (Fig. 4) indicate that when pure molasses was tested as a culture medium, H2 production rates were below the previous assays in which additional carbon sources and chemical compounds as enzyme cofactors were supplemented. Nevertheless, when Na2MoO4, FeSO4 and cysteine were added to the molasses medium, the H2 production yield reached the highest rates obtained in the present study, with 24.64 mmol H2/L for B. unamae and 17.55 mmol H2/L for B. tropica. Finally, when mineral salts were added to the molasses medium with Na2MoO4, FeSO4 and cysteine, H2 production did not show a significant difference, indicating that the addition of mineral salts was ineffective.
3.5.
Influence of Ar suppression on H2 production
Comparing the two treatments with the highest H2 production rates (molasses medium enriched with Na2MoO4, FeSO4 and cysteine and the mineral salt medium supplemented with succinic acid, sucrose and Na2MoO4, FeSO4 and cysteine), a higher H2 production rate was obtained when Ar was used in
Fig. 4 e Summary of the H2 production rates obtained under the influence of chemical compounds as enzyme cofactors and molasses treatments by B. unamae and B. tropica. Different letters indicate statistically significant differences (P £ 0.0001, a [ 0.05). Vertical bars correspond to standard errors from the mean for at least three independent experiments. CV [ 44.77%.
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both treatments (Fig. 5). However, for the treatment with the highest H2 yield (molasses with Na2MoO4, FeSO4 and cysteine), statistical analysis showed no significant difference between the two atmospheric conditions. Use of Ar does not appear to be strictly necessary for the molasses medium at least. In Table 1, we present an overview of the different treatments evaluated, indicating the culture medium and the atmospheric composition along with the H2 yield obtained in each H2 measurement assay. In Fig. 6, we present two representative chromatograms obtained by GC-TCD, corresponding to two different H2 measurement assays. In those chromatograms, the increase in H2 throughout the experimental procedure is clearly shown by the size of the H2 peaks obtained.
4.
Discussion
This report represents a comprehensive study of H2 production using the nitrogenase complex of diazotrophic bacteria and it presents the utilization of Burkholderia species (B. unamae and B. tropica) to produce H2, which has not yet been explored in the literature, presenting a different approach in biological H2 production. The manipulation of the different culture conditions tested allowed a better understanding of the metabolic process and optimal conditions for H2 production by the evaluated N2-fixing species. However, this report offers a brief outlook toward the BNF system to produce H2 but the feasibility of the process remains uncertain. When atmospheric conditions were tested, it was clear that a critical issue for the cell was the protection of the nitrogenase complex from O2 inactivation. Incubation of cells under a low O2 environment provided the highest rates of H2 production, which is corroborated by Min and Sherman [35]. In this case, for B. unamae and B. tropica, a final atmosphere of Ar 20% v/v was optimal for H2 production. Cells obtained the necessary O2 concentration for respiration but it was not high enough to trigger nitrogenase inactivation. The cellular
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mechanism for O2 protection in the N2-fixing Burkholderia species remains unclear and should be further studied. Regarding the influence of different carbon sources, the predilection of the Burkholderia strains towards sucrose as a carbon source may be related to their origin (sugarcane crops), where sucrose is the predominant carbohydrate [20]. We cannot explain why succinic acid was the single carbon source with the second highest H2 production rate, but it may be related to the metabolic role of succinic acid in the citric acid cycle and energy-production process [40]. The predilection for sucrose was also consistent when molasses was tested as the primary carbon source, assuming that the main component of molasses is sucrose (w35% wt/v) [28e30]. However, the H2 production rates in both species were significantly higher when molasses was used. This not only has to do with the difference in sucrose concentration in both media, but with the rich composition of molasses, which includes a variety of carbohydrates (sucrose, glucose and fructose) and different molecules including minerals, amino acids and vitamins necessary for cell growth [30]. Although there are several studies referring to the utilization of molasses as a culture medium for H2 production [28,29], little is known about its use in N2-fixing bacteria. Although the composition of molasses is highly heterogeneous, depending on environmental factors and its production process, it represents a potential element for the culture of a wide range of microorganisms and should be further assessed, possibly testing different specific molecules as supplements for the medium enrichment. The influence of the chemical compounds tested as stimulators of nitrogenase activity represents a major issue in the results of this study. Taking into account that the metalloprotein nitrogenase contains and needs enzyme cofactors to carry out its metabolic functions, the addition of the chemical compounds serving as enzyme cofactors to the different culture medium tested resulted in an exponential increase of the H2 production rates. The only metal ion tested that did not
Fig. 5 e Comparison of the H2 production rates by B. unamae and B. tropica using the two culture media with the highest H2 yield yet obtained, under two different atmospheric conditions, in the presence of Ar (vacuum D 80% Ar v/v) and without it (partial vacuum). Different letters indicate statistically significant differences (P £ 0.0001, a [ 0.05). Vertical bars correspond to standard errors from the mean for at least three independent experiments. CV [ 24.94%.
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Table 1 e Summary of the different culture conditions evaluated (culture medium and atmospheric condition) in this study and the H2 yield obtained in each case for both strains. Primary culture medium composition
Enzyme cofactors
Atmospheric condition (v/v)
Molasses Molasses
Na2MoO4 þ FeSO4 þ cysteine Na2MoO4 þ FeSO4 þ cysteine
Molasses þ Mineral salts Mineral salts þ succinic acid þ sucrose Mineral salts þ succinic acid þ sucrose
Na2MoO4 þ FeSO4 þ cysteine Na2MoO4 þ FeSO4 þ cysteine Na2MoO4 þ FeSO4 þ cysteine
Mineral salts þ Mineral salts þ Mineral salts þ Molasses Mineral salts þ Mineral salts þ
Na2MoO4 Cysteine FeSO4
Ar 80% Partial vacuum (air 20%) without Ar Ar 80% Ar 80% Partial vacuum (air 20%) without Ar Ar 80% Ar 80% Ar 80% Ar 80% Ar 80% Ar 80%
succinic acid þ sucrose succinic acid þ sucrose succinic acid þ sucrose succinic acid þ sucrose succinic acid þ sucrose
e e V2O5
produce a H2 increase was vanadium oxide (V2O5). In fact, both species exhibited a considerable decrease in the H2 yield when V2O5 was added to the primary culture medium composition (mineral salts þ sucrose þ succinic acid) without any enzyme
H2 production (mmol/L) B. unamae
B. tropica
24.64 3.36 23.81 3.69
17.16 8.91 14.98 3.28
23.23 1.51 17.61 2.36 10.18 1.31
14.89 7.50 12.35 6.25 7.62 1.67
4.02 3.69 2.76 2.00 1.92 1.40
0.89 1.62 1.24 0.46 0.58 0.45
3.56 2.81 2.19 1.45 1.33 1.25
1.98 2.17 1.82 0.53 0.93 0.79
cofactor (Table 1). This may be due to the high oxidation state of V2O5, acting both as an amphoteric oxide and an oxidizing agent. It may also be related to its reported capacity to produce by-products that can inhibit enzymes that process phosphate
Fig. 6 e Examples of chromatograms obtained by the GC-TCD with two different culture condition treatments. a) Mineral salts D succinic acid D sucrose, b) Molasses D FeSO4 D cysteine D Na2MoO4. In both cases, the atmospheric condition consisted in vacuum D 80% Ar. H2 peaks are observed at a retention time of w1.5 min, while the other peak represents N2.
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(PO3 4 ) [41,42]. In any case, the results allow us to discard the potential presence of alternative vanadium (V)-nitrogenase [8] in the Burkholderia strains tested. Comparison of the H2 production rates by the Burkholderia species tested versus others obtained by biological processes such as dark fermentation or indirect bio-photolysis [1] provides a useful benchmark for future improvements. The peak H2 production rates obtained are still lower than those obtained in those systems [1,3,9,10]; nevertheless, this study represents the first data using Burkholderia N2-fixing bacteria as H2 producers and opens a range of possibilities to evaluate different diazotrophic species in the future. Moreover, alternatives and variants for the optimization of culture conditions remain unexplored. Currently, among the known biological processes for H2 generation, dark fermentative H2 production is considered the most attractive, with the highest yields reported [1,9,10]. However, fermentative H2 production processes, either conducted via mixed or pure cultures, still have certain limitations that could be crucial for scaling up. Inefficient substrate conversion, formation of by-products, low resistance to high H2 partial pressures and oxygen are only some of the limitations. Above all those, comes the limiting upper value of H2 yields which is 4 mol of H2/mol hexose consumed [27]. Besides this, the gas produced in these processes is a mixture of primarily H2 and carbon dioxide (CO2), but may also contain other gases such as methane (CH4), hydrogen sulfide (H2S), or ammonia (NH4). For the scaling up of such a process, the raw material cost is considered to be among the major limitations. This particular limitation is minimized in the present approach, considering that the optimal culture medium tested for H2 production (molasses medium) is an abundant and relatively inexpensive agro-industrial waste product (cost lower than 0.5 US dollars per L) [30] and with the elimination of Ar, the production process tested is economically promising in relation to other known processes such as indirect biophotolysis and fermentation [3,7]. On the other hand, the main limitation of this nitrogenase BNF reaction approach is that this process is thermodynamically uphill and requires ATP hydrolysis. Two molecules of ATP have to be hydrolyzed for every electron made available for H2 production, making the reaction highly ATP-dependent [10]. If biohydrogen is to be produced by whole microorganisms, it must be taken into account that H2 output represents an energy loss for the cell. Regulatory networks control the biosynthesis and activity of the involved enzymes; therefore, it is necessary to understand how they function before being able to maximize the production of H2. Also, in some legume nodules, the bacterial symbiont produces an uptake hydrogenase (HUP) enzyme that oxidizes the H2 evolved, recovering a portion of the energy used in its production. However, many of the most productive N2-fixing associations lack HUP activity and the H2 diffuses into the soil where it is consumed [43]. In response to some of the mentioned limitations above, several researchers have proposed the use of genetically modified microorganisms with advanced selected properties, constructed either via mutagenesis or through genetic engineering. H2 producing bacteria can be genetically modified in several ways to increase H2 synthesis, including (1) over-
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expression of certain enzymes that can maximize substrate availability; (2) elimination of uptake hydrogenases; (3) overexpression of H2 evolving hydrogenases that have been modified to be O2 tolerant; and (4) elimination of metabolic pathways that compete for reducing equivalents required for H2 synthesis [1,27]. As mentioned before, further research and development is needed to improve H2 production rates through this novel approach. Several experimental parameters should be optimized to manipulate the nitrogenase BNF reaction to get a higher H2 yield in these two Burkholderia species. These parameters include factors which affect the distribution of electrons between H2 production and N2 reduction within nitrogenase, such as, temperature, pH, the ratio of the two component proteins of nitrogenase and the ATP concentration [44]. Those parameters do not only determine the growth of the microorganisms, but also have a crucial role on the metabolic path that the microorganisms will follow, severely affecting the final H2 yield observed. Other factors to examine further include cell concentration, combinations of species, and utilization of different organic waste products. Furthermore, future experiments will involve a careful simultaneous analysis of both H2 production and cell growth over extended periods of time to determine the best conditions for each specific application. Although it is well known that one reason for the tremendous H2 production potential of nitrogenase is that there is a vast amount of enzyme present in the cell [37], both processes need to be experimentally differentiated and independently valued.
5.
Conclusions
The hydrogen-production capacity of the diazotrophic bacteria B. unamae and B. tropica was confirmed, with B. unamae exhibiting the highest production rates in most of the conditions tested. Although the peak H2 production rates obtained are lower than the rates reported for other biological systems, the present study is innovative as it represents the first contribution of an optimization process for H2 production by N2-fixing Burkholderia species. Utilization of the chemical compounds Na2MoO4, FeSO4 and cysteine as stimulators of nitrogenase activity has been demonstrated, increasing the H2 production rates where supplemented. Moreover, using molasses as a carbohydraterich waste is a promising approach to developing a costeffective system for biohydrogen production. Finally, it is widely believed that H2 will play a crucial role in the future energy economy, but considerable research and developmental studies are needed to improve H2 production rates and to reduce production costs in order to make biohydrogen a tangible reality that is capable of meeting the increasing need for a substitute for fossil fuels.
Acknowledgments The authors are thankful to Dra. Rocı´o Castro Gonza´lez for providing the bacterial strains used and to Dra. Alina
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Juantorena Uga´s for general support. We also acknowledge the technical assistance of Tech. Vicente Betancourt and Tech. Jose´ Antonio Nopaltitla. Horacio is thankful to the Consejo Nacional de Ciencia y Tecnologı´a (CONACYT) for the graduate fellowship.
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[39] Salih F, Maleek M. Influence of metal ions on hydrogen production by photosynthetic bacteria grown in Escherichia coli pre-fermented cheese whey. J Environ Prot 2010;1:426e30. [40] Zeikus JG, Jain MK, Elankovan P. Biotechnology of succinic acid production and markets for derived industrial products. Appl Microbiol Biotechnol 1999;51:545e52. [41] Bauer G, Gu¨ther V, Hess H, Otto A, Roidl O, Roller H, et al. Vanadium and vanadium compounds. In: Ullmann’s encyclopedia of industrial chemistry, Electronic Release. 6th rev. ed., vol. 38. Weinheim: Wiley-VCH; 2003. pp. 1e21.
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Available online at www.sciencedirect.com
ScienceDirect journal homepage: www.elsevier.com/locate/he
Evaluation of the potential hydrogen production by diazotrophic Burkholderia species Horacio Terrazas-Hoyos a, Enrique Portugal-Marı´n b, Enrique Sa´nchez-Salinas a, Ma. Laura Ortiz-Herna´ndez a,* a
Laboratorio de Investigaciones Ambientales, Centro de Investigacio´n en Biotecnologı´a, Universidad Auto´noma del Estado de Morelos, Av. Universidad 1001, Col. Chamilpa, C.P. 62210 Cuernavaca, Mor, Mexico b Laboratorio de Geotermia, Instituto de Investigaciones Ele´ctricas, Reforma 113, Col. Palmira, C.P. 62490 Cuernavaca, Mor, Mexico
article info
abstract
Article history:
Hydrogen (H2) is considered one of the most promising fuels for sustainable energy.
Received 14 September 2013
Because nitrogenase produces H2 as a normal by-product, we tested the N2-fixing bacterial
Received in revised form
strains Burkholderia unamae and Burkholderia tropica to determine their H2 production ca-
16 November 2013
pacities. To maximize H2 production, several culture conditions were tested and optimized,
Accepted 6 December 2013
including atmospheric conditions, carbon sources and chemical compounds such as
Available online 4 January 2014
enzyme cofactors and sugar cane molasses. The results showed that both strains were capable of H2 production. The culture medium with the highest H2 yield was composed of
Keywords:
1% v/v molasses enriched with Na2MoO4 (0.2 g/L), FeSO4 (0.2 g/L) and cysteine (0.02 g/L)
Biohydrogen
under a partial vacuum (air 20% v/v) without Ar final atmosphere. Under these conditions,
Diazotrophic
the highest H2 production rate obtained was 24.64 mmol H2/L for B. unamae. The present
Burkholderia
study contributes an optimization process for H2 production in N2-fixing Burkholderia
Molasses
species. We propose further research and development to improve H2 production rates in order to make biohydrogen a tangible reality. Copyright ª 2013, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved.
1.
Introduction
World energy is almost completely dependent on fossil fuels. However, the intensive use of these fuels generates severe negative impacts on the global environment, contributing to global climate change, environmental degradation, health problems, and the rapid depletion of fossil resources [1,2]. Hydrogen (H2) is considered one of the most promising fuels for sustainable energy, mainly because it has an energy yield of 122 kJ/g, which is 2.75 times greater than that of hydrocarbon fuels [1,2]. Its use has a minimal environmental
impact because its utilization via combustion or fuel cells produces pure water [3]. At present, most H2 produced globally is produced through steam methane reforming of natural gas, which is a non-renewable process [4]. Thus, the biological production of H2 (biohydrogen) is potentially regarded as one of the most promising alternatives and an exciting new area of technology development that offers the potential production of usable H2 from a variety of renewable sources [2,5,6]. Biological systems provide a wide range of approaches to for generating H2, including direct bio-photolysis by green algae, indirect bio-photolysis by cyanobacteria, photo-
* Corresponding author. Tel.: þ52 777 329 7057; fax: þ52 777 329 7030. E-mail address: [email protected] (Ma.L. Ortiz-Herna´ndez). 0360-3199/$ e see front matter Copyright ª 2013, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.ijhydene.2013.12.049
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fermentation by anaerobic photosynthetic bacteria and dark fermentation by anaerobic fermentative bacteria [1,7]. All biological processes for H2 generation depend on enzymes such as hydrogenase and nitrogenase [8]. Although dark fermentation and photo-fermentation are considered the most promising processes with the highest yields reported [1,3,9], no biological process for H2 production has been found to be completely viable for various reasons. The profitability of solar-powered biohydrogen production processes is dependent on the light to H2 conversion efficiency. In fermentative and anoxygenic processes, the limiting step in terms of efficiency is the conversion of light to sugars [10]. Due to the above, in this report, we explore the manipulation of the biological nitrogen fixation (BNF) reaction as an alternative approach for H2 generation. BNF is performed by a variety of microorganisms and is a key process in which dinitrogen is reduced to form ammonia, a molecule used by living systems for the synthesis of many organic compounds [11]. N2 is an essential element for all living organisms. Although N2 gas is abundant in the atmosphere, it cannot be readily used by most organisms. The nitrogenase complex is the enzyme responsible for BNF in all nitrogen (N2)-fixing prokaryotes [8]. Nitrogenase is a metalloprotein containing iron/sulfur (Fe/S) clusters. It is composed of both an Fe-binding subunit and a molybdenumiron (MoeFe)-binding subunit to which the Fe subunit transfers electrons that are used to reduce substrates [11,12]. The Fe protein, also called component II or dinitrogenase reductase, transfers one electron at a time from a [4Fe4S] cluster to the MoeFe protein, otherwise termed ‘component I’ or ‘dinitrogenase’ [11]. Cycles of association and dissociation between these two proteins, coupled to the hydrolysis of MgATP, drive this enzymatic reaction forward [8]. Nitrogenase catalyzes the reduction of dinitrogen to ammonia, using electrons from low-potential reductants (ferredoxina or flavodoxin), according to the following reaction [11e13]:
N2 þ 8Hþ þ 8e þ 16ATP / 2NH3 þ H2 þ 16ADP þ 16Pi As seen in the reaction above, H2 is an obligate by-product of the nitrogenase reaction, reducing protons (Hþ) to H2 [14]. The key point of this study lies in the fact that this reaction is susceptible to manipulation, given that in the absence of N2, the total electron flux is directed to protons, producing only H2. Thus, H2 production by nitrogenase is not dependent upon the reduction of N2; in an argon (Ar) atmosphere, nitrogenase produces only H2 [15]. The reaction is irreversible and can provide continuous production of H2 even under an atmosphere of 100% H2 [13] according to the following reaction:
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H2 is evolved per N2 reduced, H2/N2 is more than 1.0 even under 50 atm of N2 [16]. Hence, another major unresolved problem is the ratio of H2 evolved per N2 reduced [12]. For all of these reasons, we believe that the nitrogenase BNF reaction should be further studied and manipulated in order to determine its maximal potential for H2 generation. To that end, in the present study, we tested N2-fixing bacteria from Burkholderia genus in different culture conditions to evaluate their H2 production capacity [17,18]. More than 60 bacterial species presently classified as Burkholderia are widely distributed in the natural environment [19]. In recent years, this genus has emerged as an important plant-associated bacterial species [20,21]. Until recently, among Burkholderia species, only Burkholderia vietnamiensis was recognized as a N2-fixing organism [22,19]. In 2002, novel N2-fixing Burkholderia species, including Burkholderia unamae and Burkholderia tropica, were isolated from the rhizosphere and inner tissues of field-grown sugar cane varieties [23,24]. Further studies have revealed that these diazotrophic species exhibit some plant growth-promoting activities, as well as potential for biocontrol and bioremediation processes [25]. This study aimed to determine whether the N2-fixing Burkholderia species B. unamae and B. tropica (which are phylogenetically distant from the pathogenic Burkholderia cepacia complex) are capable of producing H2 in sufficient amounts to consider using them a viable process. Several culture conditions were tested and optimized, including atmospheric conditions and the influence of different carbon sources and chemical compounds such as enzyme cofactors and sugar cane molasses in order to improve the H2 yield [26]. To make biological H2 production an economically viable process, the substrates used need to be abundant, low-cost resources that require minimal pretreatment [3]. Pure carbohydrate sources are expensive raw materials for real scale H2 production, which can only be viable when based on renewable and low-cost sources [27]. Thus, the emphasis has shifted from laboratory based pure substrates to cheap and easily available natural substrate and waste materials [9] such agroindustrial waste that could contain substrates and optimal nutrients for biohydrogen production. In the last few years, there has been an increasing interest in working with sugar cane molasses due to the possibility of using it as a costefficient carbon and energy source to cultivate microorganisms [28,29]. Sugar cane molasses is a by-product of the sugar cane industry. It is a viscous liquid rich in noncrystallized carbohydrates (sucrose, glucose and fructose). Currently this by-product is mainly used for animal feed, fermentation to ethanol and yeast production [30].
2.
Materials and methods
2.1.
N2-fixing Burkholderia strains used
2Hþ þ 2e þ 4ATP / H2 þ 4ADP Although it is well known that FeMo-co is the active center of nitrogenase [11,12], the exact sites for N2 and Hþ binding and reduction, the physiological substrates of nitrogenase, are still unknown. Additionally, the overall stoichiometric equation is still ambiguous. Although in the accepted equation one
Two Burkholderia strains were used in this study, B. unamae and B. tropica. The strains were previously isolated by Castro et al. [21] from two sugar cane crops in Mexico (in the states of Oaxaca and Morelos). Previous analyses of the N2-fixing abilities of these strains were based on acetylene reduction activity (ARA) and the presence of nifH genes [21].
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2.2.
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Initial culture medium
The previously described N2-free semisolid BAz medium (composition in g/L: azelaic acid, 5.0; K2HPO4, 0.4; KH2PO4, 0.4; MgSO4$7H2O, 0.2; CaCl2, 0.02; Na2MoO4$H2O, 0.002; FeCl3, 0.01; and agar, 2.3) was used as an enrichment culture for N2-fixing Burkholderia species. Additionally, a BAc medium (in wt/v; 0.2% azelaic acid, 0.02% L-citrulline, 0.04% K2HPO4, 0.04% KH2PO4, and 0.02%, MgSO4$7H2O) was used as a selective medium to maintain the viability of pure cultures [20].
2.3.
pH optimization for cell growth
In order to determine the optimal pH values for growth of the diazotrophic strains tested, cell growth was tested at pH 5.0, 5.5, 6.0, 6.5, 7.0 and 7.5 based on the original soil pH where the bacterial strains were isolated. pH was adjusted using potassium hydroxide (KOH), and the N2-free BAz medium was used. Bacterial growth was determined by measuring the OD610 every 12 h over a total incubation period of 96 h.
2.4.
H2 measurement
A gas chromatograph equipped with a thermal conductivity detector (GC-TCD) (Agilent Technologies 7890A) and a megabore column (0.53 mm 30 m) was used for H2 detection in cultivated samples. After initial screening experiments, analytical conditions for the GC-TCD were determined as follows: temperatures of the injection port, oven and detector were 150 C, 40 C and 150 C, respectively. N2 was used as carrier gas with a flow rate of 3 mL/min. Calibration curves were constructed with a standard gas mixture (PRAXAIR: 30% N2; 30% CH4; 25% H2; 5% He; 5% Ar and 5% CO mol/mol) before the injection of the microbiological samples. The mathematical method for the H2 yield calculation was implemented according to the proposal by Juantorena et al. [31,32]. Statistical analysis consisted of analysis of
variance (ANOVA) and Tukey’s test (a ¼ 0.05) using the statistical package SAS 9.1 [33] as reported by Burrows et al. [4].
2.5. Bioreactors used for cell cultivation and H2 measurement assays Gas sampling flaks (Fig. 1a) with a total volume of 310 mL equipped with Teflon valves [34] were used as bioreactors throughout all H2 measurement assays, providing microenvironments isolated from the external environment. The bioreactors used had the following characteristics: fully sealed, resistant to sudden temperature changes and easy to use, handle and transport. The bioreactors had a connector valve of 9 mm in thickness for proper coupling to the injection system for subsequent gas quantification. These bioreactors attached to the GC-TCD for gas detection are shown in Fig. 1b.
2.6.
We subdivided this step based on the culture condition(s) evaluated. Each stage is comprised of various H2 measurement assays described below. Based on previous results (data not shown), the incubation times of all H2 measurement assays were divided into two different phases [31]. The first phase (aerobic phase) occurred prior to atmospheric manipulation, exclusively promoting biomass production, and was set at two days. The second phase (microaerobic phase) refers to the incubation of the strains after the atmospheric manipulation (vacuum and Ar injection) and was set at three days. Bioreactors containing 200 mL of the appropriate medium were inoculated at an OD610 of 0.5 with an aliquot of 2 mL of a 24 h culture of each selected strain [35] in BAz medium. Each assay was carried out in triplicate. Based on previous results, all H2 measurement assays were executed at a pH of 6.0 and a temperature of 35 C. It is worth mentioning that during the first set of results, a bacterial consortium consisting of a mixed culture of the two Burkholderia species was evaluated on its H2 production capacity along with the same strains tested individually. However, contrary to the expected, the H2 production rates did not show a significant increase (data not shown).
2.6.1.
Fig. 1 e Gas sampling flasks used as bioreactors: (a) unattached bioreactor and its length, (b) bioreactor attached to the GC-TCD for gas detection.
Optimization of culture conditions for H2 production
Atmospheric conditions
Oxygen (O2) concentration plays a crucial role in obligate and facultative aerobic N2-fixing bacteria like Burkholderia sp. because they have to tolerate a phenomena previously described as the “O2 paradox”. In these organisms, a minimal O2 concentration is necessary to support aerobic respiration and ATP synthesis. However, when O2 is present at a concentration that is too high, the nitrogenase can be irreversibly damaged [36,37]. Therefore, it was necessary to work at microaerophilic conditions. After each bioreactor was inoculated, a vacuum was generated for a total time of 4 min, with the culture stirred for 3 s every minute to eliminate the O2. Based on the results (data not shown) of testing a wide range of different argon (Ar) concentrations (25, 50, 60, 70, 80 and 90% (v/v)), Ar flux was set to 3 0.1 mL/s at different times [31,32]. The highest H2 quantifiable detections were obtained under an 80% Ar
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atmosphere. Based on these results, an atmospheric condition of 80% v/v Ar was used for further testing.
2.6.2.
Carbon sources
The carbon sources selected and their concentration were based on previous reports in the literature using the same diazotrophic species [20,24]. Pure carbon sources tested were as follows: succinic acid, mannitol, fructose, sucrose and glucose. The culture medium was composed of the mineral salts present in the BAz medium, substituting the azelaic acid with the individual carbon source to be tested at a concentration of 0.5%. Based on prior results, an 80% v/v Ar atmosphere was used. Following the first set of results, on a separate experiment, the two carbon sources with the highest H2 yield (sucrose and succinic acid) were combined at 0.5% wt/ v each (final carbon source-concentration of 1.0% wt/v) and supplemented to the mineral salts medium (BAz) and tested for H2 production.
2.6.3.
Chemical compounds as enzyme cofactors
Because nitrogenase is a metalloprotein, different chemical compounds were tested [38] to evaluate their influence as stimulators of nitrogenase activity and subsequent H2 production [39]. The following chemical compounds were selected: ferrous sulfate (FeSO4), 0.2 g/L; vanadium oxide (V2O5), 0.02 g/L and the amino acid cysteine, 0.02 g/L. The original Na2MoO4 concentration in BAz was also modified from 0.002 g/L to 0.2 g/L. The carbon sources in the medium were sucrose and succinic acid. Based on the first set of results, the chemical compounds with the highest H2 yield were combined and tested.
2.6.4.
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production, including the cost of using Ar. Given that Ar is used to promote anaerobic conditions, it was inferred that there should be no significant difference between using it and making a vacuum for the final anaerobic atmospheric concentration.
3.
Results
3.1.
pH optimization for cell growth
The results show that the optimal pH for cell growth for both B. unamae and B. tropica is close to 6.0 (Fig. 2), where highest absorbance values were obtained. In addition, for both strains, the lowest absorbance values were obtained at a pH of 5.0. Based on these results, all further H2 measurement assays were executed at a pH of 6.0.
3.2.
H2 yield with different carbon sources
After evaluating the five different carbon sources separately, the results obtained (Fig. 3) indicate that the individual carbon source with the highest H2 production was sucrose, with rates of 0.34 mmol H2/L for B. unamae and 0.49 mmol H2/L for B. tropica. After sucrose, succinic acid showed the highest H2 production rates, with 0.17 mmol H2/L for B. unamae and 0.12 mmol H2/L for B. tropica. The carbon source with the lowest H2 yield was mannitol.
Molasses as a culture medium
Sugar cane molasses from a local factory located in Zacatepec, Morelos, Mexico, was used. In laboratory conditions, we determined its moisture percentage at 46% and pH at 6.4. The composition of the original sugar cane molasses was not determined. It is well known that the composition highly varies depending on cane varieties, soil composition and climate. However, different studies in Mexico using sugar cane molasses show an average carbon source composition of: sucrose, w68%; glucose, w13% and fructose, w18% [30]. In this report, we evaluated three different treatments using molasses as a culture medium at a constant concentration of 1% v/v (based on previous tests where different concentrations were evaluated; data not shown). The first treatment consisted of using only molasses as the culture medium. For the second treatment, the pure molasses medium was supplemented with Na2MoO4 (0.2 g/L), FeSO4 (0.2 g/ L) and cysteine (0.02 g/L) to stimulate nitrogenase activity. The third treatment composition was identical to the second with the addition of the original mineral salts in BAz.
2.6.5.
Variation of atmospheric condition: Ar suppression
This final stage consisted of testing the two H2 measurement assays with the highest H2 yield under a different atmospheric condition, eliminating the Ar injection and under a partial vacuum (air 20% v/v). This variation was assessed due to the economic aspects of maintaining a cost-effective process for biohydrogen
Fig. 2 e Growth of bacterial strains tested at different pH values. a) B. unamae, b) B. tropica.
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tested, three (Na2MoO4, FeSO4 and cysteine) resulted in an increase in H2 production rates compared to previous assays. The Na2MoO4 (0.2 g/L) showed the highest increase in H2 production, with a rate of 4.02 mmol H2/L for B. unamae and 3.56 mmol H2/L for B. tropica. Treatment with V2O5 showed no significant increase in H2 production rates in any of the two strains tested. However, the combination of the three chemical compounds (Na2MoO4, FeSO4 and cysteine) exhibited a surprisingly high H2 production rate, significantly superior to all previous culture conditions tested. For B. unamae, H2 production was 17.61 mmol H2/L and for B. tropica, 12.35 mmol of H2/L.
Fig. 3 e H2 production rates of B. unamae and B. tropica under the influence of different carbon sources. Error bars indicate standard errors from the mean for at least three independent experiments.
Furthermore, the combination of sucrose and succinic acid (individual carbon sources with the highest H2 yield), exhibited significantly higher H2 production rates of 1.22 mmol H2/L for B. unamae and 1.03 mmol H2/L for B. tropica. This significant increase is partly due to the final carbon source-percentage increase (from 0.5% when tested individually to 1.0% wt/v when combined), although the final H2 production rates exceeded the rates expected when combining the two carbon sources.
3.3. H2 yield with the addition of different chemical compounds as enzyme cofactors The results obtained at this stage represent a critical point for the purposes of this study, revealing the positive impact of most of the chemical compounds tested. The results (Fig. 4) indicate that out of the four different chemical compounds
3.4.
H2 yield with molasses as a culture medium
After evaluating the influence of the three different treatments using molasses, the results obtained (Fig. 4) indicate that when pure molasses was tested as a culture medium, H2 production rates were below the previous assays in which additional carbon sources and chemical compounds as enzyme cofactors were supplemented. Nevertheless, when Na2MoO4, FeSO4 and cysteine were added to the molasses medium, the H2 production yield reached the highest rates obtained in the present study, with 24.64 mmol H2/L for B. unamae and 17.55 mmol H2/L for B. tropica. Finally, when mineral salts were added to the molasses medium with Na2MoO4, FeSO4 and cysteine, H2 production did not show a significant difference, indicating that the addition of mineral salts was ineffective.
3.5.
Influence of Ar suppression on H2 production
Comparing the two treatments with the highest H2 production rates (molasses medium enriched with Na2MoO4, FeSO4 and cysteine and the mineral salt medium supplemented with succinic acid, sucrose and Na2MoO4, FeSO4 and cysteine), a higher H2 production rate was obtained when Ar was used in
Fig. 4 e Summary of the H2 production rates obtained under the influence of chemical compounds as enzyme cofactors and molasses treatments by B. unamae and B. tropica. Different letters indicate statistically significant differences (P £ 0.0001, a [ 0.05). Vertical bars correspond to standard errors from the mean for at least three independent experiments. CV [ 44.77%.
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both treatments (Fig. 5). However, for the treatment with the highest H2 yield (molasses with Na2MoO4, FeSO4 and cysteine), statistical analysis showed no significant difference between the two atmospheric conditions. Use of Ar does not appear to be strictly necessary for the molasses medium at least. In Table 1, we present an overview of the different treatments evaluated, indicating the culture medium and the atmospheric composition along with the H2 yield obtained in each H2 measurement assay. In Fig. 6, we present two representative chromatograms obtained by GC-TCD, corresponding to two different H2 measurement assays. In those chromatograms, the increase in H2 throughout the experimental procedure is clearly shown by the size of the H2 peaks obtained.
4.
Discussion
This report represents a comprehensive study of H2 production using the nitrogenase complex of diazotrophic bacteria and it presents the utilization of Burkholderia species (B. unamae and B. tropica) to produce H2, which has not yet been explored in the literature, presenting a different approach in biological H2 production. The manipulation of the different culture conditions tested allowed a better understanding of the metabolic process and optimal conditions for H2 production by the evaluated N2-fixing species. However, this report offers a brief outlook toward the BNF system to produce H2 but the feasibility of the process remains uncertain. When atmospheric conditions were tested, it was clear that a critical issue for the cell was the protection of the nitrogenase complex from O2 inactivation. Incubation of cells under a low O2 environment provided the highest rates of H2 production, which is corroborated by Min and Sherman [35]. In this case, for B. unamae and B. tropica, a final atmosphere of Ar 20% v/v was optimal for H2 production. Cells obtained the necessary O2 concentration for respiration but it was not high enough to trigger nitrogenase inactivation. The cellular
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mechanism for O2 protection in the N2-fixing Burkholderia species remains unclear and should be further studied. Regarding the influence of different carbon sources, the predilection of the Burkholderia strains towards sucrose as a carbon source may be related to their origin (sugarcane crops), where sucrose is the predominant carbohydrate [20]. We cannot explain why succinic acid was the single carbon source with the second highest H2 production rate, but it may be related to the metabolic role of succinic acid in the citric acid cycle and energy-production process [40]. The predilection for sucrose was also consistent when molasses was tested as the primary carbon source, assuming that the main component of molasses is sucrose (w35% wt/v) [28e30]. However, the H2 production rates in both species were significantly higher when molasses was used. This not only has to do with the difference in sucrose concentration in both media, but with the rich composition of molasses, which includes a variety of carbohydrates (sucrose, glucose and fructose) and different molecules including minerals, amino acids and vitamins necessary for cell growth [30]. Although there are several studies referring to the utilization of molasses as a culture medium for H2 production [28,29], little is known about its use in N2-fixing bacteria. Although the composition of molasses is highly heterogeneous, depending on environmental factors and its production process, it represents a potential element for the culture of a wide range of microorganisms and should be further assessed, possibly testing different specific molecules as supplements for the medium enrichment. The influence of the chemical compounds tested as stimulators of nitrogenase activity represents a major issue in the results of this study. Taking into account that the metalloprotein nitrogenase contains and needs enzyme cofactors to carry out its metabolic functions, the addition of the chemical compounds serving as enzyme cofactors to the different culture medium tested resulted in an exponential increase of the H2 production rates. The only metal ion tested that did not
Fig. 5 e Comparison of the H2 production rates by B. unamae and B. tropica using the two culture media with the highest H2 yield yet obtained, under two different atmospheric conditions, in the presence of Ar (vacuum D 80% Ar v/v) and without it (partial vacuum). Different letters indicate statistically significant differences (P £ 0.0001, a [ 0.05). Vertical bars correspond to standard errors from the mean for at least three independent experiments. CV [ 24.94%.
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Table 1 e Summary of the different culture conditions evaluated (culture medium and atmospheric condition) in this study and the H2 yield obtained in each case for both strains. Primary culture medium composition
Enzyme cofactors
Atmospheric condition (v/v)
Molasses Molasses
Na2MoO4 þ FeSO4 þ cysteine Na2MoO4 þ FeSO4 þ cysteine
Molasses þ Mineral salts Mineral salts þ succinic acid þ sucrose Mineral salts þ succinic acid þ sucrose
Na2MoO4 þ FeSO4 þ cysteine Na2MoO4 þ FeSO4 þ cysteine Na2MoO4 þ FeSO4 þ cysteine
Mineral salts þ Mineral salts þ Mineral salts þ Molasses Mineral salts þ Mineral salts þ
Na2MoO4 Cysteine FeSO4
Ar 80% Partial vacuum (air 20%) without Ar Ar 80% Ar 80% Partial vacuum (air 20%) without Ar Ar 80% Ar 80% Ar 80% Ar 80% Ar 80% Ar 80%
succinic acid þ sucrose succinic acid þ sucrose succinic acid þ sucrose succinic acid þ sucrose succinic acid þ sucrose
e e V2O5
produce a H2 increase was vanadium oxide (V2O5). In fact, both species exhibited a considerable decrease in the H2 yield when V2O5 was added to the primary culture medium composition (mineral salts þ sucrose þ succinic acid) without any enzyme
H2 production (mmol/L) B. unamae
B. tropica
24.64 3.36 23.81 3.69
17.16 8.91 14.98 3.28
23.23 1.51 17.61 2.36 10.18 1.31
14.89 7.50 12.35 6.25 7.62 1.67
4.02 3.69 2.76 2.00 1.92 1.40
0.89 1.62 1.24 0.46 0.58 0.45
3.56 2.81 2.19 1.45 1.33 1.25
1.98 2.17 1.82 0.53 0.93 0.79
cofactor (Table 1). This may be due to the high oxidation state of V2O5, acting both as an amphoteric oxide and an oxidizing agent. It may also be related to its reported capacity to produce by-products that can inhibit enzymes that process phosphate
Fig. 6 e Examples of chromatograms obtained by the GC-TCD with two different culture condition treatments. a) Mineral salts D succinic acid D sucrose, b) Molasses D FeSO4 D cysteine D Na2MoO4. In both cases, the atmospheric condition consisted in vacuum D 80% Ar. H2 peaks are observed at a retention time of w1.5 min, while the other peak represents N2.
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(PO3 4 ) [41,42]. In any case, the results allow us to discard the potential presence of alternative vanadium (V)-nitrogenase [8] in the Burkholderia strains tested. Comparison of the H2 production rates by the Burkholderia species tested versus others obtained by biological processes such as dark fermentation or indirect bio-photolysis [1] provides a useful benchmark for future improvements. The peak H2 production rates obtained are still lower than those obtained in those systems [1,3,9,10]; nevertheless, this study represents the first data using Burkholderia N2-fixing bacteria as H2 producers and opens a range of possibilities to evaluate different diazotrophic species in the future. Moreover, alternatives and variants for the optimization of culture conditions remain unexplored. Currently, among the known biological processes for H2 generation, dark fermentative H2 production is considered the most attractive, with the highest yields reported [1,9,10]. However, fermentative H2 production processes, either conducted via mixed or pure cultures, still have certain limitations that could be crucial for scaling up. Inefficient substrate conversion, formation of by-products, low resistance to high H2 partial pressures and oxygen are only some of the limitations. Above all those, comes the limiting upper value of H2 yields which is 4 mol of H2/mol hexose consumed [27]. Besides this, the gas produced in these processes is a mixture of primarily H2 and carbon dioxide (CO2), but may also contain other gases such as methane (CH4), hydrogen sulfide (H2S), or ammonia (NH4). For the scaling up of such a process, the raw material cost is considered to be among the major limitations. This particular limitation is minimized in the present approach, considering that the optimal culture medium tested for H2 production (molasses medium) is an abundant and relatively inexpensive agro-industrial waste product (cost lower than 0.5 US dollars per L) [30] and with the elimination of Ar, the production process tested is economically promising in relation to other known processes such as indirect biophotolysis and fermentation [3,7]. On the other hand, the main limitation of this nitrogenase BNF reaction approach is that this process is thermodynamically uphill and requires ATP hydrolysis. Two molecules of ATP have to be hydrolyzed for every electron made available for H2 production, making the reaction highly ATP-dependent [10]. If biohydrogen is to be produced by whole microorganisms, it must be taken into account that H2 output represents an energy loss for the cell. Regulatory networks control the biosynthesis and activity of the involved enzymes; therefore, it is necessary to understand how they function before being able to maximize the production of H2. Also, in some legume nodules, the bacterial symbiont produces an uptake hydrogenase (HUP) enzyme that oxidizes the H2 evolved, recovering a portion of the energy used in its production. However, many of the most productive N2-fixing associations lack HUP activity and the H2 diffuses into the soil where it is consumed [43]. In response to some of the mentioned limitations above, several researchers have proposed the use of genetically modified microorganisms with advanced selected properties, constructed either via mutagenesis or through genetic engineering. H2 producing bacteria can be genetically modified in several ways to increase H2 synthesis, including (1) over-
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expression of certain enzymes that can maximize substrate availability; (2) elimination of uptake hydrogenases; (3) overexpression of H2 evolving hydrogenases that have been modified to be O2 tolerant; and (4) elimination of metabolic pathways that compete for reducing equivalents required for H2 synthesis [1,27]. As mentioned before, further research and development is needed to improve H2 production rates through this novel approach. Several experimental parameters should be optimized to manipulate the nitrogenase BNF reaction to get a higher H2 yield in these two Burkholderia species. These parameters include factors which affect the distribution of electrons between H2 production and N2 reduction within nitrogenase, such as, temperature, pH, the ratio of the two component proteins of nitrogenase and the ATP concentration [44]. Those parameters do not only determine the growth of the microorganisms, but also have a crucial role on the metabolic path that the microorganisms will follow, severely affecting the final H2 yield observed. Other factors to examine further include cell concentration, combinations of species, and utilization of different organic waste products. Furthermore, future experiments will involve a careful simultaneous analysis of both H2 production and cell growth over extended periods of time to determine the best conditions for each specific application. Although it is well known that one reason for the tremendous H2 production potential of nitrogenase is that there is a vast amount of enzyme present in the cell [37], both processes need to be experimentally differentiated and independently valued.
5.
Conclusions
The hydrogen-production capacity of the diazotrophic bacteria B. unamae and B. tropica was confirmed, with B. unamae exhibiting the highest production rates in most of the conditions tested. Although the peak H2 production rates obtained are lower than the rates reported for other biological systems, the present study is innovative as it represents the first contribution of an optimization process for H2 production by N2-fixing Burkholderia species. Utilization of the chemical compounds Na2MoO4, FeSO4 and cysteine as stimulators of nitrogenase activity has been demonstrated, increasing the H2 production rates where supplemented. Moreover, using molasses as a carbohydraterich waste is a promising approach to developing a costeffective system for biohydrogen production. Finally, it is widely believed that H2 will play a crucial role in the future energy economy, but considerable research and developmental studies are needed to improve H2 production rates and to reduce production costs in order to make biohydrogen a tangible reality that is capable of meeting the increasing need for a substitute for fossil fuels.
Acknowledgments The authors are thankful to Dra. Rocı´o Castro Gonza´lez for providing the bacterial strains used and to Dra. Alina
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Juantorena Uga´s for general support. We also acknowledge the technical assistance of Tech. Vicente Betancourt and Tech. Jose´ Antonio Nopaltitla. Horacio is thankful to the Consejo Nacional de Ciencia y Tecnologı´a (CONACYT) for the graduate fellowship.
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