Conversion of H₂ and CO₂ to CH₄ and acetate in fed-batch biogas reactors by mixed biogas community: a novel route for the power-to-gas concept

Background

Applications of the power-to-gas principle for the handling of surplus renewable electricity have been proposed. The feasibility of using hydrogenotrophic methanogens as CH₄ generating catalysts has been demonstrated. Laboratory and scale-up experiments have corroborated the benefits of the CO₂ mitigation via biotechnological conversion of H₂ and CO₂ to CH₄. A major bottleneck in the process is the gas–liquid mass transfer of H₂.

Results

Fed-batch reactor configuration was tested at mesophilic temperature in laboratory experiments in order to improve the contact time and H₂ mass transfer between the gas and liquid phases. Effluent from an industrial biogas facility served as biocatalyst. The bicarbonate content of the effluent was depleted after some time, but the addition of stoichiometric CO₂ sustained H₂ conversion for an extended period of time and prevented a pH shift. The microbial community generated biogas from the added α-cellulose substrate with concomitant H₂ conversion, but the organic substrate did not facilitate H₂ consumption. Fed-batch operational mode allowed a fourfold increase in volumetric H₂ load and a 6.5-fold augmentation of the CH₄ formation rate relative to the CSTR reactor configuration. Acetate was the major by-product of the reaction.

Conclusions

Fed-batch reactors significantly improve the efficiency of the biological power-to-gas process. Besides their storage function, biogas fermentation effluent reservoirs can serve as large-scale bio CH₄ reactors. On the basis of this recognition, a novel concept is proposed, which merges biogas technology with other means of renewable electricity production for improved efficiency and sustainability.

Pressing deterioration of the global climate by human activities demands the large-scale replacement of fossil fuels with renewable energy carriers [1]. The most rapidly developing and spreading renewable technologies worldwide include the conversion of wind energy and direct solar energy (photovoltaics) to electricity. In view of the discontinuous electricity production by these technologies, coupled with fluctuating utilization, severe electricity storage problems arise, which are already apparent in countries where the implementation of renewables is well advanced. A likely solution of this emerging setback is conversion of electricity to alternative energy carriers [2] or chemicals [3]. Hydrogen (H₂) can be generated via electrolysis of water, a well-known and efficient process [4]; however, technologies to store and transport H₂ are underdeveloped at present. Methane (CH₄) is an obvious next candidate. CH₄ can be transported and stored conveniently in the existing natural gas grid and can be used in all applications where fossil natural gas is employed today. Biogenic CH₄ production takes place during anaerobic degradation of organic matter in biogas reactors, swamps, ruminants, termites, etc. [2]. The last step of these complex microbiological metabolic pathways is biogas formation by methanogens. These microbes are strict anaerobes belonging in the kingdom Archaea. Some methanogens split acetate and release a mixture of CH₄ and CO₂ (acetotrophic methanogens), while others form CH₄ by reducing CO₂ with H₂ (hydrogenotrophic methanogens) and there are methanogens which are able to carry out both reactions.

An additional advantage of the biological conversion of electricity to CH₄ is offered by coupling the process with CO₂ mitigation. CO₂ can be transformed by catalytic reduction using chemical reactions [5, 6], photosynthesis [7], bioelectrochemical processes [8–10], or methanogenesis [2].

Three main ingredients should be present to form biogenic CH₄ from CO₂: hydrogenotrophic methanogens, CO₂, and a suitable reductant. Recent metagenomic studies have revealed that hydrogenotrophic methanogens predominate among Archaea in most biogas microbial communities [11–17].

CO₂ can originate from flue gas [18] or from the biogas itself [19–22]. In the latter approach, a significant upgrading of the produced biogas has been achieved. In some cases, the anaerobic degradation of the biomass has provided the electron source [18, 23]; in other studies, H₂ gas has been employed [19, 20, 22, 24]. These experiments have revealed that the poor solubility of H₂ limits the yield and rate of CH₄ formation. In these configurations, H₂ is injected into a methanogenic reactor filled with a microbial consortium.

In the present study, fed-batch fermentation systems with the daily dispensing of H₂ gas were employed in order to partially overcome the H₂ solubility problem. Several operational conditions (see “Methods” section) were tested under mesophilic conditions and efficient CH₄ productivity was attained. Moreover, at the appropriate combination of CO₂ and H₂, the simultaneous formation of acetate and CH₄ as main products was observed.

Effect of mixing

Given the experimental conditions (see “Methods” section) and the poor solubility of H₂ in the aqueous phase, the optimal mixing conditions yielding the most efficient delivery of H₂ from the gas phase had to be determined. The reaction vessels were incubated in an orbital shaker at various mixing speeds (rpm). Figure 1 indicates that there is an optimum value for this parameter; in our arrangement, it was 150–160 rpm. It is noteworthy that at higher mixing rates CH₄ production decreased sharply in contrast to earlier observations at thermophilic temperature [19]. In all subsequent experiments, the shaker was set at 160 rpm. It is evident that this mixing rate is valid under our conditions and henceforth was applied consistently in order to limit the number varying parameters. In other systems, the optimal mixing conditions should be determined individually. The main conclusion from these experiments was that the mixing that yields optimal H₂ utilization may not be the maximum achievable mixing rate.

Dependence of the CH₄ production on shaking speed

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Optimization of H₂ dosage

Next the optimal daily H₂ dosage was established. Various volumes of H₂ were therefore injected into the batch reactors, which were treated identically in all other known aspects. The batch fermentations were started by adding 0.3 g of α-cellulose as substrate for AD according to the VDI (Verein Deutscher Ingenieure, protocol [25]. H₂ gas was injected every day and the consumption of H₂ was followed by gas chromatography. Cumulative CH₄ evolution curves are plotted in Fig. 2. CH₄ production proceeded steadily for 7–8 days in the control reactors, which received no daily H₂ dosage, but from day 12 practically no gas evolved. In total, 6.2 ± 0.54 mmol of CH₄ was generated from the residual biogas potential of the sludge and added α-cellulose substrate. 1.62 mmol of this quantity originated from the sludge and 4.58 mmol from the α-cellulose substrate. The biochemical CH₄ potential of α-cellulose is 4.71 [26] and therefore all of the added substrate was consumed by the community and was converted to CH₄. Addition of a daily 0.81 ± 0.16 mmol of H₂ gas into the headspace of the batch reactors dramatically increased the CH₄ production (Fig. 2). The GC measurements revealed that all of the injected H₂ was completely consumed by the microbes within 24 h. In separate experiments, it was established more precisely that under these conditions all the H₂ had vanished from the headspace after 16 h and CH₄ evolution started at hour 2 following H₂ injection (data not shown). A new dosage of H₂ was dispensed consistently every 24 h. Increasing the total H₂ load to 43.00 ± 1.43 mmol resulted in a somewhat faster initial CH₄ production, but the cumulative-specific CH₄ production was lower than in the case of adding 24.42 ± 0.81 mmol of H₂ in the same period of time. In line with this observation, H₂ started to accumulate in the headspace on day 14 and from day 17–18 CH₄ production ceased. On further increase of the overall H₂ injection volume to 55.69 ± 1.85 mmol, i.e., 1.86 ± 0.38 mmol H₂ day⁻¹, even less cumulative-specific CH₄ was yielded. In these reactors, H₂ build-up in the headspace started sooner, i.e., on day 10 and CH₄ evolution stopped completely on day 13. Overall, these results indicated that the system utilized the α-cellulose substrate within 7–8 days and the microbial community sustained its H₂ conversion activity for an extended period of time if the daily H₂ injection did not exceed 0.81 ± 0.16 mmol of H₂ (Table 1). The concentrations of organic acids were determined every week. Acetate levels increased significantly by the end of the experimental period. 3.43 mM acetate accumulated by the end of the experiment in the reactors receiving 55.69 ± 1.85 mmol of H₂, which exceeded the recommended threshold, but apparently this alone did not explain why CH₄ evolution stopped in the reactors loaded with higher daily H₂ injections (Fig. 3). The pH had increased considerably by the end of the 4-week experiments (Fig. 4), indicating a severe loss of the bicarbonate buffering capacity of the inoculum sludge. It is noteworthy that the pH also shifted by 1.1 units in the control reactors which were not fed with H₂. In order to employ the same protocol, these vessels were also degassed and filled with N₂ gas every day. It is therefore likely that the daily replacement of the headspace prompted a gradual desorption and loss of dissolved CO₂ and caused a shift in the bicarbonate buffering system [27, 28]. The pH increased even further, i.e., beyond pH = 9, which is a critical upper limit for the methanogenesis [29]. A similar exhaustion of the buffering capacity upon H₂ addition was noted in previous reports [19, 20]. The system could apparently tolerate high pH fairly well when 0.81 ± 0.16 mmol of H₂ was the daily dosage, but started to inhibit CH₄ biosynthesis on day 13 and 10 upon addition of daily 1.43 ± 0.28 or 2.86 ± 0.38 mmol of H₂, respectively. In this experimental set-up, it was not possible to determine the time points when the inhibitory pH range was attained. The results indicated that the likely reason for the obstruction of CH₄ formation was the limiting buffering capacity of the system due to the low bicarbonate concentration. The optimal amount of daily H₂ dosage in this system was within the range of 0.8–1.5 mmol of H₂; further experiments should determine the exact value.

Cumulative CH₄ production from H₂. α-cellulose (0.3 g) was added as substrate at the start of the experiment. H₂ was injected into the reactor headspace daily, following flushing with N₂. Green: no H₂ added, red: 0.79 mmol H₂ day⁻¹, black 1.57 mmol H₂ day⁻¹, blue: 2.36 mmol H₂ day⁻¹ added

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Levels of volatile organic acids in the reactors at the beginning (0 weeks = inoculum and after week 1 and week 4), respectively. The reactors received daily injections of H₂ gas: 0.0 (green), 0.79 (red), 1.57 (black), and 2.36 (blue)

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The initial (red) and final (blue) pH in the liquid phase of the reactors received mmole, respectively. α-cellulose (0.3 g) was added as substrate at the start of the experiment

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Effect of CO₂ addition

In the next series of batch fermentations, the inoculum originated from the same mesophilic industrial biogas plant, but at different points of time, and therefore small fluctuations of organic total solid content and microbial community composition should be taken into account when the results are subjected to direct comparison. The initial addition of α-cellulose was omitted in order to avoid any disturbing effect of the CH₄ generation from the substrate. The duration of these fermentations was extended to 80 days to test for sustainable CH₄ production. The reactors were supplied with the optimal daily dosage of 0.81 mmol of H₂ in order to check if the CO₂/bicarbonate buffering capacity was indeed the major limiting factor in the previous experiments [28, 30]. The daily CH₄ volumes measured in the headspace are plotted in Fig. 5. CH₄ evolution progressed steadily until day 28, but dropped sharply afterwards. A warning sign of system failure was noticed already on day 27, when measurable residual H₂ was detected in the headspace (Fig. 5; Table 2). As shock therapy, massive CO₂ injection (25 mL) was dispensed into the reactors following the daily dosage of H₂ on day 31 (Fig. 6). All of this CO₂ disappeared from the gas phase within 24 h, indicating that the system was indeed severely depleted of CO₂/bicarbonate. The same CO₂ treatment was repeated next day, which apparently restored the functional state of the system signaled by the build-up of residual CO₂ in the headspace (Fig. 6). The daily CO₂ dose was then gradually decreased to the stoichiometric volume, i.e., approximately 0.25 mol of CO₂/mol of H₂ per day. The system responded positively, as exhibited by the restoration of CH₄ production on day 32 accompanied by a gradual decrease of residual CO₂ levels in the gas phase. Daily CO₂ injection was stopped on day 41. H₂ accumulation commenced again almost immediately and was accompanied by the loss of CH₄-evolving ability from day 43, and therefore CO₂ injection (25 mL) was resumed on day 47. Detectable remaining CO₂ was noticed already on the next day and from this time on a daily dosage of 0.25 mol of CO₂/mol of H₂ of CO₂ was maintained until the end of the experiment. CH₄ production returned to the previous level, all of the injected daily H₂ and CO₂ were consumed within 24 h and this continued for an additional month. It is noteworthy that, except for pH bursts on days 31 and 45, the pH in both the control and H₂-fed reactors remained within the acceptable limit of pH ≤8.5 throughout the investigated period (data not shown).

The amount of daily H₂ injection (dotted yellow), CH₄ production (red) and residual H₂ (dotted dark blue). CH₄ production by the control reactor (no H₂ injected) is shown in light blue. The reactors received no external substrate

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The amount of CO₂ injected daily (red) and the residual CO₂ concentration in the headspace after 24 h (dashed blue). The reactors received no external substrate. Experimental conditions as in Fig. 5

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Several deductions could be drawn from this series of tests. First, the system becomes depleted of CO₂ if semi-continuous H₂ feeding and daily degassing are administered to the fed-batch system. This phenomenon was manifested after about 1 month in our arrangement, where daily degassing and replacement of the headspace were included to retain the same protocol in the control and experimental reactors. Clearly daily degassing is not necessary in industrial setting. Second, the residual H₂ accumulation in the gas phase is a good early warning sign of upcoming system failure due to CO₂ exhaustion. Third, the microbial community participating in the CH₄ generation process recuperates quickly and completely even after repeated system failure if the process control is alerted in time. Fourth, the microbial community supplied only with H₂ and CO₂ upholds the pH within the normal operating range. Finally, stoichiometric administration of H₂ and CO₂ yields a practically complete conversion to pure CH₄ within 24 h under mesophilic conditions.

Effect of additional substrate addition

Next, it was tested whether the addition of α-cellulose affected the CH₄ productivity from H₂. Two series of experiments were designed and the duration of the experimental period was shortened in order to avoid any complication due to CO₂ depletion and concomitant pH elevation. In the first set of batch fermentations (Fig. 7), various amounts of α-cellulose were added only at the start of the experiments, and in the second series (Fig. 8) the addition of the same amount of α-cellulose was repeated every week. Daily replacement of the headspace with N₂ and the injection of 0.81 mmol of H₂ was maintained in all reactors.

Cumulative CH₄ production at various initial α-cellulose loadings: 0 g (green), 0.3 g (red), and 0.6 g (blue)

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Cumulative CH₄ production at various weekly α-cellulose loadings: 0 g (green), 0.3 g (red), and 0.6 g (blue)

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There was no significant difference between the CH₄ productions from H₂ in the reactors receiving the substrate quantity recommended by the VDI [25] protocol as compared with those without substrate, i.e., the difference between the green and red curves in Fig. 7 correspond solely to the CH₄ produced from α-cellulose. This suggests that the addition of substrate at the beginning of the fermentation does not assist CH₄ evolution from H₂. Moreover, an inhibition of CH₄ productivity from H₂ was noted when the substrate load was doubled, i.e., upon the addition of 0.6 g substrate, 3.47 ± 0.08 mmol of CH₄ was formed from α-cellulose instead of the theoretical potential of 9.42 mmol of CH₄. It should be noted that the H₂ consumption rate remained unaffected by the substrate loading, i.e., the injected H₂ disappeared from the headspace within 24 h. The conversion efficiency of CH₄ formation from H₂ was estimated from the daily CH₄ levels in the headspace. The day-to-day values fluctuated considerably during the experimental period and achieved an average of 72 ± 25 %. The remainder of the H₂ may have been metabolized in alternative pathways, which are the subject of future studies.

In the next set of experiments, the reactors were fed with the same amount (0.3 g) of α-cellulose every week and the daily H₂ injection (0.81 mol H₂) was maintained. The aim was to test whether the microbial community remained intact for an extended period of time after the expiration of its residence time in the industrial AD facility and to see whether the metabolically active community facilitated the bioconversion of H₂ to CH₄. The cumulative CH₄ production increased almost linearly and the amount formed suggested an unchanged reaction rate for both α-cellulose and H₂ when the VDI protocol [25] was followed (Fig. 8). It is noteworthy that increasing the weekly α-cellulose load prompted a strong inhibitory effect. The collapse of the CH₄-forming activity was not associated with changes in pH. Without α-cellulose, the daily dosage of H₂ caused an increase of the pH into the dangerous zone, as observed earlier (Fig. 4), due to the depletion of the buffering capacity. Weekly supply of the substrate balanced the pH; the degradation of the α-cellulose apparently yielded enough CO₂ to maintain stable operation. Too much substrate, e.g., 0.6 g α-cellulose/week, shifted the pH to lower values, although it did not fall below 6.5, which is usually considered detrimental [29]. The accumulation of acetate increased dramatically upon substrate overloading (data not shown). This might have been the likely reason for the process inhibition. It is important to note that the H₂ conversion yields in this series of experiments were close to 100 %, which emphasizes the importance of the inoculum quality.

Storage of surplus electricity is a growing demand in renewable energy technology, with the generation of electricity in an inherently fluctuating mode of operation, such as wind and direct solar, gaining a rapidly increasing market share. In a popular strategy, electricity is used to split water and generate H₂ and O₂. There are no mature technologies available for handling H₂ today, and its conversion to CH₄ therefore seems preferable. In this scheme, electricity is transformed into CH₄, which is then stored and transported easily via the existing natural gas grids. Chemical methods to reduce CO₂ with H₂ have been known for some time and earned the Nobel prize for Paul Sabatier in 1912 [31]. The process requires high temperature, high pressure, and metal catalysts. In alternative electrochemical means of CO₂ mitigation, electrical energy input is the driving force [3, 9, 30]. Biological systems can solve the same task under mild conditions in an environmentally friendly manner. The life of hydrogenotrophic methanogens, an odd group of Archaea, relies on the same reaction, which is catalyzed by enzymes at ambient temperature and pressure. The biological route of the power-to-gas process, which is here named as power-to-biomethane (P2B), has been recognized and tested in laboratory and scale-up works [19, 22, 24, 32]. These studies have established that microbes are exceedingly efficient catalysts for the P2B process. Hydrogenotrophic methanogens are difficult to cultivate in pure culture, but they are readily available in the mixed culture of effluents from the anaerobic degradation of organic matter, i.e., the fermentation effluent of biogas plants. The rate-limiting step in the work of CH₄-forming microbial cell factories is the low solubility of H₂ in the aqueous environment. In previous studies [19, 22, 24, 32], continuously-operating fermentation systems were employed as a rule, which offer several advantageous features for process control and management, but allow short residence time for the injected H₂ gas.

In our approach, the fed-batch fermentation technique was selected to increase the contact interaction between the gaseous substrate and the biocatalyst methanogens. It was established that an optimal mixing rate has to be upheld in any P2B system in order to facilitate the dissolution of H₂ into the aqueous phase where the microbes and dissolved CO₂ reside.

Although CO₂ is readily soluble in the aqueous medium, it may become an overall limiting factor if removed from the system either by vigorous reaction with H₂ or by degassing the reactors. Depletion of CO₂ was accompanied by the elevation of pH, which might be precarious for the activity of hydrogenotrophic methanogens.

CO₂ is supplied by the biogas-generating process itself [19, 22] or can be provided from outside sources, e.g., flue gas from internal combustion engines. Consumption of the greenhouse gas CO₂ by the process is an additional benefit of the P2B technology from an environmental point of view. Addition of an organic substrate may revitalize the entire biogas microbial community, which generates additional CO₂ and thereby stabilizes the pH, but does not facilitate the conversion of H₂ to CH₄. A proper feeding routine in the fed-batch system leads to a sustained high rate of CH₄ formation and the process may operate efficiently for an extended period of time.

Comparison with previous works

Our approach to improve the P2B principle attempts to counteract the low solubility of H₂ in the aqueous environment by increasing the contact time of the gas and aqueous phases in a fed-batch fermentation arrangement. This has not been tested earlier.

There are four previous reports available to measure up against this approach. Lee et al. [24] used a fixed-bed reactor, while Reuter [32] developed several versions of a continuous stirred tank reactor (CSTR) design and scaled up the process to an industrial level. Both studies concluded that hydrogenotrophic methanogens in pure or mixed culture were markedly efficient catalysts and converted H₂ and CO₂ to CH₄ in surprisingly high yields and rates. Unfortunately, the published results from those studies contain limited data on process parameters to compare with the fed-batch system examined in the present study.

Two recent papers from the Angelidaki team [19, 22] also used CSTR reactors and reported promising results. Their thoughtfully designed and thoroughly documented reports provided data allowing the assessment with our study. Table 2 summarizes the results.

Besides the use of distinct reactor arrangements and sizes, i.e., fed-batch versus CSTR, several operational parameters differed in those studies from our set-up, e.g., inoculum composition and quality, substrate used for CH₄ generation, stirring mode and speed. Therefore, only the major tendencies and not the exact values are suitable for a rigorous comparison.

It was found that at high shaking speed the H₂ conversion process may not be limited by the gas–liquid mass transfer [19] at thermophilic temperature. In our experience, this observation could not be repeated under mesophilic conditions, and above 160 rpm CH₄ formation was inhibited (Fig. 1). It was concluded that the process in our system was critically limited by the mass transfer of H₂ at the gas–liquid interface. Hydrogenotrophic methanogens utilized the dissolved H₂ at a high rate, and therefore a concentration gradient developed between the liquid and gas phases, driving H₂ into the liquid compartment from the headspace as time advanced. It is likely that the fed-batch operation optimized the condition where the amount of H₂ transferred into the liquid phase was close to the amount consumed by the microbes. The data presented in Table 2 clearly indicate that this was indeed the case.

In the CSTR work, H₂ was dosed on the basis of the available CO₂ from the coupled biogas production [22]. Although significant upgrading of the biogas was achieved, this stipulation limited the rate and amount of H₂ injection into the system. The goal in these investigations was to achieve maximal H₂ conversion yield. H₂ bubbles are difficult to retain in the aqueous system, and diffusers and very low purging rates therefore had to be applied to facilitate the dissolution of H₂ and its conversion to CH₄ during the short residence time of the gaseous substrate in the reactor. In the fed-batch configuration, the H₂ loading rate could be increased to 4 times that of the CSTR operational mode without the loss of H₂ (Table 2).

In the present study, mesophilic conditions were maintained. Bassani et al. [22] carried out their experiments at 35 and 55 °C under otherwise identical conditions. A significant improvement in CH₄ formation rate was noted at higher temperature. A similar effect can be expected in the fed-batch system; this will have to be established in future studies. A comparison between our mesophilic data with those obtained at thermophilic temperature indicates a 2.0 [19] and 2.7 [22] times higher CH₄ production rate from H₂ in the mesophilic fed-batch reactors as compared with the thermophilic CSTR, respectively.

The mesophilic process performance parameters of Bassani et al. [22] can be compared directly with our results reported under the “Effect of CO₂ addition” subtitle above. Two sections of stable operation in our experimental period were taken into account, i.e., the initial phase without external CO₂ addition between days 2 and 28 and the part when stoichiometric CO₂ and H₂ were injected daily (days 50–80) (Figs. 5, 6). To make a fair assessment, the residual CH₄ production in the control reactors (no H₂ added) should be taken into account.

The control samples in our work started at an unusually low CH₄/CO₂ ratio (Table 2), which could be due to the residual biogas potential of the inoculum and the fact that all H₂ was removed during initial degassing of the reactors. Therefore, the activity of the hydrogenotrophic methanogens was severely restricted until some H₂ became available from the fermentation of the residual, small amount of biomass. The situation changed dramatically in the reactors receiving H₂ injections and the system produced bio CH₄ of high purity, i.e., containing only 17.71 % CO₂.

There was a 6.5-fold increase in CH₄ yield from H₂ in the fed-batch system relative to the mesophilic CSTR experiments if a stoichiometric amount of CO₂ was added to both systems together with the H₂ (Table 2). Moreover, the fed-batch system operated at a 4-times higher H₂ load than the CSTR reactor. The H₂ consumption was above 90–100 % in both systems, indicating that the reaction was carried out very efficiently in both systems. The CSTR operation mode has its benefits and advantages, but apparently does not help overcome the low H₂ solubility problem, which seems to be the major bottleneck in the accomplishment of the P2B principle at mesophilic temperature.

As an added value, it should be noted that in the fed-batch system a considerable accumulation of acetate takes place without any observable sign of acidosis-related process failure (Fig. 9). The accumulation of acetate was probably due to the inhibition of acetoclastic methanogenesis and syntrophic acetate oxidation [33] by the high H₂ doses. Acetate is a valuable commodity [30, 34] and, if acetate can be recovered by a suitable technology from the reaction mixture, it would be a useful side-product of the fed-batch fermentation-based P2B technology.

Distribution of volatile fatty acids in the control (blue) and (H₂ + CO₂)-fed (red) reactors. Experimental conditions as in Fig. 5

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A general strategy can be proposed on the basis of the results reported above to utilize the microbial community formed in the biogas reactor for the efficient conversion of H₂ to CH₄ as part of the P2B principle. Previous studies [19, 22, 24, 32] and the present work unambiguously corroborated that microbiological cell factories are very efficient catalysts to combine H₂ and CO₂ to CH₄, a renewable energy carrier that has already been in use in human practice for many years as fossil natural gas. The suitable microbial community is freely available in the effluent of anaerobic fermentation at the biogas plants operating word-wide in millions of installations at various levels of sophistication.

At the center of the projected strategic alliance comprising either of the methods yielding renewable electricity and biogas technology (Fig. 10) are the hydrogenotrophic methanogens present in the biogas effluents. They convert H₂, which is produced from excess electricity by electrolysis, to CH₄. BioCH₄ is relatively easily stored and transported with minor loss in the natural gas grids over large distances and used as energy carrier, biofuel or basic commodity [35], and several technological improvements of bioCH₄ production [36] have been therefore developed. The proposed novel strategy places biogas technology into the hub of the renewable energy production and utilization network. The biogas effluent reservoir, which forms part of most industrial-scale biogas facilities and stores the digested material until its utilization as organic fertilizer, acquires an entirely new function by becoming a bioreactor to transform green electricity-derived H₂ into bioCH₄. The gas to liquid volumetric ratio is lower in industrial-scale effluent reservoirs than the ratio used in our experiments, and installation of a gas recirculation system may therefore be required in the large-scale applications.

Proposed novel P2B scheme involving the AD fermentation residue storage tank as bio CH₄ reactor, which converts CO₂ from biogas or flue gas and H₂ from electrolysis by renewable electricity

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The potential economic advantages consequent from the scheme recommended in Fig. 10 are numerous. First, the microbial community present in the biogas effluent can be directly exploited for the efficient conversion of H₂ and CO₂ to CH₄. Second, this biological catalyst is continuously formed at the biogas plants at no additional cost. Third, the microbial community participating in the process is well organized and able to carry out the task under various environmental conditions very efficiently. Fourth, the process is easily manageable, and the microbial community flexibly tolerates the “turn-on” and “turn-off” situations. Fifth, the product is practically pure bioCH₄ needing no further purification. Sixth, the process also accomplishes a CO₂ sink and therefore directly contributes to CO₂ mitigation.

The biogas installations may therefore complement their current operation by becoming bioCH₄ producers and improve the economy of their technology without substantial additional investments.

The batch fermentation system

The total volume of the reactors was 160 mL (Wheaton glass serum bottle, Z114014 Aldrich). All the samples were run in 3 parallel biological replicates. The reactors routinely contained 40 mL inoculum from the mesophilic industrial biogas plant Zöldforrás Ltd., Szeged, Hungary. The main substrates at Zöldforrás are maize and sweet sorghum silage and pig manure in 80:20 total organic solid ratio. The inoculum was sieved on a 1 mm filter in order to remove the larger particles and was used without further treatment according to the VDI protocol [25]. In each set of experiments, three control reactors containing only the inoculum were included. The calculated amount of solid α-cellulose (C8002 Sigma) was added into the reactors when needed (Table 3). 0.3 g of α-cellulose was routinely added as substrate, as described in the VDI protocol [25]. The daily H₂ dosage was 0.81 ± 0.16 mmol, unless indicated otherwise. The reactors were sealed with butyl septa and aluminum crimps and were made anaerobic by N₂ gas exchange of the headspace (5 min). Following the daily gas composition analysis by gas chromatography (GC), the gas phases of the reactors were degassed by purging with N₂ (Messer nitrogen 4.5) for 5 min and the internal pressure was adjusted to atmospheric level. H₂ and CO₂ were injected manually and daily into the gas phase with disposable plastic syringes according to the experimental protocol (Table 3). The amount of the injected gas was verified by GC. The reactors were incubated in a rotary shaker at 37 °C.

Organic acid analysis

Samples for organic acid analysis were taken from the liquid phase of the reactors. The samples were centrifuged (13,000 rpm for 10 min,) and the supernatant was filtered through PES centrifugal filter (PES 516-0228, VWR) at 14,000 rpm for 20 min. The concentrations of volatile organic acids were measured with HPLC (Hitachi LaChrome Elite) equipped with refractive index detector L2490. The separation was performed on an ICSep ICE-COREGEL—64H column. The temperature of the column and detector was 50 and 41 °C, respectively. The eluent was 0.01 M H₂SO₄ (0.8 mL min⁻¹).

Gas composition analysis

The gas composition of the reactor headspace was measured every day by GC. The CH₄ and H₂ contents were determined with an Agilent 6890 N GC (Agilent Technologies) equipped with an HP Molesive 5 Å (30 m × 0.53 mm × 25 µm) column and a TCD detector. The temperature of the injector was 150 °C and application was made in split mode 0.2:1. The column temperature was maintained at 60 °C. The carrier gas was Linde HQ argon 5.0, with the flow rate set at 16.8 mL min⁻¹.

The amount of CO₂ was determined with a Shimadzu GC 2010 (Shimadzu Corporation) equipped with a TCD detector and a HP PlotQ (30 m × 0.5 mm × 40 µm) column. The chromatograph was applied in split injection mode (rate 0.5:1). The temperature of the inlet was 200 °C. The column and the detector temperature were maintained at 90 and 150 °C, respectively. The applied carrier gas was Messer nitrogen 4.5 at 8.4 mL min⁻¹. The samples were injected with the help of a gastight microsyringe (Hamilton). The conversion efficiency of H₂ to CH₄ was calculated by the modified theoretical equation [15].

where “A” is the experimental reactor and η = conversion efficiency of H₂ to CH₄ (%) \( {\text{r}}_{{CH_{4} A}} \) = CH₄ production of reactor A (mL L⁻¹ h⁻¹) \( {\text{r}}_{{CH_{4} B}} \) = CH₄ production of control reactor (mL L⁻¹ h⁻¹) \( {\text{r}}_{{H_{2} A}} \) = the added amount of H₂ to reactor A (mL L⁻¹ h⁻¹) \( {\text{r}}_{{H_{2} D}} \) = the residual amount of H₂ in reactor A (mL L⁻¹ h⁻¹).

oDM: The organic dry matter content was quantified by drying the biomass at 105 °C overnight and weighing the residue, giving the dry mass content. Further heating of this residue at 550 °C provided the organic dry matter (oDM) content.

pH: The value of the pH was measured with a Radelkis OP-211/2 equipped with an OP-0808P pH electrode immediately after the daily GC analysis.

CSTR:

continuous stirred tank reactor

P2B:

power to biomethane concept

Rpm:

revolution per minute

GC:

gas chromatograph

HPLC:

high-pressure liquid chromatography

TCD:

thermal conductivity detector

oDM:

organic dry matter content

MSz carried out most of the experiments and evaluated the data. NÁ and RT participated in the laboratory work and data handling. ZB took part in the experimental work and its design. AB contributed in developing the concept summarized in Fig. 10. GR participated in data interpretation. KLK conceived the study, participated in its design and compiled the manuscript. All the authors have read and approved the final manuscript.

Acknowledgements

The authors thank Ms. Netta Bozóki for technical assistance. The support and advices by Professor János Minárovits and Dean Kinga Turzó (Faculty of Dentistry, University of Szeged) are gratefully acknowledged. This work was supported by the domestic Grant GOP-1.1.1-11-2012-0128 and the EU Horizon 2020 research and innovation programme, BIOSURF project (contract number 646533).

Competing interests

The authors declare that they have no competing interests.

Funding

This work was supported by the domestic grant GOP-1.1.1-11-2012-0128 and the EU Horizon 2020 research and innovation programme, BIOSURF project (contract number 646533).

Affiliations

1. Department of Biotechnology, University of Szeged, Közép fasor 52, Szeged, 6726, Hungary

   • Márk Szuhaj
   • , Norbert Ács
   • , Attila Bodor
   • , Gábor Rákhely
   • , Kornél L. Kovács
   •  & Zoltán Bagi

2. Institute of Biochemistry, Biological Research Center, Hungarian Academy of Sciences, Temesvári krt. 62, Szeged, 6726, Hungary

   • Roland Tengölics

3. Department of Oral Biology and Experimental Dental Research, University of Szeged, Tisza L. krt. 64, Szeged, 6720, Hungary

   • Kornél L. Kovács

4. Institute of Biophysics, Biological Research Centre, Hungarian Academy of Sciences, Temesvári krt. 62, Szeged, 6726, Hungary

   • Gábor Rákhely
   •  & Kornél L. Kovács

Corresponding author

Correspondence to Kornél L. Kovács.

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