Electrofermentation of Lactic Acid Bacteria: History
Please note this is an old version of this entry, which may differ significantly from the current revision.
Contributor: , , ,

Microbial electrosynthesis is the process of supplying electrons to microorganisms to reduce CO2 and yield industrially relevant products. Such systems are limited by their requirement for high currents, resulting in challenges to cell survival. Electrofermentation is an electron-efficient form of microbial electrosynthesis in which a small cathodic or anodic current is provided to a culture to alter the oxidation–reduction potential of the medium and, in turn, alter microbial metabolism. This approach has been successfully utilised to increase yields of diverse products including biogas, butanediol and lactate. Biomass conversion to lactate is frequently facilitated by ensiling plant biomass with homofermentative lactic acid bacteria. Although most commonly used as a preservative in ensiled animal feed, lactate has diverse industrial applications as a precursor for the production of probiotics, biofuels, bioplastics and platform chemicals. Lactate yields by lactic acid bacteria (LAB) are constrained by a number of redox limitations which must be overcome while maintaining profitability and sustainability. 

  • lactic acid bacteria
  • LAB
  • electrofermentation
  • ensiling

1. History of Microbial Electrosynthesis

The earliest exploration of microbial electrosynthesis was the use of microbial fuel cells (MFCs) to produce electricity from the microbial oxidation of organic matter, which aimed to scavenge and transfer free electrons produced by electrogenic microbes to an anode. Although small, the resulting currents could theoretically be applied in contexts such as renewable electric car batteries if the yield could be enhanced, the cost decreased and dangerous by-products suppressed [1]. The first discovery of the microbial generation of electrical currents occurred in 1912 in Saccharomyces cerevisiae, but it attracted limited interest. Early research on MFCs suggested that mediators, or electron shuttles, between cell and anode were necessary for current biogenesis but this was later deemed unnecessary if using microorganisms with trans-membrane, redox-active proteins. Since then, the potential of MFCs for remote- and low-power-usage applications has been investigated, such as in conservation monitoring [2], as has the idea of using MFCs for larger-scale power generation. Translation from laboratory-scale experiments to 1–1000 L-scale pilot plants has led to the identification of issues with current generation due to decreased chemical oxygen demand [3] and difficulties in the construction of larger-volume reactors [4]. Utilising smaller interlinked stacked MFCs has been shown to reliably provide power density of 150–200 mA/m3 [5]; however, this technology has still not been adopted at any widespread scale [6].
Microbial electrolysis cells (MECs) are essentially the converse of MFCs; an electrical current is supplied to a culture of electroactive microbes, with the goal of producing H2 at the cathode [7]. While H2 evolution from most typical metabolic end-products is thermodynamically unfavourable (positive ΔG), the addition of energy to the system via a current allows these unfavourable reactions to proceed [7]. In MECs, some of the electrical potential required for the reduction of protons to H2 is supplied by the electrogenic microorganisms, so the input of power required is lower than that of hydrolysis (generally 0.2–1 V) compared to the 2 V required for alkaline water hydrolysis [8]. The increasing use of H2 as a direct fossil fuel replacement and the possibilities offered by using wastewater as a low-cost carbon source for the generation of H2 have therefore attracted renewed interest in MECs [7].

2. Scalability of Bioelectrochemical Systems: Lessons from MECs

Although it is industrially promising at the laboratory scale, EF research is still in its infancy. Despite overcoming some of the fundamental limitations of MESs, its scalability has not been extensively tested. Scalable setups are essential to assessing the industrial favourability of a process and will therefore be a major challenge in the future of EF research. Currently, the most relevant pilot-scale experiments have been carried out in wastewater treatment MECs which produce H2 or methane. Understanding the relevant pitfalls of moving up from the laboratory scale can help to predict the most industrially realistic conditions for a potential future EF scale-up.
Certain limitations of MECs which arise during pilot-scale experiments may be overcome by inherent features of EF, while others may continue to present issues. Relevant pitfalls of the scale-up are derived from oxygen requirements, the complexity of natural communities and substrates, and size-related system overpotentials. Aeration is a major cost associated with traditional wastewater treatments [9]. At scale, ensuring sufficient aeration can complicate reactor design and often impacts space needs and efficiency in traditional wastewater treatment plants [9][10]. MECs theoretically operate under anaerobic conditions. This hypothetically removes the requirement for aeration, making the technique industrially promising. However, natural microbial communities present in wastewater often include methanogens whose growth thrives under these reducing anaerobic conditions [11]. These organisms metabolise H2 (the intended product) to produce methane, reducing reactor productivity. The laboratory solution to this problem is to expose cultures to oxygen between batches to suppress this [12]. However, pilot-scale MECs must operate with continuous flow, rather than in batches, invalidating this approach [13][14]. In multiple pilot-scale MECs using wastewater, product rerouting towards methanogenesis led to methane being the dominant gaseous product [13][15], ultimately leading the focus of MECs to shift from H2 to biogas production as adding an inhibitor is economically unfeasible [16]. This case highlights a complication arising both from the scale-up from synthetic to natural substrate and its associated microbes, as well as from scaling operations from batch to continuous flow. The diversity of natural microbial communities is a consistent challenge in MEC scale-up, resulting in unpredictable synergy between species and metabolic bottlenecks [17].
The changes from synthetic to natural substrates and communities, and from batch to continuous flow when shifting from laboratory to pilot scale, are important considerations in designing EF systems for initial prospecting. Laboratory experiments with natural communities are difficult to design; however, there are natural fermentative contexts in which single metabolic groups of organisms dominate (e.g., lactic acid bacteria in silage). Where this is the case in the testing of EF with intended application in contexts such as ensilage, the challenges associated with avoiding certain metabolic outputs may be minimised. As EF fundamentally varies from MEC in that its intended product is not H2, hydrogenotrophic methanogenesis is not a primary concern. Finally, EF has been successfully tested in semi-continuous flow reactors, supporting that the shift from batch to continuous flow reactors is less likely to present a major issue in EF compared to MECs [18].
Bioelectrochemical systems need to compensate for the cost of electrons, so the lower the current for the desired effect, the better. Pilot-scale MECs highlight a further consideration: system overpotentials, which scale with the size of the reactor. As reactor size increases, the difference between the thermodynamically determined reduction potential and the experimental reduction potential also increases [19]. Essentially, the voltage required to supply the energy for the desired reaction to proceed increases. For example, in a pilot-scale, batch-operated MEC using urban wastewater, a potential loss of 0.5 V was observed, requiring the applied voltage to be increased accordingly [20]. Overpotentials increase the cost of MEC operation, and the potential losses will increase as these systems are scaled up further [21]. EF may suffer less from these potential losses than MECs, as one of the key features of EF is that it is very electron-efficient, requiring low potentials to alter the redox potential of the medium rather than being used to directly driving reactions [22].

3. LAB Ensiling and the Potential for Efficiency Gains

The presence of LAB in plant microbiota is essential for the preservation of food and crop nutrients during fermentation [23]. The fermentative preservation of foodstuffs has been a dietary staple for millennia. In the 19th century, this method was scaled to the preservation of seasonal crops for year-round animal feed provision through ensiling. This method was further developed in the 20th century, with innovations by Artturi Virtanen earning him the Nobel Prize in Chemistry in 1945 [24]. More recently, this method has been co-opted in biorefineries, both to reduce biomass loss during feedstock preservation and to generate large volumes of platform chemicals. Homofermentative LAB are therefore vital components of traditional ensiling and the renewable chemical industry, but their metabolism has biochemical limits which are the targets of yield improvement research.
Homofermentative LAB are central to the traditional ensiling process, converting soluble crop carbohydrates to lactic acid, thereby reducing pH and preventing the growth of spoilage microbes such as Clostridium species which convert both free sugar and lactic acid into butyric acid [25]. Additionally, Clostridia are often proteolytic and can convert crop proteins and amino acids into ammoniacal compounds, preventing crop preservation [26]. However, spoilage organisms typically grow above pH 5, while the endpoints of LAB fermentations are usually lower than pH 4 [27]. Successful ensiling therefore requires the rapid establishment of a sufficiently low pH to suppress the establishment, growth and metabolism of spoilage microbes. Homofermentative LAB are well suited to this application due to the large volumes of lactate and relatively small volumes of alternative products produced by their metabolism, which results in rapid pH decrease [28]. Their aerotolerance is also beneficial as this property can improve the aerobic stability of the silage after the opening of silos for feeding [29].
Ensiling has expanded beyond its initial use in animal feedstocks to allow year-round production in green biorefineries that aim to completely convert biomass into a variety of industrially valuable products, including platform chemicals, biofuels, biogas, animal feed pellets and fertiliser [23]. The biorefinery process is intended to mimic that of an oil refinery. Fresh or ensiled wet biomass is separated into solid and liquid fractions through pressing. The solid fraction consists of a press cake which contains lignocellulosic matter (LCM). As is, LCM can either be wrapped and ensiled again to produce animal feed, dried to produce feed pellets, or used to produce fibrous building materials such as insulation [23]. Ideally, however, LCM could be used to generate higher-value products such as biofuels or biogas. LAB are ideal biocatalysts for these processes as organic acids such as lactate can accelerate biogas production at the acetogenesis stage of methane production [30] and, as discussed previously, can also act as an intermediate in biofuel production [31]. The major challenge to this process is the breakdown of LCM, for which an efficient and cost-effective solution has not yet been found. Physico-chemical pre-treatment often produces toxic phenolic compounds, enzymatic breakdown is subject to end-product inhibition, syntrophic co-culture with cellulolytic organisms is high-maintenance at industrial scale, and heterologous cellulase expression in LAB has toxicity issues as well as limited secretion capacity [32].
Currently, higher-value products mainly come from the “green juice” fraction of fresh, plant biomass. In this fraction, which is rich in easily fermentable water-soluble carbohydrates and proteins, it is important to ensure preservation quickly for the same reasons as in ensiling. LAB have multiple functions in this case: to prevent contamination, produce lactic acid, recover proteins through biomass integration, or precipitate proteins through the subsequent pH decrease [33]. The resulting liquid can be separated into a protein concentrate, which can be used as feed supplement, and a “brown juice” which can be treated and used as a further fermentation broth, fed into biogas reactors or used as a fertiliser [23]. At present, many biorefineries focus on the production of polylactic acid, with outputs such as 140,000 tons of bioplastics per year [33]. Yields of PLA would, of course, be amplified if lignocellulosic waste could be efficiently broken down and converted to lactate, but this should also be complemented by attempting to increase yields by increasing the production of lactic acid from soluble carbohydrates.
There are a number of biochemical limitations which govern the conversion efficiency of sugars into lactic acid. As mentioned previously, mixed-acid fermentation can occur in homofermentative LAB under certain pH and temperature conditions, as well as conditions of low glycolytic flux [34]. Mixed- acid fermentation is a detrimental process when lactic acid is the desired output, but in the context of industrial feedstocks, it is common. Mixed-acid production is catalysed by pyruvate formate lyase, and the activity of this enzyme is allosterically and transcriptomically regulated by oxygen, glycolytic flux, pH and temperature [26][35]. Bioreactor conditions must therefore be carefully monitored to prevent significant redirection of pyruvate to acetic and formic acid through this pathway. Natural feedstocks are inherently limiting in this sense, as non-glucose sugars such as galactose can alter glycolytic flux, reducing the inhibition of PFL by GAP and DHAP and resulting in up to 27% of pyruvate being redirected towards formate, ethanol and acetate [35]. The highest outputs of lactic acid are usually found with pure starting sugars, which is irreconcilable with unrefined feedstocks. High glucose metabolism can also inhibit the metabolism of other sugars into lactic acid through catabolite repression [36].
Redox balance is also a factor which constrains LA production by LAB. Due to their obligately anaerobic lifestyle, LAB metabolism is governed by the presence of different (non-O2) electron acceptors and the redox state of the cell [37]. During homolactic fermentation, the energy-production stage occurs via the oxidation of carbohydrates to pyruvate [38]. The subsequent production of lactic acid from pyruvate is carried out to offset the redox debt generated during sugar oxidation, where NAD+ is consumed. Pyruvate is the primary electron acceptor in LAB, being reduced to form lactate, and in turn allowing NAD+ regeneration [37]. Other electron acceptors can also be used, varying based on species differences and environmental availability. Some species generate pyruvate without this NAD+ redox debt through citrate utilisation.
One obvious consequence of metabolism whose endpoint is a large volume of acid is the constant pH decrease in an enclosed reactor. Although LAB are largely acid-tolerant, most grow optimally between pH 4–6, while LA fermentations can drive reactor pH below 4 [26]. This is part of the reason why ensiling techniques are so successful for biomass preservation. To this end, LA-producing bioreactors currently require the addition of buffering materials such as calcium carbonate to allow continued growth and LA production. This has a number of downsides. Many dissociated (lactate-) forms of lactic acid, such as calcium lactate and ammonium lactate, which are produced when buffering agents are added, still have significant inhibitory effects [39]. Additionally, these compounds add cost to the industrial process, which needs to be low (< USD 0.8 per kg LA for polylactic acid) for LA production to be profitable [32]. Various chemical, synthetic biology and experimental evolution methods have been utilised to improve acid tolerance in LAB [40]. These methods have mostly been in the context of improving the probiotic properties of LAB. Ultimately, on a green biorefinery scale, highly productive mutant strains or constant LA removal/neutralisation are needed to allow the optimal growth-promoting and contamination-preventing conditions around pH 4–5, posing a challenge to the maintained profitability of the industry.
Although lactic acid metabolism is central to traditional ensiling, as well as the biofuel and biomaterial refinery industries, their potential in the latter is largely conceptual at this point. Key issues to be addressed in the coming decade are the breakdown of lignocellulosic material into usable sugars, limiting mixed-acid product formation, and maintaining a constant productive pH to maximise lactic acid production. These will require a combination of synthetic biology and bioprocess engineering to maximise the productivity and maintain the profitability of green bioreactors over fossil-fuel-based refineries.

4. Electrofermentation in Ensiling for Increased Platform-Chemical Yields from Biomass

LAB produce 90% of lactic acid globally [34]. Key factors affecting the scale-up of LA production for green biorefinery applications are the selection of appropriate biocatalysts, bioreactor conditions and the maintenance and suppression of other metabolic pathways. With the decreasing costs of sustainable electricity globally [41], the potential has arisen to integrate bioelectrochemical approaches into bioreactors to both guide biocatalyst metabolism and maintain optimal reactor conditions. As discussed previously, EF is a cost- and electron-efficient technique which can increase metabolic outputs many-fold through changes to the redox potential of the growth media [42][43], making EF an attractive candidate for increasing LA yield by LAB. Additionally, a major challenge for LA-based biorefineries is maintaining high productivity with constant acidification of the media. Cathodic electrofermentation approaches can increase the pH of the media, possibly presenting a solution to this issue. Ultimately, EF should be tested in industrial media over longer timeframes, combined with synthetic biology approaches, and tested in single-electrode reactors to optimise use in LA fermentations. In order to allow this potential to be realised, EF must first be tested in pure LAB cultures to identify the biological and economic viability and potential metabolic targets of the approach.
EF has never been tested in pure cultures of LAB, but its effects on lactic acid production have been observed in a few mixed-culture systems such as wastewater extractions. These studies demonstrated increased lactic acid production in cathodic EF treatments [44] but struggled to differentiate between community selection effects and biochemical mechanisms. Anodic EF experiments often focus on aerobic metabolism with the aim of supplying a non-O2 renewable terminal electron acceptor, but the potential to enhance the production of more oxidised products in anaerobic metabolism has also been proposed [45]. This mechanism has been explored in glycerol fermentations by Escherichia coli and Propionibacterium freudenreichii, where anodic potentials drive the production of more oxidised products [46][47]. In contrast to lactate-specific studies, which are rare, anodic and cathodic EF have been used extensively to alter the production of butanol, ethanol, 1,3-propanediol, glutamate, acetic acid and butyric acid [43], mostly in Gram-negative or mixed-culture study systems [45]. Given that LAB may play a large role in the carboxylate platform especially [48], this potential bioelectrochemical enhancement should be investigated.
An important limitation to exploring EF in LAB is their lack of natural electroactivity [1]. However, microbial electroactivity is not essential to the success of bioelectrochemical systems when mediators are present. Even the original demonstration of electrogenic microbial activity in 1910 used S. cerevisiae and E. coli, which are not considered electroactive but may have benefited from the presence of flavins and B-vitamins in the media, which potentially acted as redox shuttles [1]. As previously mentioned, mediators are molecules which can be reduced at the electrode and shuttle electrons into cells. They can be present in low amounts in media which contains yeast extract, such as the MRS media used for the lab cultures of many LAB. However, it is generally believed that mediators must either be produced by the microbes or added to the fermentation mix separately. Examples of weak mediators produced by fermentative metabolism include formate and acetate [22], both of which are produced via mixed acid fermentation in homofermentative LAB. The ability of heterofermentative products to act as mediators may be of value in avoiding the need for the addition of expensive external mediators; however, in bioreactors where product purity is highly desired, these benefits may be outweighed by reduced yields of the desired output, or the cost of additional purification or product separation. While artificial mediator addition theoretically makes EF possible in a more diverse range of microbial species, the added cost constrains industrial applications. This is especially true when mediators cannot be recycled and need to be continuously added, as is the case for the commonly utilised mediator methyl viologen. Neutral red is a recyclable alternative which has risen in popularity in recent years [49]. It is also worth noting, however, that in many mixed-culture electrofermentations, cathodic biofilms are highly diverse and not usually dominated by electroactive species [50]. This may point to electron shuttles in the media playing a larger role than previously thought.
Although LAB have a variety of industrial applications, the most immediately feasible combination with EF would be deriving platform chemicals from soluble carbohydrates in organic material, including silage. In this context, lactate is currently the main compound of interest. To this end, types of LAB should be selected which are naturally successful in ensiling environments, both as potential inoculants and as a part of the natural microbiome. Success can be measured by high lactate titres, high acid and other stress tolerance, and competitive traits. Morphology may contribute to the success of bioelectrochemical systems [51], and to this end, species should be selected to span the morphological diversity of LAB, including homo- and heterofermentative species, bacilli, coccobacilli and cocci. Other potentially interesting features could be genetic tractability to widen the scope of metabolites which could be produced, and possibly also the applicability of the species to different lactofermenting industries such as food or medicine.

This entry is adapted from the peer-reviewed paper 10.3390/en15228638

References

  1. Logan, B.E.; Hamelers, B.; Rozendal, R.; Schröder, U.; Keller, J.; Freguia, S.; Aelterman, P.; Verstraete, W.; Rabaey, K. Microbial Fuel Cells: Methodology and Technology. Environ. Sci. Technol. 2006, 40, 5181–5192.
  2. Knight, C.; Cavanagh, K.; Munnings, C.; Moore, T.; Cheng, K.Y.; Kaksonen, A.H. Application of microbial fuel cells to power sensor networks for ecological monitoring. In Wireless Sensor Networks and Ecological Monitoring; Springer: Berlin/Heidelberg, Germany, 2013; pp. 151–178.
  3. Erable, B.; Etcheverry, L.; Bergel, A. From microbial fuel cell (MFC) to microbial electrochemical snorkel (MES): Maximizing chemical oxygen demand (COD) removal from wastewater. Biofouling 2011, 27, 319–326.
  4. Blatter, M.; Furrer, C.; Cachelin, C.P.; Fischer, F. Phosphorus, chemical base and other renewables from wastewater with three 168-L microbial electrolysis cells and other unit operations. Chem. Eng. J. 2020, 390, 124502.
  5. Blatter, M.; Delabays, L.; Furrer, C.; Huguenin, G.; Cachelin, C.P.; Fischer, F. Stretched 1000-L microbial fuel cell. J. Power Sources 2021, 483, 229130.
  6. Janicek, A.; Fan, Y.; Liu, H. Design of microbial fuel cells for practical application: A review and analysis of scale-up studies. Biofuels 2014, 5, 79–92.
  7. Logan, B.E.; Call, D.; Cheng, S.; Hamelers, H.V.M.; Sleutels, T.H.J.A.; Jeremiasse, A.W.; Rozendal, R.A. Microbial Electrolysis Cells for High Yield Hydrogen Gas Production from Organic Matter. Environ. Sci. Technol. 2008, 42, 8630–8640.
  8. Lu, L.; Hou, D.; Fang, Y.; Huang, Y.; Ren, Z.J. Nickel based catalysts for highly efficient H2 evolution from wastewater in microbial electrolysis cells. Electrochim. Acta 2016, 206, 381–387.
  9. Drewnowski, J.; Remiszewska-Skwarek, A.; Duda, S.; Łagód, G. Aeration Process in Bioreactors as the Main Energy Consumer in a Wastewater Treatment Plant. Review of Solutions and Methods of Process Optimization. Processes 2019, 7, 311.
  10. Leicester, D.; Amezaga, J.; Heidrich, E. Is bioelectrochemical energy production from wastewater a reality? Identifying and standardising the progress made in scaling up microbial electrolysis cells. Renew. Sustain. Energy Rev. 2020, 133, 110279.
  11. Vítězová, M.; Kohoutová, A.; Vítěz, T.; Hanišáková, N.; Kushkevych, I. Methanogenic Microorganisms in Industrial Wastewater Anaerobic Treatment. Processes 2020, 8, 1546.
  12. Call, D.; Logan, B.E. Hydrogen Production in a Single Chamber Microbial Electrolysis Cell Lacking a Membrane. Environ. Sci. Technol. 2008, 42, 3401–3406.
  13. Cusick, R.D.; Bryan, B.; Parker, D.S.; Merrill, M.D.; Mehanna, M.; Kiely, P.D.; Liu, G.; Logan, B.E. Performance of a pilot-scale continuous flow microbial electrolysis cell fed winery wastewater. Appl. Microbiol. Biotechnol. 2011, 89, 2053–2063.
  14. Rader, G.K.; Logan, B.E. Multi-electrode continuous flow microbial electrolysis cell for biogas production from acetate. Int. J. Hydrog. Energy 2010, 35, 8848–8854.
  15. Carmona-Martínez, A.A.; Trably, E.; Milferstedt, K.; Lacroix, R.; Etcheverry, L.; Bernet, N. Long-term continuous production of H2 in a microbial electrolysis cell (MEC) treating saline wastewater. Water Res. 2015, 81, 149–156.
  16. Sun, R.; Zhou, A.; Jia, J.; Liang, Q.; Liu, Q.; Xing, D.; Ren, N. Characterization of methane production and microbial community shifts during waste activated sludge degradation in microbial electrolysis cells. Bioresour. Technol. 2015, 175, 68–74.
  17. Ferrera, I.; Sánchez, O. Insights into microbial diversity in wastewater treatment systems: How far have we come? Biotechnol. Adv. 2016, 34, 790–802.
  18. Ma, H.; Wu, W.; Yu, Z.; Zhao, J.; Fu, P.; Xia, C.; Lam, S.S.; Wang, Q.; Gao, M. Medium-chain fatty acid production from Chinese liquor brewing yellow water by electro-fermentation: Division of fermentation process and segmented electrical stimulation. Bioresour. Technol. 2022, 360, 127510.
  19. Clauwaert, P.; Aelterman, P.; Pham, T.H.; De Schamphelaire, L.; Carballa, M.; Rabaey, K.; Verstraete, W. Minimizing losses in bio-electrochemical systems: The road to applications. Appl. Microbiol. Biotechnol. 2008, 79, 901–913.
  20. Baeza, J.A.; Martínez-Miró, À.; Guerrero, J.; Ruiz, Y.; Guisasola, A. Bioelectrochemical hydrogen production from urban wastewater on a pilot scale. J. Power Sources 2017, 356, 500–509.
  21. Slate, A.J.; Whitehead, K.A.; Brownson, D.A.C.; Banks, C.E. Microbial fuel cells: An overview of current technology. Renew. Sustain. Energy Rev. 2019, 101, 60–81.
  22. Moscoviz, R.; Toledo-Alarcón, J.; Trably, E.; Bernet, N. Electro-Fermentation: How To Drive Fermentation Using Electrochemical Systems. Trends Biotechnol. 2016, 34, 856–865.
  23. Lübeck, M.; Lübeck, P.S. Application of lactic acid bacteria in green biorefineries. FEMS Microbiol. Lett. 2019, 366, fnz024.
  24. NobelPrize. The Nobel Prize in Chemistry 1945. Nobel Prize Outreach AB. 2022. Available online: https://www.nobelprize.org/prizes/chemistry/1945/summary/ (accessed on 13 October 2022).
  25. Guo, X.S.; Ke, W.C.; Ding, W.R.; Ding, L.M.; Xu, D.M.; Wang, W.W.; Zhang, P.; Yang, F.Y. Profiling of metabolome and bacterial community dynamics in ensiled Medicago sativa inoculated without or with Lactobacillus plantarum or Lactobacillus buchneri. Sci. Rep. 2018, 8, 357.
  26. Fitzsimons, A.; Duffner, F.; Curtin, D.; Brophy, G.; O’Kiely, P.; O’Connell, M. Assessment of Pediococcus acidilactici as a Potential Silage Inoculant. Appl. Environ. Microbiol. 1992, 58, 3047–3052.
  27. Vargas-Ramirez, J.M.; Haagenson, D.M.; Pryor, S.W.; Wiesenborn, D.P. Beet tissue ensiling: An alternative for long-term storage of sugars in industrial beets for nonfood use. Biomass Bioenergy 2016, 85, 135–143.
  28. Schmidt, J.; Sipocz, J.; Kaszás, I.; Szakács, G.; Gyepes, A.; Tengerdy, R.P. Preservation of sugar content in ensiled sweet sorghum. Bioresour. Technol. 1997, 60, 9–13.
  29. Danner, H.; Holzer, M.; Mayrhuber, E.; Braun, R. Acetic Acid Increases Stability of Silage under Aerobic Conditions. Appl. Environ. Microbiol. 2003, 69, 562–567.
  30. Klang, J.; Theuerl, S.; Szewzyk, U.; Huth, M.; Tölle, R.; Klocke, M. Dynamic variation of the microbial community structure during the long-time mono-fermentation of maize and sugar beet silage. Microb. Biotechnol. 2015, 8, 764–775.
  31. Petrus, L.; Noordermeer, M.A. Biomass to biofuels, a chemical perspective. Green Chem. 2006, 8, 861–867.
  32. Tarraran, L.; Mazzoli, R. Alternative strategies for lignocellulose fermentation through lactic acid bacteria: The state of the art and perspectives. FEMS Microbiol. Lett. 2018, 365, fny126.
  33. Kamm, B.; Schönicke, P.; Hille, C. Green biorefinery—Industrial implementation. Food Chem. 2016, 197, 1341–1345.
  34. Mazzoli, R.; Bosco, F.; Mizrahi, I.; Bayer, E.A.; Pessione, E. Towards lactic acid bacteria-based biorefineries. Biotechnol. Adv. 2014, 32, 1216–1236.
  35. Melchiorsen, C.V.K.J.R. The level of pyruvate-formate lyase controls the shift from homolactic to mixed-acid product formation in Lactococcus lactis. Appl. Microbiol. Biotechnol. 2002, 58, 338–344.
  36. Yun, J.-S.; Ryu, H.-W. Lactic acid production and carbon catabolite repression from single and mixed sugars using Enterococcus faecalis RKY1. Process. Biochem. 2001, 37, 235–240.
  37. Hansen, E.B. Redox reactions in food fermentations. Curr. Opin. Food Sci. 2018, 19, 98–103.
  38. Gänzle, M.G. Lactic metabolism revisited: Metabolism of lactic acid bacteria in food fermentations and food spoilage. Curr. Opin. Food Sci. 2015, 2, 106–117.
  39. Hetényi, K.; Németh, Á.; Sevella, B. Role of pH-regulation in lactic acid fermentation: Second steps in a process improvement. Chem. Eng. Process. Process. Intensif. 2011, 50, 293–299.
  40. Wang, C.; Cui, Y.; Qu, X. Mechanisms and improvement of acid resistance in lactic acid bacteria. Arch. Microbiol. 2018, 200, 195–201.
  41. International Renewable Energy Agency IRENA. Renewable Power Generation Costs in 2019; International Renewable Energy Agency IRENA: Abu Dhabi, United Arab Emirates, 2020.
  42. Choi, O.; Kim, T.; Woo, H.M.; Um, Y. Electricity-driven metabolic shift through direct electron uptake by electroactive heterotroph Clostridium pasteurianum. Sci. Rep. 2014, 4, 6961.
  43. Schievano, A.; Pepé Sciarria, T.; Vanbroekhoven, K.; De Wever, H.; Puig, S.; Andersen, S.J.; Rabaey, K.; Pant, D. Electro-Fermentation—Merging Electrochemistry with Fermentation in Industrial Applications. Trends Biotechnol. 2016, 34, 866–878.
  44. Xue, G.; Lai, S.; Li, X.; Zhang, W.; You, J.; Chen, H.; Qian, Y.; Gao, P.; Liu, Z.; Liu, Y. Efficient bioconversion of organic wastes to high optical activity of l-lactic acid stimulated by cathode in mixed microbial consortium. Water Res. 2018, 131, 1–10.
  45. Vassilev, I.; Gießelmann, G.; Schwechheimer, S.K.; Wittmann, C.; Virdis, B.; Krömer, J.O. Anodic electro-fermentation: Anaerobic production of L-Lysine by recombinant Corynebacterium glutamicum. Biotechnol. Bioeng. 2018, 115, 1499–1508.
  46. Emde, R.; Schink, B. Oxidation of glycerol, lactate, and propionate by Propionibacterium freudenreichii in a poised-potential amperometric culture system. Arch. Microbiol. 1990, 153, 506–512.
  47. Emde, R.; Swain, A.; Schink, B. Anaerobic oxidation of glycerol by Escherichia coli in an amperometric poised-potential culture system. Appl. Microbiol. Biotechnol. 1989, 32, 170–175.
  48. Agler, M.T.; Wrenn, B.A.; Zinder, S.H.; Angenent, L.T. Waste to bioproduct conversion with undefined mixed cultures: The carboxylate platform. Trends Biotechnol. 2011, 29, 70–78.
  49. Choi, O.; Um, Y.; Sang, B.-I. Butyrate production enhancement by Clostridium tyrobutyricum using electron mediators and a cathodic electron donor. Biotechnol. Bioeng. 2012, 109, 2494–2502.
  50. Wrighton, K.C.; Virdis, B.; Clauwaert, P.; Read, S.T.; Daly, R.A.; Boon, N.; Piceno, Y.; Andersen, G.L.; Coates, J.D.; Rabaey, K. Bacterial community structure corresponds to performance during cathodic nitrate reduction. ISME J. 2010, 4, 1443–1455.
  51. Jia, X.; Li, M.; Wang, Y.; Wu, Y.; Zhu, L.; Wang, X.; Zhao, Y. Enhancement of hydrogen production and energy recovery through electro-fermentation from the dark fermentation effluent of food waste. Environ. Sci. Ecotechnol. 2020, 1, 100006.
More
This entry is offline, you can click here to edit this entry!