Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1 -- 5498 2022-06-29 19:42:01 |
2 layout + 34 word(s) 5532 2022-06-30 03:52:48 | |
3 layout + 2 word(s) 5534 2022-06-30 03:54:14 | |
4 layout Meta information modification 5534 2022-06-30 03:56:35 | |
5 a brief description of biomethanation and hydrogenotrophic methanation has been added + 8 word(s) 5542 2022-06-30 10:16:21 | |
6 Format Correct Meta information modification 5542 2022-07-04 07:46:32 | |
7 Format Correct Meta information modification 5542 2022-07-11 02:09:39 | |
8 layout Meta information modification 5542 2022-07-11 04:05:14 | |
9 layout + 3 word(s) 5545 2022-07-15 10:26:04 |

Video Upload Options

Do you have a full video?


Are you sure to Delete?
If you have any further questions, please contact Encyclopedia Editorial Office.
Bellini, R.;  Bassani, I.;  Vizzarro, A.;  Azim, A.A.;  Vasile, N.S.;  Pirri, C.F.;  Verga, F.;  Menin, B. Current Advancements in Biomethanation of H2 and CO2. Encyclopedia. Available online: (accessed on 19 June 2024).
Bellini R,  Bassani I,  Vizzarro A,  Azim AA,  Vasile NS,  Pirri CF, et al. Current Advancements in Biomethanation of H2 and CO2. Encyclopedia. Available at: Accessed June 19, 2024.
Bellini, Ruggero, Ilaria Bassani, Arianna Vizzarro, Annalisa Abdel Azim, Nicolò Santi Vasile, Candido Fabrizio Pirri, Francesca Verga, Barbara Menin. "Current Advancements in Biomethanation of H2 and CO2" Encyclopedia, (accessed June 19, 2024).
Bellini, R.,  Bassani, I.,  Vizzarro, A.,  Azim, A.A.,  Vasile, N.S.,  Pirri, C.F.,  Verga, F., & Menin, B. (2022, June 30). Current Advancements in Biomethanation of H2 and CO2. In Encyclopedia.
Bellini, Ruggero, et al. "Current Advancements in Biomethanation of H2 and CO2." Encyclopedia. Web. 30 June, 2022.
Current Advancements in Biomethanation of H2 and CO2

Sustainable and renewable energy production is a global priority. Over the past decade, several Power-to-X (PtX) technologies have been proposed to store and convert the surplus of renewable energies into chemical bonds of chemicals produced by different processes. CO2 is a major contributor to climate change, yet it is also an undervalued source of carbon that could be recycled and represents an opportunity to generate renewable energy. 

Biomethanation CO2 H2 Power to X - PtX CO2 valorization hydrogenotrophic methanation

1. Biomethanation of H2 and CO2

Biological conversion of CO2 and H2 into CH4, commonly referred to as biomethanation Equation (1) has gained a lot of attention and has been widely investigated in the last 10 years.

4   H 2 ( g a s ) + C O 2 ( g a s )   C H 4 ( g a s ) + 2 H 2 O ( l i q u i d )         135.6     Δ G o     ( k J r e a c t i o n )  

Indeed this technology addresses both CO2 valorization, through its conversion into CH4, and storage of energy surplus generated by renewable sources (i.e. wind and solar power) into chemicals. Considering the H2 requirement for the reaction, the surplus of wind and solar power would be used to generate the needed H2 through water electrolysis. Although H2 could play a valuable role as clean fuel, its energy content is considerably lower (10.88 MJ/m3) than that of CH4 (36 MJ/m3) [1]. Hydrogenotrophic methanogens (HM) are the key organisms for the conversion of H2 and CO2 into CH4, and most of the knowledge regarding the hydrogenotrophic metabolism comes from pure culture studies of two major thermophilic strains Methanotermobacter thermoautotrophicum and Methanothermobacter marburgensis [2].

2. Development of In Situ and Ex Situ Strategies for Biomethanation

In the last decade, biological carbon fixation emerged as a promising technology aiming at removing and transforming CO2 and reducing GHG emissions while operating in mild temperature and pressure conditions without using chemicals, thus offering a remarkable advantage compared to traditional physical-chemical technologies for CO2 capture  [3][4][5]. Moreover, in addition to capturing CO2, this technology can also convert it into a valuable product, such as CH4. Biomethanation is an extremely versatile technology leading to the conversion of different gaseous substrates, such as rich gaseous streams of CO2, CO and H2 generated by different processes (e.g., anaerobic digestion (AD) of organic matter to biogas, thermochemical gasification of non-fermentable biomass to syngas, and natural or industrial processes). Biogas used as substrate for biomethanation can be obtained from numerous organic-matter-rich, non-food-related feedstocks, including the organic fraction of municipal solid waste, sewage sludge from wastewater treatment plants, manure from livestock, energy crops, organic industrial and commercial wastes and waste and sewage from agriculture [6]. Biogas is typically composed of mainly CH4 (50–70%) and CO2 (50–30%), while containing only residual amounts of other undesired compounds, such as N2, O2, H2S and NH3. The calorific value of CH4 stands at 36 MJ/m3-CH4; CO2 and residual compounds present in biogas lowers it to ~20 MJ/m3-biogas [7].
Similarly, syngas derived from the thermochemical gasification of lignocellulosic residues, non-fermentable by-products of bio-refineries and organic municipal wastes contains CO, CO2, CH4 and H2 in variable concentrations [7][8]. For these reasons, the biomethanation of biogas and syngas to higher CH4 content increases their calorific values, thus broadening their spectrum of potential applications and paving the way to the use of biomethane as an alternative to NG  [7][9].
As discussed in the previous section, HM requires H2 as electron donor to reduce CO2 to CH4. The use of renewable energies, such as solar and wind, is expanding worldwide, and variable weather conditions may result in an uneven distribution of energy production. Thus, the surplus of electricity can be used to hydrolyze water for the production of green H2 [10]. Alternatively, H2 can be obtained from biomass gasification, biological H2 production or residual unconverted H2 from biomethanation process [11]
Nevertheless, because of its low volumetric energy content, H2 poses some challenges related to storage and distribution [12][13]. For these reasons, the utilization of H2 generated from surplus electricity as electron donor to reduce the CO2 derived from biogas, syngas or other industrial processes offers a valuable solution to produce a clean and cheap energy carrier such as biomethane while reducing atmospheric CO2 emissions.

2.1. In Situ Biomethanation

In situ biomethanation can be achieved by injecting H2 derived from external sources directly inside a biogas reactor. Along with the CO2 produced during AD in the biogas reactor, H2 is converted into CH4 by the activity of indigenous HM (Figure 1a)  [14][15].
Figure 1. Scheme representing in situ (a) and ex situ (b) biomethanation technologies.
During biomethanation, the injection of additional H2 results in a selective pressure, leading to radical changes in the relative abundance and richness of the different microbial taxa characterizing the microbial consortium responsible for the process. Previous studies utilized comparative bioinformatic tools to elucidate the effect of the H2 on complex communities, pointing out a decrease in species involved in the fermentation and hydrolysis together with acetoclastic methanogens (AM) and a concomitant increase in the relative abundance of HM and syntrophic bacteria (e.g., SAOB or homoacetogens) [16]. For this reason, biomethanation communities are typically characterized by a very low diversity, with few main genera involved in CH4 production, i.e., MethanoculleusMethanothermobacter and Methanosarcina. Nevertheless, the coexistence of different closely related taxa able to replace the less-adapted ones has been demonstrated [17].
Previous studies [5][7][18] provided a comprehensive overview of successful in situ biogas upgrading works. In recent years, a remarkable effort has been dedicated to providing valuable solutions to major challenges related to in situ biomethanation technology.
Specifically, one of the main technical issues is the remarkable pH increase determined by CO2 removal. If the pH exceeds 8.5, it can inhibit methanogenesis [16][19][20]. CO2 dissolved in the liquid phase dissociates into H+ and HCO3, playing a fundamental role in buffering the process. Therefore, CO2 removal results in reduced H+ levels with a consequent pH increase. More details on the optimal pH range and pH control during methanogenesis will be provided in the next paragraph.
Among the works aiming at containing the increase in pH upon addition of H2, Luo and Angelidaki tested the co-digestion of manure and cheese whey, which can counteract the increase in pH and maintain it within optimal levels for methanogenesis while achieving up to 85% CO2 removal [19].
Additionally, the increase in PH2 (>10 Pa) resulting from the injection of H2 in the system could also cause methanogenesis inhibition alongside the accumulation of volatile fatty acids (VFAs) [1][20][21][22]. As it will be detailed in the next paragraph, the accumulation of VFAs could lead to process failure due to the consequent decrease in pH; thus, a careful monitoring and control of VFA levels is required to maintain the process efficiency.
Ref. [23] specifically applied isotope analysis to investigate the effect of the excess of H2 on in-batch, in situ biomethanation performance at thermophilic conditions. The results showed how the excess of H2 led to its accumulation in reactor liquid phase as dissolved H2, resulting in the inhibition of VFAs’ degradation and the stimulation of the homoacetogenic pathway for the production of acetate from CO2 and H2. Nevertheless, VFAs’ degradation and methanogenesis resumed once the excess of H2 was removed from the system. Similarly, [24] tested mesophilic batch reactors exposed to pulse H2 injections at levels exceeding stoichiometric ratio for CO2 and H2. The authors report reversible acetate accumulation, mostly attributed to homoacetogenic activity, suggesting the possibility of exploiting acetate as a temporary H2 storage before methanogenesis restoration.

2.2. Ex Situ Biomethanation

Ex situ biomethanation consists of the injection of H2 and CO2 from external sources inside an anaerobic reactor containing enriched or pure hydrogenotrophic methanogenic cultures (Figure 1b) [14][25][26][27]. Biomethanation can be decoupled from biogas production, resulting in a higher process flexibility and stability. In fact, CO2 derived from multiple sources, e.g., biogas production, biomass gasification and industrial process, can be applied to the process, ensuring a stable substrate supply independently from the biomass availability and geographical location of each. Moreover, biomethanation being carried out in a dedicated reactor, the system can benefit from higher stability, as no biomass degradation (i.e., hydrolysis and acetogenesis) is required, avoiding the technical limitations of in situ biomethanation. At the same time, the conversion efficiency of acclimatized hydrogenotrophic cultures allows for a reduction in operative costs, thanks to the possibility of providing the bioreactor with a high input of gas flow rates while keeping limited working volumes. Comprehensive overviews of tested ex situ biomethanation configurations are reported in the literature [5][7][28].
Interestingly, these studies investigated ex situ biomethanation based on biogas or CO2 as a carbon source. As mentioned above, this technology offers the possibility of sustaining the biological CO2 conversion of gas mixtures derived from different origins. Previous studies highlight the possibility of providing anaerobic reactors with syngas to biologically convert H2 and CO2 to CH4 [29]. When using mixed cultures this could be achieved by the pairing of carboxydotrophic-mediated H2 production followed by the methanogenic reduction of CO2 by HM or, alternatively, through homoacetogenesis from CO followed by acetoclastic methanogenesis or oxidation of CO to formic acid followed by its reduction to CH4 [30]. Biomethanation from syngas offers the opportunity of utilizing the fraction of organic waste remaining unused during biogas production (~50%) through gasification, followed by its reintroduction into the anaerobic reactor, with the heat produced from AD covering the power consumptions of gasification [31].
Despite the mentioned advantages associated to ex situ biomethanation technology compared to in situ biomethanation and the differences in system design and operation, both approaches are affected by a common parameter: the gas transfer to reactor liquid phase, where the biochemical reactions take place [26][30]. Because H2 is 500 times less soluble in water than CO2 [32], H2 availability for methanogens represents a remarkable limiting factor for methanogenesis [1][15][25]. Previous studies reported the existence of a correlation between H2 gas transfer coefficient (kLa), which is directly proportional to the gas–liquid mass transfer rate (rt), and in turn, the gas–liquid mass transfer depends on operational parameters such as reactor configuration, mixing speed, gas recirculation flow rate and applied gas diffusion device [7][25]. This has been demonstrated by several works aiming at optimizing H2 uptake and conversion to CH4 through the modulation of the aforementioned factors, reaching H2 conversion efficiencies close to 100% and upgrading the system CH4 content up to 98% [1][14][25][33].
A deeper insight into the rt correlation with the most important operational parameters will be provided in the next section.

2.3. Hybrid In Situ and Ex Situ Biomethanation

Recently, a new technology has been proposed integrating in situ and ex situ concepts in a single hybrid configuration, aiming at exploiting advantages of acclimatized hydrogenotrophic cultures for ex situ biomethanation and limiting the drawbacks of in situ technology, and some examples will be briefly illustrated below. Within this concept, upgraded biogas resulting from in situ H2 addition in a conventional biogas reactor is further polished to higher CH4 content by the action of acclimatized hydrogenotrophic cultures [34]. An example is the hybrid configuration presented by [35] and consisting of two separate chambers: the first dedicated to H2-assisted in situ biogas upgrading and upgrading the biogas up to 86% CH4 content, and the second receiving the biogas from the first chamber and further upgrading it to >91% CH4.
A similar concept was described by [36]  proposing a hybrid system composed of a 10 L CSTR for in situ biomethanation and a 2 L chamber filled with packing material carrying out the ex situ process. The results showed that the operation of this hybrid reactor led to a 28% higher CH4 yield and a twofold higher H2 consumption rate compared to the in situ concept alone and a 76% higher CH4 yield compared to the non-H2-fed reactor, with an overall 62% H2 consumption rate.
An alternative design has been proposed by [34], in which an H2-fed reactor for in situ biogas upgrading was supplemented by additional external biogas and H2 to demonstrate the resilience of the system to increased input gas flow rate and simulate the biomethanation of gas derived from different sources. In situ biomethanation resulted in biogas upgraded at >93% CH4 content, which also remained stable upon external gas injection, with an overall >3-fold increase in CH4 yield.

3. Characteristics and Productivity of Pure Cultures vs. Methanogenic Consortia

HMs are the key players during the biomethanation of H2 and CO2. Previous works presenting an overview of biomethanation studies carried out using hydrogenotrophic pure cultures and methanogenic microbial consortia demonstrated the feasibility of both approaches [18][37]. Both technologies have advantages and drawbacks related to the process operation, performance, operative costs and system sustainability. Specifically, the use of pure cultures in industrial applications may offer some advantages in terms of process predictability and ease of control. Conversely, enriched mixed cultures require a long adaptation time and a specific procedure. Moreover, unwanted side reactions taking place within a complex consortium could interfere with the process [37].
Pure cultures typically require more stringent conditions, in terms of nutrient content and control parameters when compared to the robustness characterizing mixed adapted cultures. In fact, during the operation of complex consortia systems, nutrients can be provided through their source substrate, without the need for sterility [38]. Moreover, in the context of industrial application, the uneven distribution of feedstock gas along the year, due to weather conditions and biomass availability, has to be considered. Recent studies [39][40] have shown the remarkable robustness and short recovery time of mixed cultures upon starvation/excess of input gas rate and oxygenation. In fact, in the presence of changing conditions, the best-adapted microorganisms will grow and become dominant. Conversly, less adapted species can survive through spore formation or utilizing residual biomass without the addition of any nutrient [38][39]. Mixed cultures are also capable of performing a variety of biochemical reactions entertaining inter-species communication, which explains the coexistence of different microbial groups in H2/CO2-fed methanogenic systems [5].
Compared to pure cultures, they also offer the possibility of polishing gas mixtures from residual components other than CO2 and H2, such as those contained in biogas or in flue gases emissions [14][37].
Additionally, mixed cultures offer advantages in terms of operative and startup costs, as specific nutrient media and stringent cultivation conditions are not required.
To improve CO2 and H2 conversion efficiency of complex methanogenic consortia, several strategies, such as bio-augmentation with pure cultures or enrichment of existing hydrogenotrophic consortia by specific nutrient or gas mixture supply, can be envisaged. However, both strategies increase operative costs, and the optimization of biochemical conditions has to be carefully considered in order to meet consortia nutrient requirements [1].

4. Critical Control Points: Physical/Chemical Parameters Affecting the Process

Despite the well-known advantages of the biological route for CO2 removal and conversion in terms of economic and environmental costs [37], an efficient biomethanation process demands a punctual setup and constant monitoring of the operational parameters [7][16]. In this section, researchers provide a list of the major factors that must be considered when designing and developing a biomethanation process and during system operation.

4.1. Temperature

In natural environments, methane formation occurs for a wide range of temperatures, going from ≤25 °C of psychrophilic methanogens to >60 °C of hyperthermophilic [41]. Nevertheless, most applications rely on mesophilic (25–45 °C) or thermophilic (45–60 °C) processes [42]. Previous studies compared reactor performances at different temperature conditions, reporting a different impact of the temperature on CH4 production and CO2 conversion efficiency.
For example, [1] demonstrated that, in batch assay, an enriched thermophilic culture resulted in >60% higher CO2 conversion compared to the mesophilic one. Conversely, tests conducted in a continuous stirred tank reactor (CSTR) showed higher CO2 conversion efficiency at 37 °C. Then, a remarkably higher CH4 production rate and yield were detected at 55 °C [16]. Similarly, [43] reported a comparable methane content in the output gas of mesophilic and thermophilic systems, despite the higher CH4 production rate detected at thermophilic conditions. Moreover, upon H2 application, several studies reported a higher diversity of methanogenic population at mesophilic conditions [39][43].
As detailed below, the higher conversion efficiency detected at mesophilic conditions can be reasonably explained by the well-known inverse correlation between gas solubility and temperature defined by Henry’s Law [44].
Finally, a study conducted at batch level, comparing process performances at thermophilic and hyperthermophilic conditions, showed that a temperature increase from 55 °C to 65 °C resulted in higher CH4 content and productivity [45].
Regarding mesophilic and thermophilic H2-adapted communities, previous studies underlined the existence of two clearly distinct populations, responding to H2 pressure in different ways, with mesophilic communities undergoing a more radical reduction in microbial diversity upon H2 exposure. Nevertheless, both adapted populations could rely on highly specialized consortia oriented towards methanogenic functions (Methanoculleus spp., Methanothermobacter spp., Methanosarcina spp.) [16][17]. Similar results were presented by [46] referring to a decrease in mesophilic population diversity and a completely different composition of the community between the two temperature conditions. According to β diversity analysis, thermophilic communities exhibiting higher CH4 production yields and conversion efficiencies were more sensitive to H2 addition. Moreover, phylogenetic analysis suggested that biomethanation occurred directly through hydrogenotrophic methanogenesis only at thermophilic conditions (Methanoculleus spp., Methanobacterium spp.), whereas homoacetogenesis and acetoclastic methanogenesis (Methanosaeta spp.) determined the major methanogenic pathway in mesophilic conditions, with SAOB only being detected in thermophilic reactors [46].
Despite the differences in process performance described here, several works reported successful biomethanation outcomes for a wide spectrum of temperatures and reactor sizes, using both pure and mixed cultures as inoculum [3][15][19][27][33][47][48][49]. Notably, among these studies, only two [27][47] were conducted at hyperthermophilic conditions, and none of them employed microbial consortia, with only the work of [45] testing hyperthermophilic conditions with mixed hydrogenotrophic culture. Specifically, they tested the activity of M. marburgensis and M. thermoautotrophicus at 65 and 60 °C in 10 and 3.5 L working volume, respectively, reaching up to 85% CH4 content, together with 950 mmol/L×h and ~50 L/L culture-day CH4 production rate.

4.2. pH

Biomethanation typically takes place in a pH range from 6.5 to 8.5 with an optimum at pH 6.5–7.5, and variations in pH were shown to directly affect archaeal growth and activity [41][50]. During the biomethanation process, the control and monitoring of the pH play a fundamental role, being the object of several studies. A specific methanogenic activity (SMA) test conducted on enriched mesophilic and thermophilic hydrogenotrophic cultures exposed to different pH conditions (from 6 to 10), pointed out that biomethanation was feasible up to pH 8.5, even though methanogenic activity was significantly reduced. Conversly, complete process inhibition was observed at pH levels >8.5 [16].
pH control strategies during biomethanation are mainly based on the injection of pH buffering solutions (NaOH and HCl) in order to stabilize the values in the theoretical optimal range for methanogenesis [21][35][43][51]. Ref. [33] report that, during continuous biomethanation of H2:CO2 at mesophilic conditions, fine control of the pH might be achieved by adjusting the CO2 flow rate in the input gas, using real time data of CO2 conversion efficiency and pH.
Among the compounds known to strongly affect the pH level of methanogenic reactors systems, VFA and ammonia (NH3) were identified, with VFA accumulation leading to reactor acidification, and NH3, generated from protein or urea degradation, resulting in higher pH [50]. Because the hydrolysis rate of organic matter increases with temperature, great attention must be paid to pH control and monitoring during methanogenesis at thermophilic or hyperthermophilic conditions.

4.3. Volatile Fatty Acids Concentration

During biomethanation, VFAs, produced from the hydrolysis of organic matter, can be utilized by AM or SAOB in syntrophic association with HM for CH4 production [50][52][53]. Inhibition of the activity of such classes of microorganisms, for instance, due to an increase in PH2 during in situ biomethanation, may lead to VFA accumulation and consequent reactor acidification. This could possibly result in process imbalance, reduced gas conversion efficiency and production rate or even system failure.
The literature offers a wide spectrum of works reporting temporary VFAs accumulation during both in situ and ex situ biomethanation, carried out at both mesophilic and thermophilic conditions [21][39][54][55]. Although the absence of organic feedstocks should keep the levels of VFAs relatively low in gas-fed biomethanation reactors, the literature reports the accumulation of VFAs, especially in thermophilic conditions. For example, during thermophilic ex situ biogas upgrading with methanogenic consortia, [14] reported a significant reduction in the activity of AM and the predominance of hydrogenotrophic taxa due to the decrease in the pH and the accumulation of acetate.

4.4. Ammonia Concentration

Ammonia concentration is another critical factor affecting the activity of the methanogenic archaea. Below a threshold concentration, NH3 ensures the buffering capacity of the reactor medium, increasing the stability of the process [56]. Nevertheless, its excess was reported as one of the main causes for process imbalance or reactor failure due to the inhibitory effect on the microbial population [57].
In anaerobic environments, ammonia is released from the hydrolysis of organic compounds, such as proteins and urea, causing an increase in the pH and counteracting the acidification induced by the acidogenesis. In aqueous solution, NH3 can be present as free un-ionized ammonia nitrogen (FAN) and ammonium nitrogen (NH4+). The dissociation balance between the two forms is strongly influenced by temperature and pH. It was reported how, at high temperature and pH, the dissociation balance tends to shift towards the FAN form [58], which is the most likely cause of process inhibition, due to FAN’s ability to permeate bacterial cell membranes [59].
AM and HM seem to respond in a different way to the stress induced by the excess of ammonia, resulting in a shift in the metabolic pathway and changes in the methanogenic population.
Ref. [53] reported that high ammonia concentrations are responsible for the inhibition of AM, resulting in competition for acetate, possibly enhancing the growth and the activity of SAOBs. SAOBs are known to form syntrophic relations with HM for the oxidation of acetate by the former and the consequent utilization of H2 and CO2 by the latter. Despite the slow SAOB growth rate, which can be a disadvantage in the competition for acetate with the AM, the high tolerance of HMs and SAOBs to ammonia favors these microbial groups at high ammonia levels [53]. Moreover, the SAO pathway is also energetically favorable at elevated temperature, which is a condition further forcing NH3 dissociation towards the FAN form [53].
While testing anaerobic digestion under different ammonia levels, [60] demonstrated that high ammonia concentrations (2.8–4.57 g NH4+/L) favor SAO and hydrogenotrophic methanogenesis (i.e., orders Methanomicrobiales and Methanobacteriales at thermophilic and mesophilic conditions, respectively). Conversely, acetoclastic methanogenesis (order Methanosaetaceae) was promoted at low ammonia levels (1.21 g NH4+/L). Similarly, it was subsequently confirmed that HM possesses higher tolerance to ammonia compared to acetoclastic methanogenic archaea [60].
These results may suggest the resilience of hydrogen-mediated methanation at high ammonia concentrations, with higher tolerance to ammonia compared to the AD process. Moreover, in gas-fed chemostats, the curtailment of organic feed should reduce the amount of NH3 present in the system, making its effect negligible. This statement is in agreement with ex situ biomethanation studies reporting a decrease in NH3 levels, following a short period of accumulation, attributable to the degradation of the residual biomass [39].

4.5. Salinity

During the last decade, the rising interest for AD from high-salt-content substrates, such as marine macroalgae, fish wastewater and brackish aquaculture sludge, has driven the development of several studies aiming at defining the range of salinity allowing for efficient biomethanation performances [61][62][63].
These studies pointed out the methanogenic inoculum adaptation to increasing salt contents as a crucial requirement for a successful AD process, with methanogens being considered as the most sensitive microbial group within the consortium [61][62][63][64]. In fact, salinity affects several biochemical processes occuring at the cellular level. For example, hyperionic and hyperosmotic stresses can cause dehydration and cell lysis. Moreover, intracellular and extracellular enzyme inhibition and cell membrane impairment may result in altered cell functioning [65][66].
Na+ has been suggested as the main methanogenesis inhibitor, compromising the process at levels as low as 6–13 g Na+/L when applied to non-acclimatized inocula [63][65][67]. More specifically, for methanogens, concentrations of 3.5–5.5 g Na+/L were reported to cause moderate inhbition, which became severe at >8 g Na+/L [68]. However, several Methanosarcina species are halotolerant, being detected at up to 18 g Na+/L and, similarly to Methanosaeta, dominating a high-salinity anaerobic digester over hydrogenotrophic methanogens [61][64][68]. In addition, Cl, which is the most common counterpart of Na+, may be responsible for plant deterioration through the corrosion of steel components [69]. Conversely, low Na+ concentrations (≤0.35 g/L) are essential for methanogens, as this ion is involved in ATP syntesis and NADH oxidation [68][70].
Consistently, during the AD of food waste leachate supplemented with 0.5 and 2 g/L NaCl (corresponding to 0.2 and 0.8 g/L Na+, respectively), a 10% higher CH4 yield was observed with 2 g/L NaCl compared to 0.5 g/L [32]. This effect was attributed to the preliminary adaptation of the methanogenic inoculum to 1.2 g/L Na+. Nevertheless, further increasing NaCl levels to 5 and 10 g/L (corresponding to 2 and 4 g/L Na+, respectively) resulted in a ~40% lower CH4 yield [71].
Among naturally high salinity substrates suitable for CH4 production, Zhang and coworkers [62] reported that the AD of marine macroalgae using an adapted inoculum was achievable at salinity levels ≤35 g/L, whereas methanogenesis was seriously affected at salt concentrations >55 g/L. Notably, they observed that the best performances were achieved at the salinity of 15 g/L, suggesting an enhancement of methanogenesis at this salt level. Regarding the methanogenic archaeal community, acetoclastic Methanosaeta and Methanosarcina were detected at remarkable relative abundance at salinity ≤35 g/L, tolerating salt levels up to 55–65 g/L and being considered as moderated halophiles. Nevertheless, the dominance of hydrogenotrophic Mehanobacterium was observed at all salt levels tested, up to 85 g/L, with a maximum at 52.65 g/L [62].
Similarly, Letelier-Gordo and colleagues [63] have recently evaluated different co-digestion scenarios of fish wastewater and manure at salinities up to 35 g/L, successfully overcoming the CH4 production rates achievable with cow manure mono-digestion and pointing out a statistical correlation between process inhibition and the level of salinity.

4.6. Nutrient Content

In addition to carbon sources, different elements are involved and required for microbial metabolisms. In fact, the main cell constituents such as C, H, O, N, P and S, along with Mg, Na, Ca and K, concurring to basic cell functions should always be available [72]. Moreover, metallic elements, such as Fe, Ni, Co, Mo, W and Se, despite being available in lower amounts, play a fundamental role as cofactors or as part of an enzyme [73].
Nutrient composition and availability depends on substrate source, with different methanogenic feedstocks being characterized by peculiar mineral compositions. The authors of [74] reported that plants fed with mixtures of animal manure and different fractions of organic waste displayed higher concentrations of mineral nutrients when compared to plants largely fed with industrial by-products (i.e., glycerol).
Ref. [75] collected many studies investigating the physiology, media demand and productivity of different methanogenic strains. In the context of anaerobic digestion (AD), it is well established that variable concentrations of TE have significant effects on the production of CH4, where the archaea community was found to be more responsive than other bacterial community members [76]. However, only few studies cover the effect of heavy metals on pure cultures of methanogens.
Ref. [77] demonstrated that Methanococcus maripaludis growth was inhibited by specific concentration of Zn ione (2.5 and 3.5 mmol/L), while Cu concentrations of 1.9 µmol/L, 4.4 and 6.3 reduced growth and delayed biomass production. More interesting is the combined effect of Zn and Cu iones, where the addition of 1 mmol/L Zn can prevent the toxicity effect of Cu.
Another important aspect lacking insight, concerns the connection between TE and the physiological and biotechnological characteristics of methanogens. It was demonstrated that TE limitation could lead to low productivity during biomethanation in mixed cultures [78], but studies on pure cultures are rare. The growth and productivity of M. marburgensis was maximized by applying the exponential feeding of TE, different medium and sulphide dilution rates and different gas inflow rates. With the right combination of these parameters, the greatest ever specific growth rate (µmax) of 0.69 h−1 and methane evolution rate (MER) of 476 mmol/L × h were achieved [79]. Other studies demonstrated that the concentration of Fe, Cu, Ni and Zn, with the exception of Co, should be increased by 100 times than the conventional method to achieve high productivity of methane with acclimated-methanogens [80][81].
Along with the mineral nutrients, vitamins such as biotin, para-aminobenzoic acid, riboflavin and different B-group vitamins were reported to be required or to stimulate the activity and growth of methanogenic archaea [82]. In a recent study [83], it was highlighted that only some methanogens, such as Methanobacteriacae and M.maripaludis, are able to grow on minimal and optimized TE solution without cysteine or vitamins, while hyperthermophylic methanogens with high MERs require a combination of a rich TE composition with additional cysteine and/or vitamins [83].

4.7. Gas Solubility and Gas Transfer Coefficient (kLa)

The solubility of gases in aqueous environments is described by Henry’s Law stating that, at constant temperature, the amount of gas that dissolves in a liquid is proportional to the partial pressure of the gas in equilibrium with the liquid. Henry’s law can be expressed as Equation (2):
C = H k × P
where C represents the gas solubility concentration at a certain temperature in a specific solvent, Hk is the Henry’s Law constant and P is the gas partial pressure at a given volume and temperature [44].
Different works underline the relationship between the Henry’s constant for a specific gas and the temperature of the system, when the partial pressure is considered in equilibrium, allowing for the calculation of the molar fraction of gas dissolved in liquid phase [44][84]. Data reported by these studies unequivocally show that the temperature increase in a specific system leads to the reduction in the solubility of the injected gases. This is mainly caused by the fact that gas solubilization is an exothermic process in which the gas dissolution releases heat to the system. A temperature rise leads to an increase in the kinetic energy of the gases’ molecules, and this may limit the formation of intermolecular bonds between the solute and the solvent [85].
As mentioned in the previous section, during biomethanation, the low solubility of H2 is one of the most relevant limiting factors for H2/CO2 conversion efficiency [5][28].
At the same time, the gas transfer coefficient (kLa) and the temperature are positively correlated [86]. Specifically, higher solvent viscosity (µa) was reported to retard diffusion of gases in Newtonian fluids [87][88]. In AD digestate, which is considered as a non-Newtonian fluid, µa changes with temperature, shear forces and solid content. While higher solids content was proven to increase µa [89], an increase in either temperature or shear stress was reported to decrease it [90].
Thus, the higher the temperature, the lower the µa and the higher the diffusivity of the gases.
Diffusivity and temperature can be correlated as per Equation (3):
D L μ / T = C o n s t a n t
where DL is the diffusivity of the solute at infinite dilution, µ is the viscosity of the solution and T is the absolute temperature. Diffusivity, temperature and viscosity are correlated and affect the gas transfer coefficient (kLa) [86], which determines the gas–liquid mass transfer rate (rt) as defined by Equation (4):
r t = 22.4   k L a * ( H 2 g H 2 l )
where 22.4 is the molar volume, kLa is the gas transfer coefficient, H2g is the H2 concentration in the gas phase and H2l is the H2 concentration in the liquid phase.
kLa comprises two other coefficients, where kL is defined as the film coefficient, depending on gas and liquid physicochemical features, and a is the interface area per unit volume of liquid [86]. Therefore, in order to take into account the dual role of temperature, the overall gas–liquid mass transfer rate should include both gas solubility and gas transfer coefficient, as defined by Equation (5) [5]:
r t = 22.4   k L a ( H k × P × H 2 g H 2 l )
Due to the lower solid content in the digestate or medium typically utilized during the ex situ biomethanation process compared to anaerobic sludge used for in situ methanogenesis, rt is expected to favor the ex situ process [37].
Despite the attempts to describe gas behavior and diffusion in a liquid [91][92], the obtained models appear too simplistic, especially when applied to a chemostat for biomethanation, in which a tri-phasic system (liquid–solid–gas) is constantly affected by the activity of the microorganisms in the developing community. In such a system, where interactions appear somewhat chaotic and hence difficult to predict, parameters can be considered individually in order to improve gas solubility.

4.8. Pressure

As described in the previous section, pressure is directly proportional to gas solubility in a liquid. The higher the volume of gas in a close system, the higher its solubility. The higher the number of gas molecules at the gas–liquid interface, the higher the interface contact; thus, the gas availability for microorganisms [86]. The resistance and improved performance and growth rate of hydrogenotrophic methanogens at extreme pressures (>100 atm) were reported [86]. A batch assay conducted on a lithotrophic strain pointed out a positive correlation between CO2 conversion efficiency and pressure, with higher conversion efficiencies being observed for a pressure increase from 1 (70 μMCH4) to 50 (3500 μMCH4) and 100 atm (7000 μMCH4). Similarly, by increasing the reactor pressure from 101 kPa to 122 kPa, CH4 production increases from 50 LCH4/Lculture/day to 65.6 LCH4/Lculture/day was reported during biomethanation using M. thermoautotrophicus pure culture [27].

4.9. Gas Hold-Up

The time a gas resides in a reactor can be defined as the gas hold-up. The longer the gas residence time, the longer the contact is between the gas and the liquid phase where metabolic reactions take place.
Gas hold-up can be modulated using gas or liquid recirculation [15][25] or through mixing speed [1], acting on the velocity of the bubbles in the reactor.
In fact, vigorous gas–liquid dispersion generated by the movement of the gas bubbles in the liquid medium results in turbulent flow, responsible for improving the gas contact with liquid phase in the reactor. Moreover, gas recirculation can enhance gas–liquid mass transfer because it increases the overall gas injection rate to the liquid, thus also increasing the gas–liquid interface area [42].


  1. Luo, G.; Angelidaki, I. Integrated Biogas Upgrading and Hydrogen Utilization in an Anaerobic Reactor Containing Enriched Hydrogenotrophic Methanogenic Culture. Biotechnol. Bioeng. 2012, 109, 2729–2736.
  2. Kletzin, A. General Characteristics and Important Model Organisms. In Archaea; ASM Press: Washington, DC, USA, 2014; pp. 14–92.
  3. Alitalo, A.; Niskanen, M.; Aura, E. Biocatalytic Methanation of Hydrogen and Carbon Dioxide in a Fixed Bed Bioreactor. Bioresour. Technol. 2015, 196, 600–605.
  4. Lee, J.C.; Kim, J.H.; Chang, W.S.; Pak, D. Biological Conversion of CO2 to CH4 Using Hydrogenotrophic Methanogen in a Fixed Bed Reactor. J. Chem. Technol. Biotechnol. 2012, 87, 844–847.
  5. Rafrafi, Y.; Laguillaumie, L.; Dumas, C. Biological Methanation of H2 and CO2 with Mixed Cultures: Current Advances, Hurdles and Challenges. Waste Biomass Valorization 2021, 12, 5259–5282.
  6. Iglesias, R.; Muñoz, R.; Polanco, M.; Díaz, I.; Susmozas, A.; Moreno, A.D.; Guirado, M.; Carreras, N.; Ballesteros, M. Biogas from Anaerobic Digestion as an Energy Vector: Current Upgrading Development. Energies 2021, 14, 2742.
  7. Angelidaki, I.; Treu, L.; Tsapekos, P.; Luo, G.; Campanaro, S.; Wenzel, H.; Kougias, P.G. Biogas Upgrading and Utilization: Current Status and Perspectives. Biotechnol. Adv. 2018, 36, 452–466.
  8. Kumar, A.; Jones, D.D.; Hanna, M.A. Thermochemical Biomass Gasification: A Review of the Current Status of the Technology. Energies 2009, 2, 556–581.
  9. Rönsch, S.; Schneider, J.; Matthischke, S.; Schlüter, M.; Götz, M.; Lefebvre, J.; Prabhakaran, P.; Bajohr, S. Review on Methanation—From Fundamentals to Current Projects. Fuel 2016, 166, 276–296.
  10. Hashimoto, K.; Kumagai, N.; Izumiya, K.; Takano, H.; Shinomiya, H.; Sasaki, Y.; Yoshida, T.; Kato, Z. The Use of Renewable Energy in the Form of Methane via Electrolytic Hydrogen Generation Using Carbon Dioxide as the Feedstock. Appl. Surf. Sci. 2016, 388, 608–615.
  11. Turner, J.; Sverdrup, G.; Mann, M.K.; Maness, P.C.; Kroposki, B.; Ghirardi, M.; Evans, R.J.; Blake, D. Renewable Hydrogen Production. Int. J. Energy Res. 2008, 32, 379–407.
  12. Jürgensen, L.; Ehimen, E.A.; Born, J.; Holm-Nielsen, J.B. Utilization of Surplus Electricity from Wind Power for Dynamic Biogas Upgrading: Northern Germany Case Study. Biomass Bioenergy 2014, 66, 126–132.
  13. Muñoz, R.; Meier, L.; Diaz, I.; Jeison, D. A Review on the State-of-the-Art of Physical/Chemical and Biological Technologies for Biogas Upgrading. Rev. Environ. Sci. Biotechnol. 2015, 14, 727–759.
  14. Kougias, P.G.; Treu, L.; Benavente, D.P.; Boe, K.; Campanaro, S.; Angelidaki, I. Ex-Situ Biogas Upgrading and Enhancement in Different Reactor Systems. Bioresour. Technol. 2017, 225, 429–437.
  15. Bassani, I.; Kougias, P.G.; Angelidaki, I. In-Situ Biogas Upgrading in Thermophilic Granular UASB Reactor: Key Factors Affecting the Hydrogen Mass Transfer Rate. Bioresour. Technol. 2016, 221, 485–491.
  16. Bassani, I.; Kougias, P.G.; Treu, L.; Angelidaki, I. Biogas Upgrading via Hydrogenotrophic Methanogenesis in Two-Stage Continuous Stirred Tank Reactors at Mesophilic and Thermophilic Conditions. Environ. Sci. Technol. 2015, 49, 12585–12593.
  17. Treu, L.; Campanaro, S.; Kougias, P.G.; Sartori, C.; Bassani, I.; Angelidaki, I. Hydrogen-Fueled Microbial Pathways in Biogas Upgrading Systems Revealed by Genome-Centric Metagenomics. Front. Microbiol. 2018, 9, 1079.
  18. Jensen, M.B.; Jensen, B.; Ottosen, L.D.M.; Kofoed, M.V.W. Integrating H2 Injection and Reactor Mixing for Low-Cost H2 Gas-Liquid Mass Transfer in Full-Scale in Situ Biomethanation. Biochem. Eng. J. 2021, 166, 107869.
  19. Luo, G.; Angelidaki, I. Co-Digestion of Manure and Whey for in Situ Biogas Upgrading by the Addition of H2: Process Performance and Microbial Insights. Appl. Microbiol. Biotechnol. 2013, 97, 1373–1381.
  20. Tao, B.; Alessi, A.M.; Zhang, Y.; Chong JP, J.; Heaven, S.; Banks, C.J. Simultaneous Biomethanisation of Endogenous and Imported CO2 in Organically Loaded Anaerobic Digesters. Appl. Energy 2019, 247, 670–681.
  21. Wang, W.; Xie, L.; Luo, G.; Zhou, Q.; Angelidaki, I. Performance and Microbial Community Analysis of the Anaerobic Reactor with Coke Oven Gas Biomethanation and in Situ Biogas Upgrading. Bioresour. Technol. 2013, 146, 234–239.
  22. Batstone, D.J.; Keller, J.; Angelidaki, I.; Kalyuzhnyi, S.V.; Pavlostathis, S.G.; Rozzi, A.; Sanders, W.T.; Siegrist, H.; Vavilin, V.A. The IWA Anaerobic Digestion Model No 1 (ADM1). Water Sci. Technol. 2002, 45, 65–73.
  23. Mulat, D.G.; Mosbæk, F.; Ward, A.J.; Polag, D.; Greule, M.; Keppler, F.; Nielsen, J.L.; Feilberg, A. Exogenous Addition of H2 for an in Situ Biogas Upgrading through Biological Reduction of Carbon Dioxide into Methane. Waste Manag. 2017, 68, 146–156.
  24. Agneessens, L.M.; Ottosen, L.D.M.; Voigt, N.V.; Nielsen, J.L.; de Jonge, N.; Fischer, C.H.; Kofoed, M.V.W. In-Situ Biogas Upgrading with Pulse H2 Additions: The Relevance of Methanogen Adaption and Inorganic Carbon Level. Bioresour. Technol. 2017, 233, 256–263.
  25. Bassani, I.; Kougias, P.G.; Treu, L.; Porté, H.; Campanaro, S.; Angelidaki, I. Optimization of Hydrogen Dispersion in Thermophilic Up-Flow Reactors for Ex Situ Biogas Upgrading. Bioresour. Technol. 2017, 234, 310–319.
  26. Díaz, I.; Pérez, C.; Alfaro, N.; Fdz-Polanco, F. A Feasibility Study on the Bioconversion of CO2 and H2 to Biomethane by Gas Sparging through Polymeric Membranes. Bioresour. Technol. 2015, 185, 246–253.
  27. Martin, M.R.; Fornero, J.J.; Stark, R.; Mets, L.; Angenent, L.T. A Single-Culture Bioprocess of Methanothermobacter Thermautotrophicus to Upgrade Digester Biogas by CO2-to-CH4 Conversion with H2. Archaea 2013, 2013, 157529.
  28. Jensen, M.B.; Ottosen, L.D.M.; Kofoed, M.V.W. H2 Gas-Liquid Mass Transfer: A Key Element in Biological Power-to-Gas Methanation. Renew. Sustain. Energy Rev. 2021, 147, 111209.
  29. Guiot, S.R.; Cimpoia, R.; Carayon, G. Potential of Wastewater-Treating Anaerobic Granules for Biomethanation of Synthesis Gas. Environ. Sci. Technol. 2011, 45, 2006–2012.
  30. Henstra, A.M.; Sipma, J.; Rinzema, A.; Stams, A.J. Microbiology of Synthesis Gas Fermentation for Biofuel Production. Curr. Opin. Biotechnol. 2007, 18, 200–206.
  31. Monlau, F.; Sambusiti, C.; Ficara, E.; Aboulkas, A.; Barakat, A.; Carrère, H. New Opportunities for Agricultural Digestate Valorization: Current Situation and Perspectives. Energy Environ. Sci. 2015, 8, 2600–2621.
  32. Ahern, E.P.; Deane, P.; Persson, T.; Ó Gallachóir, B.; Murphy, J.D. A Perspective on the Potential Role of Renewable Gas in a Smart Energy Island System. Renew. Energy 2015, 78, 648–656.
  33. Savvas, S.; Donnelly, J.; Patterson, T.; Chong, Z.S.; Esteves, S.R. Biological Methanation of CO2 in a Novel Biofilm Plug-Flow Reactor: A High Rate and Low Parasitic Energy Process. Appl. Energy 2017, 202, 238–247.
  34. Corbellini, V.; Kougias, P.G.; Treu, L.; Bassani, I.; Malpei, F.; Angelidaki, I. Hybrid Biogas Upgrading in a Two-Stage Thermophilic Reactor. Energy Convers. Manag. 2018, 168, 1–10.
  35. Wahid, R.; Horn, S.J. Impact of Operational Conditions on Methane Yield and Microbial Community Composition during Biological Methanation in in Situ and Hybrid Reactor Systems. Biotechnol. Biofuels 2021, 14, 170.
  36. Rittmann, S.K.M.R. A Critical Assessment of Microbiological Biogas to Biomethane Upgrading Systems. Adv. Biochem. Eng. Biotechnol. 2015, 151, 117–135.
  37. Rachbauer, L.; Beyer, R.; Bochmann, G.; Fuchs, W. Characteristics of Adapted Hydrogenotrophic Community during Biomethanation. Sci. Total Environ. 2017, 595, 912–919.
  38. Savvas, S.; Donnelly, J.; Patterson, T.; Chong, Z.S.; Esteves, S.R. Methanogenic Capacity and Robustness of Hydrogenotrophic Cultures Based on Closed Nutrient Recycling via Microbial Catabolism: Impact of Temperature and Microbial Attachment. Bioresour. Technol. 2018, 257, 164–171.
  39. Burkhardt, M.; Jordan, I.; Heinrich, S.; Behrens, J.; Ziesche, A.; Busch, G. Long Term and Demand-Oriented Biocatalytic Synthesis of Highly Concentrated Methane in a Trickle Bed Reactor. Appl. Energy 2019, 240, 818–826.
  40. Jensen, M.B.; Strübing, D.; de Jonge, N.; Nielsen, J.L.; Ottosen, L.D.M.; Koch, K.; Kofoed, M.V.W. Stick or Leave—Pushing Methanogens to Biofilm Formation for Ex Situ Biomethanation. Bioresour. Technol. 2019, 291, 121784.
  41. Liu, Y.; Whitman, W.B. Metabolic, Phylogenetic, and Ecological Diversity of Methanogenic Archaea. Ann. N. Y. Acad. Sci. 2008, 189, 171–189.
  42. Yun, Y.M.; Sung, S.; Kang, S.; Kim, M.S.; Kim, D.H. Enrichment of Hydrogenotrophic Methanogens by Means of Gas Recycle and Its Application in Biogas Upgrading. Energy 2017, 135, 294–302.
  43. Sander, R. Compilation of Henry’s Law Constants (Version 4.0) for Water as Solvent. Atmos. Chem. Phys. 2015, 15, 4399–4981.
  44. Guneratnam, A.J.; Ahern, E.; FitzGerald, J.A.; Jackson, S.A.; Xia, A.; Dobson, A.D.W.; Murphy, J.D. Study of the Performance of a Thermophilic Biological Methanation System. Bioresour. Technol. 2017, 225, 308–315.
  45. Zhu, X.; Chen, L.; Chen, Y.; Cao, Q.; Liu, X.; Li, D. Differences of Methanogenesis between Mesophilic and Thermophilic in Situ Biogas-Upgrading Systems by Hydrogen Addition. J. Ind. Microbiol. Biotechnol. 2019, 46, 1569–1581.
  46. Seifert, A.H.; Rittmann, S.; Herwig, C. Analysis of Process Related Factors to Increase Volumetric Productivity and Quality of Biomethane with Methanothermobacter Marburgensis. Appl. Energy 2014, 132, 155–162.
  47. Kim, S.; Choi, K.; Chung, J. Reduction in Carbon Dioxide and Production of Methane by Biological Reaction in the Electronics Industry. Int. J. Hydrogen Energy 2013, 38, 3488–3496.
  48. Burkhardt, M.; Koschack, T.; Busch, G. Biocatalytic Methanation of Hydrogen and Carbon Dioxide in an Anaerobic Three-Phase System. Bioresour. Technol. 2015, 178, 330–333.
  49. Strübing, D.; Huber, B.; Lebuhn, M.; Drewes, J.E.; Koch, K. High Performance Biological Methanation in a Thermophilic Anaerobic Trickle Bed Reactor. Bioresour. Technol. 2017, 245, 1176–1183.
  50. Gerardi, M.H. The Microbiology of Anaerobic Digesters; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2003.
  51. Sun, L.; Müller, B.; Westerholm, M.; Schnürer, A. Syntrophic Acetate Oxidation in Industrial CSTR Biogas Digesters. J. Biotechnol. 2014, 171, 39–44.
  52. Westerholm, M.; Moestedt, J.; Schnürer, A. Biogas Production through Syntrophic Acetate Oxidation and Deliberate Operating Strategies for Improved Digester Performance. Appl. Energy 2016, 179, 124–135.
  53. Rachbauer, L.; Voitl, G.; Bochmann, G.; Fuchs, W. Biological Biogas Upgrading Capacity of a Hydrogenotrophic Community in a Trickle-Bed Reactor. Appl. Energy 2016, 180, 483–490.
  54. Agneessens, L.M.; Ottosen, L.D.M.; Andersen, M.; Berg Olesen, C.; Feilberg, A.; Kofoed, M.V.W. Parameters Affecting Acetate Concentrations during In-Situ Biological Hydrogen Methanation. Bioresour. Technol. 2018, 258, 33–40.
  55. Rajagopal, R.; Massé, D.I.; Singh, G. A Critical Review on Inhibition of Anaerobic Digestion Process by Excess Ammonia. Bioresour. Technol. 2013, 143, 632–641.
  56. Hejnfelt, A.; Angelidaki, I. Anaerobic Digestion of Slaughterhouse By-Products. Biomass Bioenergy 2009, 33, 1046–1054.
  57. Fricke, K.; Santen, H.; Wallmann, R.; Hüttner, A.; Dichtl, N. Operating Problems in Anaerobic Digestion Plants Resulting from Nitrogen in MSW. Waste Manag. 2007, 27, 30–43.
  58. Müller, T.; Walter, B.; Wirtz, A.; Burkovski, A. Ammonium Toxicity in Bacteria. Curr. Microbiol. 2006, 52, 400–406.
  59. Tian, H.; Fotidis, I.A.; Kissas, K.; Angelidaki, I. Effect of Different Ammonia Sources on Aceticlastic and Hydrogenotrophic Methanogens. Bioresour. Technol. 2018, 250, 390–397.
  60. Zhang, X.; Tao, Y.; Hu, J.; Liu, G.; Spanjers, H.; van Lier, J.B. Biomethanation and Microbial Community Changes in a Digester Treating Sludge from a Brackish Aquaculture Recirculation System. Bioresour. Technol. 2016, 214, 338–347.
  61. Zhang, Y.; Alam, M.A.; Kong, X.; Wang, Z.; Li, L.; Sun, Y.; Yuan, Z. Effect of Salinity on the Microbial Community and Performance on Anaerobic Digestion of Marine Macroalgae. J. Chem. Technol. Biotechnol. 2017, 92, 2392–2399.
  62. Letelier-Gordo, C.O.; Mancini, E.; Pedersen, P.B.; Angelidaki, I.; Fotidis, I.A. Saline Fish Wastewater in Biogas Plants - Biomethanation Toxicity and Safe Use. J. Environ. Manag. 2020, 275, 111233.
  63. Wang, S.; Hou, X.; Su, H. Exploration of the Relationship between Biogas Production and Microbial Community under High Salinity Conditions. Sci. Rep. 2017, 7, 1149.
  64. Zhang, X.; Spanjers, H.; van Lier, J.B. Potentials and Limitations of Biomethane and Phosphorus Recovery from Sludges of Brackish/Marine Aquaculture Recirculation Systems: A Review. J. Environ. Manag. 2013, 131, 44–54.
  65. Vyrides, I.; Stuckey, D.C. Adaptation of Anaerobic Biomass to Saline Conditions: Role of Compatible Solutes and Extracellular Polysaccharides. Enzym. Microb. Technol. 2009, 44, 46–51.
  66. Huzir, N.M.; Mahmood, N.A.N.; Muhammad, S.A.F.A.S.; Umor, N.A.; Ismail, S. Effect of Specific Methanogenic Activity (SMA) of Anaerobic Sludge under High Salinity. J. Adv. Res. Appl. Sci. Eng. Technol. 2019, 16, 35–40.
  67. De Vrieze, J.; Hennebel, T.; Boon, N.; Verstraete, W. Methanosarcina: The Rediscovered Methanogen for Heavy Duty Biomethanation. Bioresour. Technol. 2012, 112, 1–9.
  68. Zhang, Y.; Arends, J.B.A.; Van de Wiele, T.; Boon, N. Bioreactor Technology in Marine Microbiology: From Design to Future Application. Biotechnol. Adv. 2011, 29, 312–321.
  69. Feijoo, G.; Soto, M.; Méndez, R.; Lema, J.M. Sodium Inhibition in the Anaerobic Digestion Process: Antagonism and Adaptation Phenomena. Enzym. Microb. Technol. 1995, 17, 180–188.
  70. Lee, D.H.; Behera, S.K.; Kim, J.W.; Park, H.S. Methane Production Potential of Leachate Generated from Korean Food Waste Recycling Facilities: A Lab-Scale Study. Waste Manag. 2009, 29, 876–882.
  71. Romero-Güiza, M.S.; Vila, J.; Mata-Alvarez, J.; Chimenos, J.M.; Astals, S. The Role of Additives on Anaerobic Digestion: A Review. Renew. Sustain. Energy Rev. 2016, 58, 1486–1499.
  72. Angelidaki, I.; Sanders, W. Assessment of the Anaerobic Biodegradability of Macropollutants. Re/Views Environ. Sci. Bio/Technol. 2004, 3, 117–129.
  73. Schattauer, A.; Abdoun, E.; Weiland, P.; Plöchl, M.; Heiermann, M. Abundance of Trace Elements in Demonstration Biogas Plants. Biosyst. Eng. 2011, 108, 57–65.
  74. Feng, X.M.; Karlsson, A.; Svensson, B.H.; Bertilsson, S. Industrial Waste Linking Process to Microbial Communities. FEMS Microbiol. Ecol. 2010, 74, 226–240.
  75. Ünal, B.; Perry, V.R.; Sheth, M.; Gomez-Alvarez, V.; Chin, K.J.; Nüsslein, K. Trace Elements Affect Methanogenic Activity and Diversity in Enrichments from Subsurface Coal Bed Produced Water. Front. Microbiol. 2012, 3, 175.
  76. Abdel Azim, A.; Rittmann, S.K.M.R.; Fino, D.; Bochmann, G. The Physiological Effect of Heavy Metals and Volatile Fatty Acids on Methanococcus maripaludis S2. Biotechnol. Biofuels 2018, 11, 301.
  77. Glass, J.B.; Orphan, V.J. Trace Metal Requirements for Microbial Enzymes Involved in the Production and Consumption of Methane and Nitrous Oxide. Front. Microbiol. 2012, 3, 61.
  78. Abdel Azim, A.; Pruckner, C.; Kolar, P.; Taubner, R.S.; Fino, D.; Saracco, G.; Sousa, F.L.; Rittmann, S.K.M.R. The Physiology of Trace Elements in Biological Methane Production. Bioresour. Technol. 2017, 241, 775–786.
  79. Zhang, Z.Y.; Maekawa, T. Continuous Fermentation Using Acclimated-Methanogens Feed Substrate of Mixed Carbon Dioxide on and Hydrogen. J. Soc. Agric. Struct. 1993, 24, 207–214.
  80. Zhang, Y.; Zhang, Z.; Suzuki, K.; Maekawa, T. Uptake and Mass Balance of Trace Metals for Methane Producing Bact. Biomass-Bioenergy 2003, 25, 427–433.
  81. Mauerhofer, L.M.; Zwirtmayr, S.; Pappenreiter, P.; Bernacchi, S.; Seifert, A.H.; Reischl, B.; Schmider, T.; Taubner, R.S.; Paulik, C.; Rittmann, S.K.M.R. Hyperthermophilic Methanogenic Archaea Act as High-Pressure CH4 Cell Factories. Commun. Biol. 2021, 4, 289.
  82. Jarrell, K.F.; Kalmokoff, M.L. Nutritional Requirements of the Methanogenic Archaebacteria. Can. J. Microbiol. 1988, 34, 557–576.
  83. Battino, R. The Ostwald Coefficient of Gas Solubility. Fluid Phase Equilib. 1984, 15, 231–240.
  84. Lu, J.X.; Murray, J. Biochemistry, Dissolution and Solubility; StatPearls Publishing: Treasure Island, FL, USA, 2020.
  85. Rusmanis, D.; O’Shea, R.; Wall, D.M.; Murphy, J.D. Biological Hydrogen Methanation Systems—An Overview of Design and Efficiency. Bioengineered 2019, 10, 604–634.
  86. Özbek, B.; Gayik, S. The Studies on the Oxygen Mass Transfer Coefficient in a Bioreactor. Process Biochem. 2001, 36, 729–741.
  87. Bajón Fernández, Y.; Cartmell, E.; Soares, A.; McAdam, E.; Vale, P.; Darche-Dugaret, C.; Jefferson, B. Gas to Liquid Mass Transfer in Rheologically Complex Fluids. Chem. Eng. J. 2015, 273, 656–667.
  88. Goel, R.; Komatsu, K.; Yasui, H.; Harada, H. Process Performance and Change in Sludge Characteristics during Anaerobic Digestion of Sewage Sludge with Ozonation. Water Sci. Technol. 2004, 49, 105–113.
  89. Hammadi, L.; Ponton, A.; Belhadri, M. Effects of Heat Treatment and Hydrogen Peroxide (H2O2) on the Physicochemical and Rheological Behavior of an Activated Sludge from a Water Purification Plant. Procedia Eng. 2012, 33, 293–302.
  90. Alves, S.S.; Maia, C.I.; Vasconcelos, J.M.T. Gas-Liquid Mass Transfer Coefficient in Stirred Tanks Interpreted through Bubble Contamination Kinetics. Chem. Eng. Process. Process Intensif. 2004, 43, 823–830.
  91. Montante, G.; Horn, D.; Paglianti, A. Gas-Liquid Flow and Bubble Size Distribution in Stirred Tanks. Chem. Eng. Sci. 2008, 63, 2107–2118.
  92. Leu, J.Y.; Lin, Y.H.; Chang, F.L. Conversion of CO2 into CH4 by Methane-Producing Bacterium FJ10 under a Pressurized Condition. Chem. Eng. Res. Des. 2011, 89, 1879–1890.
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to : , , , , , , ,
View Times: 689
Revisions: 9 times (View History)
Update Date: 15 Jul 2022
Video Production Service