Biotechnological Valorization of Waste Glycerol into Gaseous Biofuels—A Review

Abstract

The supply of waste glycerol is rising steadily, partially due to the increased global production of biodiesel. Global biodiesel production totals about 47.1 billion liters and is a process that involves the co-production of waste glycerol, which accounts for over 12% of total esters produced. Waste glycerol is also generated during bioethanol production and is estimated to account for 10% of the total sugar consumed on average. Therefore, there is a real need to seek new technologies for reusing and neutralizing glycerol waste, as well as refining the existing ones. Biotechnological means of valorizing waste glycerol include converting it into gas biofuels via anaerobic fermentation processes. Glycerol-to-bioenergy conversion can be improved through the implementation of new technologies, the use of carefully selected or genetically modified microbial strains, the improvement of their metabolic efficiency, and the synthesis of new enzymes. The present study aimed to describe the mechanisms of microbial and anaerobic glycerol-to-biogas valorization processes (including methane, hydrogen, and biohythane) and assess their efficiency, as well as examine the progress of research and implementation work on the subject and present future avenues of research.

1. Introduction

The supply of waste glycerol is rising steadily—spurred, in part, by the increased global production of biodiesel, which in turn is driven by biogenic fraction minimums required in conventional fuel [1]. Global biodiesel production totals about 47.1 billion liters and is a process that involves the co-production of waste glycerol, accounting for over 12% of total esters produced, regardless of the catalyst or technological process used [2]. Waste glycerol is also generated during bioethanol production and is estimated to account for 10% of the total sugar consumed on average [3]. The influx of surplus crude glycerol from the biofuels industry has led to a drastic drop in glycerol prices from 480 USD/ton in 2002 to 110 USD/ton in 2021, making it a waste product that requires significant expenditure to dispose of or reuse. This directly impacts the cost-effectiveness of biodiesel and bioethanol production [4]. Figure 1 shows the global growth of biodiesel and bioethanol production [5].

Valorization and processing of raw glycerol into value-added products is an urgent and growing priority, as well as a contributor to the viability of a biofuel-based economy [6,7]. Research on technologies for converting glycerol into value-added products has included both chemical and biological processes [8,9]. Biological pathways have several advantages over chemical conversion processes, including higher tolerance to contamination, greater environmental friendliness thanks to the lower temperatures and pressures involved, no consumption of chemicals, reduced secondary contamination, and, in many cases, a simple process design that provides versatility and ease of operation. Another added benefit is the large variety of glycerol-metabolizing microorganisms that have been identified [10].

Glycerol generates more reducing equivalents than other carbon sources (e.g., glucose or xylose), thus making it possible to produce biofuels and reduced chemicals at higher efficiency [11]. The abundance, relatively low price, and high reduction capacity of glycerol have made it an attractive feedstock to exploit for the production of various biofuels [12]. These features have significantly increased interest in developing microbial methods for the valorization of waste glycerol into gaseous biofuels, resulting in an abundance of available technologies. As shown in Figure 2, there has been a marked and fast-growing interest in research on the bioconversion of fuel to hydrogen (H₂), methane (CH₄), and, to a lesser extent, biohythane.

Although certain microorganisms that can process and metabolize glycerol into value-added products or biofuels have been selected and tested under natural conditions, their industrial applications have often been limited by relatively low yields and production rates [13]. Thus, interventions are needed to modify and increase the efficiency of natural enzymes and accelerate metabolic pathways to maximize industrial benefits [14]. Process improvement draws on research regarding metabolic engineering, genetic modification, and upgrading/optimization current processes [15,16]. The most common refinements include: eliminating transcriptional repression and inhibiting enzyme feedback; reducing the generation and accumulation of undesired by-products in the medium; enhancing and improving the medium via micro-nutrient amendment; reducing energy consumption and cost of the system; expanding the substrate and end product portfolio; and enhancing strain tolerance to inhibitors and environmental stress [17,18]. The integration of protein engineering, systems biology, and synthetic biology into metabolic engineering seems to have extended strain engineering from local modification to total system optimization. Combining insights from various omics technologies, such as genomics, transcriptomics, proteomics, and fluxomics, has provided an in-depth understanding of how glycerol metabolism proceeds in bioenergy systems and how whole microbial communities are regulated at the system level [19].

Despite the highly advanced research on waste glycerol bioconversion into gaseous biofuels and the fast proliferation of such processes, there is still a real need to seek new technologies as well as refine the existing ones. Earlier reports on the use of waste glycerol as a substrate for the production of gaseous biofuels focused mainly on methane fermentation. There are few papers analyzing the possibility of using this organic substrate in hydrogen production processes, while the biological conversion of glycerol into biohythane has not yet been presented. This study aimed to describe the mechanisms of microbial and anaerobic glycerol-to-biogas valorization processes (including CH₄, H_2, and biohythane), assess their efficiency, examine the progress of research and implementation work on the subject, and present future avenues of research. Glycerol, or propane-1,2,3-triol, also commonly known as glycerin, can be obtained through organic synthesis and biotechnological processing. Its considerable supply on the market comes from industrial biodiesel production [20].

2. Glycerol Production and Applications

Biodiesel—fatty acid methyl ester (FAME)—is an alternative fuel produced from vegetable oils, animal fats, waste fats, and other fatty feedstocks with suitable characteristics and properties. Typical feedstocks for biodiesel production are rapeseed oil, soya oil, sunflower oil, palm oil, and oil obtained from microalgae [21]. Cattle/sheep/poultry tallow, animal oil, used cooking oil, fish oil, jatropha oil, and coke have also been used [22,23].

Biodiesel is synthesized by transesterifying lipids (triacylglycerols) with a simple alcohol, such as methanol or ethanol (alcoholysis) (Figure 3). This reaction is usually catalyzed by NaOH, KOH, or sulfuric acid (VI) [24,25]. Chemical transesterification by alkaline catalysts is a fast-progressing reaction, which means that high volumes of triglycerides can be converted into their respective methyl or ethyl esters. The reaction will convert approx. 10% of the feedstock into glycerol [26,27]. However, it is difficult to separate the pure glycerol from the crude fraction due to the type and amount of impurities, such as methanol, fatty acid mono- and diacylglycerols, free fatty acids, and soaps produced during the biodiesel production process. Purified glycerol is a value-added product of great commercial and economic importance.

Glycerol is also formed during bioethanol production via alcoholic fermentation [28]. These biochemical pathways allow anaerobic microorganisms to regenerate the NAD (nicotinamide adenine dinucleotide) consumed during glycolysis. The pyruvate formed during the last step of glycolysis is then reduced to ethanol (ethyl alcohol) via a two-step process that also oxidizes the NADH (reduced form of NAD) formed during glycolysis to NAD while releasing carbon dioxide [29]. In the first step, pyruvate is converted to ethanal (acetaldehyde) and carbon dioxide by pyruvate decarboxylase. The second step reduces ethanal to ethanol by alcohol dehydrogenase (ADH), with NADH oxidized to NAD [30]. Alcoholic fermentation does not convert all of the glucose to bioethanol [31]. A portion remains unconverted to bioethanol, while at the same time some of the bioethanol is subject to further reactions, which generate glycerol as a by-product [32]. Yeast can also produce glycerol via osmotic regulation under stress [33]. It is important to note that the glycerol output of bioethanol production can vary depending on a multitude of factors, including the type of feedstock, fermentation parameters, type of yeast, etc. [34]. Bioethanol producers often strive to optimize their processes in order to minimize by-products, including glycerol [35].

The efficient use of crude glycerol is crucial for the further commercialization of biodiesel production and can significantly reduce the price of biodiesel [36]. One of the ways of neutralizing the waste glycerol fraction is to harness it for energy production, primarily by means of thermal conversion or bioconversion [37,38]. Major biofuel producers purify glycerin waste through filtration or fractional vacuum distillation [39,40], as well as by using chemical conversion or selective catalytic oxidation [41,42]. The literature also includes numerous reports on reusing the glycerin fraction in biological methods as a carbon source to fuel microbial growth. From a commercial standpoint, converting waste glycerol into gaseous biofuel via microbial bioconversion represents a major opportunity for the fuel industry. Well-selected microbial groups are capable of converting glycerol into value-added gas metabolites, and the process itself does not require that the glycerol fraction be purified [43]. The cost-effectiveness of the process can be further boosted by replacing other carbon sources with cheaper waste feedstock. In this light, crude glycerol—seen as industrial waste and available in large quantities—would seem to be a good candidate for the production of biogases with various compositions and properties [44,45]. A basic division of the methods used to valorize and reuse waste glycerol from biodiesel production is presented in Figure 4 [37,38,43].

3. Biomethane Production

A common method for neutralizing waste glycerol is by processing it via an anaerobic digestion (AD) process. AD utilizes biological conversion processes occurring under aerobic conditions, producing CH₄-rich biogas and digestate (usually utilized as organic fertilizer) [46,47,48]. Each step of the digestion is mediated by specialized microorganisms that hydrolyze polymeric substances through enzyme action [49,50,51]. They are further broken down into fatty acids, alcohols, H_2, and CO₂ [52]. AD is a process that consists of the following steps: hydrolysis, acidogenesis, acetogenesis, and methanogenesis. Hydrolysis involves the enzymatic biological degradation of complex organic compounds. Enzymes produced by anaerobic, hydrolytic microorganisms are used in this process [53]. During acidogenesis, the hydrolysates are further degraded into simpler compounds, mainly organic acids, alcohols and aldehydes, CO_2, and H₂ [54]. Acetogenesis is the process by which organic acids are converted to acetic acid, CO_2, and H₂. Acetate is synthesized via one of two pathways. The first oxidizes the fatty acids produced in the acidogenesis phase, releasing H₂; in the second phase, it uses H₂ to reduce CO₂. Acetogenesis is the most important phase of the AD process [55]. Methanogenesis is the last phase of AD. CH₄ is produced via two pathways. In the first pathway, it is produced from acetic acid, and in the second from H₂. In addition to CH₄, methanogenesis also produces CO₂, H₂S, ammonia, and water [56]. The CH₄-producing organisms involved in this process are archaea [57]. A step-by-step conversion diagram for the anaerobic digestion (AD) of glycerol (including the end products) is shown in Figure 5.

Waste glycerol is usually included as a part of a wider mix of organic feedstocks that can be readily biodegraded in anaerobic digesters. The process by which bioreactors are fed with a mixture of different feedstocks is called co-digestion [60]. Many previous studies and data from full-scale plants have shown amendment with waste glycerol to have a very positive effect on the AD process and its performance [61]. This is owing to the high energy value of the substance, which increases the organic load rate (OLR) in the digesters while also directly and significantly boosting biogas yield and quality profile [62]. Glycerol is readily biodegradable under anaerobic conditions, though it does typically require higher hydraulic retention times (HRT) in digesters [63]. In many cases, its incorporation into the organic feedstock mix results in optimal OLR values and improves the carbon-to-nitrogen (C/N) ratio, which in turn reduces the levels of ammonia—a major inhibitor of AD in bioreactors [64].

It has also been shown that waste glycerol-based processes produce digestate of better fertilizing quality [65]. If the digestate from glycerol (co-)digestion is to be reused as an organic fertilizer, humus balancing in the soil becomes an important consideration [66]. This is an important factor when growing cereals, rapeseed, root crops, and tuber crops—plantations that cause a negative organic matter balance in the soil. Such organic fertilizer can be successfully used in humus-poor, low-quality soils, including light, degraded, or marginal soils. The organic-rich digestate from glycerol digestion can be used as a first-line treatment for restoring the humus index, which has an enormous impact on soil productivity, improving the water and sorption (ion binding) capacity, structure, and biological activity [67]. The literature often references the concept of “residual effect”, which refers to the crop yield increase upon the use of organic fertilizers that cannot be achieved with mineral fertilizers. The residual effect—such as the one from digestate derived from AD of glycerol—improves the physical, chemical, and biological properties of the soil [67].

Because of the growing supply of waste glycerol and its promising properties as an AD feedstock, an increasing number of researchers and industrial-scale plant operators have been striving to optimize its use [68]. Mesophilic digestion is the most popular AD process for this purpose, as evidenced, for example, by the studies on mesophilic co-digestion of sewage sludge and glycerol [69,70]. Research has shown that the best results are achieved when glycerol accounts for 25 to 60% of the total OLR fed into the digestate. Keeping the proportions within this range can boost biogas production by 82–280%. Increasing the share of glycerol in total OLR beyond 70% inhibited the process and hamstrung the result, i.e., biogas yields, CH₄ fractions, and digestate profile [71,72]. Baba et al. [73] compared the energy balance of anaerobic digestion in a digester with an active volume of 30 m³, fed with a mix of crude glycerol and excess sludge. Once a day, 3.5 m³ of sludge and 5 to 75 L of crude glycerol were fed into the reactor. The best-performing variant provided a net energy output 106% higher than when digesting sewage sludge alone. The added benefit of the process was that the soil fertilized with the digested sludge showed improved structure and yielded 20% more grass for haylage.

Athanasoulia et al. [74] used a continuous stirred-tank reactor (CSTR) to co-digest waste glycerol and sewage sludge under mesophilic conditions. The addition of up to 4% glycerol (v/v) improved biogas production by 3.8–4.7 times. Another study investigated the impact of glycerol addition on biogas yields at a pilot scale [75] and found a low dose of glycerol (0.63% v/v) to be the optimal one. This variant had a CH₄ yield of 1.3 m³/L crude glycerol [75]. Silvestre et al. [76] studied the effect of adding crude glycerol to AD-treated sewage sludge under thermophilic and mesophilic conditions. The addition of glycerol at the thermophilic temperature range had a negative impact on the stability and performance of the process, even at low doses. In contrast, the process performed steadily within the mesophilic temperature range, with a 148% increase in CH₄ output with 1% glycerol in the influent v/v (27% of influent COD). The addition of glycerol to sewage sludge can be used to effectively balance the C/N ratio. However, no improvements in biogas yield were observed when the glycerol content exceeded 1% v/v in the influent, likely due to the high C/N [76].

These results suggest that crude glycerol can be used as a co-feedstock for the AD of sewage sludge, although various parameters should be taken into account—depending on the glycerol characteristics and operating conditions—to ensure digester stability and optimal performance [76]. Another study [77] concluded that feeding glycerol at up to 2% v/v did not enhance methane production. A detailed analysis of the process kinetics and biochemical dynamics of the microbial community found synergisms between glycerol and sewage sludge, with the result being improved biogas and CH₄ yields. Doses of up to 2% v/v glycerol provided a good balance between increasing the organic loading rate and minimizing the impact on hydraulic retention time. During continuous operation over 200 days, feeding glycerol at up to 2% v/v sharply increased the organic load by 70% and resulted in a 50% increase in CH₄ production, with nominal yields of 1248 ± 58 mL CH₄/L/d [77].

Alves et al. [78] investigated how the addition of 1 and 3% crude glycerol v/v from biodiesel production affected the AD primary sewage sludge. Process stability and performance were monitored using such parameters as pH, COD, VS, C/N, and volatile acid/alkalinity ratio. At 1% glycerol, the CH₄ yield was 223.8 mL CH₄/g VS, whereas at 3% glycerol, the CH₄ output was 368.8 mL CH₄/g VS. This translates to an increase in CH₄ yield of 61% and 167%, respectively, against the sludge-only control (138.2 mL CH₄/g VS). Peak daily biogas production (56.8 mL CH₄/g VS) was achieved with the highest glycerol fraction. However, an initial instability period (with methanogenesis inhibition) of 5.8 days was noted, likely attributable to the accumulation of intermediate volatile acids [78]. Another study by Alves et al. [79] aimed to evaluate the production of biogas (including CH₄) from the anaerobic co-digestion of primary sludge (PS) from sewage treatment, food waste (FW), and crude glycerol. To study the effect of glycerol on AD stability and performance, biochemical methane potential (BMP) tests were performed at different glycerol concentrations—1 and 3% v/v. A modified Gompertz model was used to describe biogas and CH₄ production. The study demonstrated that the small increases in the organic load from glycerol addition led to significant improvements in biogas and CH₄ production. Although methanogenesis was temporarily inhibited in the BMP tests with the highest glycerol concentration—suggesting a longer adaptation period for methanogenic Archaea—CH₄ production was later restored and even maximized under these conditions. The biogas and CH₄ production reached 432.4 mL/g VS removed and 343.3 mL/g VS at 1% glycerol, respectively, and 692.6 mL/g VS _removed and 525.7 mL CH₄/g VS at 3% glycerol, respectively. This translates to an increase in CH₄ production of 45.4% and 122.7% (for 1 and 3% glycerol, respectively) compared to the results for PS + FW [79].

Waste glycerol has also been co-digested with wastewater. Siles et al. [80] co-digested wastewater from biodiesel production with 1% v/v glycerol, obtaining 310 mL of CH₄/g COD _removed. Anaerobic co-digestion was shown to require less clean water and nutrients, resulting in commercial and environmental benefits [80]. Fountoulakis and Manios [81] tested the co-digestion of olive mill wastewater and slaughterhouse wastewater with 1% v/v glycerol. CH₄ production was 1210 ± 205 mL CH₄/d, which represents a 731 mL CH₄/d increase compared to the no-glycerol variant (479 ± 38 mL CH₄/d) [81]. The authors of [81] also co-digested the wastewater organic fraction of municipal solid waste (40% fruit, 25% potatoes, 25% vegetables, 8% bread, and 2% paper) with 1% v/v glycerol. CH₄ yield amounted to 2094 mL CH₄/d—meaning an increase of 694 mL CH₄/d compared to the no-glycerol variant (1400 mL CH₄/d).

Another common feedstock mix with waste glycerol provides for co-digestion of the glycerol with manure/slurry of various origins and properties. Astals et al. [82] demonstrated that thermophilic digestion of a feedstock mix containing pig manure and crude glycerol (3% v/v) yielded 180% more biogas (0.47 L biogas/g VS) compared with a single-feedstock process (pig manure only). The increase was mainly attributed to a balanced C/N ratio, the energy value of the organic matter fed in, and the optimal and stable OLR [82].

However, the authors emphasize that the high levels of organic biodegradable matter in the digestate preclude it from being a universal fertilizer—rather, its use is limited to light soils, mineral soils, and other soil types with a low humus content [82,83]. A high glycerol content in the organic feedstock can directly lead to high levels of organics in the digestate, thus making it difficult to stabilize [84]. The presence of biodegradable organic compounds in natural fertilizers used on agricultural soils or on degraded land under remediation can lead to sanitary difficulties, the spread of nuisance odors, or negative environmental impacts (surface runoff, water erosion, wind erosion) [85]. The quality and profile of the digestate, the options for its use as a fertilizer, and the conditions of such fertilization should always be taken into account when designing systems for the AD of glycerol [86].

A study by Usack and Angenent [87] used glycerol as a co-digestion feedstock with dairy cattle manure in a CSTR for 900 days under mesophilic conditions. The highest biogas production was noted when the ratio of dairy manure to crude glycerol (VS basis) was 62:38. The CH₄ production reached 549 ± 25 mL CH₄/g VS at 3.2 g VS/L/d OLR. The authors noted encountering some processing hurdles, primarily foaming in the anaerobic reactor, which caused limited diffusion of gas metabolites produced by anaerobic bacteria, flotation of the reactor medium, and problems with removing the digested mass from the reactor [87]. Foaming may be triggered by the presence of proteins or surfactants, which can occur in waste from dairy cow farming. These substances can originate from the milking station wash water and enter digesters with the input manure [88]. Glycerol input also significantly increases the fraction of biodegradable organic matter in the feedstock [89]. This leads to faster production of gas but also faster formation of foam, which, in the long term, can coalesce into a thick layer of scum on the surface of digesters. The scum acts as a barrier that prevents gases from diffusing at the liquid–gas interface and can lead to flow restriction or, in extreme cases, completely inhibit the flow of gaseous metabolites produced by anaerobic bacteria [90]. Ultimately, this leads to the accumulation of digestion metabolites in the medium and the inhibition of organic compound breakdown under anaerobic conditions [91].

Inhibition of anaerobic digestion of swine manure with added crude glycerol has been the subject of a study by Fierro et al. [92]. Added glycerol at over 8% v/v caused medium acidification, methanogenesis failure, and a rise in the CO₂ content in the gas metabolites. Significant increases in H₂S levels were also observed. As a result, the system had to be modified and expanded. A post-stabilization stage had to be introduced to complete the biodegradation of the feedstock organics, especially proteins and lipids [92]. The observations of Fierro et al. [92] have been reproduced by other researchers [93,94]. Glycerol is an organic substrate that, despite its positive effect on AD and its performance, should be used with caution in feedstock mixes used in bioreactors [95]. Exceeding the threshold of organic levels, and thus the OLR value, overloads the system [96,97]. In the long term, this causes volatile fatty acids (VFAs)—produced during the acidogenesis step of AD—to accumulate [98]. This, in turn, decreases pH, inhibits the metabolism of methanogenic Archaea, kills anaerobic microorganisms in the microbial community, and halts methanogenesis altogether [99].

Nevertheless, crude glycerol can be an extremely attractive candidate for an AD co-substrate due to its widely reported boosting effect on biogas production and composition [78]. It is recommended that optimization procedures and multivariate biogas tests be conducted to select the best-performing variants and identify potential problems early [86]. This is important both in terms of the process itself and the later utilization of its products. Oliveira et al. [100] co-digested Sargassum sp. macroalgae with glycerol and waste frying oil (WFO), looking for optimal CH₄ production parameters. Three variables were investigated: % of the total algal mass (%TS Sargassum sp.), type of co-feedstock (glycerol or WFO), and co-feedstock concentration (g/L). The biochemical methane potential (BMP) of Sargassum sp. was 181 ± 1 L CH₄/kg COD. Co-digestion with glycerol or WFO increased this parameter by 56% and 46%, respectively, whereas the methane production rate (k) was 38% and 19% higher for glycerol and WFO, respectively. Peak BMP (283 ± 18 L CH₄/kg COD) was obtained in the 0.5% TS, 3.0 g/L glycerol trial [100]. Optimization of anaerobic digestion has also been pursued by Takeda et al. [101]. A Central Composite Rotational Design (CCRD) was built to experimental variables such as time (16.6, 20, 25, 30, and 33.4 days), glycerol content (0.43, 0.70, 1.10, 1.50, and 1.77%), and substrate/inoculum ratio (0.23, 0.30, 0.40, 0.50, and 0.57 g COD/g VSS). The optimization was successful in maximizing the efficiency of organic matter removal (90.15%) and specific biogas production (403.15 mL/g VSS) for the parameters: 33.2 days, glycerol content of 1.71%, and substrate/inoculum ratio of 0.37 g COD/g VSS. Using a modified Gompertz model of the optimal condition performed, the specific biogas production obtained was, on average, 20.3 times higher compared to the mono-digestion of leachate [101].

All studies to date have shown that overloading a system with glycerol-derived organic compounds leads to the inhibition of their breakdown under anaerobic conditions [102], decreases CH₄ in the biogas [93], increases hydrogen sulfide levels [103], produces unstable digestate [104], and, in extreme cases, causes failures of reactors, stirrers, pumping systems, or cogeneration units [105]. Each introduction of glycerol into a tried and tested substrate mix requires systematic and skillful adaptation of the system and acclimation of the bacterial community to be used for a multi-step AD process [106].

A well-adapted bacterial community allows operators to use higher glycerol contents in the feedstock for efficient CH₄ production and value-added organic fertilizer [107]. Many have reported good results with two-step digestion processes, using physically separated sequences of acidogenic and methanogenic digestion [94]. Such separation provides better process control and reduces the impact of the intensive VFA production on the methanogenic step [108]. Researchers and operators should focus their efforts on optimizing digestion parameters and individual process steps, including C/N ratio, OLR, HRT control during hydrolysis (acidogenic and methanogenic steps), and monitoring of the bacterial community composition [109]. Examples of waste glycerol use in AD, key process parameters, and performance metrics are given in

Table 1. Comparison of the literature data on glycerol used in AD.

Base Substrate	Experimental Parameters	Glycerol Volume	Methane/Biogas Production/Yield	Ref.
70% primary sludge and 30% activated sludge	5.5 L CSTR reactor, 56 °C, 25 rpm, pH 6.8 ± 0.1, HRT 20 ± 1 days, OLR 2.5 g COD/L/d	1.5% v/v	0.25 ± 0.04 L CH4/L/d	[76]
	5.5 L CSTR reactor, 56 ± 1 °C, 25 rpm, pH 7.1 ± 0.2, HRT 22 ± 2 days, OLR 2.4 ± 0.1 g COD/L/d	1.6% v/v	0.23 ± 0.01 L CH4/L/d
	5.5 L CSTR reactor, 56 ± 6 °C, 25 rpm, pH 6.9 ± 0.1, HRT 18 ± 3 days, OLR 3.6 ± 0.6 g COD/L/d	2% v/v	0.53 ± 0.14 L CH4/L/d
	5.5 L CSTR reactor, 55 °C, 25 rpm, pH 6.4 ± 0.2, HRT 19 ± 1 days, OLR 3.4 ± 0.5 g COD/L/d	3% v/v	0.33 ± 0.15 L CH4/L/d
	5.5 L CSTR reactor, 36 ± 1 °C, 25 rpm, pH 7.6 ± 0.2, HRT 20 ± 2 days, OLR 3.0 ± 0.3 g COD/L/d	1.2% v/v	0.62 ± 0.07 L CH4/L/d
	5.5 L CSTR reactor, 36 ± 1 °C, 25 rpm, pH 7.2 ± 0.2, HRT 20 ± 1 days, OLR 3.2 ± 0.4 g COD/L/d	2.4% v/v	0.66 ± 0.09 L CH4/L/d
Waste activated sludge (WAS)	60 L CSTR, 37 °C, pH 6.8–7.2, HRT 14 days	2% v/v	100 ± 8 L biogas/d	[74]
	60 L CSTR, 37 °C, pH 6.8–7.2, HRT 16.4 days	3% v/v	90 ± 2.8 L biogas/d
	60 L CSTR, 37 °C, pH 6.8–7.2, HRT 19.7 days	4% v/v	80 ± 2.6 L biogas/d
Primary sewage sludge	50 L CSTR, 35 ± 0.1 °C, HRT 20 days	0.63% v/v	1.3 m3 CH4/L crude glycerol	[75]
Primary sewage sludge	37 °C, pH 7.24, 20 days	1% v/v	223.8 mL CH4/g VS	[78]
		3% v/v	368.8 mL CH4/g VS
Primary sewage sludge and food waste	37 °C, pH 7.48–7.13, 20 days	1% v/v	432.4 mL biogas/g VS; 343.3 mL CH4/g VS	[79]
		3% v/v	692.6 mL biogas/g VS; 525.7 mL CH4/g VS
Wastewater from the biodiesel production process	1 L Pyrex reactors, 37 °C, pH 7.38	15% v/v	310 mL CH4/g COD removed	[80]
Olive mill wastewater and slaughterhouse wastewater	4 L CSTR with 3 L working volume, 35 °C, pH 7–7.5, 46 days, HRT 23–25 d	1% v/v	1210 ± 205 mL CH4/d	[81]
Organic fraction of municipal solid waste (40% fruit, 25% potatoes, 25% vegetables, 8% bread, and 2% paper)	4 L CSTR with 3 L working volume, 35 °C, pH 7–7.5, 46 days, HRT 23–25 d	1% v/v	2094 ± 92 mL CH4/d
Landfill leachate	1 L batch reactor, 0.8 L reaction volume, 37 °C, pH 7, 33.2 days	1.71%	403.15 mL biogas/g VSS	[101]
Sargassum sp.	Work volume of 50 mL, 37 °C, pH 7.24, 42 days	3 g/L	283 ± 18 mL CH4/d	[100]
Cattle manure and noodle factorysludge	50 m3 pilot plant (30-m3 working volume), 35 °C	1 mL glycerol/L/d, (47% purity)	358 mL CH4/g COD removed; 149.1 mL CH4/g COD	[73]
Dairy manure	4.5 L reactor, 37 °C, 900 days, HRT 25 days, OLR 2.93–3.90 g VS/L/d	Dairy manure– glycerol proportion: 91:9; 75:25; 62:38; 51:49	1.4 L CH4/L/d CH4 yield: 549 L/kg VS	[87]
Pig manure	5.5 L semi-CSTR, 55 °C, 60 rpm	3% v/v	0.47 L biogas/g VS (180% higher)	[82]
Swine manure	25 L reactor, 34 °C, 80 days	2–8% v/v	1.4 L CH4/L/d CH4 yield: 380 L/kg VS	[92]
Swine effluent	5.5 L semi-continuous stirred tank reactors with a working volume of 4 L, 35 °C, pH 6.5, 60 rpm, 20 days, OLR 1.9 ± 0.1 g VS/L/d	4% v/v	0.74 ± 0.03 L biogas/g VS	[83]
	5.5 L semi-continuous stirred tank reactors with a working volume of 4 L, 35 °C, pH 6.5, 60 rpm, 20 days, OLR 1.7 ± 0.1 g VS/L/d		0.78 ± 0.02 L biogas/g VS

.

4. Biohydrogen Production

4.1. Photofermentation

H₂ meets the criteria of a low-impact energy carrier [110,111]. Its calorific value ranges from 10.8 to 12.75 MJ/Nm³, making it suitable for use in heating, power generation, and air/car transportation [112,113]. Currently, H₂ is used on a limited basis, mostly in the refining industry, fuel cells, and space technology [114]. The main barrier to use is the lack of viable methods of production and storage—ones that would be technologically feasible and cost-effective [115]. Conventional H₂ production mainly consists of thermochemical methods, including combustion, gasification, pyrolysis, thermochemical liquefaction, or water pyrolysis [116]. However, such solutions are energy-intensive, pollution-generating, and mired in high investment costs [117]. It is estimated that almost 95% of H₂ is produced by converting fossil fuels [118]. Biomass-based technologies, as well as methods that harness the biological processes of microorganisms, are becoming more and more viable as a means of H₂ production [119]. They consist mostly of organic-feedstock fermentation, carried out by specialized groups of bacteria, or biochemical processes in the cells of selected microalgae species [120]. Crude glycerol can be converted to H₂ via two different fermentation processes: photofermentation and dark fermentation. The former requires a light source to proceed.

Photofermentation is performed by anaerobic bacteria, mainly green/purple sulfur and non-sulfur bacteria such as Rhodobacter sphaeroides, Rhodobacter capsulatus, Rhodobacter sulidophilu, Rhodopseudomonas palustris, Rhodopseudomonas sphaeroides, and Halobacterium halobium [121]. These species are capable of breaking down organic acids into H₂ and CO₂ [122]. Nitrogenase—the primary photofermentative enzyme—is a metalloprotein complex responsible for the biological fixation of molecular nitrogen and commonly found in both archaea and bacteria. The nitrogen fixation process can produce H₂, thus maintaining redox homeostasis [123,124]. Nitrogenase activity can be inhibited by excessive C/N, oxygen, or ammonia, among other factors [125]. Chemical energy in the form of a proton gradient is produced from sunlight. This energy can be used to power various cellular processes, including a reduction in nitrogenase via reverse electron flow. On the other hand, reduced nitrogenase uses the hydrolysis of ATP, produced by photophosphorylation, to reduce metabolically produced protons to H₂. The normal physiological function of nitrogenase is to reduce dinitrogen to ammonia, with the reduced release of one H₂ per N_2. However, nitrogenase turnover can still occur even in the absence of other reducible substrates, reducing protons to H₂. The process follows the flowchart presented in Figure 6 [122].

The literature data indicates that Rhodopseudomonas palustris is the most popular microorganism for the photofermentation of waste glycerol, owing to its particular enzymatic activity [126]. Pott et al. [127] found that an R. palustris community fed with waste glycerol grew at a rate of 0.074 h⁻¹. In this particular process, glycerol was converted at a conversion efficiency of approximately 90% at a reaction rate of 34 mL H₂/g_dw/h. Inhibition of microbial growth and H₂ production was noted during the experiment, which was attributed to the contaminants in the crude glycerol. Saponified fatty acids were determined to be the main inhibitors of the process [127].

Sabourin-Provost and Hallenbeck [128] conducted a comparative study with both pure and crude glycerol as feedstock for photofermentative H₂ production by Rhodopseudomonas palustris. The yields were approx. 6 moles of H₂/mole glycerol across all variants. The impurities present in crude glycerol were not found to negatively affect the process of microbial growth in this case [128]. In another study, Ghosh et al. [129] endeavored to optimize a photofermentative H₂ production process by Rhodopseudomonas palustris. Response Surface Methodology (RSM) was applied to select the process conditions and parameters important for glycerol-to-H₂ conversion. This optimization protocol was used to determine the interactive effects among the process parameters, such as the concentration of crude glycerol and glutamate, light intensity, nitrogen source, and N levels, on H₂ production efficiency.

Under optimal conditions (light intensity of 175 W/m², 30 mM glycerol, and 4.5 mM glutamate), 6.69 moles H₂/mole of crude glycerol were obtained, which translates to 96% of the theoretical yield. Determination of nitrogenase activity and expression levels showed that process variables produced relatively little variation in nitrogenase protein levels, whereas nitrogenase activity varied considerably, with peak nitrogenase activity (228 nmol C₂H₄/mL/min) achieved at the optimal central point [129]. Zhang et al. [130] used modeling to simulate the entire growth phase of R. palustris in order to maximize H₂ production. Two piecewise models were designed, both fitted via batch experiments, by solving the underlying optimal control problems through stable and accurate discretization techniques. The optimal initial glutamate-to-glycerol ratio was 0.25, regardless of the initial biomass concentration. The glycerol conversion efficiency was found to correlate with the initial biomass concentration, with a computational peak of 64.4%. By optimizing a 30 day industrial batch process, an H₂ production rate of 37.7 mL g biomass/h was achieved, while the glycerol conversion efficiency was maintained at 58% [130].

Jiang et al. [131] investigated the correlation between H₂ yield improvement during photofermentation by Arundo donax L. and glycerol amendment. Glycerol concentrations of 0, 10, 15, 20, and 30 g/L were tested. The highest H₂ yield improvement (79.2 mL/g substrate) was obtained at 15 g/L added glycerol, translating to a 294% increase in production compared with the Arundo donax L. mono-substrate system. Under the optimal glycerol addition (15 g/L), process factors were as follows: glycerol/Arundo donax L. ratio: 1/1, C/N ratio: 25.1, initial medium redox potential (Eh): 57 mV, solid/liquid ratio: 1/68. A canonical correlation analysis (CCA) found that initial and final Eh had the strongest relationship with photofermentation yield improvement. Furthermore, Pearson’s correlation analysis showed that the Arundo donax L./glycerol ratio played a key role in the photofermentative H₂ production process [131].

Process parameters play a key role in photofermentation. The process requires a temperature between 30 °C and 36 °C, a near-neutral pH (6.8–7.5), and a source of bright 400–1000 nm light (6–10 klux) [112,132]. Using the right illumination intensity boosts H₂ production rates and yields. However, due to high running costs, alternating light–dark cycles are usually used, with a 12/12 h interval being the most common choice [133]. H₂ production efficiency is also directly related to the type and design of the photobioreactor (PBR) [134]. The most common are tubular, column, and flat-plate PBRs [135]. PBRs are typically closed and hermetically sealed, preventing oxygen penetration, contamination, and the growth of competing microbial species. PBRs are similar in design to reactors used for cultivating and growing microalgae [136,137]. Ross and Pott [138] have demonstrated that one way to increase H₂ production is to develop improved photobioreactor (PBR) systems. Du Toit and Pott [139] compared H₂ production from glycerol by Rhodopseudomonas palustris using planktonic cultures and immobilized cells in packed-bed or fluidized-bed reactors. The fluidized-bed PBR achieved a maximum specific H₂ production rate and substrate conversion efficiency of 15.74 ± 2.2 mL/g/h and 43%, respectively, outperforming the conventional planktonic culture and the packed-bed PBR. There have also been attempts to use transparent polyvinyl-alcohol (PVA) cryogel beads to immobilize Rhodopseudomonas palustris for long-term H₂ production [139].

4.2. Dark Fermentation

Bacteria-mediated dark fermentation is one of the most effective means of bioconverting biomass (including glycerol) to H₂ [140]. The primary sub-types of such fermentation are butyrate/butanol fermentation, conducted by the genus Clostridium sp., and mixed-acid fermentation, typical of the family Enterobacteriaceae (Enterobacter aerogenes, Escherichia coli, Vibrio cholerae, Klebsiella pneumoniae, Shigella dysenteriae), Klebsiella sp., Bacillus sp., Clostridium pasteurianum, Clostridium butyricum, Thermotoga neapolitana, or mixed microbial communities [141]. In commonly used technological processes, microbial cultures are fed with glycerol medium, containing waste glycerol, nutrients (nitrogen, potassium, and other compounds), and sometimes yeast. The process is conducted in an anaerobic environment at a temperature of approx. 30–40 °C [142].

H₂ release by bacteria results from pyruvate molecules being broken down into acetyl-CoA. This reaction is catalyzed by pyruvate-ferredoxin oxidoreductase. Ferredoxin plays a key role in electron distribution within the cell as an electron carrier. Reduced ferredoxin can transfer electrons to iron-containing hydrogenase, allowing protons to be used as a final electron acceptor, thus facilitating H₂ production. An example flowchart of H₂ production by Clostridium sp. bacteria is presented in Figure 7 [143].

Research to date has shown that the efficiency of glycerol-to-H₂ conversion via dark fermentation is not affected by impurities in the starting substrate, regardless of the degree of contamination [141]. Zahedi et al. [144] found that adding 1% v/v crude glycerol to the organic fraction of municipal waste almost doubled H₂ production in a process mediated by Eubacteria and Archaea. A trial using municipal waste as the sole feedstock resulted in a yield of 1.25 ± 0.29 LH₂/L. By comparison, H₂ production after amendment with glycerol (1% v/v) amounted to 2.32 ± 0.21 LH₂/L [144]. Trchounian K. and Trchounian A. [145] conducted a dynamic and kinetic analysis of glycerol-to-H₂ conversion by E. coli. Metabolic engineering of E. coli proved to directly improve dark fermentation efficiency. It was demonstrated that it was possible to redirect metabolic pathways, induce the DHAP production pathway, and block lactic, acetic acid, and ethanol production [145]. Hu and Wood [146] obtained an improved H₂-producing strain that yielded 0.68 mmol H₂L/hL in a glycerol medium. This represented a 20-fold increase in H₂ production compared to the precursor [146]. What is more, Sanchez-Torres et al. [147] and Tran et al. [148] identified uncharacterized genes (54% of the E. coli genome has been experimentally demonstrated; the remainder is uncharacterized or has been modeled through computational methods) that can be inactivated to boost H₂ production from glycerol. Some of the identified strains were able to produce 1.6 times more H₂ [147,148].

There have also been studies comparing E. coli with other bacteria in terms of the optimal fermentation conditions, process parameters, and H₂ yields [149]. Lo et al. [150] tested crude glycerol from the biodiesel industry for H₂ production using seven isolated H₂-producing bacterial strains (Clostridium pasteurianum, Clostridium butyricum, and Klebsiella sp.). Among the strains tested, C. pasteurianum CH₄ had the highest H₂ production performance under the optimal conditions at a temperature of 35 °C, an initial pH = 7.0, an agitation rate of 200 rpm, and a glycerol concentration of 10 g/L. When using pure glycerol as the carbon source for continuous fermentative hydrogen production, the average H₂ production rate was 103.1 ± 8.1 mL/h/L, with yields of 0.50 ± 0.02 moles H₂/mole glycerol. By comparison, when crude glycerol was used as the carbon source, the H₂ production rate and H₂ yields rose to 166.0 ± 8.7 mL/h/L and 0.77 ± 0.05 moles H₂/mole glycerol, respectively [150].

Kumar et al. [151] used Bacillus thuringiensis EGU45H-2 for fermentative hydrogen production from crude glycerol, which yielded H₂ at 0.646 mol H₂/mole glycerol consumed in a batch culture. Under continuous culture, after 2 days’ hydraulic retention time, B. thuringiensis immobilized on lignocellulosic materials (banana leaves—BL, 10% v/v) produced 0.386 mol H₂/mole glycerol consumed. The maximum yield was 0.393 mol H₂/mole glycerol consumed [151]. Ito et al. [152] evaluated the production of H₂ and ethanol from glycerol-containing biodiesel waste using Enterobacter aerogenes HU-101. H₂ and ethanol yields were inversely correlated with glycerol levels. The highest production of H₂ and ethanol—1.12 and 0.96 moles H₂/mole glycerol, respectively—was obtained at 1.7 g/L glycerol [152].

Experimental reports confirm that co-digestion of mixed carbon sources—for example, formate and glycerol—can significantly increase H₂ production. Trchounian et al. [153] found that H₂ production by E. coli BW 25113 wild type, supplemented with glycerol, reached 0.75 ± 0.03 mmol H₂/L at the end of the exponential growth phase (7 h growth). The action of formate (10 mM) and glycerol (10 g/L) at pH 6.5 led to a yield of 0.83 ± 0.05 mmol H₂/L at the early exponential phase (2 h growth). The same stimulatory effect of formate and glycerol on H₂ production was observed at pH 7.5 [153].

There have also been studies on how the type and concentration of heavy metals affect H₂ production and E. coli growth rate [154]. It was found that E. coli possessed four [Ni-Fe]-Hyd enzymes, the activity of which requires heavy metals. Furthermore, Mo is required to activate formate hydrogen lyase (FHL). It was found that low concentrations of Ni, Fe, and Mo ions, as well as combinations thereof (Ni²⁺ + Fe³⁺ (50 mM), Ni²⁺ + Fe³⁺Mo⁶⁺ (20 mM), and Fe3⁺ + Mo⁶⁺ (20 mM)), could increase bacterial biomass and H₂ production, mostly by way of glycerol fermentation under acidic conditions (pH 5.5 and pH 6.5). In contrast, Cu⁺ and Cu²⁺ (100 mM) were found to have no effect [154].

Pachapur et al. [155] verified the effect of added surface-active substances on H₂ production from glycerol in a mixed Enterobacter aerogenes and Clostridium butyricum culture. The process yielded up to 32.1 ± 0.03 mmol H₂/L medium under the optimized conditions of 17.5 g/L crude glycerol and 15 mg/L non-ionic surfactant (Tween 80). The increase in H₂ production was around 1.25-fold higher with Tween 80 than without, with a production of 25.56 ± 0.91 mmol/L [155]. Faber and Ferreira-Leitão [156] endeavored to optimize the conditions of the dark fermentation of waste glycerol. A statistical analysis showed that the optimal conditions were: pH = 5.5, glycerol concentration of 0.5 g/L, and volatile suspended solids of 8.7 g/L. H₂ production in the optimized process was 2.44 mol H₂/mole glycerol [156].

H₂ is often combined with microbial fuel cells. Sharma et al. [157] investigated the possibilities of converting glycerol into H₂ and electricity. Hydrogen bioreactors (HPB) and microbial fuel cells (MFC) were used for this purpose. A comparative analysis of pure and waste glycerol was carried out. A comparable H₂ production efficiency was achieved with 0.17–0.18 mol H₂/mol glycerol, regardless of the source of the substrate. The highest power density of 4579 mW/m³ was achieved with 2 g/L of pure glycerol. The observed power densities of waste glycerol ranged from 1614 to 2324 mW/m³ [157]. Chookaew et al. [158] conducted analog studies with a two-stage process combining dark fermentation with an MFC or a microbial electrolysis cell (MEC). The efficiency of H₂ production was 332 mL/L. 20% of the organic matter was biodegraded in the process. The two-chamber MFC produced a power density of 92 mW/m² and removed 50% of the COD. The columbic efficiency (CE) was 14%. When the cell was fed with 50% diluted fermentation product, similar power output (90 mW/m²) and COD removal (49%) values were achieved, but the CE was almost doubled (27%). When similar substrates were used to produce H₂ in two-chamber MECs, the diluted influent provided better performance, with the yield reaching a maximum of 106 mL H₂/g COD and 24% CE. Thus, dark fermentation coupled with MFC/MEC proved to be a viable option for converting waste glycerol into bioenergy [158].

Fermentative H₂ production from glycerol is influenced by many factors and system parameters, the most important being: glycerol type, origin and composition, presence of impurities, hydraulic retention time, type of digester, pH, temperature, and microbial strain [159]. Even trace amounts of oxygen in the system can inhibit hydrogenase activity when obligate anaerobes are used. Therefore, it is usually a safer choice to use facultative anaerobes, such as Clostridium sp. and Enterobacter sp., which better tolerate oxygen in the bioreactor [160]. The optimal pH range for efficient hydrogen-producing fermentation of glycerol is 5.0 to 6.0. Any increase in pH above that range can induce methanogenic bacteria to grow, consuming H₂ to produce CH₄ [161]. On the other hand, lower pH values cause microbes to switch their metabolism towards other biochemical processes. This leads to a change in the composition of the gaseous products and impaired H₂ production. Additionally, a pH below 4.0 can inhibit microbial growth [162].

Heat treatment at 80–104 °C is a common processing step used to eliminate methanogenic bacteria from the anaerobic sludge communities [163]. This conditioning of anaerobic microflora ensures the survival of hydrogenous spore-forming microbes, including Bacillus sp. and Clostridium sp. [164]. Short hydraulic retention times (HRT of 12 h or less) can be used to limit the growth of microbes that compete with H₂-producing bacteria [165].

The efficiency of H₂ production can be hampered by an excess of undissociated volatile fatty acids (VFAs) generated in the reactor [166]. Excessive molecular pressure in the gas phase of the reactor is another inhibitor of fermentative hydrogen production. Too much H₂ in the system causes the accumulation of propionic acid and butyric acid, reducing H₂ yields. Reducing the pressure—and thus the H₂ concentration—can significantly improve performance [167]. Another crucial parameter for fermentative hydrogen production is the concentration of iron (involved in hydrogenase activity), which should be in the range of 10 to 100 mg/L. Nitrogen levels are also important since N is an essential nutrient for microbes [168].

4.3. Mixed Fermentation

A combination of dark and photofermentation has been used to enhance H₂ production [169]. Dark fermentation produces alcohols and organic acids as by-products, which serve as a carbon source for the bacteria involved in photofermentative H₂ production [170]. Integrated biological processes significantly improve the ratio of energy stored in H₂ to the energy needed to maintain the culture. The literature data show that ratios of up to 3.0 are achievable [171,172]. Chookaew et al. [158,173] studied H₂ production from crude glycerol and experimented with a two-step process involving dark fermentation followed by photofermentation [174]. The strains used for the two-steps were Klebsiella sp. TR17 and Rhodopseudomonas palustris TN1, respectively. The optimal results of photofermentation were obtained with diluted wastewater, added glutamate, and no added yeast. Under these conditions, total H₂ production across the two-step process was 6.42 mmol/g COD—10.4% of the theoretical yield [174].

Rodrigues et al. [175] evaluated H₂ production in an integrated biosystem through dark fermentation followed by photofermentation, using crude glycerol from biodiesel production from waste cooking oils (WCO). A mixed Clostridiales culture was able to take up crude glycerol during dark fermentation at 37 °C and pH 5.5, generating a high H₂ output of 22.38 mmol H₂/L with yields of 1.75 mol H₂/mole glycerol (which corresponds to 24.06 mmol H₂/g COD). The dark fermentation effluent (DFE) in concentrations of 1.0, 2.0, and 3.0 g COD/L was used to grow a phototrophic bacteria culture of the order Rhizobiales and Clostridiales at 37 °C, pH 7.0, and 18.50 W/m². The optimum conditions were achieved with 1.0 g COD/L DFE, generating 3.94 mmol H₂/g COD _consumed, and removing 76.10% COD. The culture was able to absorb ethanol (76.86%), acetic acid (95.73%), butyric acid (94.76%), and particularly methanol (99.18%) [175].

Sarma et al. [176] investigated the technical and economic aspects of two-step H₂ production using dark fermentation followed by photofermentation. The authors concluded that 1 kg of crude glycerol can be converted into the equivalent of 2.56 L, but at a cost of USD 330. The media components accounted for 82% of production costs, meaning that the reduction in these costs is crucial to making the process viable. The process was also found to be very environmentally beneficial. It was demonstrated that the conversion of 1 kg of crude glycerol reduces greenhouse gases by 7.66 kg [176]. Examples of glycerol use in fermentative hydrogen production, key process parameters, and performance metrics are given in

Table 2. Use of crude glycerol for fermentative hydrogen production.

Method	Base Substrate	Experimental Parameters	Glycerol Volume	Production/Yield	Ref.
Photofermentation	R. palustris	125 mL serum bottles, 30 °C, 200 W/m2	10 mM	6.1 moles H2/mole crude glycerol (87%)	[177]
	R. palustris	125 mL serum bottles, 30 °C, 50 W halogen light	10 mM	6 moles H2/mole glycerol	[128]
	R. palustris	125 mL serum bottles, 30 °C, 200 W/m2	30 mM	6.69 moles H2/mole crude glycerol	[129]
	Arundo donax	200 mL reactor, 30 ± 1 °C, pH 7.0, incubator with 2000 Lux illumination for 72 h	10 g/L	32.14 mL/g substrate	[131]
			15 g/L	79.15 mL/g substrate
			20 g/L	51.06 mL/g substrate
			30 g/L	12.43 mL/g substrate
	R. palustris	Fluidized bed PBR (FB), immobilized, 28 ± 1 °C, pH 7, 100 W/m2	50 mM	14.7 mL/g/h	[138]
		Packed bed PBR (PB), immobilized, 28 ± 1 °C, pH 7, 100 W/m2		4.53 mL/g/h
		Column PBR, planktonic, 28 ± 1 °C, pH 7, 100 W/m2		12.68 mL/g/h
	R. palustris	500 mL bioreactors, planktonic, 35 ± 0.2 °C, pH 7, 250 ± 20 W/m2	50 mM	5.1 mL/g/h	[139]
		500 mL bioreactors, immobilized, 35 ± 0.2 °C, pH 7, 250 ± 20 W/m2		8 mL/g/h
Dark fermentation	Enterobacter aerogenes and C. butyricum	125 mL serum bottles, 36 °C, pH 6.5, 150 rpm	17.5 g/L	1.8 mmol H2/g glycerol	[155]
	Industrial and municipal solid waste (Eubacteria and Archaea)	2 L reactor with a useful volume of 1.7 L, pH 6.61 ± 0.12, 23 rpm	1% v/v	51 ± 4 LH2/kg VS	[144]
	Enterobacter aerogenes HU-101	Cylindrical glass column reactor with a working volume of 60 mL, 37 °C, pH 6.8	1.7 g/L	1.12 moles H2/mole glycerol	[152]
			3.3 g/L	0.9 moles H2/mole glycerol
			10 g/L	0.71 moles H2/mole glycerol
			25 g/L	0.71 moles H2/mole glycerol
	Bacillus thuringiensis EGU45	Batch, 37 °C, pH 7, 1% w/v NH4Cl	1% v/v	0.261 moles H2/mole glycerol	[151]
			2% v/v	0.452 moles H2/mole glycerol
			3% v/v	0.252 moles H2/mole glycerol
		Batch, 37 °C, pH 7, 1% w/v NaNO3	1% v/v	0.748 moles H2/mole glycerol
			2% v/v	0.570 moles H2/mole glycerol
			3% v/v	0.314 moles H2/mole glycerol
		Batch, 37 °C, pH 7, 1% w/v NH4NO3	1% v/v	0.547 moles H2/mole glycerol
			2% v/v	0.646 moles H2/mole glycerol
			3% v/v	0.299 moles H2/mole glycerol
		Aspirator bottles, 1.2 L capacity with working a volume of 1.0 L, 37 °C, pH 7, 1% (w/v) NH4Cl, support material: free-floating (FF), HRT 2 days	2% v/v	0.188 moles H2/mole glycerol
			5% v/v	0.046 moles H2/mole glycerol
		Aspirator bottles, 1.2 L capacity with working a volume of 1.0 L, 37 °C, pH 7, 1% (w/v) NH4Cl, support material: banana leaves (BL), HRT 2 days	2% v/v	0.273 moles H2/mole glycerol
			5% v/v	0.113 moles H2/mole glycerol
		Aspirator bottles, 1.2 L capacity with working a volume of 1.0 L, 37 °C, pH 7, 1% (w/v) NH4Cl, support material: coconut coir (CC), HRT 2 days	2% v/v	0.237 moles H2/mole glycerol
			5% v/v	0.116 moles H2/mole glycerol
		Aspirator bottles, 1.2 L capacity with working a volume of 1.0 L, 37 °C, pH 7, 1% (w/v) NH4Cl, support material: free-floating (FF), HRT 4 days	2% v/v	0.283 moles H2/mole glycerol
			5% v/v	0.146 moles H2/mole glycerol
		Aspirator bottles, 1.2 L capacity with working a volume of 1.0 L, 37 °C, pH 7, 1% (w/v) NH4Cl, support material: banana leaves (BL), HRT 4 days	2% v/v	0.410 moles H2/mole glycerol
			5% v/v	0.366 moles H2/mole glycerol
		Aspirator bottles, 1.2 L capacity with working a volume of 1.0 L, 37 °C, pH 7, 1% (w/v) NH4Cl, support material: coconut coir (CC), HRT 4 days	2% v/v	0.288 moles H2/mole glycerol
			5% v/v	0.286 moles H2/mole glycerol
	C. pasteurianum CH4	Batch, 35 °C, pH 7, 200 rpm	10 g/L of pure glycerol	1.11 ± 0.16 moles H2/mole glycerol, 256.8 ± 8.1 mL/h/L	[150]
		CSTR, HRT 12 h, 35 °C	10 g/L of pure glycerol	0.50 ± 0.02 moles H2/mole glycerol, 103.1 ± 8.1 mL/h/L
			10 g/L of crude glycerol	0.77 ± 0.05 moles H2/mole glycerol, 166.0 ± 8.7 mL/h/L
	Klebsiella sp. TR17	Immobilized, 40 °C, up-flow anaerobic sludge blanket reactor, 4 h	50% glycerol	29.00–44.27 mmol H2/g glycerol	[173]
	T. neapolitana	75 °C, pH 7.0–7.5, itaconic acid	-	-	[178]
	bd	125 mL serum bottles, 37 °C, pH 7.9	15 g/L	0.96 moles H2/mole glycerol (2.2 L/L/d) 7.92 g ethanol/L	[179]
	Enterobacteriaceae (90%), with the genera Klebsiella and Escherichia/Shigella, followed by Burkholderiaceae (10%) with the genus Cupriavidus	3 L bioreactor, 37 °C, pH 6.8, 120 rpm	15 g/L	0.9 moles H2/mole glycerol (2960 mL H2/L/d)	[180]
	bd	120 mL serum bottles, 40 °C, 150 rpm	1 g/L	1.41 moles H2/mole glycerol consumed	[181]
	bd	Up-flow column reactor, 35 °C, HRT 24 h, pH 7, OLR 29.7 ± 0.5 g COD/d/L	4.2 ± 1.2 g/L	107.3 L H2/kg waste glycerol	[182]
	bd	160 mL serum bottles, 30 °C, pH 5.5, 350 rpm, GCI/IGCT 2.5 h/20 h	7 g/L	0.75 moles H2/mole glycerol	[183]
	bd	120 mL serum bottles, 35 °C, 48 h	10 g/L	0.55 moles H2/mole glycerol (332.04 mL H2/L)	[158]
	C. pasteurianum	Batch, 37 °C, pH 7	60 g/L	0.93 moles H2/mole glycerol	[184]
	Granular sludge from a mesophilic UASB reactor used for the treatment of poultry slaughterhouse wastewater	Anaerobic fluidized bed reactor, 30 °C, pH 4.5	18 g/L	0.17 moles H2/mole glycerol	[185]
	Granular sludge from a UASB reactor from an effluent treatment plant of a soybean processing company	Semi-continuous, 35 °C, pH 5.5	50 g/L	0.01 moles H2/mole glycerol	[186]
Dark fermentation and photofermentation	Dark fermentation: Klebsiella sp., Photofermentation: Rhodopseudomonas palustris TN1	Dark fermentation: 60 mL serum bottle with a 36 mL working volume, 40 °C, pH 8; Photofermentation: 60 mL of serum bottle with a 36 mL working volume, 30 °C, pH 7, anaerobic-light (3000 lux) condition	11.14 g/L glycerol (50% purity)	6.42 mmol H2/g COD consumed (80.21% glycerol conversion rate)	[174]
	R. palustris	30 °C, 175 W/m2	85% glycerol, <0.5% methanol	6.69 moles H2/mole glycerol	[176]
	Dark fermentation: Clostridiales, Photofermentation: Rhizobiales and Clostridiales	Dark fermentation feed: peptone 5 g/L, meat extract 5 g/L, and yeast extract 5 g/L, pH 5.5, 37 °C. Photofermentation: 37 °C, pH 7.0, and 18.50 W/m2	Dark fermentation: 20 g/L COD from crude glycerol; photofermentation: 1 g/L COD from DFE	28 mmol H2/g COD consumed	[175]
			Dark fermentation: 20 g/L COD from crude glycerol; Photofermentation: 2 g/L COD from DFE	25.51 mmol H2/g COD consumed
			Dark fermentation: 20 g/L COD from crude glycerol; photofermentation: 3 g/L COD from DFE	24.91 mmol H2/g COD consumed

.

5. Biohythane Production

Coupling H₂ production with CH₄ production—a solution known as “biohythane”—can optimize the reuse of leftover organic matter from fermentative hydrogen production and thus help recover gas energy [187]. The leftover spent media from dark fermentative H₂ production contains high levels of short-chain fatty acids such as acetate, butyrate, propionate, etc., which could be a suitable feedstock for methanogens [188]. Before the spent media are subjected to biomethanation, their pH should be adjusted to a range of 7 to 7.8. Additionally, the H₂ dissolved in the media also promotes the growth of hydrogenotrophic methanogens. The recovery of gaseous energy in the form of H₂ alone might not be enough to make the process commercially viable, since only 20–30% of total energy can be recovered through H₂ production. In this light, integrated biohythane production seems to be a good choice since it provides an attractive method of converting organic residues rich in carbohydrates, lipids, and proteins for clean energy generation. On the other hand, these systems present challenges of their own, including the shading effect of pigments produced by photofermentative organisms and problems with scale-up [188].

Two-step AD has been acknowledged to be a promising method of biohythane production, as it enables reduced organic loading and increased total energy conversion efficiency by generating two gases with high fuel-burning capacity [189]. By separating the hydrolysis/acidogenesis phase from the methanogenesis phase, the stability of the whole process can be improved—controlling the acidification phase at the H₂ generation stage (first stage) can prevent the inhibition of the methanogen population during the methane generation phase (second stage) [190]. Two-step biohythane production has been carried out using feedstocks such as maize silage [191], algal biomass [192], food waste [193], dairy waste [194], organic fraction of municipal sewage wastewater [195], cassava stillage [196], tequila vinasses [197], and skim latex serum [198]. There are few reports on the use of glycerol for biohythane production, even though glycerol is a very promising substrate due to its high content of carbon and macronutrients. Jiang et al. [199] conducted a two-step AD process to degrade glycerol. The system produced H₂ and CH₄ yields of 0.026 L H₂/g COD and 0.29 L CH₄/g COD _removed, respectively. In addition, 93% COD removal was noted [199].

Co-digestion with glycerol can boost biohythane production at the optimal C/N range of 19 to 41 and the right H₂/CH₄ ratio of 0.06 to 0.3 [200]. Jehlee et al. [200] used glycerol as a co-substrate with Chlorella sp. TISTR 8411 biomass for biohythane production. Mono-digestion of Chlorella sp. TISTR 8411 biomass produced H₂ and CH₄ yields of 36.4 mL H₂/g VS and 166.18 mL CH₄/g VS, respectively. Co-digestion of Chlorella biomass brought a marked improvement in biohythane production. The optimal dose of 2% TS glycerol waste resulted in 39.8 mL H₂/g VS and 577.33 mL CH₄/g VS being produced [200]. Dounavis et al. [201] developed a continuous process for the production of biohythane from crude glycerol in a two-step reactor system. In the first step, H₂ production was tested using mixed acidogenic communities in an up-flow column bioreactor. Cylindrical ceramic beads with a porosity of 600 m²/L were used as an attachment matrix for bacterial cells. The experimental parameters were: HRT 24 h; pH 6, 6.5, and 7; and a glycerol concentration of 20 g/L. The effluent from the hydrogenic reactor was fed to a methanogenic continuous stirred reactor (CSTR), which was used to examine the effect of organic loading on CH₄ yield. The gaseous phase of the reactors was mixed to produce biohythane. At 20 g/L glycerol and 6, 6.5, and 7 feedstock pH, the H₂ output was 0.051, 0.070, and 0.094 L/g COD, respectively. CH₄ was produced at yields of 0.257 L/g COD (commercial glycerol), 0.283 L/g COD (crude glycerol), 0.198, 0.242, and 0.273 L/g COD (effluents from the hydrogenogenic stage (1st stage), diluted with water to 5, 7.5, and 10 g COD/L, respectively) [201].

Sittijunda et al. [202] produced biohythane by co-fermentation of crude glycerol and microalgae biomass. The focus was on optimizing the technological parameters that determine the efficiency of H₂ production, including the initial glycerol concentration, the amount of anaerobic bacterial inoculum used, and the microalgae biomass. The highest fermentation efficiency of 655.1 mL H₂/L was achieved with 13.83 g/L glycerol, 23.1 g VS/L algal biomass, and 10.3% (v/v) anaerobic sludge. The substrate for CH₄ production was the effluent from the bioreactor for hydrogen fermentation. The highest yield and kinetics of CH₄ production were 868.7 mL CH₄/L and 2.95 mL CH₄/L^−h, respectively. The use of a sequence of hydrogen and methane reactors proved to be a more effective solution than the single-stage process and provided a total additional energy of 1.27 kJ/g VS [202].

Silva et al. [94] studied H₂ and CH₄ production in a two-step AD process with co-digestion of food waste, sewage sludge, and glycerol under mesophilic conditions (35 °C). The study evaluated the effect of adding 1 and 3% v/v glycerol as a co-substrate in three-component mixtures, keeping the concentration of all substrates at 10 g VS/L. Glycerol was found to increase H₂ production under all tested conditions. The highest H₂ yield (179.3 mL H₂/g VS) was obtained for the 3% glycerol mixture. In terms of CH₄ output, however, the 1% glycerol batch performed the best at 342 mL CH₄/g VS generated. At 3% glycerol, abrupt reductions in biogas pH and CH₄ content were observed, stemming from unstable methanogenesis. When taking into account both the H₂ and CH₄ production stages, the three-component mixture with 1% glycerol showed the highest energy production, yielding 15.5 kJ/g VS [94].

Microbes capable of producing H₂ and CH₄ have been identified across a wide temperature range. The majority of the literature reports on H₂ and CH₄ production focus on mesophilic dark fermentation [203]. The digester operating temperature should be adapted to the microbes used. Methanogens are the most sensitive to temperature changes. Even minute temperature shifts by 2–3 °C can lead to methanogen suppression and promote acidogen formation, causing VFA accumulation in the system. This, in turn, can cause pH to drop below 7 and inhibit methanogenesis [204]. Kanchanasuta and Sillaparassamee [205] have posited that crude glycerol could be used not only for co-digestion but also as a pH-adjuster for fermentative hydrogen production (to maintain pH > 5 throughout the fermentation process). Examples of glycerol use in biohythane production, key process parameters, and performance metrics are given in

Table 3. Use of glycerol in biohythane production.

Base Substrate	Experimental Parameters (Step 1)		Glycerol Volume	H2 Production/Yield	Experimental Parameters (Step 2)		CH4 Production/Yield	Ref.
Chlorella sp. TISTR 8411	250 mL serum bottles with a working volume of 200 mL, 55 °C, pH 5.5		2%	39.8 mL/g VS	250 mL serum bottles with a working volume of 200 mL, 55 °C		577.33 mL/g VS	[200]
			4%	29.44 mL/g VS			214.98 mL/g VS
			6%	20.93 mL/g VS			130.67 mL/g VS
			8%	8.86 mL/g VS			78.11 mL/g VS
			11%	4.28 mL/g VS			54.32 mL/g VS
			15%	2.4 mL/g VS			41.02 mL/g VS
Glycerol	PVC up-flow, packed bed column bioreactor, 35 ± 0.5 °C, 600 m2/L;	pH 6	20 g/L, purity 92.2 ± 0.3%	0.051 L/g COD	Continuous Stirred Tank Reactor (CSTR), 35 °C, HRT 20 day	Effluents from the hydrogenogenic stage (1st stage) diluted with water to 5 g COD/L	0.198 L/g COD	[201]
		pH 6.5		0.070 L/g COD		Effluents from the hydrogenogenic stage (1st stage) diluted with water to 7.5 g COD/L	0.242 L/g COD
		pH 7		0.094 L/g COD		Effluents from the hydrogenogenic stage (1st stage) diluted with water to 10 g COD/L	0.273 L/g COD
Chlorella sp.	Serum bottles of 120 mL with 70 mL working volumes, 35 ± 4 °C, pH 6, 150 rpm	Inoculum concentration of 6.17% v/v, algal biomass concentration of 4.62 g VS/L	10 g/L	252.56 ± 2.27 mL H2/L	Serum bottles of 120 mL with 70 mL working volumes, pH 7.5, inoculum concentration 25% v/v	Effluent from fermentative hydrogen production at 10 g/L of glycerol	387.4 ± 20.04 mL CH4/L	[202]
		Inoculum concentration of 25% v/v, algal biomass concentration of 23.1 g VS/L	40 g/L	140.67 ± 3.41 mL H2/L		Effluent from fermentative hydrogen production at 40 g/L glycerol	428.0 ± 13.12 mL CH4/L
		Inoculum concentration of 10.31% v/v, algal biomass concentration of 23.1 g VS/L	13.83 g/L	655.12 ± 1.64 m LH2/L		Effluent from fermentative hydrogen production at 13.83 g/L glycerol	868.7 ± 19.98 mL CH4/L
Food waste, sewage sludge	250 mL glass bottles, 35 ± 1 °C, pH 5.5, 150 rpm		1% v/v	140.2 mL H2/g VS	250 mL glass bottles, 35 °C, pH 7, 150 rpm		342 mL CH4/g VS	[94]
			3% v/v	177 mL H2/g VS			224.4 mL CH4/g VS

.

6. Summary and Conclusions

Applying waste organic substrates, such as glycerol, for industrial purposes can help reduce industrial greenhouse gas emissions and significantly decrease the cost of commercial production of alternative fuels. It is also fully in line with the principles of the circular economy and the ongoing trend of limiting and fixing CO₂ emissions. Biodiesel production is one of the major sources of waste glycerol. Given that the glycerol generated in the process is very costly to purify, alternative solutions are being sought. Waste glycerol can be harnessed for bioenergy production by using it as a feedstock for the fermentative production of gas biofuels, including CH₄, H_2, and biohythane.

Though our understanding of glycerol fermentation is quite extensive and well systematized, there is still a need for further research with a view to optimizing processes, upgrading existing technologies, and improving final performance. With regard to the microbial fermentation of crude glycerol into energy products, it is crucial to develop more efficient technologies by utilizing new microbial strains and adapting existing ones, improving their selectivity and durability, and, in particular, bolstering their tolerance to high levels of volatile fatty acids (VFAs) in the medium and to impurities in the glycerol. Important considerations include selecting the best ratio of crude glycerol to other organic waste in anaerobic processing and developing innovative projects, as well as modifying operational guidelines for glycerol-to-biogas bioconversion systems. It is important to optimize operation conditions and running parameters, such as pH, temperature, substrate concentration, type of dosing/stirring, hydraulic retention time, and the organic load rate.

Another prerequisite to ensuring stability and good performance of digestion processes lies in minimizing the adverse effect of by-products/intermediates accumulating due to imbalances in anaerobic biochemical processes. Currently, the most popular solution is to adopt two-step digestion processes, hybrid systems, systems that integrate dark fermentation with photofermentation processes, and mixtures of multiple bacterial strains. Another important avenue of research is to explore the use of pre-treatment techniques to boost the performance of biological anaerobic processes, both in terms of biogas yield/quality and the degree of digestate mineralization/stabilization (an important factor for subsequent digestate reuse). An important aspect in this regard is to improve the overall cost-effectiveness of the process.

It should be emphasized that the use of waste glycerol as a substrate in methane fermentation processes is well recognized and is characterized by a high technological readiness level. This is proven by plants operating efficiently on a technical scale. In this case, the basic technological parameters of the process have already been determined, the final results obtained have already been estimated fairly accurately, and the functional features of the plant have been verified. The use of this organic substrate for the production of biohydrogen and biohythane is still at the laboratory-scale research level. Due to the environmentally friendly properties of these gaseous biofuels and, thus, their high competitiveness compared to methane, this is a promising research direction. The next step towards their realization should be to increase their level of technological maturity by starting with pilot-scale research and then developing technical and technological guidelines for large-scale plants. Raising the technological readiness level will allow an objective and reliable assessment of the technical and environmental effectiveness as well as the economic viability of these solutions.