Harnessing Biogas into High-Value Chemicals: The Role of Algal–Methanotrophic Co-Cultures

Abstract

The conversion of biogas into high-value chemicals for pharmaceutical, cosmetic, and nutraceutical markets offers an attractive alternative to conventional fossil-based production routes, enabling circular value chains with significant socio-economic impact. This study evaluated the valorization of biogas into osmolyte and carotenoid compounds with market prices ranging from 1000 to 7000 $·kg⁻¹. Specifically, an algal–methanotrophic co-culture operated under saline conditions, preventing external microbial contamination and stimulating osmolytes and carotenoids, was assessed for its capacity to simultaneously remove methane (CH₄) and carbon dioxide (CO₂), with efficiencies of 92 and 89%, respectively. while producing ectoine, hydroxyectoine, lutein, β-carotene, and astaxanthin. Shotgun metagenomic analyses identified the key microorganisms driving the process, predominantly alkaliphilic and halophilic green algae (Chlorella, Dunaliella) and cyanobacteria (Leptolyngbya), and halotolerant methanotrophs (Methylotuvimicrobium) and methylotrophs (Methylophaga). Metagenomics further revealed the presence of key metabolisms related to C1 utilization and biosynthetic genes associated with carotenoid and osmolyte production, confirming the metabolic potential of the consortium to convert biogas-derived carbon directly into high-value compounds. Overall, these results demonstrate the feasibility of an efficient, biologically driven bio-platform capable of transforming greenhouse gas-rich waste streams into economically relevant bioactive molecules, contributing to global priorities in sustainable biomass-to-biochemical innovation.

1. Introduction

Biogas, primarily composed of carbon dioxide (CO₂) and methane (CH₄), is a by-product of anaerobic digestion in wastewater treatment plants and organic waste treatment facilities. Despite its potential as a renewable energy source, the biogas-to-electricity market remains largely untapped due to several limiting factors, including the presence of trace contaminants (such as hydrogen sulfide and siloxanes) and the high associated purification costs, and competition from renewable energy sources like solar and wind power. A paradigm shift is therefore underway: instead of valorizing biogas merely as an energy carrier, it can act as a carbon and energy-rich feedstock for the sustainable production of high-value biochemicals. The biological valorization of its two main components (CH₄ and CO₂) not only creates new economic opportunities but also contributes to climate mitigation by converting two of the most potent greenhouse gases (GHGs) into valuable molecules. These dual benefits position biogas upgrading and bioconversion as an attractive strategy to unlock the environmental and economic potential of this overlooked resource.

Among biological routes, microalgae and methanotrophic bacteria represent complementary platforms capable of turning biogas into valuable biomolecules. Microalgae are particularly attractive for biorefinery applications due to their rich metabolic diversity and their ability to synthesize photoprotective and light-harvesting pigments, at concentrations often exceeding those of terrestrial plants [1]. These pigments, including carotenes, xanthophylls, chlorophylls, and phycobiliproteins [2], not only play an essential role in photosynthesis and photoprotection, but also hold significant commercial value. Driven by growing concerns regarding the environmental and health impacts of synthetic dyes, demand for microalgal natural pigments is rapidly expanding across food, cosmetic, nutraceutical, and feed sectors. Carotenoids in particular stand out for their antioxidant, nutritional, and therapeutic properties, with the global carotenoid market projected to reach nearly 3 billion dollars by 2029 [3]. More than 600 carotenoids have been identified to date, though microalgae mainly produce lycopene, β-carotene, zeaxanthin, astaxanthin, lutein, and violaxanthin, which nearly compose 90% of carotenoids within the human body and diet [4]. Within this group, astaxanthin (3,3′-dihydroxy-β,β-carotene-4,4′-dione) is one of the most valuable carotenoids, reaching market prices of 2000–7000 $·kg⁻¹ [5]. Its red-orange color, combined with exceptional antioxidant activity (up to 10 times stronger than that of lutein, zeaxanthin, β-carotene, and canthaxanthin [6]), has propelled its use in aquaculture, functional foods, nutraceuticals, cosmetics, and pharmaceuticals. Likewise, β-carotene, a vitamin A precursor, is widely utilized as a natural food colorant (e.g., soft drinks, baked goods, and margarine) and as an active ingredient in antioxidant supplements [4]. Lutein, a lipophilic tetraterpene ranging in color from red-orange to yellow, is recognized not only for its strong antioxidant activity and anticancer effects but also for its essential role in infant neural development and ocular protection [7]. Collectively, these carotenoids exemplify the biotechnological potential of microalgae as sustainable sources of natural bioactive compounds.

In parallel, methanotrophic bacteria have gained significant attention as efficient biocatalysts for the conversion of biogas into high-value products. Among them, halotolerant methanotrophs can produce osmolytes, such as ectoine and its hydroxylated derivative, hydroxyectoine, which are particularly attractive for the pharmaceutical and cosmetic industries due to their exceptional osmoprotective and stabilizing properties. With a market price of approximately 1000 $·kg⁻¹ [8], ectoine production from waste-derived CH₄ represents an economically compelling and environmentally sustainable approach.

Co-cultures of microalgae and methanotrophs have been increasingly explored as biotechnological platforms to produce valuable compounds and renewable energy from biogas (in the form of biomethane) [9]. These synergistic systems offer several advantages: dissolved oxygen (O₂) produced by photosynthetic algae can sustain methanotrophic growth, while the dissolved CO₂ generated by methanotrophs can be assimilated by algae, thus enabling in situ gas exchange and the simultaneous sequestration of both CH₄ and CO₂ [10]. Moreover, co-cultivation has been shown to increase total biomass yield and nutrient removal efficiency compared to monocultures, as the mutual exchange of gases and metabolites creates a more balanced and synergistically productive microenvironments [11]. To date, these co-cultures have been investigated as a platform for nutrient recovery from wastewater [11], biodiesel [12], biohydrogen generation [13], or ectoine production [14]. However, the functional understanding of the metabolic interactions, pathways involved and the coordinated production of high-value biomolecules, particularly carotenoids and osmolytes, remains still poorly understood.

This study aims to demonstrate the potential of algal–methanotrophic co-cultures composed of haloalkaliphilic extremophiles as an integrated platform for the conversion of biogas into osmolytes and carotenoids. By integrating physiological assays with metagenomic analyses, this study unravels the metabolic interactions and functional complementarities between the algal and methanotrophic partners, identifies the key metabolic pathways involved in dual-product formation and assesses the impact of co-cultivation on biomass productivity and bioproduct generation.

2. Results and Discussion

2.1. Gas Consumption and Growth Dynamics

The dynamics of biomass growth and gas concentrations over time across all cultures are shown in Figure 1. The algal and algal–methanotrophic cultures exhibited active growth when supplied with CH₄ and CO₂ as the sole carbon sources and light as the energy source. Likewise, the methanotrophic culture grew supplemented with CH₄ and CO₂ as the sole carbon and energy sources. Distinct growth phases and gas consumption patterns were observed across cultures.

In the algal culture, with an initial TSS of 0.08 ± 0.001 g·L⁻¹, a lag phase of 4 days was followed by exponential growth, reaching biomass concentrations of 1.21 ± 0.02 gTSS·L⁻¹ by day 28. A temporary decrease in biomass was recorded between days 11 and 16, which resulted from the collapse of one of the three biological replicates and the subsequent reinoculation, leading to a transient reduction in the mean biomass concentration before growth resumed. During the cultivation period, CO₂ concentrations in the gas phase decreased from 26.61% to 2.61%, corresponding to a CO₂ removal efficiency of 90%. Despite the high CO₂ partial pressure in the gas phase, pH values remained within a range suitable for the alkaliphilic culture due to the presence of a high-capacity buffer: the pH was initially set at 9.0, decreased moderately during the first 10 days as CO₂ dissolved into the liquid phase, and subsequently increased again as photosynthetic CO₂ assimilation intensified, stabilizing close to the initial value by the end of the experiment. Concurrently, O₂ concentrations increased from 2.65% to 38.98% as a result of photosynthetic activity. When expressed relative to the maximum theoretically possible from CO₂ consumption (i.e., 100%), this rise represents a 93% net O₂ production, reflecting an active photosynthetic metabolism under illuminated conditions (Figure 1a). These results demonstrate that the algal culture efficiently assimilated inorganic carbon even under CO₂ levels typical of raw biogas streams. The presence of CH₄ did not adversely impact photosynthetic activity, confirming their suitability for biogas upgrading applications, in agreement with previous reports [15,16]. Although a slight decrease in CH₄ concentration was observed, this reduction is mainly attributable to gas sampling losses and methodological constraints rather than biological CH₄ uptake by the algal culture.

On the other hand, the methanotrophic culture exhibited a longer adaptation period, with a 15-day lag phase likely due to the metabolic adaptation necessary for efficient CH₄ oxidation. After this phase, biomass increased steadily over a 43-day cultivation period, reaching average biomass concentrations of 0.83 ± 0.14 gTSS·L⁻¹. CH₄ removal efficiency averaged at 69%, with CO₂ accumulated as a metabolic by-product, reaching a production of 42% (Figure 1b). O₂ was supplied periodically to maintain concentrations above 15%, ensuring sustained methanotrophic activity without O₂ limitation.

Finally, in the algal–methanotrophic co-culture, methanotrophs were inoculated when O₂ concentrations reached 6.7 ± 1.2%. However, during the initial cultivation phase (first 12 days), despite clear biomass accumulation (2.41 ± 0.09 gTSS·L⁻¹), net CH₄ consumption was not observed in the co-culture (Figure 1c). This apparent lag phase can be explained by the asynchronous establishment of the mutualistic interaction between the two partners: methanotrophs require O₂ to oxidize CH₄, but at the beginning of the experiment, algal photosynthetic activity was not yet sufficient to generate the necessary O₂ levels. As a result, methanotrophs remained metabolically active at a basal level supported by transient O₂ availability, internal storage compounds, or residual organic carbon carried over with the inoculum. This would explain the observed biomass accumulation in the absence of measurable CH₄ consumption. To overcome this bottleneck, an external supply of O₂ was added on days 13 and 15, which triggered CH₄ oxidation and CO₂ fixation. From that point onward, the culture evolved towards a functionally self-sustaining state, in which algal O₂ production supported methanotrophic growth without the need for further O₂ addition, while methanotrophic-derived CO₂ met the carbon demands of photosynthetic microorganisms. The maximum biomass concentration of 5.18 ± 0.74 gTSS·L⁻¹ was achieved by day 28, after which a stationary phase was maintained (Figure 1c). By the end of the experiment, both CH₄ and CO₂ were efficiently removed from the gas phase, showing removal efficiencies of 92 and 89%, respectively. These gases would be available for microbial bioconversion into valuable bioproducts as will be discussed in Section 2.2.

Overall, the three cultures exhibited distinct growth dynamics and gas utilization patterns that reflect their metabolic characteristics and their potential for biogas valorization. The algal culture showed the fastest adaptation and the highest efficiency in CO₂ fixation, reaching substantial O₂ production rates that supported robust photosynthetic metabolism, even in the presence of CH₄. In contrast, the methanotrophic culture displayed a longer adaptation period and more moderate biomass accumulation, coupled with effective CH₄ oxidation. The algal–methanotrophic co-culture integrated the strengths of both partners, achieving the highest biomass concentrations and efficient simultaneous removal of CH₄ and CO₂ once the initial delay in establishing the mutualistic interaction was overcome. To prevent this early-stage limitation, the optimal initial proportion of both microorganisms should be optimized, as this parameter is critical for achieving efficient simultaneous sequestration of CH₄ and CO₂ [10]. Ultimately, the co-culture demonstrated the most balanced and efficient performance for GHG abatement and biocarbon valorization processes.

2.2. Ectoine and Carotenoid Production

Ectoine (ectoine and hydroxyectoine) production was observed in both algal and methanotrophic cultures, as well as in the algal–methanotrophic co-culture (Figure 2), demonstrating the potential of these microbial consortia for osmolyte synthesis. The highest concentrations of ectoine (13.2 ± 5.3 mg_ectoine·L⁻¹, corresponding to 36.1 ± 10.9 mg_ectoine·g_biomass⁻¹) and hydroxyectoine (15.8 ± 4.4 mg_{hydroxyectoine}·L⁻¹ corresponding to 70.6 ± 5.1 mg_{hydroxyectoine}·g_biomass⁻¹) were achieved in the methanotrophic culture during the exponential and stationary phase, respectively. The co-culture yielded a maximum concentration of 19.77 ± 1.37 mg_ectoine·L⁻¹ (34.7 ± 4.9 mg_ectoine·g_biomass⁻¹) and 6.5 ± 1.2 mg_{hydroxyectoine}·L⁻¹ (10.1 ± 2.4 mg_{hydroxyectoine}·g_biomass⁻¹) during the exponential phase, demonstrating that the integration of microalgae with methanotrophs still supported the accumulation of valuable osmolytes. These results are consistent with the only previously reported study on ectoine production from algal–methanotrophic co-cultures. Ruiz-Ruiz et al. [14] observed a maximum ectoine content of 51.3 mg_ectoine·gVSS⁻¹ at 4.5% NaCl, but production in that case required the application of osmotic shock (increasing NaCl from 3 to 4.5%) to trigger higher accumulation.

Ectoines were also detected in the algal culture, with maximum concentrations of 3.3 ± 1.8 mg_ectoine·L⁻¹ (7.2 ± 2.9 mg_ectoine·g_biomass⁻¹) and 6.5 ± 1.2 mg_{hydroxyectoine}·L⁻¹ (26.7 ± 5.4 mg_{hydroxyectoine}·g_biomass⁻¹). Since algae are not known to synthesize ectoines, their presence is likely attributable to bacteria present within the algal culture (see Section 2.3).

No carotenoid production was detected in the methanotrophic cultures, even though certain methanotrophic genera, such as Methylomonas, are known to synthesize C30 and C40 carotenoids [17]. However, carotenoid concentrations in methanotrophic microbiomes are typically very low, which may explain the negligible levels observed in this study. This likely reflects both the dominance of non-pigmented bacterial taxa within the enriched community and the absence of environmental cues (light exposure and oxidative stress) that generally trigger carotenoid biosynthesis in these microorganisms [18].

In contrast, the production of carotenoids, including β-carotene, astaxanthin, and lutein, was observed in both the algal culture and the co-culture (Figure 3). Notably, the co-culture exhibited significantly higher contents of β-carotene and lutein compared to the algal culture, with maximum lutein concentrations of 14.8 ± 2.9 mg·L⁻¹ (28.9 ± 1.8 mg·g_biomass⁻¹), and β-carotene concentrations of 2.6 ± 1.0 mg·L⁻¹ (5.7 ± 0.6 mg·g_biomass⁻¹), underscoring its enhanced potential for high-value carotenoid production. This enhancement can be attributed to the metabolic coupling between algae and methanotrophs, whereby methanotrophic CO₂ production provided a continuous and localized source of inorganic carbon. This constant carbon availability likely mitigated carbon limitation stress and enabled the photosynthetic algae and bacteria to redirect metabolic fluxes toward the isoprenoid biosynthetic pathway via enhanced carbon fixation under dynamic CO₂/O₂ exchange conditions [19].

Overall, the use of an algal–methanotrophic co-culture fed on biogas promoted maximum lutein contents that exceeded those reported in other studies using algal monocultures fed with CO₂ as the sole carbon source [7]. The observed stationary phase values (6.0 ± 0.6 mg·g_biomass⁻¹) were consistent with those obtained for other microalgal species [20,21,22]. These findings demonstrate that algal–methanotrophic co-cultures can act as an integrated platform for biogas valorization, simultaneously enhancing carotenoid production.

2.3. Microbial Dynamics and Metabolic Networks Involved in Biogas Valorisation

Metagenomic analyses were performed to characterize the dominant microbial communities in each assay and to elucidate the genes and taxa involved in the bioconversion of CO₂ and CH₄ present in biogas into β-carotene, astaxanthin, lutein, ectoine, and hydroxyectoine.

In the methanotrophic culture, bacteria dominated the community, representing 99.7 ± 1.7% of the total reads, consistent with the selective enrichment conditions (biogas as the sole carbon and energy source and the absence of light). The relative abundance of bacterial representatives decreased in the algal–methanotrophic co-culture (75.7 ± 7.5%) and further declined in the algal culture (63.7 ± 12.8%). In contrast, the algal culture exhibited a substantial representation of Chlorophyta (22.2 ± 8.4% of the total community), with Chlorella vulgaris as the predominant genus (86.5 ± 1.3% of the Chlorophyta fraction). Chlorophyta was also the major phototrophic group in the algal–methanotrophic co-culture, representing 17.1 ± 2.5% of the total community. However, in this case, two different species co-dominated: C. vulgaris represented 42.0 ± 5.1% of the Chlorophyta members, while Dunaliella viridis represented 32.3 ± 7.8%. In addition to green algae, Cyanobacteria belonging to the genus Leptolyngbya were detected in both the algal and co-culture assays, representing 2.5 ± 0.8 and 0.3 ± 0.05% of the total bacterial community, respectively. Other cyanobacterial genera such as Pseudoanabaena were also present in both cultures, but represented less than 0.02% of the total community.

2.3.1. CO₂ and CH₄ Metabolism

As expected, the metagenomic analysis confirmed that phototrophs were the main microorganisms involved in CO₂ fixation via the Calvin–Benson cycle. For this pathway, genes encoding the ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) large subunit (rbcL/cbbL) and small subunit (rbcS/cbbS) were targeted. Their read abundances were high in the algal culture (4118 ± 334) and the algal–methanotrophic co-culture (3399 ± 569), while they remained low in the methanotrophic culture (20 ± 6) (Figure 4).

In the algal culture, the Calvin–Benson cycle was conducted by Chlorophyta members of the genera Chlorella and Auxenochlorella, although Cyanobacteria also contributed to this pathway (Leptolyngbya and Pseudoanabaena) (Figure 5). Although several non-phototrophic bacterial taxa were present and could potentially harbor CO₂ fixation genes, no Rubisco genes were detected outside the phototrophic community. In the algal–methanotrophic co-culture, metagenomic analysis also showed that CO₂ fixation was conducted mainly by the phototrophic community, primarily Chlorella, Dunaliella, Leptolyngbya, and Pseudoanabaena. However, other prokaryotic genera harbored Rubisco genes, including Marimonas, Thioalkalivibrio, and the methylotroph Methylophaga. All these genera are known to encode Rubisco. In the case of Methylophaga, the most abundant of these taxa, carbon assimilation proceeds via the 2-keto-3-deoxy-6-phosphogluconate aldolase variant of the ribulose monophosphate pathway, which is co-driven by methanol availability [23]; thus, any carbon fixation associated with this genus likely occurred through methylotrophic cross-feeding rather than through direct autotrophic CO₂ fixation. In the case of the methanotrophic culture, Rubisco abundance was overall low and, although it was detected in Methylophaga, Paracoccus, and Salipiger, CO₂ fixation via Rubisco likely represented a minor, auxiliary carbon assimilation route leading to zero net CO₂ reduction.

The other pathway investigated for CO₂ fixation was the reductive tricarboxylic acid (rTCA) cycle, for which the target enzyme was the 2-oxoglutarate-ferredoxin/oxidoreductase subunits alpha (korA, oorA, oforA) and beta (korB, oorB, oforB). These genes were identified in several taxa across the three enrichment cultures at the same relative abundances, reflecting a diverse metabolic network combining phototrophic, heterotrophic, and C1-utilizing microorganisms (Figure 4 and Figure 5).

CH₄ oxidation was observed in the methanotrophic and algal–methanotrophic cultures, while it was almost negligible in the algal culture. The identification of the CH₄ oxidation pathway was based on the detection of two key functional genes encoding the particulate CH₄ monooxygenase (pmoCAB) and the soluble CH₄ monooxygenase (mmoXYZC) (Figure 4). The total abundance of genes belonging to the pmoCAB cluster was substantially higher in the methanotrophic culture (2195 ± 230) than in the algal–methanotrophic co-culture (21 ± 1.7) and was almost negligible in the algal culture (2 ± 1.5). Genes encoding mmoXYZC were not detected in any of the samples, most likely due to the inherent difficulty of recovering such genes from metagenomic datasets of complex communities, where their low relative abundance may have precluded their assembly into MAGs [24]. Taxonomic analysis of the assembled genomes confirmed the presence of methanotrophic organisms, which represented 22.7 ± 1.7% of the total community in the methanotrophic assay, 9.1 ± 0.6% in the mixed culture, and only 0.6 ± 0.3% in the algal assay (Figure S1, Supplementary Materials). Although additional methanotrophs were detected based on the presence of pmoCAB genes (Figure 5), Methylotuvimicrobium clearly dominated CH₄ oxidation, representing 85.9 ± 2.5%, 62.7 ± 7.2%, and 22.9 ± 7.8% of the methanotrophic fraction in the respective assays. Within this genus, Methylotuvimicrobium alcaliphilum and Methylotuvimicrobium buryatense were the most abundant species, consistent with their well-documented ability to thrive under high-salinity and alkaline conditions [25].

Methylotrophs were also abundant when CH₄ was present, comprising 26.8 ± 7.3% of the total community in the methanotrophic culture and 10.0 ± 3.3% in the co-culture (Figure S2, Supplementary Materials). This group was dominated by the genus Methylophaga in the methanotrophic assay (95.9 ± 1.1% of the methylotrophic fraction), a well-known haloalkaliphilic bacterium commonly found in marine and hypersaline environments [26]. In contrast, in the algal–methanotrophic co-culture, there were two dominant genera, Methylophaga and Methylonatrum (50:40%). A similar pattern was observed in the algal culture; although methylotrophs exhibited a lower overall abundance (3.8 ± 1.3%), the dominant methylotrophic genera were again Methylonatrum (30.1 ± 5.8%) and Methylophaga (39.4 ± 7.3%). The dominance of Methylonatrum in the presence of a different microbiome could be related to its ability to utilize formate and ethanol as carbon and energy sources, which could have been more available in this system, whereas Methylophaga competes more efficiently when methanol or methylamine predominate. In all cases, methylotrophs occurred in proportions comparable to those of methanotrophs within the same culture, suggesting a tightly coupled metabolic interaction between CH₄ oxidizers and methanol-utilizing species. This cooperation establishes a syntrophic relationship in which methanotrophs oxidize CH₄ to intermediates such as methanol and formaldehyde, which are subsequently assimilated by methylotrophs. By scavenging these potentially toxic intermediates, methylotrophic species sustain methanotrophic activity and extend overall community growth [27].

2.3.2. Ectoine and Hydroxyectoine Production

Ectoine and hydroxyectoine synthesis was investigated by targeting the genes encoding L-ectoine synthase (ectC) and ectoine hydroxylase (ectD). Both genes showed the highest total abundance in the methanotrophic culture, with ectC (6317 ± 646) and ectD (3590 ± 119), followed by the algal methanotrophic co-culture, where ectC (3960 ± 380) and ectD (2115 ± 623) were also abundant. These trends are consistent with the ectoine concentrations obtained experimentally. In the algal culture, ectC (2761 ± 142) and ectD (872 ± 44) were relatively abundant; however, ectoine accumulation was approximately fivefold lower than in the co-culture. This discrepancy suggests that ectoine may have been utilized as an external carbon source by other members of the community [28]. Several known ectoine-producing genera, including Halomonas, Rhodococcus/Pararhodococcus, and Marinomonas, were detected in all assays (Figure 5). However, based on their relative abundances, Methylophaga and Methylotuvimicrobium were the predominant putative ectoine producers in the methanotrophic culture and the algal–methanotrophic co-culture, whereas Roseinatronobacter (12.3 ± 2.5%) was the main potential producer in the algal culture. Hydroxyectoine synthesis was most likely associated with Nitratireductor, given its relative abundance and the detection of ectD genes in its genome [29], possibly supported by secondary ectoine-hydroxylating taxa (Figure 5). Although M. alcaliphilum harbors a gene annotated as ectD, previous studies have demonstrated that its native enzyme lacks confirmed catalytic activity for hydroxyectoine formation.

2.3.3. β-Carotene, Astaxanthin, and Lutein Production

Carotenoid biosynthesis was studied by targeting the genes involved in the terpenoid backbone biosynthesis pathway. For β-carotene, the cruA/B genes that codify for the lycopene cyclase and lcyB, crtL1, and crtY that codify for the lycopene beta-cyclase were analyzed. These genes were predominantly detected in photosynthetic organisms, such as the green algae Chlorella and the cyanobacteria Leptolyngbya and Pseudoanabaena, and were therefore most abundant in the algal culture (1992 ± 268) and the algal–methanotrophic co-culture (1621± 283) (Figure 4 and Figure 5). Chlorella carried the lcyB and crtY genes, while Leptolyngbya and Pseudoanabaena contained cruA/B. Interestingly, several bacterial taxa that were exclusively detected in the algal–methanotrophic co-culture, including Ilumatobacter, Gelidibacter, and Sphingopxys, also possessed the genes lcyB, crtL1, crtY, suggesting a potential contribution to carotenoid synthesis under mixed-culture conditions. In contrast, in the methanotrophic culture, the genes for β-carotene synthesis were scarcely represented (144 ± 41), with only Gordonia possessing the lycopene cyclase lycB gene. In both the algal–methanotrophic culture and the methanotrophic culture, a gene cluster related to 4,4′-diapolycopene biosynthesis [30] including diapolycopene oxygenase (crtP) and phytoene desaturase (crtI) was identified in Methylomonas and Methylotuvimicrobium. This indicates that methanotrophs may have contributed to carotenoid precursor formation, such as lycopene, although they were unlikely to be directly involved in β-carotene synthesis [31].

Astaxanthin biosynthesis was investigated by targeting the genes encoding β-carotene/zeaxanthin 4-ketolase (crtW, bkt), which catalyzes the ketolation of β-carotene or zeaxanthin to form astaxanthin as the final product. These genes were exclusively detected in the cyanobacteria Leptolyngbya and Pseudoanabaena and were present only in the algal (157.6 ± 12.3) and algal–methanotrophic (1.7 ± 0.3) cultures, being completely absent in the methanotrophic assay (Figure 4 and Figure 5). The distribution of these genes mirrored the occurrence of their corresponding cyanobacterial hosts, which were present only in the algal and co-culture assays. The absence of crtW and bkt in Chlorella and Dunaliella is consistent with its known carotenoid metabolism, as most green algae preferentially synthesize lutein and β-carotene rather than astaxanthin, unless genetically modified or exposed to extreme stress [32]. Accordingly, the experimentally measured astaxanthin concentrations were low across all assays, matching the low total abundance of crtW and bkt genes and cyanobacterial taxa. Based on these observations, astaxanthin presence in the system was tentatively associated with cyanobacteria due to their genomic potential; however, this inference is based solely on gene detection, and further targeted metabolite studies are required to confirm active astaxanthin biosynthesis and to elucidate the role of possible metabolic interactions within the microbial community.

Lutein biosynthesis was examined by targeting the genes LUT1 and CYP97C1, which encode the carotenoid epsilon-hydroxylase. This enzyme catalyzes the hydroxylation of the epsilon-ring of α-carotene, leading to the formation of lutein, one of the major xanthophylls in photosynthetic organisms. These genes were exclusively detected in the halotolerant green alga Dunaliella and appeared only in the algal–methanotrophic co-culture (670.8 ± 54.1), while they were completely absent in the methanotrophic culture and very low in the algal culture (Figure 4 and Figure 5). These findings indicate that mixed algal–bacterial consortia can stimulate xanthophyll biosynthesis through interspecies interactions and metabolic crosstalk, contributing to the overall increase in pigment yield.

Overall, the metagenomic analyses revealed a tightly interconnected microbial network underpinning biogas valorization, in which phototrophic, methanotrophic and methylotrophic microorganisms played complementary metabolic roles. CO₂ fixation was primarily driven by phototrophs via the Calvin–Benson cycle, particularly in algal cultures and co-cultures, while CH₄ oxidation was conducted by halotolerant methanotrophs, supported by syntrophic interactions with methylotrophs. These interactions facilitated efficient carbon transfer and community stability under saline conditions. The methanotrophic culture and the co-culture showed the highest potential for ectoine and hydroxyectoine synthesis, consistent with product yields and highlighting the role of C1-metabolizing bacteria in osmolyte production. In contrast, carotenoid biosynthesis was mainly associated with phototrophic taxa, with β-carotene linked to green algae and cyanobacteria, astaxanthin restricted to cyanobacteria, and lutein enhanced in the algal–methanotrophic co-culture through the activity of Dunaliella. These results highlight that algal–methanotrophic co-cultures foster metabolic complementarity and functional redundancy, enhancing biogas valorization and broadening the spectrum of recoverable bioproducts.

3. Materials and Methods

3.1. Mineral Medium and Inoculum

The mineral salt medium (MSM) was composed of (per L of solution) 0.20 g MgSO₄·7H₂O, 0.11 g CaCl₂·2H₂O, 1.0 g KNO₃, 50 mL of 1M NaHCO₃ buffer (82 g·L⁻¹), 5.0 mL 1M Na₂CO₃ buffer (106 g·L⁻¹), 1 mL of trace elements solution, and 6% w/w NaCl (based on the MSM for haloalkaliphilic methane-oxidizing bacteria proposed by Kalyuzhnaya et al. [33]). The pH was adjusted to 9 using a phosphate-buffered solution containing (per L) 14.0 g KH₂PO₄ and 30 g Na₂HPO₄·12H₂O. All reagents were purchased from Panreac Applichem (Barcelona, Spain) and COFARCAS (Burgos, Spain) with a purity > 99%.

The methanotrophic inoculum was obtained from a Taylor reactor operated at the Institute of Sustainable Processes (Valladolid, Spain) containing a halotolerant methanotrophic bacterial consortium [34]. The algal inoculum consisted of a non-axenic culture previously enriched in a photobioreactor for carotenoid production [35]. Methanotrophic and algae cultivations were conducted separately in triplicate 120 mL glass serum bottles containing 50 mL of MSM with 3% NaCl (30 g·L^–1). The bottles were closed with gastight butyl septa and aluminum caps, and synthetic biogas (70% CH₄:30% CO₂, v/v) was injected into the headspace. In the methanotrophic cultures, 20% (v/v) of the biogas headspace was replaced with pure O₂. The synthetic biogas and pure O₂ were purchased from Carburos Metálicos (Barcelona, Spain). After autoclaving, bottles were inoculated with 20 mL of each culture. Inocula were grown at 25 °C under orbital agitation (150 rpm). Algal cultures were exposed to light (700 μmol m⁻² s⁻¹), while methanotrophic cultures were maintained in the dark. When the biomass reached the exponential growth phase, each inoculum was transferred to triplicate serum bottles containing fresh MSM supplemented with 6% NaCl (60 g·L^–1), prepared in the same manner. Once active growth was achieved and biomass concentrations reached 100–400 mg·L^–1, the cultures were used as inoculum for the bioreactor experiments.

3.2. Experimental Set-Up

The experiments were conducted in triplicate using autoclaved 2.1 L gas-tight glass bioreactors, with three reactors operated for methanotrophic cultures, three for algal cultures, and three for algal–methanotrophic co-cultures. The reactor headspace was filled with biogas (70% CH₄:30% CO₂, v/v) for the algal and co-culture assays, and with biogas supplemented with O₂ (80 biogas:20 O₂, v/v) for methanotrophic assays (

Table 1. Cultivation conditions during experimental assays.

Assay	Gas Phase (v/v)	Light (μmol·m−2·s−1)	Cultivation Time (Days)	Initial Biomass (g TSS·L−1)
Algal culture	70% CH4:30% CO2	700 ± 50	28	0.08 ± 0.001
Methanotrophic culture	56% CH4:24% CO2:20% O2	0	43	0.05 ± 0.01
Algal–methanotrophic co-culture	70% CH4:30% CO2	700 ± 50	56	0.69 ± 0.001

). These gases were filtered through 0.22 μm pore size filters before being injected into the bioreactors’ headspace. Each reactor contained a working volume of 450 mL and was inoculated with 350 mL of sterile MSM (6% NaCl) and 100 mL of fresh inoculum. In the co-culture experiments, the inoculum comprised 50% algal culture and 50% methanotrophic culture (v/v). Abiotic controls, prepared identically but without inoculum, were also set up in triplicate. All cultures were incubated in a temperature-controlled room (25 °C) on a multi-position magnetic stirrer at 200 rpm (Thermo Scientific TM CimarecTM, Waltham, MA, USA). Methanotrophs were cultured in darkness, while algae and algae–methanotrophs were exposed to LED lights (AVP650 PCBA-M CLEARFLOOD 120 2D O5 740 LED PCBA, Signify, Madrid, Spain).

3.3. Analytical Methods

Liquid samples were collected regularly during cultivation to monitor cell biomass concentration, as well as the production of ectoines (ectoine and hydroxyectoine) and carotenoids. Biomass concentration was assessed by measuring optical density (OD) and total suspended solids (TSS). OD was determined by culture absorbance at a wavelength of 600 nm (OD₆₀₀) in a SPECTROstar spectrophotometer (BGM LABTECH, Ortenberg, Germany), while TSS was monitored gravimetrically following standard methods [36] using 0.22 µm nitrocellulose filters to quantify the dry biomass concentration.

Gas samples (100 µL) were collected from the headspace of the bioreactors using lock-tight syringes, and their composition was analyzed using a Bruker 430 GC-TCD gas chromatograph (Palo Alto, CA, USA). The system was equipped with CP-Molsieve 5A (15 m × 0.53 mm × 15 mm) and CP-PoraBOND Q (25 m × 0.53 mm × 10 mm) columns, with helium (13 mL/min) as the carrier gas. The column and detector temperatures were maintained at 45 °C and 200 °C, respectively.

Carotenoids were extracted from 2 mL liquid samples using preheated DMSO at 50 °C, followed by shaking in the dark at 100 rpm for 5 min. Subsequently, 300 µL of acetone was added, and the mixture was homogenized manually. Samples were centrifuged at 11,000 rpm for 5 min, and the resulting supernatants were filtered through 0.22 µm membranes. Approximately, 250 µL of each filtrate was transferred into amber vials for liquid chromatography analysis. Carotenoid composition was analyzed via High-Performance Liquid Chromatography–Ultraviolet (HPLC-UV) using a 60 min program and a C30 bonded silica-based reversed-phase column (5 µm, 250 × 4.6 mm, YMC Co., Ltd., Kyoto, Japan). The mobile phase consisted of eluent A (CH₃OH:MTBE:H₂O 81%:15%:4% v/v/v) and eluent B (CH₃OH:MTBE:H₂O 6%:90%:4% v/v/v), with detection at 254 nm. High-purity standards (>99.5%) of astaxanthin, lutein, and β-carotene were obtained from Sigma-Aldrich (Madrid, Spain). Accordingly, the chromatographic analysis was targeted exclusively to these carotenoids as part of the preliminary screening of compounds of biotechnological interest.

For ectoine and hydroxyectoine analysis, 2 mL liquid samples were centrifuged at 11,000 rpm for 9 min, and the supernatants were discarded. Pellets were resuspended in 2 mL of ultrapure water and transferred to 2 mL microcentrifuge tubes containing 25–30 mg of zirconia powder. Homogenization was performed through mechanical disruption in three cycles of 60 s each, with a 1 min pause between cycles. After a final centrifugation at 11,000 rpm for 2 min, approximately 1.5 mL of the supernatant was filtered (0.22 µm) and transferred to vials for analysis. Samples were analyzed via HPLC-UV using a 20 min program with a liquid purple amino column (3 µm, 150 × 4.6 mm). The mobile phase consisted of ACN:H₂O (75%:25% v/v), with detection at 210 and 220 nm. High-purity standards (>99.5%) for ectoine and hydroxyectoine were obtained from Sigma-Aldrich (Spain).

3.4. Metagenomic Analysis

Shotgun metagenomics analysis was performed to assess the microbial populations from each culture and the main organisms involved in CH₄ and CO₂ metabolisms, as well as those involved in ectoine and carotenoid biosynthesis in the algal culture, methanotrophic culture, and their co-culture. To this aim, triplicate biomass samples were collected from each culture at the end of the experimental period. The DNA was extracted and sent for Illumina shotgun sequencing (Novaseq pe150 platform-Illumina NovaSeq 6000) to Novogene Co. (Munich, Germany). Library construction, quality control and sequencing were developed according to Marcos-Rodrigo et al. [37]. Fastp (v. 0.23.1) (https://github.com/OpenGene/fastp (accessed on 22 July 2025)), and Bowtie2 (v. 2.5.4) software (http://bowtie-bio.sourceforge.net/bowtie2/index.shtml (accessed on 22 July 2025)) was used for preprocessing the raw data from the Illumina sequencing platform. The assembly of metagenomes was developed using MEGAHIT (v1.2.9) [38]. MetaGeneMark (v.3.38) (http://topaz.gatech.edu/GeneMark/ (accessed on 22 July 2025)) was then used to perform ORF prediction for scaftigs (≥500 bp) of each sample [39]. For the ORF prediction results, CD-HIT software (v.4.8.1) (http://www.bioinformatics.org/cd-hit/ (accessed on 22 July 2025)) was used to obtain the non-redundant initial gene catalog (the nucleic acid sequences encoded by successive non-redundant genes are called genes) [40]. Clean data of each sample were aligned to the initial gene catalog by using Bowtie2 and genes were filtered out to finally determine the gene catalog (Unigenes) for subsequent analysis [40]. Taxonomic analysis was conducted by aligning unigene sequences against bacterial, fungal, archaeal, and viral sequences from NCBI’s NR database (https://www.ncbi.nlm.nih.gov/ (accessed on 23 July 2025)) using a cut-off e-value of 10⁻⁵. MEGAN (v6.25.9) software was used to retain species annotation information during taxonomic classification. Functional annotation of assembled metagenomes was conducted using DIAMOND (v0.9.9), aligning unigenes against functional databases, including KEGG Orthology (https://www.kegg.jp/kegg/ (accessed on 23 July 2025)), eggNOG (v.5.0) (http://eggnog5.embl.de/#/app/home (accessed on 23 July 2025)), and CAZy (https://www.cazy.org/ (accessed on 23 July 2025)), with a cut-off e-value of 10⁻⁵. Functional abundance differences were assessed using ANOSIM (v. R. 4.0.3), while comparative analyses of metabolic pathways and functional differences were performed via MetaStat and LefSe. The annotated genes were used to search for the specific genes for CH₄ oxidation, CO₂ fixation, ectoine and hydroxyectoine production, and carotenoid (astaxanthin, β-carotene and lutein) biosynthesis. The specific genes associated with each pathway were then mapped to their corresponding bacterial taxa using the NCBI database (https://www.ncbi.nlm.nih.gov/ (accessed on 10 September 2025)). This whole-genome shotgun project has been deposited in GenBank under Bioproject number: PRJNA1369521.

4. Conclusions

This study demonstrates that an algal–methanotrophic co-culture composed of haloalkaliphilic extremophiles provides an effective waste-to-value biotechnological platform for the simultaneous abatement and valorization of the GHGs, CH₄ and CO₂. Under saline conditions, the co-culture achieved up to 90% removal of both gases while producing high-value osmolytes (ectoine, hydroxyectoine) and selected carotenoids (lutein, β-carotene, astaxanthin), which were targeted in this first-stage screening. Integrated physiological and metagenomic analyses revealed strong metabolic complementarity between methanotrophs and photosynthetic microorganisms, enabling the direct conversion of biogas-derived carbon into bioactive compounds. Future research efforts will focus on optimizing cultivation conditions, improving metabolic fluxes, expanding compound profiling, and scaling up production processes to maximize the yield and cost-effectiveness of ectoine and carotenoid synthesis from biogas-derived carbon.

Beyond biogas valorization, this approach is adaptable to any industrial environment generating CH₄ and/or CO₂-rich gases, expanding its relevance to broader GHG mitigation and circular bioeconomy objectives. By enabling the direct biotransformation of GHGs into commercially valuable molecules, this platform advances both environmental and economic global priorities.