Next Article in Journal
Room-Temperature Processable TiO2 Solar Paint for Dye-Sensitized Solar Cells
Previous Article in Journal
Curriculum Development of EdTech Class Using 3D Modeling Software for University Students in the Republic of Korea
Previous Article in Special Issue
The Biofactory: Quantifying Life Cycle Sustainability Impacts of the Wastewater Circular Economy in Chile
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Sustainability
Volume 15
Issue 24
10.3390/su152416607
sustainability-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Enrichment of a Mixed Culture of Purple Non-Sulfur Bacteria for Hydrogen Production from Organic Acids

by
Sean C. Smith
1,
Javiera Toledo-Alarcón
2,
María Cristina Schiappacasse
1 and
Estela Tapia-Venegas
3,4,*
1
Escuela de Ingeniería Bioquímica, Pontificia Universidad Católica de Valparaíso, Av. Brasil, 2085, Valparaíso 2340025, Chile
2
Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Av. Padre Hurtado 750, Viña del Mar 2520000, Chile
3
Departamento de Ciencias de la Ingeniería para la Sostenibilidad, Facultad de Ingeniería, Universidad de Playa Ancha, Valparaíso 2500100, Chile
4
HUB Ambiental UPLA, Universidad de Playa Ancha, Valparaíso 2500100, Chile
*
Author to whom correspondence should be addressed.
Sustainability 2023, 15(24), 16607; https://doi.org/10.3390/su152416607
Submission received: 22 September 2023 / Revised: 2 December 2023 / Accepted: 4 December 2023 / Published: 6 December 2023
(This article belongs to the Special Issue Biorefinery of Wastewater Treatment Plant: The Pathway to Sustainable Development)
Browse Figures
Versions Notes

Abstract

:
Hydrogen (H2) as a clean fuel holds global potential and can be produced through bio-processes. To enhance bioH2 yields, integrated systems have been proposed, combining dark fermentation (DF) of wastewater with a subsequent photofermentation (PF) stage involving purple non-sulfur (PNS) bacteria. Mixed cultures of PNS bacteria and their microbial ecology have been relatively understudied despite the known benefits of mixed cultures in industrial applications. The aim of this study was to obtain various mixed cultures of PNS bacteria under different environmental conditions during the enrichment stage. Four different mixed cultures were obtained (A, B, C, and D). However, in the H2 production phase, only Consortium A, which had been enriched with malic acid as the carbon source, exposed to 32 W m−2 of irradiance, and subjected to intermittent agitation, produced H2 with a yield of 9.37 mmol H2 g−1 COD. The consortia enriched were a hybrid of PF and DF bacteria. Especially in Consortium A, Rhodopseudomonas palustris was the dominant organism, and various DF bacteria were positively associated with H2 production, with their dominance comparable to that of PNS bacteria. Despite the reported low yields, optimizing environmental conditions for this culture could potentially enhance hydrogen production from DF effluents.

1. Introduction

International initiatives that focus on decarbonizing the energy sector and mitigating environmental pollution have positioned hydrogen (H2) as the ideal replacement for fossil fuels. Hydrogen, renowned for its high energy density, has the potential to power the world without emitting any pollutants, thus contributing to sustainability by providing an alternative for clean energy production [1]. Among the alternatives for biological H2 production, dark fermentation (DF) and photofermentation (PF) stand out. Despite their comparatively lower yields when compared to steam reforming and water electrolysis, these technologies have attracted considerable interest due to their ability to utilize various organic substrates, including waste materials, which enhances the sustainability of hydrogen production [1,2,3]. To increase yields, different strategies have been proposed, such as coupling biological systems to maximize the conversion of organic matter into H2. In this context, H2 can be produced in two consecutive stages of DF and PF: raw substrate is fed to a higher-yield DF reactor, which has an effluent high in volatile fatty acids (VFAs); this effluent is fed into a second stage PF reactor, where the VFAs are fully metabolized into H2 and CO2 by purple non-sulfur (PNS) bacteria [1,2,3,4,5,6]. An integrated process between DF and PF would lead to a more complete conversion of substrates into biohydrogen [7].
Several researchers have studied this DF-PF-coupled system technology with pure cultures of PNS bacteria and report yields from 6 to 85% of the theoretical maximum [8,9,10]. DF effluents are generally diluted, centrifuged or sterilized before being introduced into the PF stage to avoid adverse effects such as high biomass concentration that affects the availability of light in the photo-bioreactors or excess concentration of VFA, ammonia and/or other potential inhibitors found in DF effluents which are unfavorable to the optimal functioning of PNS bacteria [10,11]. In previous studies, it has been reported that pure cultures have higher yields compared to mixed cultures due to competition for substrate among microorganisms (negative interactions) [12]. However, mixed cultures exhibit a distinctive feature of strong interactions between dominant and subdominant microorganisms, which can also be positive, potentially leading to a broader range of metabolic pathways and increased resistance to contamination. This is particularly significant when dealing with non-sterile effluents from the DF used as feedstock [13,14,15,16,17]. These characteristics make the study of mixed cultures particularly interesting due to the economic advantages they offer, as it avoids the energy consumption associated with sterilizing the raw material used to feed these cultures [12,13,14].
Most pure-culture studies of PNS bacteria have been conducted using Rhodospirillum rubrum, Rhodobacter sphaeroides, Rhodobacter capsulatus, Rhodovulum sulfidophilum, Rhodopseudomonas faecalis, and Rhodopseudomonas palustris, as well as Rhodospirillum rubrum which have been isolated from saltwater, freshwater, or wastewater treatment plants [18]. Additionally, the environmental conditions that influence H2 production from PF are the substrate type and concentration, the nutrient composition in the culture medium, illumination conditions, temperature, pH, and the shape and mechanical configuration of the photo-bioreactor [14]. Typically, malate, lactate, succinate, acetate, and butyrate are recognized as the most typical organic acid substrates, exhibiting different impacts on the yield of hydrogen (H2) production depending on the PNS bacteria species [19]. However, acetate and butyrate have been found to promote the production of polyhydroxyalkanoic acids (PHAs) [20,21]. In addition, the ideal VFA substrate for H2 production differs by species of PNS bacteria [21]. PNS bacteria have the capability to capture and utilize both visible and infrared light (400–700 nm and 700–900 nm, respectively) using bacteriochlorophyll a or b, along with various carotenoids. Illumination is generally provided by lamps with an intensity of approximately 4 to 300 W m−2. It has been reported that butyrate consumption requires higher light intensities in comparison to acetate or propionate [9,22,23]. The culture conditions are generally a pH of 6.8–7 and a temperature of 28 to 40 °C [9,10]. However, it has been reported that with a mixed culture the optimal pH range may vary between 7 and 8 to 8 and 9 for acetate and butyrate, respectively [23]. Also, enrichments have previously been carried out to obtain mixed cultures of PNS bacteria from activated sludge found in wastewater treatment plants; the best conditions for H2 production using these consortia are not always those same enrichments [24].
The microbial enrichment phase in a PF is essential for the development of a robust mixed culture, enabling the selection of bacteria that can work together and adapt to new substrates and environmental conditions. Simultaneously, it is crucial to determine the ecological role of each bacterial species to gain a better understanding of microbial metabolic networks within mixed cultures and to promote positive interactions among them.
This study aimed to obtain different mixed cultures of PNS bacteria from a natural inoculum, using different sets of environmental conditions (enrichment stage), and to study their production of H2 from a DF effluent (H2 production stage), placing emphasis on the analysis of the microbial community developed under the experimental conditions.

2. Materials and Methods

2.1. Selection of the Purple Mixed Microbial Culture: Enrichment Phases

To obtain a H2-producing mixed culture enriched with PNS bacteria, two consecutive stages, Phase I and Phase II, of enrichment were carried out. Phase I was conducted as a one-time enrichment to isolate PNS bacteria from anaerobic sludge. Phase II was carried out in sequential batches to obtain a culture of PNS bacteria with a higher concentration of biomass. Different environmental conditions, including irradiance, agitation mode, and carbon source, were tested in accordance with the parameters outlined in Table 1 during the enrichment phases. Irradiance was evaluated at 16 (low) and 32 (high) W m−2 (as measured from the surface normal of the bottle to the source of illumination); agitation was continuous or intermittent (on/off 90 s:90 s) at 120 rpm; the source of carbon was either malic acid (3.3 g chemical oxygen demand (COD) L−1) or a VFAs mixture of 3.3 g COD L−1 with acetic, succinic and butyric acids (0.4 gL−1, 1.6 gL−1 and 0.5 gL−1, respectively). The VFA mixture was designed to replicate a synthetic DF effluent, following the description provided by Silva-Illanes et al. [25]. Essential nutrients for PNS bacteria growth were added to both culture media. The nutrient medium used was adapted from Fang et al. [26] and was composed of 500 mgL−1 KH2PO4, 400 mgL−1 MgSO4, 400 mgL−1 NaCl, 50 mgL−1 CaCl2, 3.9 mgL−1 Fe(III)-citrate; 0.3 mgL−1 HBO3, 0.03 mgL−1 Na2MoO4, 0.1 mgL−1 ZnSO4, 0.2 mgL−1 CoCl2, 0.01 mgL−1 CuCl2, 0.03 mgL−1 MnCl2, 0.02 mgL−1 NiCl2, 0.04 mgL−1 vitamin B12 and 0.589 gL−1 glutamic acid as sources of nitrogen. The experiments were conducted in closed bottles of 0.5 L useful volume. The temperature was maintained at 37 °C in a heated chamber and the initial pH was set at 7.0. Irradiance was provided by artificial illumination with a 150 W halogen lamp (2500 lx measured with a light meter, Light Probe MeterTM, EXTECH Instruments, on the surface of the bottles) placed 30 cm and 15 cm from the bottles for low and high irradiance, respectively (see Figure 1). The spatial distribution of irradiance in the reactors was determined by measuring multiple points along the irradiated surface using a spectroradiometer (LI-1400; Li-Cor, Lincoln, NE, USA). Argon gas was purged through the aqueous medium for 20 min after seeding the bottles with bacterial culture to establish and maintain anaerobic conditions. Specific information for Phases I and II is detailed below. The phases were evaluated for the formation of purple biomass and biogas production (especially H2).
Phase I—Enrichment: The sludge obtained from a granular anaerobic digester, which treated wastewater from a tobacco industrial wastewater plant, was used as the inoculum (“Anaerobic sludge (AI)”, Table 1) and introduced into the experimental bottles at a suspended solids/chemical oxygen demand of carbon source ratio (SS/COD) of 0.5 mgSS mgCOD−1. During this stage, various environmental conditions (Table 1) were implemented for a duration of 7 days.
The microbial consortium selected at this stage (“Purple inoculum” (PI)) was visually identified as a deep red biofilm forming on the bottle wall facing the light source and was collected from the bottle wall using an inoculation loop which was used as the inoculum in Phase II.
Phase II—Enrichment: This phase involved the utilization of the purple inoculum (PI) obtained in Phase I. The PI was suspended in the same culture medium and introduced into the experimental bottles with an SS/COD carbon source ratio of 0.5 mgSS mgCOD−1 (see Table 1). The enrichment was conducted in sequential batches until the carbon source was depleted by 90% (approximately 7 days). This process was repeated until a biomass concentration of 600 mg-VSS L−1 was achieved, using the same experimental conditions (see Table 1) that led to the selection of the specific consortium in Phase I. After each batch, the contents of all bottles were subjected to centrifugation for 10 min at 10,000× g (10,000 rpm). The resulting pellet was then resuspended in fresh culture media. The microbial consortium chosen during this stage was referred to as the enriched purple inoculum (EPI) and was sampled for 16S rDNA sequence analysis.

2.2. Hydrogen Production Phase: H2 Production by the Enriched Purple Inoculum (EPI)

An evaluation of H2 production in a synthetic DF effluent was performed for each selected consortium in 0.5 L (working volume) glass batch anaerobic reactors. All consortia successfully enriched in Phase II (EPI) provided biomass as a source of inoculum for the H2 production phase. After the Phase II enrichment, the bacterial suspension was centrifuged for 10 min at 10,000× g (10,000 rpm) to obtain a concentrated pellet corresponding to the EPI. In this stage of the study, each EPI was seeded in an environmental condition independent of the conditions used for their enrichment. Operational conditions were 16 W m−2 irradiance under artificial illumination, continuous agitation and a VFA mixture as the carbon source, simulating a DF effluent [25]. The initial pH in all experiments was 7.0, and the temperature was maintained for all reactors at 37 °C. Artificial illumination was provided by a halogen lamp as previously described in the enrichment phases. The mineral medium, excluding the carbon source, remained the same as in the prior enrichment phases (Phase I and Phase II). The carbon source concentration was 0.6 g chemical oxygen demand (COD L−1) and the initial inoculum concentration was added in a SS/COD ratio of 0.5. The H2 yield was measured as total moles of H2 produced divided by mg of chemical oxygen demand (COD) consumed.

2.3. Analytical Methods

Reactor samples were collected at specific time intervals from the mixed culture liquor. The collected samples were then subjected to centrifugation and filtered using a 0.22 μm filter. To determine the concentrations of fatty acids using the HPLC system, we followed the methodology of Vesga-Baron et al. [27]. The biogas flow rate was measured using the Automatic Methane Potential Test System® (BPC Instruments, Lund Sweden), which is an automated measurement tool. Gas chromatography analysis was employed to determine the composition of the biogas, which included H2, N2, CH4, and CO2, following the methodology described by Silva Illanes et al. [25]. Suspended solids and chemical oxygen demand (COD) were assessed following the methodology described in Tapia-Venegas [28].

2.4. Microbial Diversity Analyses

The microbial communities of each enriched consortium after Phase II (EPI) were characterized by collecting samples. Initially, the samples were centrifuged, and DNA extraction was performed following the methodology described by Silva-Illanes et al. [25]. The extracted DNA was resuspended in a 0.06% NaCl solution and stored at −20 °C before being used. For the amplification of the V4 variable region of the 16S rRNA gene, PCR primers 515/806 with a barcode on the forward primer were used. The PCR reaction was conducted for 28 cycles, with 5 cycles dedicated to PCR product amplification, using the HotStarTaq Plus Master Mix Kit (Qiagen, Redwood City, CA, USA). The PCR conditions consisted of an initial denaturation at 94 °C for 3 min, followed by 28 cycles of denaturation at 94 °C for 30 s, annealing at 53 °C for 40 s, and extension at 72 °C for 1 min. A final elongation step at 72 °C for 5 min was carried out. The amplification success was evaluated by visualizing PCR products, following the methodology described by Ahmad et al. [28]. The Illumina DNA library was prepared by pooling and purifying the PCR products, following the methodology described by Ahmad et al. [29]. The sequencing was carried out at MR DNA (www.mrdnalab.com, accessed on 15 July 2022; Shallowater, TX, USA), and the instrument used and the processing of the sequence data were also performed according to the methodology described by Ahmad et al. [29]. Sequencing targeted the V4 region of the 16S rRNA gene, following the protocols provided by MR DNA Laboratory. The generated sequences underwent several processing steps to ensure data quality. Barcodes and primers were removed, sequences below 150 base pairs and those with ambiguous base calls were eliminated, and homopolymer runs longer than 6 base pairs were also filtered out. Operational taxonomic units (OTUs) were defined through clustering with a 3% divergence threshold (97% similarity). The taxonomic classification of the final OTUs was performed using BLASTn against a curated database compiled from RDPII and NCBI. The obtained sequences were deposited in the NCBI database under accession number OR687728-OR689030. Only microorganisms with abundances higher than 1% were considered in the composition analysis. These microorganisms accounted for more than 90% of the total community, comprising a total of 2193 OTUs. On average, each sample exhibited a total of 476 selected OTUs.

2.5. Data Analysis and Statistical Tools

To assess the impact of the selection and enrichment operational conditions on the structure of the microbial community, Canonical Correspondence Analysis (CCA) was conducted using PAST 4.06b software (http://folk.uio.no/ohammer/past/, accessed on 10 August 2023) [30,31]. Correlations between the bacterial community structure and various (normalized) environmental variables (EVs) were investigated. The EVs were represented on the CCA plot as arrows, with their direction indicating the increasing EV gradient and their length representing the magnitude of the correlation between the EV and the ordination. The dissimilarities between Consortia A, B, C, and D were quantified using the Euclidean distance metric.

3. Results

3.1. Enrichment and Hydrogen Production Phase of the Purple Mixed Microbial Culture

Table 2 presents the qualitative (purple coloring) and quantitative results (biogas production and composition) obtained in enrichment Phases I and II for the different experimental conditions studied (Experiments 1 to 8). After 7 days of microbial growth in Phase I, purple biomass growth on the bottle walls was observed in 50% of the conditions studied (see Figure 2a). As shown in Table 2, the on/off agitation mode and an irradiance of 32 W m−2 were more common in the bottles that showed purple biomass, while there was no apparent preference among the carbon sources tested (Experiments 1, 2, 4, 5). However, Experiment 4 used an irradiance of 16 W m−2 and Experiment 5 used continuous agitation mode. Only in Experiment 1, where on/off agitation, an irradiance of 32 W m−2 and malic acid as carbon source were used, biogas accumulation associated with methane production was observed, probably due to the origin of the initial inoculum in Phase I.
According to the above results, Experiments 1, 2, 4, and 5 were selected for enrichment in Phase II, maintaining the same environmental conditions as in Phase I. After 4 weeks, only Experiment 1 accumulated biogas, producing mainly H2 (73.2%) and CO2 (26.8%). A biomass concentration of 600 mg-VSS/L was reached at 8 weeks (Figure 2b). According to Table 2, at the end of Phase II, enriched purple inocula (EPI) from Experiments 1, 2, 4, and 5 were named Consortia A, B, C and D, respectively.
For the H2 production phase, only Consortium A exhibited H2 production under the conditions studied. It had a yield of 9.37 mmol H2 g−1 COD, which could also be represented as 4.7 mmol H2 g−1 SS of the mixed culture or cumulative H2 production 5.6 mmol H2 L−1 (see Table 2). In relation to the removal of fatty acids from the synthetic medium, a 100% removal of each of them (acetic, succinic, and butyric) was observed.

3.2. Microbial Community Structure of Purple Mixed Microbial Culture

The microbial community structures at the family and genus levels, obtained at the conclusion of Phase II for each respective EPI (A, B, C, and D), are presented in Table 3. These structures are then compared to the initial inoculum of Phase I (AI). For AI only, the presence of the microorganisms found in PI enrichment is presented (its full diversity is added in Supplementary Materials Table S1). PNS bacteria enrichment was confirmed in all cases of each EPI (A, B, C, and D) by finding an increase with respect to I of the Bradyrhizobiaceae and Rhodocyclaceae families, mainly represented by the Rhodopseudomonas and Rhodocyclus genus, respectively.
In particular, Consortium A (with over 10% individual microbial diversity detected) was dominated by the families Campylobacteraceae (24.4%), Aeromonadaceae (20.1%), Bradyrhizobiaceae (19.7%), and Bacteroidaceae (10.3%), represented mainly by genera Sulfurospirillum, Tolumonas, Rhodopseudomonas, and Bacteroides, respectively. Consortium B was dominated by the families Bradyrhizobiaceae (39.9%), Rhodocyclaceae (14.6%) and Bacteroidaceae (11.7%), represented mainly by genera Rhodopseudomonas, Rhodocyclus, and Bacteroides, respectively. Consortium C was dominated by the families Bradyrhizobiaceae (32.2%), Porphyromonadaceae (13.1%), and Veillonellaceae (13.0%), represented mainly by genera Rhodopseudomonas, Parabacteroides, and Sporomusa, respectively, while Consortium D was dominated by the families Pseudomonadaceae (49.0%), Rhodocyclaceae (17.9%), and Marinilabiliaceae (9.9%), represented mainly by genera Pseudomonas, Rhodocyclus, and Alkaliflexus, respectively.
A was the only EPI that produced H2. It was observed that the subdominant genera (less than 10% and greater than 1%) in this consortium were Rhodocyclus, Parabacteroides, Desulfovibrio, and Clostridium and that they were also found in the other EPI (B, C, and D).

3.3. Influence of Enrichment Operating Conditions on Microbial Community Structure

A canonical correspondence analysis (CCA) was performed to investigate the relationship between the operational conditions and the selection and enrichment of the microbial community, as illustrated in Figure 3. The eigenvalues associated with CCA1 and CCA2 were 0.638 and 0.434, representing 51.5% and 35.0% of the total inertia, respectively. A permutation (N: 999; Trace: 1.239; p-value: 0.756) was used to determine that the data corresponded significantly and deviated from the null hypothesis of no association, according to the logic of the Pearson’s χ2 statistic [30].
Operational conditions applied developed different microbial communities in each EPI (A, B, C, and D), as evidenced by the distance between the samples in Figure 3 (yellow triangles marked A, B, C, and D). These distances were quantified using the basic Euclidean distance, where Consortium D was the most different with a distance of 0.638, 0.649 and 0.640 from Consortia A, B, and C, respectively. The most similar consortia were B and C with a distance of 0.264. Consortium A showed a distance of 0.363 and 0.414 from Consortia B and C, respectively.
PNS bacteria enrichment was strongly influenced by the operating conditions applied, as shown in Figure 3. Members of the Rhodocyclaceae family were strongly favored when using high irradiance. In addition, this family was strongly related to the Desulfovibrionaceae family enrichment. Members of the Bradyrhizobiaceae family were strongly favored when using on/off agitation and they were moderately influenced by low irradiance and VFA as a carbon source. In addition, this family was strongly related to the enrichment of Acidaminococcaceae, Acholeplasmataceae, and Clostridiaceae families.
The operational conditions applied not only significantly affected the PNS bacteria enrichment; other members of the microbial community were also strongly affected. Continuous agitation favored the selection of the Pseudomonadaceae, Marinilabiliaceae, and Hyphomicrobiaceae families, while on/off agitation favored the selection of the Clostridiaceae, Acholeplasmataceae, Acidaminococcaceae, and Bacteroidaceae families. VFA as a carbon source favored the selection of the Acidaminococcaceae, Porphyromonadaceae, Desulfomicrobiaceae, Sphingobacteriaceae, Gracilibacteraceae, and Spirochaetaceae families.
H2 production was positively associated with the Aeromonadaceae family represented by the genus Tolumonas, Campylobacteraceae represented by Sulfurospirillum and Arcobacter, Bacteroidaceae represented by the genus Bacteroides, and Clostridiaceae represented by the genus Clostridium and Geosporobacter (see Figure 3). Apparently, the abundance of PNS bacteria was not directly related to H2 production.
Figure 4 was constructed to highlight the dominance among bacteria positively associated with hydrogen production and the PNS bacteria. According to Figure 4, the relative abundance of PNS bacteria in Consortium A (26.3%), especially the genus Rhodopseudomonas (19.7%), was lower compared to Consortium B (54.5%), Consortium C (35.3%), and Consortium D (18.1%). When considering only Consortium A, the microorganisms positively related to hydrogen production that were dominantly present were Tolumonas and Sulfurospirillum at 20.1% and 22.2%, respectively. In contrast, Bacteroides were moderately dominant (10.3%), while Clostridium (4%) and Arcobacter (2.2%) had low dominance.

4. Discussion

4.1. Operational Enrichment Conditions

The different environmental conditions used to enrich mixed cultures generated consortia with diverse compositions, but also with a significant percentage of PNS bacteria (ranging from 18 to 53%) and non-PNS bacteria associated with hydrogen production, which were not dominant in all cases (1 to 59%). The only hydrogen-producing culture was obtained under the enrichment conditions of malic acid, on/off agitation conditions, and 32 Wm−2 of irradiance. This consortium had 26% PNS bacteria and 59% non-photofermentative bacteria. According to this study, PNS bacteria were enriched in both the VFA mixture and malic acid, which is not surprising. It has been reported that these bacteria can utilize a wide variety of organic carbon substrates, including acetic, propionic, butyric, malic, succinic, lactic acid, etc. [17,32,33]. Agitation has been considered a positive environmental condition because it helps improve nutrient and substrate mass transfer efficiencies, further influencing light penetration and therefore optimal light intensity [34]. In the literature, continuous stirring speeds between 100 and 150 rpm are typically used [33], and this study was within that range. In this study, it is likely that the negative influence of continuous agitation was due to its intensity, preventing purple bacteria from adhering to the reactor wall, thus hindering their visibility and easy selection. In the literature, irradiances between 0.5 and 800 W m−2 have been tested [33,35], and diverse ranges, such as 19 to 375 W m−2, have been explored, even for mixed cultures of PNS [13,36,37]. This study also used irradiances within these ranges. Although it has been demonstrated that light intensity is a crucial factor for hydrogen generation, its effectiveness depends on various conditions, such as the utilized substrates (C), the nitrogen sources (N), the C/N molar ratio, and trace elements, as well as microbial diversity [35,36]. In this context, Ren et al. [38] concluded that the H2 production yield with Rhodopseudomonas faecalis decreased with increasing light intensity, reaching the maximum value at 3000 to 5000 lx. Therefore, optimizing irradiance conditions in future studies may enhance hydrogen production.

4.2. Microbial Community Structure in the Consortia

The predominant photosynthetic organism in the sole H2-producing consortium (A) was Rhodopseudomonas palustris, constituting 19.7% of its composition. Interestingly, this species was also found in other consortia (B and C) but in higher proportions, accounting for 39.9% and 32.2% of their respective compositions. Previous studies involving mixed cultures enriched with purple non-sulfur (PNS) bacteria often identified Rhodopseudomonas as the dominant genus, reaching up to 72% in some cases [33,39]. Considering the conditions of this study, it is expected that R. palustris would exhibit a photoheterotrophic metabolism, and this metabolism was confirmed in this study through its complete consumption of the provided carbon sources. It was not surprising to find it enriched under different conditions tested in this study, because this organism is well-known for its remarkable ability to metabolize a wide variety of feedstocks [40,41]. According to Figure 3, no Rhodopseudomonas was positively associated with H2 production. Therefore, it is explained that even though there was a higher concentration of it in Consortia B and C, hydrogen production was not observed.
Other non-PNS bacteria such as Tolumonas, Sulfurospirillum, Bacteroides, Clostridium, and Arcobacter were positively associated with hydrogen production. Among these, those with the highest percentage in Consortium A were Tolumonas and Sulfurospirillum. The Tolumonas genus is noteworthy for its fermentative qualities, its isolation from a wastewater treatment plant, and its role as an electroactive bacterium [42,43]. Tolumonas auensis has also been detected within mixed cultures, contributing to H2 production during dark fermentation, particularly in the acid and H2 production phase [44]. T. auensis is recognized for its involvement in the production of acetate, ethanol, propionate, formate, and H2, particularly when cultivated in a glucose-rich environment [45]. Various species of Sulfurospirillum have demonstrated the capability to reduce sulfur, arsenate, or tetrachloroethane [46]. H2 production, along with subsequent consumption, has been well-documented during pyruvate fermentation for Sulfurospirillum spp., including Sulfurospirillum carboxydovorans [47].
Among the bacteria with medium dominance in Consortium A, Bacteroides is listed. This bacterium has been previously recognized as an anaerobic bacterium proficient in DF. It is capable of achieving high hydrogen yields while simultaneously producing acetic acid and propionic acid. Cases of symbiosis with a PNS bacterium, Rhodobacter, have been reported [48,49,50]. However, in this study, a clear relationship with Rhodopseudomonas or Rhodocyclus is not evident. Arcobacter has been identified in other studies on dark fermentation as a component of the dominant microbial community within the wastewater influent [51]. The bacteria with a lower dominance positively associated with hydrogen are Clostridium and Arcobacter. Clostridium stands out as one of the most commonly used microorganisms for hydrogen production through dark fermentation. It has been reported to achieve the highest documented yields while concurrently producing fatty acids [52,53]. In contrast, Arcobacter anaerophilus exhibits lithoautotrophic growth utilizing H2 and hydrogen sulfide [46,49]. Arcobacter is known to consume H2, and prior studies have revealed a mutualistic relationship between Lenisia limosa and Arcobacter with regards to hydrogen transfer. Notably, the hydrogenase enzyme in L. limosa can generate acetate from NAD(P)H, and its expression is observed in the presence of Arcobacter, but not in its absence [54].
The other photosynthetic organism found in the consortia was Rhodocyclus tenuis. Its presence was expected in this study under all conditions, as it has been documented to adapt to various carbon and nitrogen sources [41]. However, it only dominated in Consortium D. In turn, when R. palustris dominated, as in Consortia A, B, and C, it was in medium or low dominance, suggesting a potential competition between these PNS microorganisms. The lack of hydrogen production in Consortium D could be expected because other carbon sources, such as formate, described as inducers of maximum H2 production, were not used in this study. Ammonium chloride stood out among the nitrogen sources. In contrast, lactate and L-tyrosine have been shown to induce minimal H2 production [41]. Therefore, the conditions studied were not conducive to hydrogen generation in Consortium D.
Therefore, it is evident that only the H2-producing consortium exhibited a hybrid metabolism (PF and DF), as evidenced by the presence of other non-PNS microorganisms positively associated with hydrogen production. Furthermore, it could represent positive interactions among these [16]. According to the bacteria present in Consortium A, it is possible to find unequivocal hydrogen producers such as Tolumonas, Bacteroides and Clostridium, which are indicative of dark fermentation. On the other hand, regarding those such as the Campylobacteraceae family, it is unclear whether they could contribute to hydrogen production and/or consume the hydrogen produced. Considering the PNS bacteria observed in all consortia, known as hydrogen producers, their role in this case remains unclear. However, they do help remove the organic acids fed. Therefore, now that we have a potential hybrid microbial culture of DF and PF, such as Consortium A selected and enriched, it would be appropriate to study the conditions that enhance hydrogen production and prevent its consumption.

4.3. Hydrogen Production of the Consortia

According to the literature, the average H2 yield obtained by photofermentation is 14 mmol H2 g−1 COD feed (0.11 g COD H2 g−1 COD feed) and the maximum that has been achieved is 31 mmol H2 g−1 COD feed (0.25 g COD g COD−1 feed) depending on the conditions used (for example, the nutrients present to support growth, competing microorganisms, substrates not fully degradable, or light limitations) and the species of PNS bacteria present [33]. Hakobyan et al. [55] studied Rhodobacter sphaeroides at different iron concentrations obtaining specific yields between 0 and 11.57 mmol H2 g−1 of dry weight. If we compare with this study, yields and productions lower than the maximum reported are obtained, but within the range (9.57 mmol g−1 COD, or 5.6 mmol H2 L−1, or 4.7 mmol H2 g−1 SS). For example, considering the production of hydrogen moles per volume, Ghimire et al. [7] used a photoheterotrophic pure culture of Rhodobacter sphaeroides with a mixture of VFAs with a high concentration of butyric acid, followed by acetic and propionic in a range of 3.5 to 2.1 g COD L−1, an illuminance of approximately 4000 lx (20 W compact fluorescent light) and positioned on a continuous stirrer (250 rpm) in 500 mL bottles, obtaining a cumulative H2 production of 168.7 mL H2 L−1 (approximately 7 mmol L−1). Other studies showed that Rhodobacter sphaeroi fed with acetic acid (1.9 g COD L−1) have produced 2310 mL H2 L−1 (96 mmol H2 L−1) [56]. Sun et al. [57] studied the production of H2 for Rhodopseudomonas palustris at 35 °C, with lighting provided by an incandescent lamp, glucose as the carbon source, and stirring between 0 and 240 rpm, obtaining an accumulated production of H2 between 32 and 92.7 mmol H2 L−1 of cultivation.
Montiel-Corona et al. [24], in their study on H2 production from a mixed photoheterotrophic inoculum enriched from activated sludge, achieved a production of 1478 mL L−1 (61 mmol H2 L−1) when fed with a higher concentration of VFA mixture (24.8 g COD L−1). In the same study, they compared it to a pure culture of Rhodobacter capsulatus and obtained similar results in hydrogen production. Additionally, in this case, only one condition was used for enrichment (3 kLux, 30 °C, without stirring reported), and microbial diversity was not reported. Therefore, it remains unclear whether it was a hybrid inoculum consisting of both PF and DF microorganisms.
Other studies with hybrid cultures of PF and DF include, for example, a co-culture of Rhodobacter sphaeroides and Clostridium acetobutylicum. It was observed that at pH levels lower than 6.5, the H2 production rates in the co-culture were lower than those in dark fermentation alone and photofermentation alone, where yields were highest at pH levels above 7.0 [58]. However, in this study, an initial pH of 7.0 was used; therefore, improving pH control could enhance hydrogen yields. Several co-cultures of Enterobacter cloacae and PNS bacteria, such as Rhodobacter capsulatus or Rhodopseudomonas palustris, fed with glucose and lactose were also studied in a batch system working at 32 ± 2 °C, with stirring at 130 rpm, a luminosity of 5 kLux, and a pH of 6.25. The co-culture of E. cloacae and R. capsulatus achieved the highest hydrogen production compared to other hybrid cultures and dark fermentation alone [59]. A reported mixed consortium of purple bacteria contained R. palustris with 72% diversity, followed by Rubrivivax gelaninosus (14%), Pseudonomonas citronellolies (8%), Rhodobacter sphaeroides (2%), Pseudomonas aeruginosa (1%), Dygonomonas sp (1%), among others making up 2%. This consortium was immobilized (7 kLux at 32 °C), achieving yields of 14 and 20 mL H2 g−1 VS h−1 with an initial pH of 6.8 [39].
Although the yields and production obtained are low, it is known that environmental conditions could improve this production, such as adjusting the pH. Nevertheless, it will be necessary to study the changes in the diversity of the enriched mixed culture inoculum since, in this study, the selection and enrichment of these cultures under environmental conditions are associated with significant changes in enriched microbial diversity. It is clear that if one intends to work under non-sterile conditions, a hybrid mixed culture of PF and DF will be obtained, leading to economic savings by eliminating the need for raw material sterilization. Furthermore, this hybrid culture of PF and DF has the potential to increase overall hydrogen production from the effluent of an initial dark fermentation. In parallel, PHA production could be quantified, since PNS bacteria are receiving attention due to their ability to produce them simultaneously with H2, which could make the prospect of their cultivation more attractive [60].

5. Conclusions

Th environmental conditions for the enrichment of a promising mixed culture of PNS bacteria for a two-stage system of H2 production were malic acid as the carbon source, 32 W m−2, and discontinuous agitation.
The enriched mixed culture of PNS bacteria was characterized by H2 production fed with fatty acids that imitated an effluent from a DF-H2-producing system obtaining a H2 yield of 9.37 mmol H2 g−1 COD.
The consortia of purple non-sulfur bacteria for hydrogen production were hybrid cultures of PF and DF bacteria. These latter ones were associated with hydrogen production as they were found in greater quantities than in the other consortia. The predominant photosynthetic organism in the sole hydrogen-producing consortium was Rhodopseudomonas palustris. The dark fermentation bacteria positively associated with hydrogen production included Tolumonas, Sulfurospirillum, Bacteroides, Clostridium, and Arcobacter.
Although the reported yield in this study was low, the importance of optimizing environmental conditions to enhance the performance of the hybrid mixed culture of PF and DF is emphasized. This optimization could increase the overall hydrogen production from the effluent of an initial dark fermentation process. Simultaneously, the use of a hybrid cultivation of PF and DF can reduce energy consumption by avoiding the sterilization of the feedstock.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/su152416607/s1, Table S1: Characterization of the bacterial communities at family level initial inoculum initial inoculum of Phase I (I). Illumina 16S rRNA gene analysis.

Author Contributions

Conceptualization, E.T.-V., M.C.S. and S.C.S.; methodology, E.T.-V., M.C.S. and S.C.S.; formal analysis, E.T.-V., M.C.S. and S.C.S.; investigation, E.T.-V. and S.C.S.; data curation, E.T.-V., J.T.-A. and S.C.S.; writing—original draft preparation, E.T.-V., J.T.-A. and S.C.S.; writing—review and editing, E.T.-V., J.T.-A. and S.C.S.; supervision, E.T.-V. and M.C.S.; funding acquisition, E.T.-V. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by FONDECYT project N° 11200211. S. Smith’s master studies was funded by Rotary Global Grants scholarship from the Rotary International Foundation.

Institutional Review Board Statement

Ethical review and approval were not applicable for this study.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study were obtained through laboratory analyses and were not available in public databases.

Acknowledgments

This work is dedicated in honor of Gonzalo Ruiz-Filippi, our respected colleague who will always be remembered.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Kumar, R.; Kumar, A.; Pal, A. An overview of conventional and non-conventional hydrogen production methods. Mater. Today Proc. 2021, 46, 5353–5359. [Google Scholar] [CrossRef]
  2. Mishra, P.; Krishnan, S.; Rana, S.; Singh, L.; Sakinah, M.; Ab Wahid, Z. Outlook of fermentative hydrogen production techniques: An overview of dark, photo and integrated dark-photo fermentative approach to biomass. Energy Strategy Rev. 2019, 24, 27–37. [Google Scholar] [CrossRef]
  3. Tapia-Venegas, E.; Ramirez-Morales, J.E.; Silva-Illanes, F.; Toledo-Alarcón, J.; Paillet, F.; Escudie, R.; Marone, A. Biohydrogen production by dark fermentation: Scaling-up and technologies integration for a sustainable system. Rev. Environ. Sci. Bio/Technol. 2015, 14, 761–785. [Google Scholar] [CrossRef]
  4. Verma, D.; Ram Kumar, N.; Subudhi, S. Isolation and characterization of a novel photoheterotrophic strain ‘Rubrivivax benzoatilyticus TERI-CHL1’: Photo fermentative hydrogen production from spent effluent. Int. J. Hydrogen Energy 2020, 45, 14245–14254. [Google Scholar] [CrossRef]
  5. Moscoviz, R.; Trably, E.; Bernet, N.; Carrere, H. The environmental biorefinery: State-of-the-art on the production of hydrogen and value-added biomolecules in mixed-culture fermentation. Green. Chem. 2018, 20, 3159–3179. [Google Scholar] [CrossRef]
  6. Anam, K.; Habibi, M.S.; Harwati, T.U.; Susilaningsih, D. Photofermentative hydrogen production using Rhodobium marinum from bagasse and soy sauce wastewater. Int. J. Hydrogen Energy 2012, 37, 15436–15442. [Google Scholar] [CrossRef]
  7. Ghimire, A.; Trably, E.; Frunzo, L.; Pirozzi, F.; Lens, P.N.L.; Esposito, G.; Escudié, R. Effect of total solids content on biohydrogen production and lactic acid accumulation during dark fermentation of organic waste biomass. Bioresour. Technol. 2018, 248, 180–186. [Google Scholar] [CrossRef]
  8. Uyar, B.; Eroglu, I.; Yücel, M.; Gündüz, U. Photofermentative hydrogen production from volatile fatty acids present in dark fermentation effluents. Int. J. Hydrogen Energy 2009, 34, 4517–4523. [Google Scholar] [CrossRef]
  9. Nath, K.; Muthukumar, M.; Kumar, A.; Das, D. Kinetics of two-stage fermentation process for the production of hydrogen. Int. J. Hydrogen Energy 2008, 33, 1195–1203. [Google Scholar] [CrossRef]
  10. Laurinavichene, T.V.; Belokopytov, B.F.; Laurinavichius, K.S.; Khusnutdinova, A.N.; Seibert, M.; Tsygankov, A.A. Towards the integration of dark- and photo-fermentative waste treatment. 4. Repeated batch sequential dark- and photofermentation using starch as substrate. Int. J. Hydrogen Energy 2012, 37, 8800–8810. [Google Scholar] [CrossRef]
  11. Singh, L.; Wahid, Z.A. Methods for enhancing bio-hydrogen production from biological process: A review. J. Ind. Eng. Chem. 2015, 21, 70–80. [Google Scholar] [CrossRef]
  12. Monroy, I.; Buitrón, G. Production of polyhydroxybutyrate by pure and mixed cultures of purple non-sulfur bacteria: A review. J. Biotechnol. 2020, 317, 39–47. [Google Scholar] [CrossRef] [PubMed]
  13. Lazaro, C.Z.; Vich, D.V.; Hirasawa, J.S.; Varesche, M.B.A. Hydrogen production and consumption of organic acids by a phototrophic microbial consortium. Int. J. Hydrogen Energy 2012, 37, 11691–11700. [Google Scholar] [CrossRef]
  14. Tawfik, A.; El-Bery, H.; Kumari, S.; Bux, F. Use of mixed culture bacteria for photofermentive hydrogen of dark fermentation effluent. Bioresour. Technol. 2014, 168, 119–126. [Google Scholar] [CrossRef] [PubMed]
  15. Cardeña, R.; Moreno, G.; Valdez-Vazquez, I.; Buitrón, G. Optimization of volatile fatty acids concentration for photofermentative hydrogen production by a consortium. Int. J. Hydrogen Energy 2015, 40, 17212–17223. [Google Scholar] [CrossRef]
  16. Cabrol, L.; Marone, A.; Tapia-Venegas, E.; Steyer, J.P.; Ruiz-Filippi, G.; Trably, E. Microbial ecology of fermentative hydrogen producing bioprocesses: Useful insights for driving the ecosystem function. FEMS Microbiol. Rev. 2017, 41, 158–181. [Google Scholar] [CrossRef] [PubMed]
  17. Argun, H.; Kargi, F. Photo-fermentative hydrogen gas production from dark fermentation effluent of ground wheat solution: Effects of light source and light intensity. Int. J. Hydrogen Energy 2010, 35, 1595–1603. [Google Scholar] [CrossRef]
  18. Adessi, A.; De Philippis, R. Photobioreactor design and illumination systems for H2 production with anoxygenic photosynthetic bacteria: A review. Int. J. Hydrogen Energy 2014, 39, 3127–3141. [Google Scholar] [CrossRef]
  19. Sivaramakrishnan, R.; Shanmugam, S.; Sekar, M.; Mathimani, T.; Incharoensakdi, A.; Kim, S.; Parthiban, A.; Geo, V.; Brindhadevi, K.; Pugazhendhi, A. Insights on biological hydrogen production routes and potential microorganisms for high hydrogen yield. Fuel 2021, 291, 120136. [Google Scholar] [CrossRef]
  20. Ghosh, S.; Dairkee, U.K.; Chowdhury, R.; Bhattacharya, P. Hydrogen from food processing wastes via photofermentation using Purple Non-sulfur Bacteria (PNSB)—A review. Energy Convers. Manag. 2017, 141, 299–314. [Google Scholar] [CrossRef]
  21. Zhang, M.; Wang, S.; Ji, B.; Liu, Y. Towards mainstream deammonification of municipal wastewater: Partial nitrification-anammox versus partial denitrification-anammox. Sci. Total Environ. 2019, 692, 393–401. [Google Scholar] [CrossRef]
  22. Nath, K.; Das, D. Effect of light intensity and initial pH during hydrogen production by an integrated dark and photofermentation process. Int. J. Hydrogen Energy 2009, 34, 7497–7501. [Google Scholar] [CrossRef]
  23. Dong, L.; Zhenhong, Y.; Yongming, S.; Xiaoying, K.; Yu, Z. Hydrogen production characteristics of the organic fraction of municipal solid wastes by anaerobic mixed culture fermentation. Int. J. Hydrogen Energy 2009, 34, 812–820. [Google Scholar] [CrossRef]
  24. Montiel-Corona, V.; Revah, S.; Morales, M. Hydrogen production by an enriched photoheterotrophic culture using dark fermentation effluent as substrate: Effect of flushing method, bicarbonate addition, and outdoor–indoor conditions. Int. J. Hydrogen Energy 2015, 40, 9096–9105. [Google Scholar] [CrossRef]
  25. Silva-Illanes, F.; Tapia-Venegas, E.; Schiappacasse, M.; Trably, E.; Ruiz-Filippi, G. Impact of hydraulic retention time (HRT) and pH on dark fermentative hydrogen production from glycerol. Energy 2017, 141, 358–367. [Google Scholar] [CrossRef]
  26. Fang, H.H.P.; Liu, H.; Zhang, T. Phototrophic hydrogen production from acetate and butyrate in wastewater. Int. J. Hydrogen Energy 2005, 30, 785–793. [Google Scholar] [CrossRef]
  27. Vesga-Baron, A.; Etchebehere, C.; Schiappacasse, M.C.; Chamy, R.; Tapia-Venegas, E. Controlled oxidation-reduction potential on dark fermentative hydrogen production from glycerol: Impacts on metabolic pathways and microbial diversity of an acidogenic sludge. Int. J. Hydrogen Energy 2021, 46, 5074–5084. [Google Scholar] [CrossRef]
  28. Tapia-Venegas, E.; Cabrol, L.; Brandhoff, B.; Hamelin, J.; Trably, E.; Steyer, J.P.; Ruiz-Filippi, G. Adaptation of acidogenic sludge to increasing glycerol concentrations for biohydrogen production. Appl. Microbiol. Biotechnol. 2015, 99, 8295–8308. [Google Scholar] [CrossRef]
  29. Ahmad, H.; Ramli, R.; Ismail, N.N.; Aidit, S.N.; Yusoff, N.; Samion, M.Z. Passively mode locked thulium and thulium/holmium doped fiber lasers using MXene Nb2C coated microfiber. Sci. Rep. 2021, 11, 11652. [Google Scholar] [CrossRef]
  30. Hammer, Ø.; Harper, D.R.P. PAST: Paleontological Statistics Software Package for Education and Data Analysis. Palaeontologia Electronica 2001, 4, 1–9. Available online: http://www.sciepub.com/reference/139944 (accessed on 8 January 2023).
  31. Legendre, P.; Legendre, L.F.J.; Numerical Ecology. Elsevier Science. 1998. Available online: https://books.google.fr/books?id=KBoHuoNRO5MC (accessed on 8 January 2023).
  32. Okubo, Y.; Futamata, H.; Hiraishi, A. Characterization of Phototrophic Purple Nonsulfur Bacteria Forming Colored Microbial Mats in a Swine Wastewater Ditch. Appl. Environ. Microbiol. 2006, 72, 6225–6233. [Google Scholar] [CrossRef]
  33. Capson-Tojo, G.; Batstone, D.J.; Grassino, M.; Vlaeminck, S.E.; Puyol, D.; Verstraete, W.; Hülsen, T. Purple phototrophic bacteria for resource recovery: Challenges and opportunities. Biotechnol. Adv. 2020, 43, 107567. [Google Scholar] [CrossRef]
  34. Tiang, M.F.; Hanipa, M.A.F.; Abdul, P.M.; Jahim, J.M.; Mahmod, S.S.; Takriff, M.S.; Lay, C.-H.; Reungsang, A.; Wu, S.-Y. Recent advanced biotechnological strategies to enhance photo-fermentative biohydrogen production by purple non-sulphur bacteria: An overview. Int. J. Hydrogen Energy 2020, 45, 13211–13230. [Google Scholar] [CrossRef]
  35. Li, S.; Tabatabaei, M.; Li, F.; Ho, S.H. A review of green biohydrogen production using anoxygenic photosynthetic bacteria for hydrogen economy: Challenges and opportunities. Int. J. Hydrogen Energy, 2022; in press. [Google Scholar] [CrossRef]
  36. Cerruti, M.; Stevens, B.; Ebrahimi, S.; Alloul, A.; Vlaeminck, S.E.; Weissbrodt, D.G. Enrichment and Aggregation of Purple Non-sulfur Bacteria in a Mixed-Culture Sequencing-Batch Photobioreactor for Biological Nutrient Removal from Wastewater. Front. Bioeng. Biotechnol. 2020, 8, 557234. [Google Scholar] [CrossRef]
  37. Hülsen, T.; Barry, E.M.; Lu, Y.; Puyol, D.; Keller, J.; Batstone, D.J. Domestic wastewater treatment with purple phototrophic bacteria using a novel continuous photo anaerobic membrane bioreactor. Water Res. 2016, 100, 486–495. [Google Scholar] [CrossRef]
  38. Ren, N.Q.; Liu, B.F.; Ding, J.; Xie, G.J. Hydrogen production with R. faecalis RLD-53 isolated from freshwater pond sludge. Bioresour. Technol. 2009, 100, 484–487. [Google Scholar] [CrossRef]
  39. Guevara-Lopez, E.; Buitrón, G. Evaluation of different support materials used with a photo-fermentative consortium for hydrogen production. Int. J. Hydrogen Energy 2015, 40, 17231–17238. [Google Scholar] [CrossRef]
  40. Brown, B.; Wilkins, M.; Saha, R. Rhodopseudomonas palustris: A biotechnology chassis. Biotechnol. Adv. 2022, 60, 108001. [Google Scholar] [CrossRef]
  41. Munjam, S.; Rani, K. Enhanced bioproduction of hydrogen by alginate immobilized cells of a purple non-sulphur bacterum Rhodocyclus tenuis KU 017. Int. J. Pharma Bio Sci. 2014, 5, B887–B897. [Google Scholar]
  42. Thapa, S.; Kim, T.; Pandit, S.; Song, Y.; Afsharian, Y.; Rahimnejad, M.; Kim, J.; Oh, S. Overview of electroactive microorganisms and electron transfer mechanisms in microbial electrochemistry. Bioresour. Technol. 2022, 347, 126579. [Google Scholar] [CrossRef] [PubMed]
  43. Paquete, C.M. Electroactivity across the cell wall of Gram-positive bacteria. Comput Struct Biotechnol. J. 2020, 18, 3796–3802. [Google Scholar] [CrossRef] [PubMed]
  44. Khanthong, K.; Purnomo, C.W.; Daosud, W.; Laoong-u-thai, Y. Microbial diversity of marine shrimp pond sediment and its variability due to the effect of immobilized media in biohydrogen and biohythane production. J. Environ. Chem. Eng. 2021, 9, 106166. [Google Scholar] [CrossRef]
  45. Fischer-Romero, C.; Tindall, B.J.; Jüttner, F. Tolumonas auensis gen. nov., sp. nov., a toluene-producing bacterium from anoxic sediments of a freshwater lake. Int. J. Syst. Bacteriol. 1996, 46, 183–188. [Google Scholar] [CrossRef] [PubMed]
  46. Gill, C.O.; Youssef, M.K. MICROBIOLOGICAL SAFETY OF MEAT|Emerging Pathogens. In Encyclopedia of Meat Sciences, 2nd ed.; Dikeman, M., Devine, C., Eds.; Academic Press: Cambridge, MA, USA, 2014; pp. 340–344. [Google Scholar] [CrossRef]
  47. Kruse, S.; Goris, T.; Westermann, M.; Adrian, L.; Diekert, G. Hydrogen production by Sulfurospirillum species enables syntrophic interactions of Epsilonproteobacteria. Nat. Commun. 2018, 9, 4872. [Google Scholar] [CrossRef]
  48. Luan, J.; Xu, Y. The bio-hydrogen production by klebsiella, bacteroides and ruminococcus from wastewater which contained coconut oil, hydraulic oil or peanut oil. J. Biotechnol. 2014, 185, S124. [Google Scholar] [CrossRef]
  49. Zhou, A.; Liu, H.; Varrone, C.; Shyryn, A.; Defemur, Z.; Wang, S.; Liu, W.; Yue, X. New insight into waste activated sludge acetogenesis triggered by coupling sulfite/ferrate oxidation with sulfate reduction-mediated syntrophic consortia. Chem. Eng. J. 2020, 400, 125885. [Google Scholar] [CrossRef]
  50. Wang, J.; Yin, Y. Clostridium species for fermentative hydrogen production: An overview. Int. J. Hydrogen Energy 2021, 46, 34599–34625. [Google Scholar] [CrossRef]
  51. McLellan, S.L.; Roguet, A. The unexpected habitat in sewer pipes for the propagation of microbial communities and their imprint on urban waters. Curr. Opin. Biotechnol. 2019, 57, 34–41. [Google Scholar] [CrossRef]
  52. Roalkvam, I.; Drønen, K.; Stokke, R.; Daae, F.L.; Dahle, H.; Steen, I.H. Physiological and genomic characterization of Arcobacter anaerophilus IR-1 reveals new metabolic features in Epsilonproteobacteria. Front. Microbiol. 2015, 6, 987. [Google Scholar] [CrossRef]
  53. Hamann, E.; Gruber-Vodicka, H.; Kleiner, M.; Tegetmeyer, H.E.; Riedel, D.; Littmann, S.; Strous, M. Environmental Breviatea harbour mutualistic Arcobacter epibionts. Nature 2016, 534, 254–258. [Google Scholar] [CrossRef] [PubMed]
  54. Zhao, L.; Zhang, J.; Xu, Z.; Cai, S.; Chen, L.; Cai, T.; Ji, X.M. Bioconversion of waste activated sludge hydrolysate into polyhydroxyalkanoates using Paracoccus sp. TOH: Volatile fatty acids generation and fermentation strategy. Bioresour. Technol. 2022, 363, 127939. [Google Scholar] [CrossRef] [PubMed]
  55. Hakobyan, L.; Gabrielyan, L.; Trchounian, A. Bio-hydrogen production and the F0F1-ATPase activity of Rhodobacter sphaeroides: Effects of various heavy metal ions. Int. J. Hydrogen Energy 2012, 37, 17794–17800. [Google Scholar] [CrossRef]
  56. Hustede, E.; Steinbüchel, A.; Schlegel, H.G. Relationship between the photoproduction of hydrogen and the accumulation of PHB in non-sulphur purple bacteria. Appl. Microbiol. Biotechnol. 1993, 39, 87–93. [Google Scholar] [CrossRef]
  57. Sun, M.; Lv, Y.; Liu, Y. A New Hydrogen-Producing Strain and Its Characterization of Hydrogen Production. Appl. Biochem. Biotechnol. 2015, 177, 1676–1689. [Google Scholar] [CrossRef] [PubMed]
  58. Zagrodnik, R.; Łaniecki, M. Hydrogen production from starch by co-culture of Clostridium acetobutylicum and Rhodobacter sphaeroides in one step hybrid dark- and photofermentation in repeated fed-batch reactor. Bioresour. Technol. 2017, 224, 298–306. [Google Scholar] [CrossRef]
  59. Moreira, F.S.; Rodrigues, M.S.; Sousa, L.M.; Batista, F.R.X.; Ferreira, J.S.; Cardoso, V.L. Single-stage repeated batch cycles using co-culture of Enterobacter cloacae and purple non-sulfur bacteria for hydrogen production. Energy 2022, 239, 122465. [Google Scholar] [CrossRef]
  60. Montiel-Corona, V.; Buitrón, G. Polyhydroxyalkanoates from organic waste streams using purple non-sulfur bacteria. Bioresour. Technol. 2021, 323, 124610. [Google Scholar] [CrossRef]
Sustainability 15 16607 g001
Figure 1. Schematic diagram of the experimental setup illustrating the spatial arrangement of the experiment. The distance (d) between the lamp and the bottles set at either 15 cm or 30 cm, which alters the irradiance between 32 Wm−2 and 16 Wm−2, respectively.
Figure 1. Schematic diagram of the experimental setup illustrating the spatial arrangement of the experiment. The distance (d) between the lamp and the bottles set at either 15 cm or 30 cm, which alters the irradiance between 32 Wm−2 and 16 Wm−2, respectively.
Sustainability 15 16607 g001
Sustainability 15 16607 g002
Figure 2. Cultivation of Purple Non-Sulfur (PNS) Bacteria: (a) In Phase II—Enrichment after one week. The numbers correspond to experiments conducted in this phase, where the presence of purple biomass was noted in Experiments 1, 2, 4, and 5. No purple biomass was observed in Experiment 3. (b) In Phase II—Enrichment after 4 weeks. The numbers represent the experiments conducted in this phase (1, 2, 4 and 5).
Figure 2. Cultivation of Purple Non-Sulfur (PNS) Bacteria: (a) In Phase II—Enrichment after one week. The numbers correspond to experiments conducted in this phase, where the presence of purple biomass was noted in Experiments 1, 2, 4, and 5. No purple biomass was observed in Experiment 3. (b) In Phase II—Enrichment after 4 weeks. The numbers represent the experiments conducted in this phase (1, 2, 4 and 5).
Sustainability 15 16607 g002
Sustainability 15 16607 g003
Figure 3. Canonical correspondence analysis (CCA) triplot using type II scaling, performed with data on enrichment operating conditions and microbial community structure.
Figure 3. Canonical correspondence analysis (CCA) triplot using type II scaling, performed with data on enrichment operating conditions and microbial community structure.
Sustainability 15 16607 g003
Sustainability 15 16607 g004
Figure 4. Bacteria associated with H2 production according to canonical analysis of Enriched Purple Inocula (EPI) and PNS bacteria at species level. Illumina 16S rRNA gene analysis.
Figure 4. Bacteria associated with H2 production according to canonical analysis of Enriched Purple Inocula (EPI) and PNS bacteria at species level. Illumina 16S rRNA gene analysis.
Sustainability 15 16607 g004
Table 1. Experimental conditions used during enrichment phases applied for enrichment of a purple non-sulfur bacteria mixed culture. In Phase 1, all experimental conditions from experiments 1 to 8 were evaluated. In Phase 2, only the experimental conditions that resulted in the production of purple biomass, referred to as Purple Inoculum (PI), were assessed.
Table 1. Experimental conditions used during enrichment phases applied for enrichment of a purple non-sulfur bacteria mixed culture. In Phase 1, all experimental conditions from experiments 1 to 8 were evaluated. In Phase 2, only the experimental conditions that resulted in the production of purple biomass, referred to as Purple Inoculum (PI), were assessed.
Operational ConditionsN° Experiment
12345678
Agitation modeOn/OffOn/OffOn/OffOn/OffContinuousContinuousContinuousContinuous
Irradiance
(μmolm−2s−1)		3232161632321616
Carbon sourceMalic acidVFAsMalic acidVFAsMalic acidVFAsMalic acidVFAs
Table 2. Qualitative and quantitative results from Phase I and Phase II of enrichment.
Table 2. Qualitative and quantitative results from Phase I and Phase II of enrichment.
N° Experiment12345678
Phase I: EnrichmentPurple coloring *++−++−−−
Accumulated biogas **+−−−−−−−
% CH4 ***+−−−−−−−
% H2 ***−−−−−−−−
Phase II: EnrichmentAccumulated biogas **12.4 ± 0.5−NE−−NENENE
%H2 ***73.2 ± 2.0−NE−−NENENE
H2 yield (mmol H2 g−1 COD)0.23 ± 0.3−NE−−NENENE
Enriched Purple Inocula (EPI) name AB−CD−−−
Hydrogen production phaseH2 yield (mmol H2 g−1 COD)9.37 ± 1.0-NE--NENENE
+: Positive result and −: negative result. * Positive when purple biofilm was observed. ** Positive when biogas production was >10 mmol CH4 or H2 L−1. *** Positive when% H2 or CH4 in biogas > 0. NE: not executed.
Table 3. Characterization of the bacterial communities at family and genus level of Enriched Purple Inocula (EPI) compared to the initial inoculum of Phase I (AI). Illumina 16S rRNA gene analysis. The shown community represents over 90% of the total community (2193 OTUs), though only including sequence groups with relative abundance over 1%.
Table 3. Characterization of the bacterial communities at family and genus level of Enriched Purple Inocula (EPI) compared to the initial inoculum of Phase I (AI). Illumina 16S rRNA gene analysis. The shown community represents over 90% of the total community (2193 OTUs), though only including sequence groups with relative abundance over 1%.
FamilyGenusAI * (%)EPI (%)
ABCD
PseudomonadaceaePseudomonas 0.60.70.149.0
Other genus0.10.00.00.00.0
BradyrhizobiaceaeRhodopseudomonas 19.739.932.20.2
Other genus0.00.00.00.00.0
CampylobacteraceaeSulfurospirillum 22.21.30.00.0
Arcobacter 2.22.40.00.0
Other genus0.10.00.00.00.0
AeromonadaceaeTolumonas 20.10.70.00.0
Other genus0.00.00.00.00.0
RhodocyclaceaeRhodocyclus 6.013.02.617.5
Other genus0.21.61.60.50.4
PorphyromonadaceaeParabacteroides 1.54.811.40.0
Other genus0.20.10.21.73.6
VeillonellaceaeSporomusa 0.00.07.92.6
Other genus0.20.91.45.12.1
BacteroidaceaeBacteroides 10.311.71.00.1
Other genus10.30.00.00.00.0
MarinilabiliaceaeAlkaliflexus 0.00.00.08.4
Other genus0.20.40.80.61.5
DesulfomicrobiaceaeDesulfomicrobium 0.32.39.00.0
Other genus0.00.00.00.00.0
SphingobacteriaceaeSolitalea 0.00.06.40.0
Other genus0.10.00.30.00.0
SpirochaetaceaeTreponema 0.00.05.60.0
Other genus2.90.00.00.20.0
AcholeplasmataceaeAcholeplasma 0.15.80.20.0
Other genus0.00.00.00.00.0
DesulfovibrionaceaeDesulfovibrio 1.83.30.15.8
Other genus0.20.00.00.00.0
ClostridiaceaeGeosporobacter 0.00.02.30.1
Clostridium 4.02.21.90.7
Other genus1.80.51.80.01.6
RuminococcaceaeRuminococcus 0.00.02.10.4
Other genus0.370.00.30.90.9
GracilibacteraceaeGracilibacter 0.00.02.30.1
Other genus0.00.00.00.00.0
AcidaminococcaceaePhascolarctobacterium 0.00.02.20.1
Succinispira 1.10.00.00.0
Other genus0.00.00.60.00.0
HyphomicrobiaceaeBlastochloris 0.10.40.01.7
Other genus0.10.00.00.00.3
Others families < 2.0%84.36.44.53.62.9
(*) Only the presence of the microorganisms found in EPI is presented for initial inoculum of Phase I (AI). The remaining information is found in Supplementary Materials.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Smith, S.C.; Toledo-Alarcón, J.; Schiappacasse, M.C.; Tapia-Venegas, E. Enrichment of a Mixed Culture of Purple Non-Sulfur Bacteria for Hydrogen Production from Organic Acids. Sustainability 2023, 15, 16607. https://doi.org/10.3390/su152416607

AMA Style

Smith SC, Toledo-Alarcón J, Schiappacasse MC, Tapia-Venegas E. Enrichment of a Mixed Culture of Purple Non-Sulfur Bacteria for Hydrogen Production from Organic Acids. Sustainability. 2023; 15(24):16607. https://doi.org/10.3390/su152416607

Chicago/Turabian Style

Smith, Sean C., Javiera Toledo-Alarcón, María Cristina Schiappacasse, and Estela Tapia-Venegas. 2023. "Enrichment of a Mixed Culture of Purple Non-Sulfur Bacteria for Hydrogen Production from Organic Acids" Sustainability 15, no. 24: 16607. https://doi.org/10.3390/su152416607

APA Style

Smith, S. C., Toledo-Alarcón, J., Schiappacasse, M. C., & Tapia-Venegas, E. (2023). Enrichment of a Mixed Culture of Purple Non-Sulfur Bacteria for Hydrogen Production from Organic Acids. Sustainability, 15(24), 16607. https://doi.org/10.3390/su152416607

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop