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Article

Enhanced Growth and Productivity of Arthrospira platensis H53 in a Nature-like Alkalophilic Environment and Its Implementation in Sustainable Arthrospira Cultivation

by
Kittipat Chotchindakun
1,2,
Songphon Buddhasiri
3 and
Panwong Kuntanawat
4,*
1
Institute of Research and Development, Suranaree University of Technology, Nakhon Ratchasima 30000, Thailand
2
Biomedical Engineering and Innovation Research Center, Chiang Mai University, Chiang Mai 50200, Thailand
3
Research Center for Veterinary Biosciences and Veterinary Public Health, Faculty of Veterinary Medicine, Chiang Mai University, Chiang Mai 50100, Thailand
4
School of Biotechnology, Institute of Agricultural Technology, Suranaree University of Technology, Nakhon Ratchasima 30000, Thailand
*
Author to whom correspondence should be addressed.
Sustainability 2024, 16(19), 8627; https://doi.org/10.3390/su16198627 (registering DOI)
Submission received: 5 August 2024 / Revised: 1 October 2024 / Accepted: 2 October 2024 / Published: 5 October 2024
Browse Figures
Versions Notes

Abstract

:
Synthetic culture media, such as Zarrouk’s medium (ZM), are widely used in industrial Arthrospira cultivation but rely heavily on chemical fertilizers, raising concerns over cost and environmental impact. In natural habitats where Arthrospira blooms, the macronutrient concentrations are much lower than those provided by synthetic media. We hypothesized that natural growth may be facilitated by a microbial consortium. To test this, we developed a lab-scale Arthrospira platensis H53 cultivation system using a newly developed organic compost medium (OCM), designed to mimic the natural nutrient composition and microbial interactions. Compared to ZM, A. platensis H53 grown in OCM exhibited elevated growth by day 7. The specific growth rate in OCM was 0.20 day−1, higher than that of 0.17 day−1 in ZM, with optical density values reaching 1.57, compared to 1.13 in ZM. A 1.63-fold increase in biomass was observed in OCM, despite lower initial macronutrient concentrations. Nutrient use efficiency (NUE) in OCM was significantly improved, with nitrate (NO3−) and phosphate (PO43−) utilization up to 5.8-fold higher. Additionally, A. platensis H53 filaments in OCM were more tightly coiled, indicating a physiological change in response to lowered macronutrient concentrations. Microbial composition analysis using 16S rRNA gene amplicon sequencing revealed the presence of growth-promoting bacteria, including Pontibacter spp., Brevundimonas spp., and Aliihoeflea spp., likely contributing to nutrient cycling and enhanced growth. These findings suggest potential symbiotic interactions between cyanobacteria and non-cyanobacteria in the OCM system, promoting increased growth and productivity. This study is the first to propose such symbiosis in an extremely alkalophilic environment, offering another sustainable alternative to traditional chemical-based Arthrospira cultivation methods.

1. Introduction

Arthrospira is a cyanobacterium recognized as food for the future, due to its substantial nutritional composition and bioactive compounds. Arthrospira has been part of the human diet since at least the 13th century [1], when the Aztecs harvested it from natural cyanobacterial mats in Lake Texcoco, Mexico [2]. Similarly, the Kanembu tribe in Central Africa has used dried Arthrospira from Lake Chad as food for many generations [3]. The widespread global presence of Arthrospira, from America through Africa to Asia, is attributed to its ability to adapt to habitats unfavorable for most microalgae and other microorganisms, such as alkaline lakes, brackish waters, and saline environments [4]. This remarkable adaptation has facilitated Arthrospira mass production and utilization for human consumption, biomass feedstocks, and high-value ingredients [5].
Since Arthrospira has been commercially cultured in industrial production, its culture media have been extensively developed, often based on synthetic culture media which are convenient for preparation and scaling up. Zarrouk’s medium (ZM), the first synthetic medium formulated for Arthrospira cultivation [6], has been widely used for its ability to support optimal growth and biomass production [7]. However, the mass production of Arthrospira on a large scale depends heavily on the substantial input of synthetic chemicals, resulting in high production costs [8,9]. The high macronutrient concentrations in ZM, particularly nitrate (NO3−) and phosphate (PO43−), are the main factors limiting the maximum potential profit for industrial production. Additionally, the manufacturing of chemical fertilizers consumes significant energy and emits greenhouse gases, raising environmental concerns and impacting long-term sustainability [10,11]. These fertilizers are often produced in geopolitically sensitive regions [12], creating supply chain vulnerabilities and potential price volatility for industries reliant on their import. Moreover, crops cultivated with chemical inputs are often of lower nutritional quality compared to their naturally harvested counterparts [13].
Numerous strategies have been investigated to reduce chemical usage and costs. These include lowering the concentration of ZM [14], altering the nitrogen sources, and optimizing their quantity [14,15,16]. Other approaches have explored the use of complex fertilizers such as NPK, which is cheaper [17], and locally sourced or natural substitutes. However, these substitutes often still require chemical fertilizer supplementation to achieve adequate growth [18,19]. In addition, some of the natural substitutes have a limited use, confined to only certain regions where they are locally found. Wastewaters have also been proposed as potential alternatives [16,20]. Nevertheless, few wastewater sources are suitable for Arthrospira cultivation intended for human consumption. Their use is restricted by fluctuations in quality, potential contamination risks, and the need for pre-treatment, which may still require the use of additional synthetic chemicals [16]. These challenges highlight the need for sustainable alternatives that minimize chemical dependency while maintaining productivity.
We believe that one of the best ways to tackle this problem, when sustainability is the main issue, is to learn from nature. In Arthrospira’s natural habitat, macronutrient concentrations can be up to a thousand times lower than those in synthetic media used in the industry. Arthrospira blooms have been reported globally, including in Central America, Africa, Asia, and Europe. The macronutrient concentrations in these blooms range from 0.55 to 16.1 mg L−1 for NO3− and from 0.01 to 0.88 mg L−1 for PO43− [4,21,22,23,24,25,26], which are lower compared to the standard ZM medium levels of 1822.5 mg L−1 and 272.5 mg L−1, respectively [6]. This observation suggests that successful Arthrospira culture may not require a high macronutrient supply. However, this observation has not yet been systematically verified in the laboratory setting, nor it has been explained how the less-chemically supplied system can sustain growth and what factors are involved.
Naturally, cyanobacteria coexist with other microorganisms, particularly bacteria, forming a niche microenvironment called the cyanosphere. This microbial consortium demonstrates unique interactions ranging from mutualism to parasitism. It has been reported that cyanobacteria–bacteria interactions facilitate the circulation of macronutrients such as nitrogen, phosphate, and carbon, as well as of micronutrients like vitamin B12 and plant growth-promoting factors secreted by certain microorganisms, significantly promoting cyanobacterial growth [27]. These microbial relationships are typically dependent on specific cyanobacterial species and their habitats. There are no reports of such interactions in the case of Arthrospira thriving in a highly alkaline environment. We hypothesize that microbial interactions may be the key factors assisting cyanobacterial growth in low-nutrient natural environments.
Traditionally, Arthrospira cultivation has relied on monoculture cultivation, viewing existing bacteria as contaminants. However, in recent decades, the synergistic effects between cyanobacteria and bacteria have been found to benefit cyanobacterial and algal physiology and biomass production [28]. Understanding the natural system where Arthrospira growth is associated with microbial symbionts may pave the way for more sustainable cyanobacteria cultivation using fewer chemicals.
The present study aimed to develop a lab-scale growth system for Arthrospira that mimics the natural physicochemical and microbial profile of Arthrospira’s habitat using Arthrospira platensis H53 as a model organism. The system was used to test previous observations that Arthrospira blooms in much lower macronutrient concentrations than those commonly used. In comparison to the synthetic medium, we investigated the evolution of A. platensis H53 population growth, filament morphology, and nutrient utilization in the nature-like system using a newly invented organic compost medium (OCM) derived from agricultural leftovers. To systematically explore and observe the possible factors that may facilitate cyanobacterial growth in the systems, the physicochemical and microbial community profiles of both systems were tracked over the cultivation period.

2. Materials and Methods

2.1. Microorganisms and Cultivation Systems

The Arthrospira platensis strain H53, hereafter referred to as A. platensis H53, was obtained from the Algae and Natural Product Technology Laboratory (Suranaree University of Technology, Nakhon Ratchasima, Thailand). This strain was isolated from a water sample collected from a treated water pool at a cassava processing facility (14°34′46.9″ N; 102°11′59.1″ E), Nakhon Ratchasima, Thailand. This pool is an outdoor, man-made lake surrounded by abundant greenery. The water is directly exposed to sunlight and has high alkalinity. Arthrospira was the dominant phytoplankton genus in the lake. The average temperature of the region is 27 °C. This monoclonal culture, derived from a single ancestral filament, was not axenic as it contained coexisting bacteria, though these did not significantly affect Arthrospira cultivation in an industrial context. All procedures involving live A. platensis H53 were conducted aseptically. The OCM was prepared using AgentWild™, a processed organic compost containing mixed soil, organic matters, and a natural pH enhancer, purchased from Siambiota Co., Ltd. (Nakhon Ratchasima, Thailand). To simulate Arthrospira’s aquatic habitat where human contact is minimal, AgentWild™ was suspended in deionized water, agitated at 25 °C using a magnetic stirrer, and allowed to settle for 24 h to maximize nutrient extraction. The compost amount and incubation time, leading to macronutrient saturation as measured by electroconductivity, were determined in our separate study. The clear supernatant, filtered through a cellophane membrane, was stored at 4 °C until use. An A. platensis H53 inoculum was prepared using sterile ZM [6]. The inoculum was transferred to either sterile ZM or non-sterile OCM systems. Each system consisted of a 500 mL Erlenmeyer flask, plugged with cotton wool, containing 150 mL of the respective medium (n = 3). The final medium compositions and physicochemical characteristics are detailed in Table 1. The initial optical density at 560 nm (OD560) was approximately 0.1, and the pH was adjusted to between 9.20 and 9.25 for all experiments. The systems were maintained on a shaking platform at 150 rpm, with continuous fluorescent light (60 µmol photons m−2 s−1) at 25 °C for 14 days. The macronutrient concentrations of NO3− and PO43− were measured using the nitrate TNT-plus Vial and PhosVer 3 Test kits (HACH, Loveland, CO, USA). All chemicals used were of reagent grade (Kemaus, Cherrybrook, Australia).

2.2. Growth Monitoring and Biomass Production Analysis

The growth patterns of both systems were assessed by measuring the optical density at 560 nm [29], targeting the phycocyanin peak specific to Arthrospira. This wavelength was chosen to minimize interference from non-cyanobacterial materials, such as bacteria and particulate residues. Each day, 1 mL of culture suspension was sampled. Then, 1 mL of deionized water was added to the system to maintain the initial volume. The optical density was measured using an Ultrospec 2000 UV/VIS spectrophotometer (GE Healthcare, Chicago, IL, USA). The pH of the same culture suspension was then measured with a pH meter (Thermo Scientific, Waltham, MA, USA). Dry biomass yield was quantified through a gravimetric analysis method [30]. Briefly, 20 mL of the culture suspension was filtered using pre-weighed GF/C Whatman filter paper (Whatman, Maidstone, UK). The cells were then rinsed with deionized water and dried in a 60 °C oven until a constant weight was achieved. Sterile and non-sterile OCM without inoculation were used as controls. The specific growth rate (µ) was calculated using a previously described equation [31]. Dry biomass yield was expressed in mg L−1.

2.3. Calculation of Nutrient Use (NU) and Nutrient Use Efficiency (NUE)

The characteristics of A. platensis H53’s macronutrient utilization in both ZM and OCM systems were expressed as nutrient use (NU) and nutrient use efficiency (NUE). NUE refers to the dry biomass produced per unit of a given macronutrient used. NU and NUE were calculated by adapting a previously described method [32] as follows:
N U = N u t r i e n t ( t 1 ) − N u t r i e n t ( t 2 )
N U E = B i o m a s s N U
where Nutrient(t1) and Nutrient(t2) represent the concentrations (mg L−1) of NO3− or PO43− in the system before and after A. platensis H53 cultivation, respectively, and Biomass represents the dry biomass yield (mg L−1).

2.4. Microscopic Observations

The morphological features of A. platensis H53 filaments, including length, thickness, helix diameter, and helix pitch, quantified as shown in Figure 1, were investigated at three different intervals (Day 0, Day 5, and Day 10). A total of hundred filaments (n = 100) from each system were examined using a Leica Microsystems 10,447,197 EZ4 Stereo Microscope (Leica, Nanterre, France).

2.5. DNA Extraction and Amplicon Sequencing

Bacterial genomic DNA was extracted from both groups of A. platensis H53 cultures using the QIAamp PowerFecal Pro DNA Kit (Qiagen, Redwood, CA, USA). The V4-V5 region of the bacterial 16S rRNA genes was amplified by PCR using the 515F (5′−GTGCCAGCMGCCGCGGTAA−3′) and 907R (5′−CCGTCAATTCCTTTGAGTTT−3′) primers [33]. DNA library preparation and sequencing were performed by NovogeneAIT Genomics Singapore Pte. The samples were subjected to 250 bp paired-end sequencing using the NovaSeq 6000 Sequencer (Illumina, Inc., San Diego, CA, USA).

2.6. Microbiota Analysis

The sequencing data were processed using the Quantitative Insights into Microbial Ecology 2 (QIIME2, version 2022.8) pipeline [34]. Paired-end sequences were denoised and merged using the DADA2 plugin, and amplicon sequence variants (ASVs) were aligned with the mafft plugin. Taxonomic assignment of the 16S rRNA sequences was performed using the Silva 138.99% taxonomy classifier [35]. Alpha diversity was assessed by calculating observed ASVs, Shannon, and Simpson’s indices. Differential abundance analysis was conducted using linear discriminant analysis (LDA) effect size (LEfSe) to identify significant bacterial taxa among groups [36]. The relative abundances of the taxa were compared. The microbiota results were analyzed using the Kruskal–Wallis test and visualized with the microeco R package version 1.9.1 [37].

2.7. Statistical Analysis

All experimental data are reported as the mean ± standard deviation of at least three independent experimental replicates. Statistical differences between groups were analyzed using Student’s t-test with IBM SPSS Statistics (Version 21.0, SPSS Inc., Chicago, IL, USA). Statistical significance was defined as p < 0.05.

3. Results and Discussion

3.1. Growth Profile and pH Determination

The growth profiles of A. platensis H53 cultured in the ZM and OCM systems over a 14-day period are presented in Figure 2. Initially, both systems exhibited similar growth patterns. However, by day 7, the OCM system showed a significantly higher OD560, reaching a peak of 1.566 ± 0.087 on day 14, which was a 1.38-fold increase compared to the growth measured in the ZM system (1.134 ± 0.032) (p < 0.05) (Figure 2a). This increase correlated with a 14.5% higher maximum specific growth rate (µmax·day−1) in the OCM system compared to the ZM system (Table S1). The pH trends in both systems were consistent with the growth patterns. Statistical analysis revealed a significant pH increase in the OCM system starting from day 7, with a final pH of 11.55 ± 0.22 and a rise rate of 0.2 unit·day−1 (p < 0.01). In contrast, the ZM system showed a more gradual pH increase, ending at 10.37 ± 0.03 (Figure 2a). While the OCM system showed higher final OD560 and pH than the ZM system, the pattern of pH elevation during the cultivation of A. platensis was consistent with that of previous studies using synthetic media and agro-industrial wastewater media [38,39].
The pH increase correlates with A. platensis H53 growth driven by photosynthesis, where CO2 is converted into organic compounds. In our present study, sodium bicarbonate (NaHCO3) was used as the carbon source in both systems. Dissolved HCO3− can be readily converted to carbonic acid (H2CO3) and hydroxide ions (OH−), with subsequent dissociation of H2CO3 producing free CO2 utilized by cyanobacteria. The remaining OH− increases the pH [20]. It should be noted that A. platensis cultivation is generally performed in a pH range from 9.5 to 11.0, with an optimal pH of around 10.5 for effective growth and pathogen development prevention [40]. A pH above 11.0 could disrupt the H2CO3 dissociation equilibrium, converting dissolved carbon into unavailable CO32−, which limits free CO2 availability and impairs photosynthesis [41]. Despite the same NaHCO3 concentration in both systems, the OCM system achieved a significantly higher pH above the optimal range (≈11.0) by the final stage, while the ZM system’s pH plateaued. This suggests that A. platensis H53 growth in the OCM system, though possibly slowing, remained photosynthetically active unlike in the ZM system. The question arises as to how A. platensis H53 continued to grow and photosynthesize effectively in the unfavorable pH environment.
The OCM system, rich in organic matter, likely supported heterotrophic bacteria from either the A. platensis H53 inoculum or the non-sterile compost. This may have established a cyanobacteria–bacteria consortium, called cyanosphere, where each group took part in the mutualistic interaction [42]. We hypothesize that the coexisting microorganisms in the OCM system continually produced an amount of CO2 that sufficiently supported the cyanobacterial CO2 uptake rate but did not lower the overall pH significantly. However, since the cyanobacteria and bacteria were in proximity, the produced CO2 was localized in the microenvironment and was readily available for cyanobacterial assimilation. In addition, this could drop the pH in the microenvironment and shifted the local H2CO3-HCO3−-CO32− equilibrium, turning more CO32− into H2CO3 and CO2, respectively. This localized ‘CO2 unlocking’ likely took place transiently, since CO2 tends to be dissolved and diffuse with ease in a highly alkaline macroenvironment. The repeated CO2 unlocking by bacteria might explain the sustained utilization of CO2 and the ongoing pH rise in the OCM system.
To validate this hypothesis, the presence and role of the symbiotic microbial consortium need further investigation. Previous literature suggests that cyanobacteria–bacteria mutualistic interactions exist, where cyanobacteria provide dissolved organic carbon (DOC) through released organic matter or dead cells [43]. This DOC benefits heterotrophic bacteria, which in turn supply CO2 for cyanobacterial photosynthesis. Additionally, heterotrophic microorganisms can produce essential growth compounds, including fixed nitrogen and plant growth hormones [27,44,45], which potentially explains the enhanced growth in the OCM system. Future studies should focus on quantifying total free CO2 and HCO3− and verifying microbial interactions to clarify these mechanisms.

3.2. Cellular Morphological Changes in A. platensis H53

The characteristics of A. platensis H53 filaments, including length, thickness, helix diameter, and helix pitch, were quantified as shown in Figure 2. The initial inoculation was transferred from the standard ZM cultivation to the ZM and OCM systems and exhibited a similar filament morphology Figure 3a,d. Figure 3b,e illustrate the morphological changes in A. platensis H53 filaments in both systems after a 5-day cultivation period. After 14 days, significant overall-filament transformations were observed in the OCM system (Figure 3c), with decreased filament length and helix pitch and increased thickness and helix diameter (p < 0.05) compared to day 0 (Table 2). In contrast, no filament transformations were found in the ZM system (Figure 3f).
Arthrospira’s helical structure facilitates its spiral movement through fluid environments [46]. The filament shape can be altered in response to specific stimuli and environmental factors, including physical and chemical factors. For example, the coils may become tightened or loosened to self-shade or to be more exposed to light, depending on light intensity [47]. Additionally, several factors including light spectrum [48], temperature [49,50], and salinity [51,52] can change the spiral characteristics.
In our experiment, the chemical composition of the culturing systems was one of the independent variables. The transformed helical characteristics of A. platensis H53 in the OCM system might be due to the macronutrient concentrations. The less concentrated NO3− formula, shown in Table 1, might be responsible for the lowered average A. platensis H53 filament pitch in the OCM system. It was previously reported that Arthrospira’s helical tightening (lowered helical pitch) is associated with a decrease in NO3− concentration, and helical loosening is observed when NO3− concentration is elevated [53].
Biological compounds, particularly phytohormones, have been found to facilitate the morphological adaptation and physiological functions of cyanobacteria, including Arthrospira [54,55,56]. Principal growth phytohormones, such as salicylic acid (SA) and 1-aminocyclopropane-1-carboxylic acid (ACC), are recognized for their ability to induce helix tightening [57]. On the other hand, helix loosening has been linked to the presence of indole-3-acetic acid (IAA) and trans-zeatin (tZ), a cytokinin [58]. Although the present study did not incorporate these phytohormones into the systems, it is possible they were internally supplied by specific microorganisms, especially actinobacteria [59,60,61], if they coexisted in the culturing system. OCM was prepared using organic compost as the main component, whose dissolved compounds may facilitate the establishment of a microbial consortium comprising plant growth-promoting bacteria. An in-depth study of the in situ microbial composition was conducted to elucidate this hypothesis and is reported in the following section. The transformation of Arthrospira morphology suggests physiological adaptation of the cells. It can be concluded that the chemical composition of the OCM affects A. platensis H53′s cellular physiology, likely as an adaptation to changes in macronutrient concentrations. The microbial consortium may play a part in this process and requires further investigation to confirm its presence and potential functions, including 16S rRNA amplicon sequencing, as discussed in the following section.

3.3. Dry Biomass Yield, Nutrient Use, and Nutrient Use Efficiency

The dry biomass yield, nutrient use (NU), and nutrient use efficiency (NUE) of A. platensis H53 cultivated in the ZM and OCM systems are presented in Table 3. The OCM system exhibited a dry biomass yield of 711.73 ± 61.46 mg L−1, which was 1.63-fold greater than that of the ZM system (p < 0.01). Another separate experiment on incubating OCM without A. platensis H53 inoculation (see in Supplementary Materials) demonstrated that the bacteria could not be harvested in a significant quantity using our methodology. This implies that the measured dry biomass in our study solely belonged to A. platensis H53. This superior performance in the OCM system was also reflected in a significantly higher biomass productivity (mg L−1 day−1) (p < 0.01) (Table S2). However, the NUs of both NO3− and PO43− were much lower in the OCM system, with reductions of 3.58-fold and 1.70-fold, respectively, compared to the ZM system (p < 0.01) (Table 3). The higher NUEs of both NO3− and PO43− in the OCM system (Table 3) suggest a more efficient conversion of over 80% of nutrients into the system, with a portion contributing to A. platensis H53 biomass. These values are significantly higher than those reported in the literature, e.g., by Markou et al. [62], who observed up to 50% NO3− conversion in mixotrophic cultures of A. platensis using olive mill wastewater pre-treated with sodium hypochlorite. Similarly, Papadopoulos et al. [63] reported a maximum of 28.3% PO43− conversion in A. platensis cultures grown in a brewery wastewater-based medium supplemented with NaHCO3 and NaCl.
This finding indicates that greater biomass production and efficient nutrient utilization can be achieved in an A. platensis H53 cultivation system with less concentrated nutrients, with concentrations up to 90% lower than the typical concentrations. This also explains the occurrence of Arthrospira blooms in less concentrated natural environments. Previous studies similarly found a positive correlation between NUE and nutrient dilution rates, particularly for NO3− [32], and demonstrated enhanced Arthrospira biomass production in less concentrated ZM medium [14,64,65].
However, it should be noted that the initial macronutrient concentrations (NO3− and PO43−) in the OCM were 228.15 and 85.00 mg L−1, respectively (Table 1). These values are much lower than the final concentrations of NO3− and PO43− in the ZM system when Arthrospira growth began to decelerate (approximately 1301.98 mg L−1 and 119.07 mg L−1, respectively, based on Table 1 and Table 3). When other environmental factors such as pH, light intensity, and temperature are optimal, the macronutrient concentrations become the key factor influencing the growth deceleration phase in microbial cultures, including Arthrospira cultures [66]. Thus, the initial concentration of OCM appeared too low to support a highly productive A. platensis H53 culture.
The OCM system contains organic matter that can be converted into cyanobacterial macronutrients by specific groups of heterotrophic bacteria. We hypothesize that, in addition to the macronutrients supplied externally at the start of the cultivation, bacterial activity may generate an internal macronutrient supply that helps sustain cyanobacterial growth and productivity. It is important to note that the reported NUs were calculated based on the initial external macronutrient concentrations and the apparent final concentrations. The actual amount of nutrients used in the OCM system might be higher than that displayed in Table 3, as extra macronutrients might be produced and utilized within the system during cultivation. Therefore, the NUs and NUEs values in Table 3 should be regarded as those of the externally supplied nutrients. Further microbial consortium analysis in the OCM compared to the ZM system would help support this hypothesis.

3.4. Microbial Diversity between ZM and OCM Systems

A total of 1,120,963 DNA sequences were denoised into 966 amplicon sequence variants (ASVs). The alpha diversity between the standard ZM and OCM systems is depicted in Figure 4a–c. The OCM system exhibited the highest number of unique ASVs, corresponding to 664, while the ZM system showed 121 ASVs. Only 181 ASVs were shared between the two groups, indicating that 302 ASVs, along with those unique to ZM and shared between two groups, originated from the Arthrospira stock culture. A total of 966 ASVs were obtained from both systems, spanning various taxonomic levels including phyla, orders, families, and genera (Figure 4a). A statistical analysis revealed a substantial difference in microbial diversity between the two systems. The OCM system exhibited higher microbial diversity than the ZM system, with significant differences in microbiota richness and evenness (observed ASVs p < 0.05, Shannon p < 0.05, evenness p < 0.05). The OCM system’s natural organic matter content likely contributed to the presence of unique microorganisms absent in the ZM system. Our findings indicate a different abundance ratio of shared taxa between the two groups, prompting further investigation into the prokaryotic composition of both systems to understand the influence of OCM on the microbial profile’s evolution.

3.5. Composition of the Microbial Community in the ZM and OCM Systems

The prokaryotic community composition in A. platensis H53 cultures for both ZM and OCM systems is presented in Figure 5a–c. The ten dominant ASVs are shown in terms of phylum composition (Figure 5a). Cyanobacteria was the dominant phylum in both groups, with a relative abundance ranging from 41.1% to 68.0%, followed by Pseudomonadota (17.1% ± 4.2) and Bacteroidota (4.3% ± 2.7). However, significant differences in microbial composition were observed at the phylum level (p < 0.01). The phylum Actinomycetota particularly stood out in the OCM system (12.1% ± 3.3), including the families Micrococcaceae, Nocardioidaceae, and Nitriliruptoraceae, with a relative abundance that was 12.8-fold higher than in the ZM system (0.9% ± 0.2). Conversely, the phylum Verrucomicrobiota was more prevalent in the ZM system (16.9% ± 2.1), consisting solely of the Kiritimatiellae family, with an almost 3-fold higher abundance than in the OCM system (5.7% ± 0.6).
Further identification at the genus level, as shown in the microbial heat map, (Figure 5b) revealed that the cyanobacterial Arthrospira sp. PCC7345 was a major component in both systems. Additionally, the unculturable haloalkaliphilic bacterium WCHB1-41 and the halo-tolerant Salinarimonas spp. (4.2% ± 2.4) and alkali-tolerant Roseibaca spp. (3.8% ± 1.5) were common genera. These findings are in agreement with the high pH (10.4–11.5) and salinity (18–22 parts per thousand; ppt) of both systems. However, the OCM system showed different relative abundances of some bacterial genera compared to the ZM system, indicating a unique microbial community and specific relationships.
To clarify the microbial characteristics of each system, biomarker discovery using linear discriminant analysis (LEfSe) was used to identify unique microbial taxa with LDA scores ≥ 3.5 (Figure 5c). Twenty-four core bacterial taxa were found in the ZM and OCM systems (p < 0.05). WCHB1-41 (family Kiritimatiellae) and Salinarimonas spp. (family Beijerinckiaceae) were biomarkers of the ZM system. The OCM system, however, had a higher ratio of nutrient-cycling and plant growth-promoting microorganisms. In particular, the unclassified Micrococcaceae (phylum Actinomycetota) were identified as a biomarker. Micrococcaceae are widely distributed in various environments, including soil and freshwater [67], and play important roles in nutrient circulation [68]. The high relative abundance of Micrococcaceae in the OCM system coincided with lower nutrient concentrations, especially of NO3− and PO43−. Previous studies reported that nitrogen and phosphorus limitations can promote Micrococcaceae dominance in terms of community abundance [69]. Although most Micrococcaceae were unclassified, the classified genera (≤1%) included Paenarthrobacter, Arthrobacter, and Nesterenkonia, known for their effects on microbial nutrient metabolism and plant growth promotion [69,70,71,72]. Moreover, the diazotrophic bacterium Pontibacter (belonging to the family Hymenobacteraceae), the phosphate-solubilizing bacterium Brevundimonas (belonging to the family Caulobacteraceae), and the arsenic-degrading bacterium Aliihoeflea (belonging to the family Rhizobiaceae) were also identified as biomarkers in the OCM system. They were found to enhance nutrient utilization and reduce toxic contamination in previous studies [73,74,75].
Although two unique, most prevalent microbes were identified for the OCM and ZM systems, the most common microbes in each system were also present in the other, but in minor proportions. This implies that these consortia coexisted in the A. platensis H53 stock culture, rather than solely originating from the cultivation media. Our preliminary study demonstrated that conditions provided by the natural habitat-like OCM system can leverage the abundance of indigenous microorganisms in A. platensis H53 cultivation. However, while our findings confirmed their presence and potential functions, their exact roles remain unverified. The metanalytic results support the hypothesis that bacteria could assist in A. platensis H53 growth and productivity through mechanisms like temporary pH regulation, carbon dioxide unlocking, production of plant growth promoters, and internal macronutrient supply (Figure 6). Further systematic studies on this matter to clarify the roles of these microbes and their mechanism in assisting Arthrospira’s growth are required, and we will perform them in the future.
In conclusion, the OCM system, representing Arthrospira’s natural habitat, can improve biomass production using very low chemical concentrations. Despite using compost materials and bacteria, the system does not pose microbiological hazards. Food-borne pathogens [76], such as Bacillus cereus, Escherichia coli, Staphylococcus aureus, Clostridium spp., Salmonella spp., and Vibrio spp., were not detected. The high alkalinity and salinity in the OCM system naturally prevent pathogen growth, ensuring that Arthrospira biomass can be safely used for food production. This system has the potential to be expanded for producing high-value bioactive compounds, such as phycocyanin, PUFAs, and carotenoids [77,78], or even third-generation feedstock [79,80] in a more sustainable manner.

4. Conclusions

We successfully reconstructed the aquatic conditions in which natural Arthrospira blooms may occur. Our system enabled more systematic observation and exploration of the key factors influencing cyanobacterial growth in nature. Using this system, we observed higher A. platensis H53 growth and productivity at low nutrient concentrations, with high nutrient use (NU) and nutrient use efficiency (NUE), calculated from the initial exogenous macronutrient levels and apparent final concentrations, compared to those obtained in the ZM. We propose that bacterial-assisted Arthrospira growth mechanisms, such as transient pH balancing, carbon supply, internal macronutrient supply, and the presence of growth-promoting molecules, are linked to the existence of a beneficial microbial consortium in the highly alkaline OCM system. A microbial consortium analysis revealed that the OCM system is highly biodiverse and contains significant populations of beneficial bacteria, including Micrococcaceae, Pontibacter spp., Brevundimonas spp., and Aliihoeflea spp., which are known to contribute to nutrient cycling and the production of plant growth promoters. These beneficial bacteria are also present in the ZM system but not as the dominant microbial residents, suggesting that the OCM conditions favor their growth. This study demonstrates that sustainable Arthrospira production with lower chemical input is achievable by mimicking the natural conditions. However, the system’s current limitations should be considered. This study was conducted with A. platensis H53, and further research on other strains is necessary to confirm the broader applicability of the system. Additionally, the preparation of OCM relies on compost for organic matter, making consistent sourcing and processing of materials essential for maintaining chemical and biological consistency in OCM quality. To apply this method at a larger scale, industrial-scale preparation of OCM must be developed. Scaling up processes like mixing, medium incubation, and filtration will be crucial for optimizing the efficiency of OCM production. Future studies will focus on further elucidating the roles of the microbial consortia and improving scalability for less chemical-dependent Arthrospira cultivation.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/su16198627/s1, Table S1: The raw data of specific growth rate of A. platensis H53 cultured in the ZM and OCM systems; Table S2: The raw data of dry biomass yield and biomass productivity of A. platensis H53 cultured in the ZM and OCM systems at day 14; Table S3: The raw data of OD560 of A. platensis H53 cultured in the ZM and OCM systems; Table S4: The raw data of pH of A. platensis H53 cultured in the ZM and OCM systems; Table S5: The raw data of the morphology of A. platensis H53 filaments in the ZM and OCM system, including length, thickness, helix diameter, and helix pitch; Table S6: The raw data of nutrient use and nutrient use efficiency of A. platensis H53 cultured in the ZM and OCM systems; Table S7: The raw data of OD560 in control experiments using non-sterile and sterile OCM without A. platensis H53 inoculation; Table S8: The raw data of weight in the control experiments with non-sterile and sterile OCM without A. platensis H53 inoculation.

Author Contributions

Conceptualization, P.K.; methodology, K.C.; software, K.C. and S.B.; validation, K.C.; formal analysis, K.C., S.B. and P.K.; investigation, K.C.; resources, S.B. and P.K.; data curation, K.C.; writing—original draft preparation, K.C.; writing—review and editing, P.K.; visualization, K.C. and S.B.; supervision, P.K.; project administration, P.K.; funding acquisition, P.K. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Science, Research and Innovation Fund 179312.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Further information supporting the conclusions of this article will be made available by the authors on request. The microbiome data of this study are available in the GenBank database with BioProject ID: PRJNA1140430, https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1140430, accessed on 4 August 2024.

Acknowledgments

This work was supported by the following funding entities: (i) Suranaree University of Technology (SUT), (ii) Thailand Science Research and Innovation (TSRI), and (iii) National Science, Research and Innovation Fund (NSRF) (project code 179312). We are thankful to Thanathorn Thongtan for the substantial help provided and to Thanyalak Pimsalee for coordinating with Siambiota Co., Ltd. All authors reviewed the manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. The morphological features of A. platensis H53 filament include its length, thickness, helix diameter, and helix pitch.
Figure 1. The morphological features of A. platensis H53 filament include its length, thickness, helix diameter, and helix pitch.
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Figure 2. Growth profile of A. platensis H53 in ZM and OCM systems over a 14-day period, showing (a) optical density at 560 nm (OD560) and (b) pH levels.
Figure 2. Growth profile of A. platensis H53 in ZM and OCM systems over a 14-day period, showing (a) optical density at 560 nm (OD560) and (b) pH levels.
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Figure 3. The morphological transformation of A. platensis H53 cultured in the ZM and OCM systems, observed over a period of 10 days.
Figure 3. The morphological transformation of A. platensis H53 cultured in the ZM and OCM systems, observed over a period of 10 days.
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Figure 4. Microbial diversity between ZM and OCM systems was calculated to assess alpha diversity by (a) observed ASVs (b) Shannon’s indices, and (c) Simpson’s indices. * indicate p-value < 0.05.
Figure 4. Microbial diversity between ZM and OCM systems was calculated to assess alpha diversity by (a) observed ASVs (b) Shannon’s indices, and (c) Simpson’s indices. * indicate p-value < 0.05.
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Figure 5. Microbiome analysis in A. platensis H53 cultures in ZM and OCM systems. (a) Histogram of relative abundance (%) of the 10 most abundant bacteria phyla; (b) the heatmaps show the top 20 microorganisms based on their relative abundance at the genus level for the microbial metagenomes of the ZM and OCM systems; (c) LefSe analysis: taxa with significant differences that had an LDA score larger than the threshold value of 3.5; letters in front of the taxa represent the taxonomic level (p = phylum, c = class, o = order, f = family, g = genus).
Figure 5. Microbiome analysis in A. platensis H53 cultures in ZM and OCM systems. (a) Histogram of relative abundance (%) of the 10 most abundant bacteria phyla; (b) the heatmaps show the top 20 microorganisms based on their relative abundance at the genus level for the microbial metagenomes of the ZM and OCM systems; (c) LefSe analysis: taxa with significant differences that had an LDA score larger than the threshold value of 3.5; letters in front of the taxa represent the taxonomic level (p = phylum, c = class, o = order, f = family, g = genus).
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Figure 6. Schematic overview of the proposed cyanobacteria–bacteria mutualistic interactions in the OCM system stimulating A. platensis H53 growth through macronutrient conversion and CO2 unlocking.
Figure 6. Schematic overview of the proposed cyanobacteria–bacteria mutualistic interactions in the OCM system stimulating A. platensis H53 growth through macronutrient conversion and CO2 unlocking.
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Table 1. The medium composition and physicochemical characteristics of the ZM and OCM systems used in the experiments.
Table 1. The medium composition and physicochemical characteristics of the ZM and OCM systems used in the experiments.
ZM System
Medium CompositionAmount (g L−1)Physiochemical CharacteristicsValue
NaHCO3 16.8NO3− (mg L−1)1989.07
NaNO32.5PO43− (mg L−1)291.00
NaCl1.0pH9.26
K2SO41.0
K2HPO40.5
MgSO4·7H2O0.2
FeSO4·7H2O0.01
CaCl2·2H2O0.04
a C10H14N2Na2O8·2H2O0.08
b trace element mixture A51 mL
c trace element mixture B61 mL
OCM System
Medium CompositionAmount (g L−1)Physiochemical CharacteristicsValue
AgentWild™350.0NO3− (mg L−1)228.15
NaHCO316.8PO43− (mg L−1)85.00
pH9.24
a ethylenediaminetetraacetic acid disodium salt dihydrate (EDTA disodium salt dihydrate); b trace element mixture A5 (mg L−1): H3BO3, 2.86; MnCl2·4H2O, 1.81; ZnSO4·4H2O, 0.222; Na2MoO4, 0.0177; CuSO4·5H2O, 0.079; c trace element mixture B6 (mg L−1): NH4VO3, 22.9; NiSO4·7H2O, 47.8; NaWO2, 17.9; Ti2(SO4)3·6H2O and Co(NO3)2·6H2O, 4.4.
Table 2. The morphology of A. platensis H53 filaments cultured in the ZM and OCM systems, including length, thickness, helix diameter, and helix pitch.
Table 2. The morphology of A. platensis H53 filaments cultured in the ZM and OCM systems, including length, thickness, helix diameter, and helix pitch.
Morphological FeaturesDay 0Day 5Day 10
Length (µm)
ZM618.7 ± 60.7607.8 ± 77.6617.3 ± 74.8
OCM616.3 ± 57.3529.6 ± 74.0352.3 ± 28.4
p-Valuens******
Thickness (µm)
ZM7.4 ± 1.18.3 ± 0.77.6 ± 1.1
OCM7.5 ± 1.110.4 ± 1.411.6 ± 1.6
p-Valuens******
Helix diameter (µm)
ZM29.8 ± 3.230.2 ± 0.931.2 ± 3.2
OCM30.5 ± 3.137.4 ± 5.442.7 ± 5.7
p-Valuens******
Helix pitch (µm)
ZM77.2 ±5.782.2 ± 1.981.1 ± 4.9
OCM78.5 ± 8.971.4 ± 8.164.9 ± 7.4
p-Valuens******
*** Significant morphological difference for A. platensis H53 filaments at p < 0.001; ns = not significant (p > 0.05).
Table 3. Dry biomass yield, nutrient use, and nutrient use efficiency of A. platensis H53 in ZM and OCM systems.
Table 3. Dry biomass yield, nutrient use, and nutrient use efficiency of A. platensis H53 in ZM and OCM systems.
SystemsDry Biomass Yield (mg L−1)Nutrient Use (mg L−1)Nutrient Use Efficiency
NO3−PO43−NO3−PO43−
ZM436.67 ± 4.99687.09 ± 62.62119.70 ± 3.200.64 ± 0.063.65 ± 0.09
OCM713.33 ± 61.46191.91 ± 2.1870.60 ± 2.413.72 ± 0.3110.09 ± 0.56
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Chotchindakun, K.; Buddhasiri, S.; Kuntanawat, P. Enhanced Growth and Productivity of Arthrospira platensis H53 in a Nature-like Alkalophilic Environment and Its Implementation in Sustainable Arthrospira Cultivation. Sustainability 2024, 16, 8627. https://doi.org/10.3390/su16198627

AMA Style

Chotchindakun K, Buddhasiri S, Kuntanawat P. Enhanced Growth and Productivity of Arthrospira platensis H53 in a Nature-like Alkalophilic Environment and Its Implementation in Sustainable Arthrospira Cultivation. Sustainability. 2024; 16(19):8627. https://doi.org/10.3390/su16198627

Chicago/Turabian Style

Chotchindakun, Kittipat, Songphon Buddhasiri, and Panwong Kuntanawat. 2024. "Enhanced Growth and Productivity of Arthrospira platensis H53 in a Nature-like Alkalophilic Environment and Its Implementation in Sustainable Arthrospira Cultivation" Sustainability 16, no. 19: 8627. https://doi.org/10.3390/su16198627

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