Next Article in Journal
Research on the Water Entry of the Fuselage Cylindrical Structure Based on the Improved SPH Model
Previous Article in Journal
Chemical Composition Analysis of Lilium brownii var. Viridulum Baker and the Effect of Postharvest Primary Processing on Its Quality
Previous Article in Special Issue
The Effect of Methyl Jasmonate-Doped Nano-Particles and Methyl Jasmonate on the Phenolics and Quality in Monastrell Grapes during the Ripening Period
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Applied Sciences
Volume 13
Issue 19
10.3390/app131910796
applsci-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

Application of Extracellular Polymeric Substances during Cultivation of Microalgae Biomass

by
Paulina Rusanowska
,
Marcin Zieliński
*,
Magda Dudek
and
Marcin Dębowski
Department of Environmental Engineering, Faculty of Geoengineering, University of Warmia and Mazury in Olsztyn, 10-720 Olsztyn, Poland
*
Author to whom correspondence should be addressed.
Appl. Sci. 2023, 13(19), 10796; https://doi.org/10.3390/app131910796
Submission received: 29 August 2023 / Revised: 23 September 2023 / Accepted: 27 September 2023 / Published: 28 September 2023
(This article belongs to the Special Issue Knowledge-Based Biotechnology for Food, Agriculture and Fisheries)
Browse Figures
Versions Notes

Abstract

:

Featured Application

The biomass of microalgae/cyanobacteria selected in this study had different characteristics and applications and included Chlorella sp. (bio-oil production), Tetraselmis subcordiformis (bio-hydrogen production), and Arthrospira platensis (phycocyanin production). A critical, well-diagnosed, and proven problem of microalgae/cyanobacteria cultivation on an industrial scale is the lack of a simple and effective technology for separating the obtained biomass from the culture medium. The study aimed to solve the problem of biomass harvesting using extracellular polymeric substances extracted from activated sludge.

Abstract

Extracellular polymeric substances (EPS) produced by microorganisms contain polymers that are used for the bioflocculation of microalgae; however, these polymers are also organic compounds that might be used as carbon sources. The study analyzed two strategies for the introduction of EPS for Tetraselmis subcordiformis, Chlorella sp., and Arthrospira platensis biomass harvesting. In the first variant, EPS in the dose of 100 mg TOC/g were added to the photobioreactor every other day from the beginning of the cultivation, while in the second variant, EPS in the two doses of 100 mg TOC/g and 300 mg TOC/g were only added at the end of cultivation. In the first variant, the results proved that microalgae/cyanobacteria can use the EPS as external carbon sources. The cultures were characterized by a faster increase in biomass concentration, which contained less chlorophyll. However, the EPS content did not change. In the second variant, the addition of EPS did not affect the EPS content and the sedimentation of the Chlorella sp. biomass. The biomass of T. subcordiformis was characterized by a much better sedimentation coefficient. The greatest differences were observed in the A. platensis culture: the biomass concentration increased from 1.2 ± 0.2 g/L to 1.9 ± 0.2 g/L, EPS content increased by 16%, and sedimentation efficiency increased to 72%.

1. Introduction

EPS secreted by microorganisms are complex compositions of biopolymers with a high molecular weight, such as polysaccharides, proteins, humic substances, nucleic acids, and lipids [1]. EPS serve to shield cells from environmental stresses, such as dehydration, external heavy metals, and toxic compounds, and helps to maintain matrix stability [2]. Polymers included in EPS retain stable matrix structures and form a network for cells to interact with each other and mediate their adhesion to surfaces.
One of the possible applications of extracellular polymers is the bioflocculation of microalgae biomass. Technologies using microalgae biomass are solutions with significant potential that can be applied in broadly understood field of environmental engineering. Research carried out so far has proved that it is possible to use systems for the intensive cultivation of microalgae biomass in wastewater and leachate treatment, waste and sewage sludge management, carbon dioxide biosequestration, or flue gas purification. An important problem of microalgae cultivation on an industrial scale is the lack of simple and effective technology for the separation of the biomass from the medium.
Centrifugation involves using a rotational force that accelerates the mobility and detachment of particles by different densities of the particle and the medium. This process, because of its high efficiency and simplicity, is the most commonly employed technology for industrial microalgae harvesting [3]. However, it is not recommended for certain species due to shear stress. The disc stack centrifuges enable the separation of microalgae of small sizes (3–30 mm) with low cell densities. However, the high capital costs and considerable energy requirements make this technology unprofitable [4].
Filtration is the separation of solid and liquid via a membrane with pores that allow microalgae medium to pass through while retaining cells [5]. Membrane fouling is the biggest issue with using this technology, and it usually requires various improvement strategies to increase its efficiency [6].
Flocculation is a process that requires the presence of flocculating agents inducing microalgal cells to aggregate, which facilitates easier and faster sedimentation. Flocculants can be inorganic, organic, or have biological origin. Currently, research has focused on developing integrated systems for simultaneous microalgae culture and cell harvesting, such as the co-culture of microalgae and fungi, the co-culture of microalgae and bacteria, using microalgae biofilms, etc. These systems show a high density of microalgae cells and efficient cell separation from the medium realized in the same reactor [7]. However, most of these approaches are still in the early stages of development, and future research should focus on improving and optimizing these integrated culture and harvesting systems.
Bioflocculation is initiated by extracellular polymers (EPS) produced by well-settling microorganisms (microalgae, bacteria, fungi, or yeast). The advantage of bioflocculation is its non-toxic nature and potential as an economic method for the concentration of microalgae biomass. EPS added for bioflocculation determine the settling properties and dewatering of flocs. The bioflocculant produced by Solibacilus silbestris was used to concentrate microalgae Nannochloropsis oceanica [8]. The bioflocculant did not require the addition of other coagulants and was not toxic to microalgae cells. In addition, the research presented the possibility of its recovery after applying filtration and evaporation. The results indicate that the bioflocculation efficiency of the recovered bioflocculant dropped by only 3%, which significantly reduces the costs of biomass concentration. However, these results are only attractive for small-scale processes. In addition, the presented approach is connected to the necessity of purchasing a specific bacterial species, its cultivation, and the isolation of the bioflocculant in a separate reactor, which may pose technological difficulties and significantly increase the costs of biomass production.
There are many studies of the use of different substances to improve the sedimentation properties of microalgae. However, according to the authors’ knowledge, EPS extracted from activated sludge from wastewater treatment plants have never been used for this purpose. However, as EPS are rich in organic carbon, they can also play a role as extracellular energy and carbon sinks. They are an abundant source of structurally and compositionally diverse biopolymers. Therefore, EPS are important carbon sources for different organisms in the food chain.
The separation method for microorganisms’ EPS can, in turn, influence the compositions, contents, structure, properties, and functions of EPS [9,10]. Commonly applied methods, e.g., high-temperature or alkaline environments, could markedly change the properties and compositions of the extracted EPS. The cation-exchange resin method was selected as the most effective for the extraction of EPS, which prevents external forces from causing chemical contamination and changing the characteristics and properties of EPS [11,12].
The presence of both proteins and polysaccharides in the extracted EPS could stimulate the aggregation of cells or production of EPS by microalgae, which can contribute to the better sedimentation properties of microalgae. Thus, the aim of this study was to assess the possibility of the utilization of extracellular polymeric substances (EPS) produced by the biomass of bacteria used in wastewater treatment (activated sludge) to densify the biomass of selected species of microalgae and cyanobacteria. The research analyzed the impact of isolated EPS on the cultivation of microalgae Chlorella sp., Tetraselims subcordiformis, and cyanobacteria Arthrospira platensis.

2. Materials and Methods

2.1. EPS Origin

EPS were extracted from activated sludge using an aerobic reactor with a volume of 20 L, which was fed with model municipal wastewater with a high load of organic compounds. The basic technological parameters of the bioreactor operation were as follows: the concentration of activated sludge in the chamber was 5200 mg/L, load of organic compounds was 0.7 kg COD/(m3∙d), and oxygen content was 2 mg/L. During the operation of the aerobic bioreactor, the sedimentation time of the activated sludge was successively shortened in the range from 15 to 5 min. Then, EPS were extracted and characterized. The EPS content was expressed as mg of total organic carbon (TOC)/g biomass. The extracted EPS were characterized by 429 ± 34 mg TOC/g VS, 310 ± 18 mg of proteins (PN)/g biomass, and 61 ± 9 mg of carbohydrates (PS)/g biomass.

2.2. EPS Extraction

EPS were isolated using ion exchange resins [13]. In brief, EPS extraction was performed using a Dowex 50 × 8, Na+ Form cation-exchange resin (Fluka). The ratio of the rinsed cation-exchange resin to VS of biomass was 35 g to 0.5 g. The prepared samples were stirred at 750 rpm for 4 h in the dark at 4 °C. The blank sample was prepared with PBS buffer and cation-exchange resin. Then, the samples were centrifuged at 10,000 rpm for 1 min and again centrifuged at 10,000 rpm for 10 min in new clean tube. Afterwards, the samples were filtered using the membrane filter (0.22 µm pore size). The total quantity of extracted EPS was measured using an Elementar High TOC analyzer (Shimadzu TOC-L, Kyoto, Japan).

2.3. Microalgae Cultivation

Microalgae cultures were carried out in tubular reactors with an active volume of 2.5 L, equipped with a lighting and air mixing system. Based on the experience gained during the implementation of previous research, the cultures of Chlorella sp. and A. platensis were illuminated in a continuous mode, while T. subcordiformis was illuminated in the mode of 14 h of light/10 h of darkness. The cultures were illuminated by cool-white light at 700 lx using a T8 Sylvania Gro-Lux 18 W fluorescent tube. The temperature in the photobioreactors was kept at 22 ± 1.0 °C for Chlorella sp. and T. subcordiformis, and A. platensis was kept at 30 ± 2.0 °C. The composition of the T. subcordiformis medium was as described elsewhere [14], and the A. platensis medium was as described by [15], while for the cultivation of Chlorella sp., a modified BBM medium (bold basal medium) was used, the composition of which was described in [16]. The air introduced into the photobioreactors from the bottom ensured the mixing of the cultures and was a carrier of carbon dioxide. Each experiment was performed in triplicate. The taxonomic biomass changes were controlled at microscope total magnifications of 100× or 400× using the algae analyzer BBE (Moldaenke, Schwentinental, Germany).

2.4. Bioflocculation

This study analyzed two variants used to introduce EPS into the photobioreactor for the cultivation of microalgae/cyanobacteria. In the first variant, EPS were introduced into the photobioreactor every other day from the beginning of the culture (V1), while in the second variant, EPS were only introduced at the end of microalgae/cyanobacteria growth (on 7th day). In the first variant, EPS in the amount of 100 mg TOC/g of microalgae were introduced into the culture. In the second variant, EPS in the amounts of 100 mg TOC/g (V2) and 300 mg TOC/g (V3) of microalgae were introduced into the culture (Figure 1).
The biomass characteristics included the determination of the contents of total solids (TS), volatile solids (VS), and chlorophyll. The medium characteristics included the determination of total nitrogen (Ntot), phosphorus (Ptot), and organic compounds (COD). The EPS characteristics included the determination of polysaccharides and proteins.

2.5. Analytical Methods

The TS and VS concentrations were assessed via a gravimetric method. The samples of nutrients and organic analyses in the medium were first centrifuged via a laboratory centrifuge MPW-251 (Donserv, Warsaw, Poland) with a rotational speed of 5000 rpm for 10 min. The COD, Ntot, and Ptot concentrations were measured via LCK cuvette tests (Hach-Lange, Berlin, Germany) using the DR 5000 spectrophotometer with the HT 200 s mineralizer (Hach-Lange, Berlin, Germany). EPS were isolated from the biomass using ion exchange resins collected at the end of the experiments. The total quantity of extracted EPS was measured using an Elementar High TOC analyzer (Shimadzu TOC-L, Kyoto, Japan). Lowry’s method [17] was used to measure the amount of protein, relative to a standard curve created using bovine serum albumin (BSA). Anthrone method [18] was used to measure the amount of carbohydrates, relative to a standard curve created using glucose. The Assay kits were used for the specific measurement of monosaccharides, disaccharides, and trisaccharides (arabinose, fructose, glucose, gluconic acid, fucose, glucosamine, glucuronic acid, galactouronic acid, lactose, galactose, sucrose, maltose, raffinose, rhamnose, trehalose, and xylose) (Megazyme Ltd., Bray, Ireland). Only the content of carbohydrates higher than 1 mg/g VS was presented in the results. The content of chlorophyll a was measured via extraction with acetone and spectrophotometrically determined at 661.7 nm with a calibration curve using chlorophyll a as standard (Sigma-Aldrich, Burlington, MA, USA) [19,20]. The sedimentation efficiency was measured in cylinders with a capacity of 100 cm3 after 30 min. The cylinders had a reference scale to record the results in millimeters.
After testing for the homogeneity of variance via Levene’s test, the significance of the differences between the variants was tested via Tukey’s HSD test. Differences were considered significant at p < 0.05.

3. Results and Discussion

The study tested different strategies used for the possible application of EPS for microalgae/cyanobacteria biomass. In strategy V1, the biomass was characterized by lower chlorophyll content compared to the control cultivation (Table 1). In this mixotrophic cultivation (V1), the chlorophyll content was significantly reduced by 72.5%, 71.4%, and 50% in the biomass of T. subcordiformis, Chlorella sp., and A. platensis, respectively (p < 0.05). The introduction of EPS during the cultivation in V1 caused microalgae/cyanobacteria to use polymers as an external carbon source.
Photoautotrophic microorganisms use light as a source of energy for CO2 assimilation and store light energy in the chemical bonds of organic molecules. Some of these microorganisms also have a capacity for heterotrophic growth or mixing heterotrophic and photoautotrophic growth in metabolic traits called photomixotrophic traits. The strategy for changing metabolic traits is used for fast acclimation to the environmental conditions of aquatic microorganisms. The environmental conditions that could modulate the metabolic traits and, therefore, growth of microorganisms include light intensity, temperature, and concentrations of different forms of carbon. The ability of microorganisms to change metabolic traits is poorly described in the literature and has remained rather undefined. It was found that among microalgae species, Chlorella sp. is capable of mixotrophic growth [21,22]. The studies proved that higher biomass concentrations were observed under mixotrophic growth than under only photoautotrophic or heterotrophic growth. The presence of organic and inorganic carbon sources during mixotrophic conditions results in combining photosynthetic activity with high intracellular inorganic carbon concentrations created by glucose breakdown and the release of CO2, despite the direct utilization of glucose-derived carbon skeletons for growth [22,23].
During the proteomic analysis under different metabolic conditions in Chlorella sp. cultivation, it was observed that after feeding the photoautotrophic culture with glucose to start the mixotrophic metabolism, the chlorophyll content in biomass decreased immediately, and this decline lasted 8 h. During the next 24 h, the transition from the mixotrophic conditions to the photoautotrophic conditions occurred [22]. The present study showed slightly different results: the chlorophyll content in biomass did not increase in the two days after the addition of organic carbon (EPS). These differences might be the result of different sources and concentrations of organic carbon being added to cultivation. EPS consist of polysaccharides, proteins, nucleic acids, and lipids, which could exert a longer effect on cell metabolism than simple compounds, such as glucose. The presence of organic carbon sources inhibits the synthesis of pigments at the start of mixotrophy, causing the lower chlorophyll content [24]. Moreover, the reduction in the abundance of proteins from the thylakoid membranes and the downregulation of photosynthesis-related genes were reported under mixotrophic conditions [25]. Similar observations of lower chlorophyll content in the cultures with organic carbon addition were observed during cultivation of Arthrospira platensis [26] and Tetraselmis subcordiformis [14,27].
The biomass growth in the control and until 10th day of microalgae/cyanobacteria cultivation in V2 and V3 was the same since on the 10th day after EPS were added to the control cultivation. The biomass concentration during strategy V1 was higher than during the control for all tested microalgae/cyanobacteria. During V1, the biomass concentration of T. subcordiformis and Chlorella sp. was similar, and the growth rate was around 600 mg/(L∙d) in the first 5 days of cultivation and around 100 mg/(L∙d) in the next 5 days (Figure 2). The A. platensis growth rate was steady during cultivation, being around 150 mg/(L∙d).
Microalgae growth generally can be divided into six phases during batch cultivation: the lag phase, exponential phase, linear phase, declining growth phase, stationary phase, and death phase. In the lag phase, the growth of the cell is delayed due to the presence of non-viable cells or adaptation to new environmental conditions (the addition of organic carbon). In the present study, the lag phase was not observed for all tested strategies. Similar observations for Chlorella vulgaris were noted during transcriptomic studies of the transition of photoautotrophic to mixotrophic metabolism, in which biomass after the addition of glucose started an exponential phase [22]. Additionally, the studies of terrestrial cyanobacteria cells under heterotrophic and mixotrophic conditions did not show a lag phase [28]. The lag phase for C. sorokiniana during mixotrophic cultivation with different volatile fatty acids lasted less than 1 day, which can also be the explanation for observations in the present study because the measurements were taken once a day [29]. The above-mentioned explanations support the statement of mixotrophic metabolism observed during EPS addition in strategy V1.
The mixotrophic cultivation did not increase the sedimentation efficiency of T. subcordiformis and Chlorella sp. biomass (Table 1). The biomass of A. platensis was characterized by a 5% better efficiency of sedimentation after mixotrophic cultivation than autotrophic cultivation. The introduction of EPS at the end of cultivation (V2 and V3) also lowered chlorophyll content in biomass; however, the reduction was not significant (Table 1). In V3, the chlorophyll content was lower than in V2. The higher concentrations of source of organic carbon available for biomass, the greater the reduction in the chlorophyll content in biomass. In V2 and V3, the chlorophyll content was reduced by about 25%, 60%, and 38% compared to autotrophic cultivation in the biomass of T. subcordiformis, Chlorella sp., and A. platensis, respectively. The sedimentation efficiency was not improved in Chlorella sp. when EPS was introduced at the end of cultivation, independently of the EPS concentration. However, the sedimentation efficiency of T. subcordiformis and A. platensis were improved. In the case of T. subcordiformis, the better sedimentation properties were not connected to higher EPS content, because in all variants of the experiments, the EPS content was similar. However, in A. platensis biomass, the EPS content increased.
The EPS content in the control biomass of microalgae/cyanobacteria and the biomass from the microalgae/cyanobacteria culture to which EPS of the activated sludge were added during cultivation did not differ. The use of mixotrophic conditions did not affect the improvement in the sedimentation properties and EPS content. It is well known that increased carbon availability during cultivation will increase the biomass concentration and decrease the chlorophyll content [30]. Moreover, mixotrophic cultivation also often has a positive effect on the “production capacity” of microalgal cells, e.g., Chlorella sp. often has more oil in the cells, while A. platensis has more phycocyanin in the cells [31].
In this study, in the EPS samples, the protein content, sugar content, and carbohydrate composition were determined (Table 2). In the EPS extracted from T. subcordiformis and Chlorella sp., the protein content was similar in variants of experiments. However, in the EPS extracted from A. platensis, the protein content increased. The positively charged amino groups in proteins can neutralize the negative charges of polysaccharides, uronic acid, and carboxylic acid in DNA, phosphate groups, etc.; thus, proteins might effectively change the surface properties of the cell, inducing the formation of aggregates [13,32,33]. The higher content of proteins in A. platensis could improve the sedimentation properties of biomass.
In the EPS extracted from T. subcordiformis, sugar content has significantly increased in the variants of experiments where EPS were added at the end of microalgae growth (V2 and V3) compared to autotrophic cultivation (Table 2). Among the measured carbohydrates, the highest concentration was glucose, the concentration of which also increased in V2 and V3; however, it was not a significant increase. The higher content of sugars in the EPS might be caused by some other exopolysaccharide that also influenced the better sedimentation of T. subcordiformis (Table 1). Exopolysaccharide produced by Bacillus subtilis was used for harvesting Nannochloropsis oculata and Botryococcus braunii biomass [34]. The authors obtained the best flocculation with the dose of 5% v/v of exopolysaccharide and pointed out that harvesting efficiency and flocculant concentration are not linearly dependent. Polysaccharides were also connected to the formation of algal-bacterial bioflocs. The sedimentation properties of bioflocs were better with a higher content of polysaccharides [35]. Similarly, polysaccharides were responsible for cell capture and aggregation during the rapid biogranulation of the microalgae Ankistrodesmus falcatus var. acicularis [36]. The polysaccharide derived from T. subcordiformis should be further investigated via extraction and analyses to determine its influence on the sedimentation properties.
In the EPS extracted from Chlorella sp., the sugar content increased in the variants to which EPS were added at the end of microalgae growth (V2 and V3) compared to autotrophic cultivation (Table 2). The main components of carbohydrates in the EPS extracted from Chlorella sp. were glucose and mannose. Despite the increase in sugar content, the EPS content and sedimentation efficiency did not improve. Based on the results of the EPS content and composition and sedimentation properties of T. subcordiformis and Chlorella sp., it might be concluded that similar flocculants induce the production of different exopolysaccharides with different properties.
The applied strategy and flocculant did not improve the sedimentation properties of Chlorella sp. Similar observations were made by [37]. The authors tested the settling properties of two species of microalgae, namely Scenedesmus sp. and Chlorella vulgaris, with and without cultures of the bacterium Burkholderia cepacia and bacterial exudates. C. vulgaris’ settling properties were not improved by applying flocculants at any growth stage. In contrast, glutathione significantly increased the EPS content in Chlorella pyrenoidosa and the settling properties above 72% [38]. The biomass of Chlorella vulgaris could even have 94% harvesting efficiency when the culture was contaminated with bacteria [39]. The stress conditions that could include bacteria presence, nitrogen depletion, etc., could induce the autoflocculation of microalgal cells and improve sedimentation [40]. The stress and autoflocculation might also be the results of changes in cell surface properties. It was proved that the cell surface charge of Chlorella sp. is different during the growth phases of biomass. These changes are related to the formation of a protective layer at the cell wall during late cultivation stages [41]. However, in the present study, the EPS added to the microalgae culture probably did not cause stress conditions for the cells of Chlorella sp. and was only a good source of external carbon for changing the metabolism for mixotrophic cultivation.
In the EPS extracted from A. platensis, the sugar content increased. The carbohydrates detected in all variants of experiments were glucose, rhamnose, and galactose. The addition of EPS at the end of the growth phase (V2 and V3) only increased the content of rhamnose in the EPS extracted from A. platensis. In the comparison of the mixotrophic and heterotrophic cultivation of A. platensis with whey as an external carbon source, the growth rate and protein and lipid contents were higher under heterotrophic cultivation; however, chlorophyll-a and total carbohydrates were higher in the mixotrophic culture [30]. In the present study, the cultures of A. platensis with the addition of EPS at the end of the growth phase were also characterized by higher protein content that, together with the carbohydrates, increased the sedimentation properties of this microalgae.
The differences in the bioflocculation of different species might be attributed to differences in cell size and shape, culture density, and EPS produced [20]. Chlorella sp. has the smallest cells, with a diameter range from 2 to 10 μm. The other important factor seems to be type of EPS produced by microalgae. As mentioned above, well sedimentation properties of A. platensis were connected to exopolysaccharide extracted by microalgae.

4. Conclusions

The study can be summarized based on the following conclusions:
-
Extracellular polymeric substances from bacteria can be used as an external carbon source by microalgae/cyanobacteria;
-
Extracellular polymeric substances can be used for harvesting Tetraselmis subcordiformis and Arthrospira platensis;
-
The sugar content in Tetraselmis subcordiformis biomass was responsible for the higher sedimentation coefficient.

Author Contributions

Conceptualization, P.R. and M.D. (Marcin Dębowski); methodology, P.R.; software, P.R.; validation, P.R.; formal analysis, P.R.; investigation, P.R. and M.D. (Magda Dudek); resources, P.R.; data curation, P.R.; writing—original draft preparation, P.R.; writing—review and editing, M.Z., M.D. (Magda Dudek) and M.D. (Marcin Dębowski); supervision, M.Z.; project administration, P.R.; funding acquisition, M.D. (Marcin Dębowski). All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Felz, S.; Vermeulen, R.; van Loosdrecht, M.C.M.; Lin, Y.M. Chemical characterization methods for the analysis of structural extracellular polymeric substances (EPS). Water Res. 2019, 157, 201–208. [Google Scholar] [CrossRef] [PubMed]
  2. Schambeck, C.M.; Magnus, B.S.; de Souza, L.C.R.; Leite, W.R.M.; Derlon, N.; Guimarães, L.B.; da Costa, R.H.R. Biopolymers recovery: Dynamics and characterization of alginate-like exopolymers in an aerobic granular sludge system treating municipal wastewater without sludge inoculum. J. Environ. Manag. 2020, 263, 110394. [Google Scholar] [CrossRef] [PubMed]
  3. Zhang, H.Y.; Zhang, X.Z. Microalgal harvesting using foam flotation: A critical review. Biomass Bioenergy 2019, 120, 176–188. [Google Scholar] [CrossRef]
  4. Dassey, A.J.; Theegala, C.S. Harvesting economics and strategies using centrifugation for cost effective separation of microalgae cells for biodiesel applicationsc. Bioresour. Technol. 2013, 128, 241–245. [Google Scholar] [CrossRef]
  5. Liu, Z.; Hao, N.; Hou, Y.; Wang, Q.; Liu, Q.; Yan, S.; Chen, F.; Zhao, L. Technologies for harvesting the microalgae for industrial applications: Current trends and perspectives. Bioresour. Technol. 2023, 387, 129631. [Google Scholar] [CrossRef]
  6. Zhao, Z.Y.; Ilyas, A.; Muylaert, K.; Vankelecom, I.F.J. Optimization of patterned polysulfone membranes for microalgae harvesting. Bioresour. Technol. 2020, 309, 123367. [Google Scholar] [CrossRef]
  7. Huang, K.X.; Vadiveloo, A.; Zhou, J.L.; Yang, L.; Chen, D.Z.; Gao, F. Integrated culture and harvest systems for improved microalgal biomass production and wastewater treatment. Bioresour. Technol. 2023, 376, 128941. [Google Scholar] [CrossRef]
  8. Wan, C.; Zhao, X.Q.; Guo, S.L.; Asraful Alam, M.; Bai, F.W. Bioflocculant production from Solibacillus silvestris W01 and its application in cost-effective harvest of marine microalga Nannochloropsis oceanica by flocculation. Bioresour. Technol. 2013, 135, 207–212. [Google Scholar] [CrossRef]
  9. Takahashi, E.; Ledauphin, J.; Goux, D.; Orvain, F. Optimising extraction of extracellular polymeric substances (EPS) from benthic diatoms: Comparison of the efficiency of six EPS extraction methods. Mar. Freshw. Res. 2009, 60, 1201–1210. [Google Scholar] [CrossRef]
  10. Frolund, B.; Palmgren, R.; Keiding, K.; Nielsen, P.H. Extraction of extracellular polymers from activated sludge using a cation exchange resin. Water Res. 1996, 30, 1749–1758. [Google Scholar] [CrossRef]
  11. Sheng, G.P.; Yu, H.Q.; Li, X.Y. Extracellular polymeric substances (EPS) of microbial aggregates in biological wastewater treatment systems: A review. Biotechnol. Adv. 2010, 28, 882–894. [Google Scholar] [CrossRef] [PubMed]
  12. Flemming, H.C.; Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 2010, 8, 623–633. [Google Scholar] [CrossRef] [PubMed]
  13. McSwain, B.S.; Irvine, R.L.; Hausner, M.; Wilderer, P.A. Composition and distribution of extracellular polymeric substances in aerobic flocs and granular sludge. Appl. Environ. Microbiol. 2005, 71, 1051–1057. [Google Scholar] [CrossRef] [PubMed]
  14. Dudek, M.; Dębowski, M.; Zieliński, M.; Nowicka, A.; Rusanowska, P. Water from the Vistula Lagoon as a medium in mixotrophic growth and hydrogen production by Platymonas subcordiformis. Int. J. Hydrog. Energy 2018, 43, 9529–9534. [Google Scholar] [CrossRef]
  15. Aiba, S.; Ogawa, T. Assessment of growth yield of a blue-green alga: Spirulina platensis in axenic and continuous cultur. J. Gen. Microbiol. 1977, 102, 179–182. [Google Scholar] [CrossRef]
  16. Szwarc, K.; Szwarc, D.; Zieliński, M. Removal of biogenic compounds from the post-fermentation effluent in a culture of Chlorella vulgaris. Environ. Sci. Pollut. Res. 2020, 27, 111–117. [Google Scholar] [CrossRef]
  17. Lowry, O.H.; Rosebrough, N.J.; Farr, A.L.; Randall, R.J. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 1951, 193, 265–275. [Google Scholar] [CrossRef]
  18. Gaudy, A.F. Colorimetric determination of protein and carbohydrate. Ind. Water Wastes 1962, 7, 17–22. [Google Scholar]
  19. Johan, F.; Jafri, M.Z.; Lim, H.S.; Wan Maznah, W.O. Laboratory measurement: Chlorophyll-a concentration measurement with acetone method using spectrophotometer. In Proceedings of the 2014 IEEE International Conference on Industrial Engineering and Engineering Management, Selangor, Malaysia, 9–12 December 2014; pp. 744–748. [Google Scholar]
  20. Sasadara, M.M.V.; Nayaka, N.M.D.M.; Yuda, P.E.S.K.; Dewi, N.K.L.A.; Cahyaningsih, N.; Wirawan, I.G.P.; Silalahi, D. Optimization of chlorophyll extraction solvent of bulung sangu (Gracilaria sp.) seaweed. IOP Conf. Ser. Earth Environ. Sci. 2021, 913, 012073. [Google Scholar] [CrossRef]
  21. Heredia-Arroyo, T.; Wei, W.; Ruan, R.; Hu, B. Mixotrophic cultivation of Chlorella vulgaris and its potential application for the oil accumulation from non-sugar materials. Biomass Bioenergy 2011, 35, 2245–2253. [Google Scholar] [CrossRef]
  22. Vidotti, A.D.S.; Riaño-Pachón, D.M.; Mattiello, L.; Giraldi, L.A.; Winck, F.V.; Franco, T.T. Analysis of autotrophic, mixotrophic and heterotrophic phenotypes in the microalgae Chlorella vulgaris using time-resolved proteomics and transcriptomics approaches. Algal Res. 2020, 51, 102060. [Google Scholar] [CrossRef]
  23. Liang, Y.; Sarkany, N.; Cui, Y. Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnol. Lett. 2009, 31, 1043–1049. [Google Scholar] [CrossRef] [PubMed]
  24. Roth, M.S.; Gallaher, S.D.; Westcott, D.J.; Iwai, M.; Louie, K.B.; Mueller, M.; Walter, A.; Foflonker, F.; Bowen, B.P.; Ataii, N.N.; et al. Regulation of oxygenic photosynthesis during trophic transitions in the green alga Chromochloris zofingiensis. Plant Cell 2019, 31, 579–601. [Google Scholar] [CrossRef] [PubMed]
  25. Xiang, T.; Jinkerson, R.E.; Clowez, S.; Tran, C.; Krediet, C.J.; Onishi, M.; Cleves, P.A.; Pringle, J.R.; Grossman, A.R. Glucose-induced trophic shift in an endosymbiont dinoflagellate with physiological and molecular consequences. Plant Physiol. 2018, 176, 1793–1807. [Google Scholar] [CrossRef]
  26. Narayan, M.S.; Manoj, G.P.; Vatchravelu, K.; Bhagyalakshmi, N.; Mahadevaswamy, M. Utilization of glycerol as carbon source on the growth, pigment and lipid production in Spirulina platensis. Int. J. Food Sci. Nutr. 2005, 56, 521–528. [Google Scholar] [CrossRef]
  27. Xie, J.; Zhang, Y.; Li, Y.; Wang, Y. Mixotrophic cultivation of Platymonas subcordiformis. J. Appl. Phycol. 2001, 13, 343–347. [Google Scholar] [CrossRef]
  28. Schwarz, A.; Walther, J.; Geib, D.; Witthohn, M.; Strieth, D.; Ulber, R.; Muffler, K. Influence of heterotrophic and mixotrophic cultivation on growth behaviour of terrestrial cyanobacteria. Algal Res. 2020, 52, 102125. [Google Scholar] [CrossRef]
  29. Lacroux, J.; Seira, J.; Trably, E.; Bernet, N.; Steyer, J.; Van Lis, R. Mixotrophic Growth of Chlorella sorokiniana on Acetate and Butyrate: Interplay between Substrate, C:N Ratio and pH. Front. Microbiol. 2021, 12, 703614. [Google Scholar] [CrossRef]
  30. Tosuner, Z.V.; Ürek, R.Ö. Cultivation of Arthrospira platensis in heterotrophic and mixotrophic conditions with different concentrations of whey. Aquat. Res. 2022, 5, 146–153. [Google Scholar] [CrossRef]
  31. Caporgno, M.P.; Haberkorn, I.; Böcker, L.; Mathys, A. Cultivation of Chlorella protothecoides under different growth modes and its utilisation in oil/water emulsions. Bioresour. Technol. 2019, 288, 121476. [Google Scholar] [CrossRef]
  32. Wang, Z.; Liu, L.; Yao, J.; Cai, W. Effects of extracellular polymeric substances on aerobic granulation in sequencing batch reactors. Chemosphere 2006, 63, 1728–1735. [Google Scholar] [CrossRef] [PubMed]
  33. Campo, R.; Corsino, S.F.; Torregrossa, M.; Di Bella, G. The role of extracellular polymeric substances on aerobic granulation with stepwise increase of salinity. Sep. Purif. Technol. 2018, 195, 12–20. [Google Scholar] [CrossRef]
  34. Lutfi, M.; Nugroho, W.A.; Fridayestu, W.P.; Susilo, B.; Pulmar, C.; Sandra, S. Bioflocculation of Two Species of Microalgae by Exopolysaccharide of Bacillus subtilis. Nat. Environ. Pollut. Technol. 2019, 18, 167–173. [Google Scholar]
  35. Wang, H.; Qi, B.; Jiang, X.; Jiang, Y.; Yang, H.; Xiao, Y.; Jiang, N.; Deng, L.; Wang, W. Microalgal interstrains differences in algal-bacterial biofloc formation during liquid digestate treatment. Bioresour. Technol. 2019, 289, 121741. [Google Scholar] [CrossRef]
  36. Wang, Q.; Shen, Q.; Wang, J.; Zhao, J.; Zhang, Z.; Lei, Z.; Yuan, T.; Shimizu, K.; Liu, Y.; Lee, D.J. Insight into the rapid biogranulation for suspended single-cell microalgae harvesting in wastewater treatment systems: Focus on the role of extracellular polymeric substances. Chem. Eng. J. 2022, 430, 132631. [Google Scholar] [CrossRef]
  37. Manheim, D.; Nelson, Y. Settling and bioflocculation of two species of algae used in wastewater treatment and algae biomass production. Environ. Prog. Sustain. Energy 2013, 32, 946–954. [Google Scholar] [CrossRef]
  38. Hao, N.; Liu, Z.; Hou, Y.; Fan, Z.; Li, Y.; Chen, F.; Zhao, L. Small peptide glutathione-induced bioflocculation for enhancing the food application potential of Chlorella pyrenoidosa. Bioresour. Technol. 2022, 365, 128138. [Google Scholar] [CrossRef]
  39. Liber, J.A.; Bryson, A.E.; Bonito, G.; Du, Z.-Y. Harvesting Microalgae for Food and Energy Products. Small Methods 2020, 4, 2000349. [Google Scholar] [CrossRef]
  40. Nguyen, T.D.; Frappart, M.; Jaouen, P.; Pruvost, J.; Bourseau, P. Harvesting Chlorella vulgaris by natural increase in pH: Effect of medium composition. Environ. Technol. 2014, 35, 1378–1388. [Google Scholar] [CrossRef]
  41. Gerken, H.G.; Donohoe, B.; Knoshaug, E.P. Enzymatic cell wall degradation of Chlorella vulgaris and other microalgae for biofuels production. Planta 2013, 237, 239–253. [Google Scholar] [CrossRef]
Applsci 13 10796 g001
Figure 1. The scheme of the experiment.
Figure 1. The scheme of the experiment.
Applsci 13 10796 g001
Applsci 13 10796 g002aApplsci 13 10796 g002b
Figure 2. Microalgae/cyanobacteria growth during different strategies of EPS addition to cultivation: (A) biomass concentration of T. subcordiformis; (B) biomass concentration of Chlorella sp.; (C) biomass concentration of A. platensis.
Figure 2. Microalgae/cyanobacteria growth during different strategies of EPS addition to cultivation: (A) biomass concentration of T. subcordiformis; (B) biomass concentration of Chlorella sp.; (C) biomass concentration of A. platensis.
Applsci 13 10796 g002aApplsci 13 10796 g002b
Table 1. Biomass concentrations, chlorophyll content, EPS content, and sedimentation efficiency obtained in the variants of experiments during the cultivation of T. subcordiformis, Chlorella sp., and A. platensis.
Table 1. Biomass concentrations, chlorophyll content, EPS content, and sedimentation efficiency obtained in the variants of experiments during the cultivation of T. subcordiformis, Chlorella sp., and A. platensis.
Variant of ExperimentBiomass Concentration (mg/L)Chlorophyll Content (mg/g VS)EPS Content (mg TOC/g VS)Sedimentation Efficiency (%)
Tetraselmis subcordiformisC3621 ± 17440 ± 1195 ± 1140
V14109 ± 16011 ± 497 ± 742
V23698 ± 10232 ± 997 ± 2555
V33856 ± 11128 ± 1297 ± 1066
Chlorella sp.C3920 ± 28928 ± 860 ± 2032
V14210 ± 3208 ± 661 ± 931
V24290 ± 25513 ± 562 ± 1234
V34321 ± 26210 ± 261 ± 1936
Arthrospira platensisC1213 ± 18216 ± 481 ± 649
V11512 ± 1918 ± 483 ± 954
V21760 ± 19511 ± 691 ± 1268
V31912 ± 1509 ± 496 ± 272
Table 2. EPS composition obtained in the variants of experiments during the cultivation of T. subcordiformis, Chlorella sp., and A. platensis.
Table 2. EPS composition obtained in the variants of experiments during the cultivation of T. subcordiformis, Chlorella sp., and A. platensis.
Variant of ExperimentProtein Content (mg/g VS)Sugar Content (mg/g VS)Carbohydrates
Glucose (mg/g VS)Mannose (mg/g VS)Rhamnose (mg/g VS)Galactose (mg/g VS)
Tetraselmis subcordiformisC27.4 ± 6.721.5 ± 4.515.2 ± 5.4--1.3 ± 0.5
V128 ± 4.825.3 ± 5.621 ± 1.1--1.9 ± 0.4
V227.9 ± 3.532 ± 324.3 ± 2.7--1.9 ± 1.5
V328.3 ± 9.241 ± 522.5 ± 3.2--2.1 ± 0.8
Chlorella sp.C32.9 ± 4.741.1 ± 417.8 ± 3.212.9 ± 5.2-1.1 ± 0.2
V133.1 ± 8.850.3 ± 8.223.8 ± 2.519.4 ± 3.8-1.7 ± 0.1
V233.7 ± 5.353.7 ± 9.124.1 ± 5.320.1 ± 2.7-1.7 ± 0.5
V335.6 ± 4.956.3 ± 7.723.9 ± 4.521.1 ± 3.2-2.2 ± 1.1
Arthrospira platensisC45 ± 10.337.8 ± 6.411.4 ± 1.2-25.1 ± 3.210.9 ± 1.4
V159.1 ± 9.142.8 ± 9.810.5 ± 5-26.7 ± 1.98.6 ± 3.1
V257 ± 13.245.2 ± 8.111.9 ± 2.1-29.6 ± 0.59.1 ± 4
V360.1 ± 1148 ± 10.112.4 ± 2.9-30.4 ± 39.0 ± 3.2
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

Rusanowska, P.; Zieliński, M.; Dudek, M.; Dębowski, M. Application of Extracellular Polymeric Substances during Cultivation of Microalgae Biomass. Appl. Sci. 2023, 13, 10796. https://doi.org/10.3390/app131910796

AMA Style

Rusanowska P, Zieliński M, Dudek M, Dębowski M. Application of Extracellular Polymeric Substances during Cultivation of Microalgae Biomass. Applied Sciences. 2023; 13(19):10796. https://doi.org/10.3390/app131910796

Chicago/Turabian Style

Rusanowska, Paulina, Marcin Zieliński, Magda Dudek, and Marcin Dębowski. 2023. "Application of Extracellular Polymeric Substances during Cultivation of Microalgae Biomass" Applied Sciences 13, no. 19: 10796. https://doi.org/10.3390/app131910796

APA Style

Rusanowska, P., Zieliński, M., Dudek, M., & Dębowski, M. (2023). Application of Extracellular Polymeric Substances during Cultivation of Microalgae Biomass. Applied Sciences, 13(19), 10796. https://doi.org/10.3390/app131910796

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