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Review

Recent Advances of Microalgae Exopolysaccharides for Application as Bioflocculants

1
Laboratory of Microbiology and Biochemistry, College of Chemistry and Food Engineering, Federal University of Rio Grande, P.O. Box 474, Rio Grande 96203-900, RS, Brazil
2
Laboratory of Biochemical Engineering, College of Chemistry and Food Engineering, Federal University of Rio Grande, P.O. Box 474, Rio Grande 96203-900, RS, Brazil
3
Department of Bioprocess Engineering and Biotechnology, Federal University of Paraná, P.O. Box 19011, Curitiba 81531-990, PR, Brazil
*
Author to whom correspondence should be addressed.
Submission received: 29 January 2022 / Revised: 4 March 2022 / Accepted: 4 March 2022 / Published: 8 March 2022
(This article belongs to the Collection Current Opinion in Polysaccharides)

Abstract

:
Microalgae are used in flocculation processes because biopolymers are released into the culture medium. Microalgal cell growth under specific conditions (temperature, pH, luminosity, nutrients, and salinity) provides the production and release of exopolysaccharides (EPS). These biopolymers can be recovered from the medium for application as bioflocculants or used directly in cultivation as microalgae autoflocculants. The optimization of nutritional parameters, the control of process conditions, and the possibility of scaling up allow the production and industrial application of microalgal EPS. Therefore, this review addresses the potential use of EPS produced by microalgae in bioflocculation. The recovery, determination, and quantification techniques for these biopolymers are also addressed. Moreover, other technological applications of EPS are highlighted.

Graphical Abstract

1. Introduction

Microalgae are photosynthetic microorganisms cultivated in marine, hypersaline, brackish, freshwater, or wastewater for the production of high value-added compounds (pigments, proteins, lipids, polyunsaturated fatty acids, and intracellular and extracellular polysaccharides) [1,2,3]. Thus, the biomass of these microorganisms is of industrial interest in the development of pharmaceuticals, nutraceuticals, cosmetics, and food/feed [4]. However, the recovery of microalgal biomass is costly (20–30% of total production costs), limiting the commercialization of these bioproducts on a large scale [5].
Moreover, there is growing interest in alternative methods of harvesting microalgae biomass at low cost and energy. Microalgae exopolysaccharides (EPS) have been highlighted for promoting autoflocculation or acting as bioflocculants. Thus, these compounds can act in the process of microalgal biomass recovery and treatment of industrial effluents, with the additional advantage of low energy consumption, low environmental impact, and reduced production of toxic compounds [6,7,8,9].
In this sense, Yang et al. [10] reported bioflocculant activity of EPS from Scenedesmus acuminatus in the recovery of the biomass of this same microalga. The results showed that the addition of microalgae EPS (3.2 mg g−1) significantly reduced the use of aluminum coagulant (Al3+) from 77.6 to 4.5 mg g−1. The authors noted that this result potentially reduced the chemical cost by up to 75%. Aljuboori, Uemura, and Thanh [11] reported that EPS from Scenedesmus quadricauda showed bioflocculant activity in the biomass recovery of this same microalga with flocculation efficiency of up to 86.7%.
Microalgal EPS production depends on the microalgae species, cultivation conditions, or nutrient limitations such as nitrogen and phosphorus deprivation [12,13,14]. Furthermore, polysaccharides produced by microalgae can be classified based on their functionality: (i) structure of cell walls, (ii) storage (intracellular), and (iii) extracellular or exopolysaccharides, which are released into the environment. In addition to being a flocculating agent, microalgal EPS may have immunomodulatory [15], antioxidant [16], anti-inflammatory [17], anticoagulant [18], antibacterial [19], and anticancer properties [20], and may act as a gelling and thickening agent [12].
The most prominent families of microalgae in the production of EPS were Desmidiaceae, Chlamydomonadaceae, Chlorellaceae, Porphyridiaceae, and Glaucosphaeraceae [1]. However, there are few studies about other microalgae diversities in the production of this metabolite and its flocculating efficiency. In this context, this review reports the potential use of EPS produced by microalgae in bioflocculation. The recovery, determination, and quantification techniques concerning these biopolymers are also addressed. Moreover, other technological applications of EPS are highlighted.

2. Potentiality of Microalgal Polysaccharides in Bioflocculation

The use of autoflocculating microalgae to induce flocculation of non-flocculating species is considered one of the most promising methods in bioflocculation [21]. EPS can create a viscous coating around cells [12]. This coating in bioflocculation allows the formation of an aggregate consisting of microalgae–microalgae, or microalgae with another microorganism, leading to adherence of microalgae to the flocculent moss surface [22] or flocculent sludge surface [23,24].
Wang et al. [25] examined the algal-bacterial bioflocculation induced by strains of Scenedesmus obliquus, Botryococcus braunii, Chlorella sp. BWY-1, Haematococcus pluvialis, Dictyosphaerium ehnenbergianum, and Chlorella vulgaris. The microalgae Scenedesmus obliquus and Botryococcus braunii did not allow flocculation. Chlorella sp. BWY-1, Haematococcus pluvialis, and Dictyosphaerium ehnenbergianum showed high flocculation activity with 67.8, 50.9, and 43.2%, respectively. The concentration of chlorophyll a was approximately 5.5 mg L−1 for Chlorella sp. BWY-1, 3.0 mg L−1 for Haematococcus pluvialis, and 4.9 mg L−1 for Dictyosphaerium ehnenbergianum. Chlorella vulgaris exhibited the best flocculation activity (86.6%). However, the concentration of chlorophyll a decreased rapidly, reaching the lowest value in the experiment (1.6 mg L−1). The authors attributed these results to the high energy required for the production of EPS.
EPS represent carbon and energy reserves for cells and are often excreted by microalgae as part of physiological processes or under stress conditions [12,21,26,27,28], such as light intensity and light continuity, temperature [29,30], pH [31,32], excess, limitation or absence of nutrients, such as carbon, nitrogen, phosphorus, sulfur, sodium, potassium, iron, magnesium and calcium [14,33,34], and toxic substances [35,36], such as cadmium [37], copper, lead, chromium, nickel [31,38], and silver [39].
Koçer et al. [40] investigated the potential for EPS production using the microalgae Chlorella minutissima, Chlorella sorokiniana, and Botryococcus braunii. The authors analyzed the effects of nitrogen and carbon concentrations in the culture medium and light intensity on EPS production. Chlorella minutissima produced the highest concentration of EPS (0.245 ± 0.003 g L−1) compared to Chlorella sorokiniana (0.163 ± 0.002 g L−1) and Botryococcus braunii (0.117 ± 0.001 g L−1). Regarding the effects of nitrogen (NaNO3) and carbon (Na2CO3) concentration in the BG-11 medium and lighting time on EPS production, the best conditions for three microalgae were nitrogen reduction (0.2 g L−1) and carbon (0.02 g L−1) and 12 h of lighting time. Under these conditions, Chlorella sorokiniana, Botryococcus braunii, and Chlorella minutissima produced 0.183, 0.120, and 0.215 g L−1 EPS, respectively. Thus, the authors observed an inverse relationship between the supply of these nutrients and the concentration of EPS produced.
Surendhiran and Vijay [41] analyzed the flocculation efficiency of the Chlorella salina using a microbial flocculant. The authors found that flocculation was improved with zinc chloride (ZnCl2) as a cationic inducer. Moreover, the flocculation obtained maximum efficiency (98.6%) with the following conditions: temperature (30.6 °C), pH (10.4), flocculation time (6.2 h), the volume of bioflocculant (0.34 mL), and cationic inductor concentration (0.031 mM).
Thus, in addition to contributing to biomass recovery and mitigation of industrial effluents, the use of microalgae for the production of EPS proves to be an efficient and low environmental impact way to reduce costs in the flocculation process.

3. Recent Advances in Harvesting Algae and Pretreatments for the Extraction of Cell-Bound EPS

Studies on optimization strategies for the recovery of microalgae biomass are increasing since it demands high energy and operating costs (20 to 30% of the total production cost) [42,43,44]. In this way, it is necessary to define the recovery method to process high biomass production (Table 1). Thus, physical and chemical characteristics of the culture medium, such as pH, salinity, and cellular structure of microorganisms, must be analyzed and linked to the chosen method [44,45,46].
Traditionally, methods used in biomass recovery include coagulation, flocculation, flotation, gravity sedimentation, and centrifugation [45,46,48]. In flocculation methods, chemical compounds such as sodium hydroxide (NaOH), magnesium sulfate (MgSO4), magnesium chloride (MgCl2), calcium chloride (CaCl2), sodium alginate (NaC6H7O6), tannin, and other polymers can be used (Table 2). However, these can be combined to optimize the processes in the recovery of larger volumes of biomass [43,45,49]. In recent years, combined methods such as sedimentation–flocculation–coagulation, flocculation–centrifugation, and electrocoagulation–flotation have been used and show promise concerning cost and energy efficiency [43,44,45,46,49,50]. Moreover, natural (including EPS) and synthetic flocculating agents are applied in microalgal recovery [42,51].
Nguyen et al. [42] develop cationic polymers (poly[2(acryloyloxy) ethyl]trimethylammonium chloride and poly(3acrylamidopropyl) trimethylammonium chloride) for the harvest of Chlorella vulgaris and Porphyridium purpureum. The polymers show excellent flocculation performance for both microalgae with stable floc formation. Similar recovery strategies were also observed by Zhu et al. [51] when they analyzed three types of sulfates (aluminum sulfate, aluminum potassium sulfate, and ferric sulfate) as flocculants for harvesting Chlorella vulgaris. The results showed the flocculate potential of the chemical agents at a dosage of 2.5 g L−1 and speeds for coagulation and flocculation (150 and 25 rpm), and time of 10 min. The biomass recovery efficiency found ranged from 83 to 90%.
After recovery, it is important to pretreat the biomass to obtain EPS bound to microalgal cells [12,60]. Researchers describe that up to 50% of the total EPS can remain bound to the cell of these microorganisms. However, there are no standard methods for this extraction. The use of chemical reagents such as formaldehyde, ethylenediaminetetraacetic acid, sodium hydroxide, as well as sonication, heating, and washing with distilled water and/or complexation/treatment with ionic resins, were performed to recover these polysaccharides from the surfaces of microalgal cells [12,60,61].
Furthermore, the method used to break EPS and cell wall interactions must not promote cell lysis to avoid contamination by intracellular compounds and compromise the entire EPS recovery process [60,61]. Thus, some chemical agents such as formaldehyde and glutaraldehyde were used to protect the microalgal cell from lysis during EPS isolation [60,61]. These fixing agents chemically react with hydroxyl, sulfhydryl, carbonyl, or amino groups present in microalgae cell membranes and prevent cell lysis during EPS extraction. However, they can compromise the method if they react with the extracted EPS [12,60,61]. The washing of microalgae cells with water demands temperature (30–95 °C) and time (1–4 h), which can promote cell lysis and consequent contamination with intracellular constituents [60]. In this sense, in most studies, microalgae EPS were isolated without biomass treatment since these treatments add a high cost to the processes [12,60,61].

4. Techniques for Recovery/Identification of Microalgae Polysaccharides

The recovery of intracellular and extracellular compounds from microalgae cultures is the bottleneck to applying this sustainable technology [62]. Recently, several studies have investigated microalgae recovery and sedimentation methods from flocculants as an alternative with high energy efficiency (Table 3) [43].
Several species of algae can act as flocculants, where the process allows the advantage of recycling the medium [67]. The flocculation capacity of autoflocculating microalgae is closely related to EPS secretion [6]. With the optimization of cultivation conditions, the extraction of EPS becomes advantageous since it promotes higher productivity of the biomass and biocompound. In this way, increases in the extraction yield and sustainability of the process are reached. The implementation of recovery and identification protocols varies according to the location of the polysaccharides in the culture [68,69].
Among the classic methods of extracting EPS are centrifugation and microfiltration. These procedures separate the biomass from the EPS-constituted precipitate [20]. After this step, the centrifuged material must be precipitated using methanol, alcohol, ethanol, or isopropanol. With this method, the selective concentration of EPS is possible [70]. As an alternative to the classical methods described, the recovery of EPS can be carried out during the downstream and upstream processes, without the need for chemical additives [12,71]. Methods such as sonication and heating are also used to extract microalgal EPS [72]. Filtration modules from 1 kDa to 500 kDa have been used for the concentration of extracellular compounds present in the culture medium. The ultrafiltration technique can be performed in the following forms: rotating devices, tubular, flat, or spiral plate and hollow fiber, where the liquid medium flows parallel to the ultrafiltration surface and the fraction of interest is permeated through the membrane [12,69].
To increase the performance of filtration techniques, the use of synthetic material is necessary, such as nanocomposite membranes consisting of nanoparticles in a polymeric membrane (SiO2, TiO2) [73]. The identification of EPS can be performed through Fourier transform infrared spectroscopy from functional compound determination [68]. Gas chromatography with mass spectrometry has shown excellent results in the identification of microalgae EPS. Other techniques, such as ion-exchange chromatography, size exclusion chromatography, and affinity chromatography, are widely used to purify and fractionate microalgal polysaccharides [68,69].

5. Application of Microalgal Bioflocculants in Microalgae Harvesting

5.1. Bioflocculation

Bioflocculation is considered a sustainable method that occurs from the aggregation of microalgal cells in the presence of biopolymers synthesized by microorganisms. Biopolymers are mainly composed of extracellular polymeric substances, which contain polysaccharides, proteins, lipids, and nucleic acids in their structure [21,24,74,75].
In addition to the presence of metabolites synthesized by microorganisms, the bioflocculation processes of non-flocculating microalgae can occur in the presence of other microorganisms, such as fungi, bacteria, and other microalgae [21,75]. This process was demonstrated by Guo et al. [7], using supernatant and cell suspension from the autoflocculating Scenedesmus obliquus AS-6-1 culture for the recovery of non-flocculating microalgae biomass. Furthermore, to increase the efficiency of bioflocculation processes, other flocculants such as Al3+ and Fe3+ can be added together with extracellular polysaccharides [7,10,76]. According to Yang et al. [76], the extracellular polymeric substance co-extracted in the Al3+ recovery process after the primary flocculation step contributed to the clotting process of Scenedesmus acuminatus. There was an increase in the process’ efficiency when extracellular substances (≥0.430 mg L−1) were added.
Bioflocculation has been considered a promising strategy for cost reduction in the recovery of microalgal biomass. Among the advantages of this method, there is the absence of chemical flocculants, ease of operation, and an ecologically correct and sustainable approach [24].

5.2. Autoflocculation

Unlike bioflocculation processes, autoflocculation can occur naturally from cell adhesion and aggregation. The autoflocculation of microalgae cells is a phenomenon caused by the secretion of flocculating substances (e.g., glycosides or polysaccharides) which adhere to the microalgal cells. Under alkalinity conditions, autoflocculation occurs from positive precipitates formed by calcium and magnesium ions that neutralize the negative charge of microalgal cells. The other mechanism is related to the EPS produced by microalgae during their physiological activities, which induce flocculation [74]. The autoflocculation process is dependent on the cellular characteristics of the microalgae and other factors such as the composition of available nutrients (e.g., the concentration of Ca, Mg, N, and P), type and concentration of precipitates formed, and pH value [21,24,74,77]. Some autoflocculating species have been reported, such as Scenedesmus rubescens SX [68], Scenedesmus obliquus, Chlorella vulgaris, Ettlia texensis, Ankistrodesmus falcatus [78], and Neocystis mucosa SX [60], among others. Although the mechanisms of autoflocculation are still not well understood, it has been shown that extracellular polymeric substances can influence the autoflocculating capacity of microalgae [21,77]. According to Wan et al. [79], autoflocculation can occur when flocculants produced by the microalgae neutralize charges, forming bridges or patching adjacent cells. Additionally, the hydroxyl and carboxyl groups in the polysaccharide are strongly related to microalgae flocculation. They serve as binding sites during this process. These characteristics were demonstrated in studies by Alam et al. [9], Guo et al. [7], and Lv et al. [60].
Some microalgae produce extracellular polymeric substances in significant amounts during physiological activities, especially at the end of the growth phase when the extracellular polymer acts as a flocculant [74,80]. In these cases, the parameters used in cultivation tend to influence this process, as they affect the production and composition of extracellular polymeric substances [81].
Guo et al. [7] determined that the autoflocculant activity of Scenedesmus obliquus AS-6-1 occurred until the end of the exponential phase and increased with the time of cultivation and the cell concentration of the medium. Autoflocculation occurred from the presence of extracellular biopolymers, which formed a membrane on the cell surface, forming aggregates and sedimenting. Alam et al. [9] studied the spontaneous flocculation of Chlorella vulgaris JSC-7 and the addition of medium from this strain in non-flocculent microalgae. According to the authors, spontaneous microalgae flocculation was associated with an extracellular polysaccharide composed of glucose, mannose, and galactose. Chlorella vulgaris JSC-7 was also able to improve the biomass recovery of the other microalgae tested. In another study, polymeric substances synthesized by Chlorella vulgaris (FACHB-31) and bound to the cell were responsible for increasing autoflocculation. The production of polymeric substances was influenced by glycine added in the medium with light intensity and mixing time. As the concentration of polymeric substances is higher at the end of the cultivation, this period was also responsible for the higher solid concentration rates achieved in the flocculation. However, cultivation time is a parameter that must be considered, as it can increase the costs of harvesting microalgae [80]. Table 4 presents some studies on the production and potential application of extracellular polymeric substances in microalgae harvesting. According to Ummalyma et al. [24], more studies are needed to understand the mechanisms involved in microalgae autoflocculation. The development of research based on the mechanisms of autoflocculation will contribute to cost reduction, ensuring sustainability in downstream processes.

6. Other Applications of Microalgal EPS

EPS produced by microalgae have specific structural and physicochemical characteristics that allow industrial and environmental application (Figure 1). The use of these biopolymers as biosurfactants and heavy metal biosorbents is an innovation in environmental biotechnology. These approaches are economically and ecologically sound strategies for reducing environmental pollution [85]. EPS are also crucial for biological soil crust (biofilm) development. This application reduces water infiltration into the soil by inducing surface sealing and clogging of the pores. Therefore, there is an increase in the availability of nutrients and improvement in the soil’s aggregate stability [86,87]. In addition, the different biological activities presented by EPS, such as antiviral and antibacterial [88], antioxidant [89], anti-inflammatory [90], immunomodulatory [91], and anticancer, indicate the potential of these compounds for application in various sectors such as food, cosmetics, pharmaceuticals, and biomaterials [14,20].

7. Conclusions

Microalgae exhibit rapid growth to produce metabolites under specific cultivation conditions, contributing to a more ecological approach to biomass and EPS production. In addition, to improve the economic competitiveness of innovative products derived from microalgae, industries must seek in scientific research the effectiveness and advantages of using these biotechnological processes. Microalgal EPS have been explored in the field of flocculation due to the need for new products obtained from sustainable alternatives to petroleum-based compounds. The main future challenges in the bioremediation sector will be EPS production on an industrial scale. In addition, the cost reduction of the identification and recovery processes of these biopolymers also deserves further investigation. However, the structural diversity of EPS produced by microalgae provides different properties that imply alternative and integrative applications. Moreover, EPS have antioxidant, anti-inflammatory, anticancer, antiviral, antimicrobial, and immunomodulatory activities, which boost the development of natural pharmaceuticals and nutraceuticals.

Author Contributions

Conceptualization, J.B.M., S.G.K., P.Q.M.B., A.P.A.C., M.Z. and J.L.V.S.; writing—original draft preparation, J.B.M., S.G.K., P.Q.M.B., A.P.A.C., M.Z. and J.L.V.S.; writing—review and editing, J.B.M., S.G.K., P.Q.M.B., A.P.A.C., M.Z. and J.L.V.S.; visualization, J.B.M., S.G.K., P.Q.M.B., A.P.A.C., M.Z., J.L.V.S., J.A.V.C. and M.G.M.; supervision, J.B.M., J.A.V.C. and M.G.M.; project administration, J.B.M., J.A.V.C. and M.G.M.; funding acquisition, J.B.M., J.A.V.C. and M.G.M. All authors have read and agreed to the published version of the manuscript.

Funding

This study was financed in part by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior—Brasil (CAPES)—Finance Code 001. This research was developed within the scope of the Capes-PrInt Program (Process # 88887.310848/2018-00).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available on request from the corresponding author.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Gaignard, C.; Laroche, C.; Pierre, G.; Dubessay, P.; Delattre, C.; Gardarin, C.; Gourvil, P.; Probert, I.; Dubuffet, A.; Michaud, P. Screening of marine microalgae: Investigation of new exopolysaccharide producers. Algal Res. 2019, 44, 101711. [Google Scholar] [CrossRef]
  2. Bezerra, P.Q.M.; Moraes, L.; Cardoso, L.G.; Druzian, J.I.; Morais, M.G.; Nunes, I.L.; Costa, J.A.V. Spirulina sp. LEB 18 cultivation in seawater and reduced nutrients: Bioprocess strategy for increasing carbohydrates in biomass. Bioresour. Technol. 2020, 316, 123883. [Google Scholar] [CrossRef] [PubMed]
  3. Bezerra, P.Q.M.; Moraes, L.; Silva, T.N.M.; Cardoso, L.G.; Druzian, J.I.; Morais, M.G.; Nunes, I.L.; Costa, J.A.V. Innovative application of brackish groundwater without the addition of nutrients in the cultivation of Spirulina and Chlorella for carbohydrate and lipid production. Bioresour. Technol. 2022, 345, 126543. [Google Scholar] [CrossRef] [PubMed]
  4. Borowitzka, M.A. High-value products from microalgae—Their development and commercialisation. J. Appl. Phycol. 2013, 25, 743–756. [Google Scholar] [CrossRef]
  5. Cuevas-Castillo, G.A.; Navarro-Pineda, F.S.; Baz Rodríguez, S.A.; Sacramento Rivero, J.C. Advances on the processing of microalgal biomass for energy-driven biorefineries. Renew. Sustain. Energy Rev. 2020, 125, 109606. [Google Scholar] [CrossRef]
  6. Salim, S.; Kosterink, N.R.; Tchetkoua Wacka, N.D.; Vermuë, M.H.; Wijffels, R.H. Mechanism behind autoflocculation of unicellular green microalgae Ettlia texensis. J. Biotechnol. 2014, 174, 34–38. [Google Scholar] [CrossRef]
  7. Guo, S.L.; Zhao, X.Q.; Wan, C.; Huang, Z.Y.; Yang, Y.L.; Asraful Alam, M.; Ho, S.H.; Bai, F.W.; Chang, J.S. Characterization of flocculating agent from the self-flocculating microalga Scenedesmus obliquus AS-6-1 for efficient biomass harvest. Bioresour. Technol. 2013, 145, 285–289. [Google Scholar] [CrossRef]
  8. Salim, S.; Bosma, R.; Vermuë, M.H.; Wijffels, R.H. Harvesting of microalgae by bio-flocculation. J. Appl. Phycol. 2011, 23, 849–855. [Google Scholar] [CrossRef] [Green Version]
  9. Alam, M.A.; Wan, C.; Guo, S.L.; Zhao, X.Q.; Huang, Z.Y.; Yang, Y.L.; Chang, J.S.; Bai, F.W. Characterization of the flocculating agent from the spontaneously flocculating microalga Chlorella vulgaris JSC-7. J. Biosci. Bioeng. 2014, 118, 29–33. [Google Scholar] [CrossRef]
  10. Yang, L.; Zhang, H.; Cheng, S.; Zhang, W.; Zhang, X. Enhanced Microalgal Harvesting Using Microalgae-Derived Extracellular Polymeric Substance as Flocculation Aid. ACS Sustain. Chem. Eng. 2020, 8, 4069–4075. [Google Scholar] [CrossRef]
  11. Aljuboori, A.H.R.; Uemura, Y.; Thanh, N.T. Flocculation and mechanism of self-flocculating lipid producer microalga Scenedesmus quadricauda for biomass harvesting. Biomass Bioenergy 2016, 93, 38–42. [Google Scholar] [CrossRef]
  12. Delattre, C.; Pierre, G.; Laroche, C.; Michaud, P. Production, extraction and characterization of microalgal and cyanobacterial exopolysaccharides. Biotechnol. Adv. 2016, 34, 1159–1179. [Google Scholar] [CrossRef] [PubMed]
  13. Boonchai, R.; Kaewsuk, J.; Seo, G. Effect of nutrient starvation on nutrient uptake and extracellular polymeric substance for microalgae cultivation and separation. Desalin. Water Treat. 2014, 55, 360–367. [Google Scholar] [CrossRef]
  14. Costa, J.A.V.; Lucas, B.F.; Alvarenga, A.G.P.; Moreira, J.B.; de Morais, M.G. Microalgae Polysaccharides: An Overview of Production, Characterization, and Potential Applications. Polysaccharides 2021, 2, 759–772. [Google Scholar] [CrossRef]
  15. Bae, S.Y.; Yim, J.H.; Lee, H.K.; Pyo, S. Activation of murine peritoneal macrophages by sulfated exopolysaccharide from marine microalga Gyrodinium impudicum (strain KG03): Involvement of the NF-κB and JNK pathway. Int. Immunopharmacol. 2006, 6, 473–484. [Google Scholar] [CrossRef]
  16. Chen, B.; You, W.; Huang, J.; Yu, Y.; Chen, W. Isolation and antioxidant property of the extracellular polysaccharide from Rhodella reticulata. World J. Microbiol. Biotechnol. 2009, 26, 833–840. [Google Scholar] [CrossRef]
  17. Guzmán, S.; Gato, A.; Lamela, M.; Freire-Garabal, M.; Calleja, J.M. Anti-inflammatory and immunomodulatory activities of polysaccharide from Chlorella stigmatophora and Phaeodactylum tricornutum. Phytother. Res. 2003, 17, 665–670. [Google Scholar] [CrossRef]
  18. Majdoub, H.; Mansour, M.B.; Chaubet, F.; Roudesli, M.S.; Maaroufi, R.M. Anticoagulant activity of a sulfated polysaccharide from the green alga Arthrospira platensis. Biochim. Biophys. Acta 2009, 1790, 1377–1381. [Google Scholar] [CrossRef]
  19. Najdenski, H.M.; Gigova, L.G.; Iliev, I.I.; Pilarski, P.S.; Lukavský, J.; Tsvetkova, I.V.; Ninova, M.S.; Kussovski, V.K. Antibacterial and antifungal activities of selected microalgae and cyanobacteria. Int. J. Food Sci. Technol. 2013, 48, 1533–1540. [Google Scholar] [CrossRef]
  20. Zhang, J.; Liu, L.; Chen, F. Production and characterization of exopolysaccharides from Chlorella zofingiensis and Chlorella vulgaris with anti-colorectal cancer activity. Int. J. Biol. Macromol. 2019, 134, 976–983. [Google Scholar] [CrossRef]
  21. Babiak, W.; Krzemińska, I. Extracellular Polymeric Substances (EPS) as Microalgal Bioproducts: A Review of Factors Affecting EPS Synthesis and Application in Flocculation Processes. Energies 2021, 14, 4007. [Google Scholar] [CrossRef]
  22. Li, Y.; Chen, Y.F.; Chen, P.; Min, M.; Zhou, W.; Martinez, B.; Zhu, J.; Ruan, R. Characterization of a microalga Chlorella sp. well adapted to highly concentrated municipal wastewater for nutrient removal and biodiesel production. Bioresour. Technol. 2011, 102, 5138–5144. [Google Scholar] [CrossRef] [PubMed]
  23. Chandra, R.; Iqbal, H.M.N.; Vishal, G.; Lee, H.S.; Nagra, S. Algal biorefinery: A sustainable approach to valorize algal-based biomass towards multiple product recovery. Bioresour. Technol. 2019, 278, 346–359. [Google Scholar] [CrossRef] [PubMed]
  24. Ummalyma, S.B.; Gnansounou, E.; Sukumaran, R.K.; Sindhu, R.; Pandey, A.; Sahoo, D. Bioflocculation: An alternative strategy for harvesting of microalgae—An overview. Bioresour. Technol. 2017, 242, 227–235. [Google Scholar] [CrossRef]
  25. 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]
  26. Shi, Y.; Huang, J.; Zeng, G.; Gu, Y.; Chen, Y.; Hu, Y.; Tang, B.; Zhou, J.; Yang, Y.; Shi, L. Exploiting extracellular polymeric substances (EPS) controlling strategies for performance enhancement of biological wastewater treatments: An overview. Chemosphere 2017, 180, 396–411. [Google Scholar] [CrossRef]
  27. Naveed, S.; Li, C.; Lu, X.; Chen, S.; Yin, B.; Zhang, C.; Ge, Y. Microalgal extracellular polymeric substances and their interactions with metal(loid)s: A review. Crit. Rev. Environ. Sci. Technol. 2019, 49, 1769–1802. [Google Scholar] [CrossRef]
  28. Xiao, R.; Zheng, Y. Overview of microalgal extracellular polymeric substances (EPS) and their applications. Biotechnol. Adv. 2016, 34, 1225–1244. [Google Scholar] [CrossRef]
  29. Kumar, D.; Kvíderová, J.; Kaštánek, P.; Lukavský, J. The green alga Dictyosphaerium chlorelloides biomass and polysaccharides production determined using cultivation in crossed gradients of temperature and light. Eng. Life Sci. 2017, 17, 1030–1038. [Google Scholar] [CrossRef] [Green Version]
  30. Li, W.; Xu, X.; Fujibayashi, M.; Niu, Q.; Tanaka, N.; Nishimura, O. Response of microalgae to elevated CO2 and temperature: Impact of climate change on freshwater ecosystems. Environ. Sci. Pollut. Res. Int. 2016, 23, 19847–19860. [Google Scholar] [CrossRef]
  31. Zhou, Y.; Xia, S.; Zhang, J.; Nguyen, B.T.; Zhang, Z. Insight into the influences of pH value on Pb(II) removal by the biopolymer extracted from activated sludge. Chem. Eng. J. 2017, 308, 1098–1104. [Google Scholar] [CrossRef]
  32. Raungsomboon, S.; Chidthaisong, A.; Bunnag, B.; Inthorn, D.; Harvey, N.W. Production, composition and Pb2+ adsorption characteristics of capsular polysaccharides extracted from a cyanobacterium Gloeocapsa gelatinosa. Water Res. 2006, 40, 3759–3766. [Google Scholar] [CrossRef] [PubMed]
  33. Patwal, T.; Baranwal, M. Scenedesmus acutus extracellular polysaccharides produced under increased concentration of sulphur and phosphorus exhibited enhanced proliferation of peripheral blood mononuclear cells. 3 Biotech 2021, 11, 171. [Google Scholar] [CrossRef] [PubMed]
  34. Liu, L.; Pohnert, G.; Wei, D. Extracellular metabolites from industrial microalgae and their biotechnological potential. Mar. Drugs 2016, 14, 191. [Google Scholar] [CrossRef] [PubMed]
  35. Shen, L.; Li, Z.; Wang, J.; Liu, A.; Li, Z.; Yu, R.; Wu, X.; Liu, Y.; Li, J.; Zeng, W. Characterization of extracellular polysaccharide/protein contents during the adsorption of Cd(II) by Synechocystis sp. PCC6803. Environ. Sci. Pollut. Res. Int. 2018, 25, 20713–20722. [Google Scholar] [CrossRef]
  36. Wei, L.; Li, Y.; Noguera, D.R.; Zhao, N.; Song, Y.; Ding, J.; Zhao, Q.; Cui, F. Adsorption of Cu2+ and Zn2+ by extracellular polymeric substances (EPS) in different sludges: Effect of EPS fractional polarity on binding mechanism. J. Hazard. Mater. 2017, 321, 473–483. [Google Scholar] [CrossRef]
  37. Ozturk, S.; Aslim, B.; Suludere, Z. Cadmium(II) sequestration characteristics by two isolates of Synechocystis sp. in terms of exopolysaccharide (EPS) production and monomer composition. Bioresour. Technol. 2010, 101, 9742–9748. [Google Scholar] [CrossRef]
  38. Pereira, S.; Zille, A.; Micheletti, E.; Moradas-Ferreira, P.; De Philippis, R.; Tamagnini, P. Complexity of cyanobacterial exopolysaccharides: Composition, structures, inducing factors and putative genes involved in their biosynthesis and assembly. FEMS Microbiol. Rev. 2009, 33, 917–941. [Google Scholar] [CrossRef]
  39. Miao, A.J.; Schwehr, K.A.; Xu, C.; Zhang, S.J.; Luo, Z.; Quigg, A.; Santschi, P.H. The algal toxicity of silver engineered nanoparticles and detoxification by exopolymeric substances. Environ. Pollut. 2009, 157, 3034–3041. [Google Scholar] [CrossRef]
  40. Koçer, A.T.; İnan, B.; Kaptan Usul, S.; Özçimen, D.; Yılmaz, M.T.; Işıldak, İ. Exopolysaccharides from microalgae: Production, characterization, optimization and techno-economic assessment. Braz. J. Microbiol. 2021, 52, 1779–1790. [Google Scholar] [CrossRef]
  41. Surendhiran, D.; Vijay, M. Influence of bioflocculation parameters on harvesting Chlorella salina and its optimization using response surface methodology. J. Environ. Chem. Eng. 2013, 4, 1051–1056. [Google Scholar] [CrossRef]
  42. Nguyen, L.N.; Vu, H.P.; Fu, Q.; Johir, M.A.H.; Ibrahim, I.; Mofijur, M.; Labeeuw, L.; Pernice, M.; Ralph, P.J.; Nghiem, L.D. Synthesis and evaluation of cationic polyacrylamide and polyacrylate flocculants for harvesting freshwater and marine microalgae. Chem. Eng. J. 2022, 433, 133623. [Google Scholar] [CrossRef]
  43. Choi, O.K.; Hendren, Z.; Kim, G.D.; Dong, D.; Lee, J.W. Influence of activated sludge derived-extracellular polymeric substance (ASD-EPS) as bio-flocculation of microalgae for biofuel recovery. Algal Res. 2020, 45, 101736. [Google Scholar] [CrossRef]
  44. Pandey, A.; Pathak, V.V.; Kothari, R.; Black, P.N.; Tyagi, V.V. Experimental studies on zeta potential of flocculants for harvesting of algae. J. Environ. Manag. 2019, 231, 562–569. [Google Scholar] [CrossRef] [PubMed]
  45. Khan, S.; Naushad, M.; Iqbal, J.; Bathula, C.; Sharma, G. Production and harvesting of microalgae and an efficient operational approach to biofuel production for a sustainable environment. Fuel 2022, 311, 122543. [Google Scholar] [CrossRef]
  46. Ghazvini, M.; Kavosi, M.; Sharma, R.; Kim, M. A review on mechanical-based microalgae harvesting methods for biofuel production. Biomass Bioenergy 2022, 158, 106348. [Google Scholar] [CrossRef]
  47. Valdovinos-García, E.M.; Barajas-Fernández, J.; Olán-Acosta, M.d.l.A.; Petriz-Prieto, M.A.; Guzmán-López, A.; Bravo-Sánchez, M.G. Techno-Economic Study of CO2 Capture of a Thermoelectric Plant Using Microalgae (Chlorella vulgaris) for Production of Feedstock for Bioenergy. Energies 2020, 13, 413. [Google Scholar] [CrossRef] [Green Version]
  48. Xu, K.; Zou, X.; Chang, W.; Qu, Y.; Li, Y. Microalgae harvesting technique using ballasted flotation: A review. Set. Purif. Tecnol. 2021, 276, 119439. [Google Scholar] [CrossRef]
  49. Parmentier, D.; Manhaeghe, D.; Baccini, L.; Meirhaeghe, R.V.; Rousseau, D.P.L.; Hulle, S.V. A new reactor design for harvesting algae through electrocoagulation-flotation in a continuous mode. Algal Res. 2020, 47, 101828. [Google Scholar] [CrossRef]
  50. Min, K.H.; Kim, D.H.; Ki, M.-R.; Pack, S.P. Recent progress in flocculation, dewatering, and drying technologies for microalgae utilization: Scalable and low-cost harvesting process development. Bioresour. Technol. 2022, 344, 126404. [Google Scholar] [CrossRef]
  51. Zhu, L.; Hu, T.; Li, S.; Nugroho, Y.K.; Li, B.; Cao, J.; Show, P.-L.; Hiltunen, E. Effects of operating parameters on algae Chlorella vulgaris biomass harvesting and lipid extraction using metal sulfates as flocculants. Biomass Bioenergy 2020, 132, 105433. [Google Scholar] [CrossRef]
  52. Mayers, J.J.; Landels, A.R.; Allen, M.J.; Albers, E. An energy and resource efficient alkaline flocculation and sedimentation process for harvesting of Chromochloris zofingiensis biomass. Bioresour. Technol. Rep. 2020, 9, 100358. [Google Scholar] [CrossRef]
  53. Smith, B.T.; Davis, R.H. Sedimentation of algae flocculated using naturally-available, magnesium-based flocculants. Algal Res. 2012, 1, 32–39. [Google Scholar] [CrossRef]
  54. Yang, F.; Xiang, W.; Fan, J.; Wu, H.; Li, T.; Long, L. High pH-induced flocculation of marine Chlorella sp. for biofuel production. J. Appl. Phycol. 2016, 28, 747–756. [Google Scholar] [CrossRef]
  55. Vandamme, D.; Beuckels, A.; Markou, G.; Foubert, I.; Muylaert, K. Reversible Flocculation of Microalgae using Magnesium Hydroxide. Bioenergy Res. 2015, 8, 716–725. [Google Scholar] [CrossRef] [Green Version]
  56. Mayers, J.J.; Vaiciulyte, S.; Malmhäll-Bah, E.; Alcaide-Sancho, J.; Ewald, S.; Godhe, A.; Ekendahl, S.; Albers, E. Identifying a marine microalgae with high carbohydrate productivities under stress and potential for efficient flocculation. Algal Res. 2018, 31, 430–442. [Google Scholar] [CrossRef]
  57. Cancela, Á.; Sánchez, Á.; Álvarez, X.; Jiménez, A.; Ortiz, L.; Valero, E.; Varela, P. Pellets valorization of waste biomass harvested by coagulation of freshwater algae. Bioresour. Technol. 2016, 204, 152–156. [Google Scholar] [CrossRef]
  58. Zhu, L.; Li, Z.; Hiltunen, E. Microalgae Chlorella vulgaris biomass harvesting by natural flocculant: Effects on biomass sedimentation, spent medium recycling and lipid extraction. Biotechnol. Biofuels 2018, 11, 183. [Google Scholar] [CrossRef]
  59. Vu, H.P.; Nguyen, L.N.; Emmerton, B.; Wang, Q.; Ralph, P.J.; Nghiem, L.D. Factors governing microalgae harvesting efficiency by flocculation using cationic polymers. Bioresour. Technol. 2021, 340, 125669. [Google Scholar] [CrossRef]
  60. Lv, J.; Zhao, F.; Feng, J.; Liu, Q.; Nan, F.; Xie, S. Extraction of extracellular polymeric substances (EPS) from a newly isolated self-flocculating microalga Neocystis mucosa SX with different methods. Algal Res. 2019, 40, 101479. [Google Scholar] [CrossRef]
  61. Huang, R.; He, Q.; Ma, J.; Ma, C.; Xu, Y.; Song, J.; Sun, L.; Wu, Z.; Huangfu, X. Quantitative assessment of extraction methods for bound extracellular polymeric substances (B-EPSs) produced by Microcystis sp. and Scenedesmus sp. Algal Res. 2021, 56, 102289. [Google Scholar] [CrossRef]
  62. Kong, W.; Yang, S.; Wang, H.; Huo, H.; Guo, B.; Liu, N.; Zhang, A.; Niu, S. Regulation of biomass, pigments, and lipid production by Chlorella vulgaris 31 through controlling trophic modes and carbon sources. J. Appl. Phycol. 2020, 32, 1569–1579. [Google Scholar] [CrossRef]
  63. Balti, R.; Le Balc’h, R.; Brodu, N.; Gilbert, M.; Le Gouic, B.; Le Gall, S.; Sinquin, C.; Massé, A. Concentration and purification of Porphyridium cruentum exopolysaccharides by membrane filtration at various cross-flow velocities. Process Biochem. 2018, 74, 175–184. [Google Scholar] [CrossRef] [Green Version]
  64. García-Cubero, R.; Wang, W.; Martín, J.; Bermejo, E.; Sijtsma, L.; Togtema, A.; Barbosa, M.J.; Kleinegris, D.M.M. Milking exopolysaccharides from Botryococcus braunii CCALA778 by membrane filtration. Algal Res. 2018, 34, 175–181. [Google Scholar] [CrossRef]
  65. Zaouk, L.; Massé, A.; Bourseau, P.; Taha, S.; Rabiller-Baudry, M.; Jubeau, S.; Teychené, B.; Pruvost, J.; Jaouen, P. Filterability of exopolysaccharides solutions from the red microalga Porphyridium cruentum by tangential filtration on a polymeric membrane. Environ. Technol. 2020, 41, 1167–1184. [Google Scholar] [CrossRef] [PubMed]
  66. Gaignard, C.; Macao, V.; Gardarin, C.; Rihouey, C.; Picton, L.; Michaud, P.; Laroche, C. The red microalga Flintiella sanguinaria as a new exopolysaccharide producer. J. Appl. Phycol. 2018, 30, 2803–2814. [Google Scholar] [CrossRef]
  67. Ummalyma, S.B.; Supriya, R.D.; Sindhu, R.; Binod, P.; Nair, R.B.; Pandey, A.; Gnansounou, E. Biological pretreatment of lignocellulosic biomass—Current trends and future perspectives. In Second and Third Generation of Feedstocks: The Evolution of Biofuels; Basile, A., Dalena, F., Eds.; Elsevier: Amsterdam, The Netherlands, 2019; pp. 197–212. [Google Scholar]
  68. Lv, J.; Guo, B.; Feng, J.; Liu, Q.; Nan, F.; Liu, X.; Xie, S. Integration of wastewater treatment and flocculation for harvesting biomass for lipid production by a newly isolated self-flocculating microalga Scenedesmus rubescens SX. J. Clean. Prod. 2019, 240, 118211. [Google Scholar] [CrossRef]
  69. Zhu, L.; Zhou, J.; Lv, M.; Yu, H.; Zhao, H.; Xu, X. Specific component comparison of extracellular polymeric substances (EPS) in flocs and granular sludge using EEM and SDS-PAGE. Chemosphere 2015, 121, 26–32. [Google Scholar] [CrossRef]
  70. De Brouwer, J.F.C.; Wolfstein, K.; Ruddy, G.K.; Jones, T.E.R.; Stal, L.J. Biogenic stabilization of intertidal sediments: The importance of extracellular polymeric substances produced by benthic diatoms. Microb. Ecol. 2005, 49, 501–512. [Google Scholar] [CrossRef]
  71. Charcosset, C. Membrane processes in biotechnology: An overview. Biotechnol. Adv. 2006, 24, 482–492. [Google Scholar] [CrossRef]
  72. Mishra, A.; Jha, B. Isolation and characterization of extracellular polymeric substances from micro-algae Dunaliella salina under salt stress. Bioresour. Technol. 2009, 100, 3382–3386. [Google Scholar] [CrossRef] [PubMed]
  73. Jhaveri, J.H.; Murthy, Z.V.P. A comprehensive review on anti-fouling nanocomposite membranes for pressure driven membrane separation processes. Desalination 2016, 379, 137–154. [Google Scholar] [CrossRef]
  74. Li, T.; Hu, J.; Zhu, L. Self-flocculation as an efficient method to harvest microalgae: A mini-review. Water 2021, 13, 2585. [Google Scholar] [CrossRef]
  75. Barros, A.I.; Gonçalves, A.L.; Simões, M.; Pires, J.C.M. Harvesting techniques applied to microalgae: A review. Renew. Sustain. Energy Rev. 2015, 41, 1489–1500. [Google Scholar] [CrossRef] [Green Version]
  76. Yang, L.; Wang, L.; Zhang, H.; Li, C.; Zhang, X.; Hu, Q. A novel low cost microalgal harvesting technique with coagulant recovery and recycling. Bioresour. Technol. 2018, 266, 343–348. [Google Scholar] [CrossRef] [PubMed]
  77. Ogbonna, C.N.; Nwoba, E.G. Bio-based flocculants for sustainable harvesting of microalgae for biofuel production. A review. Renew. Sustain. Energy Rev. 2021, 139, 110690. [Google Scholar] [CrossRef]
  78. Salim, S.; Vermuë, M.H.; Wijffels, R.H. Ratio between autoflocculating and target microalgae affects the energy-efficient harvesting by bio-flocculation. Bioresour. Technol. 2012, 118, 49–55. [Google Scholar] [CrossRef]
  79. Wan, C.; Alam, M.A.; Zhao, X.-Q.; Zhang, X.-Y.; Guo, S.-L.; Ho, S.-H.; Chang, J.-S.; Bai, F.-W. Current progress and future prospect of microalgal biomass harvest using various flocculation technologies. Bioresour. Technol. 2015, 184, 251–257. [Google Scholar] [CrossRef]
  80. Shen, Y.; Fan, Z.; Chen, C.; Xu, X. An auto-flocculation strategy for Chlorella vulgaris. Biotechnol. Lett. 2015, 37, 75–80. [Google Scholar] [CrossRef]
  81. González-Fernández, C.; Ballesteros, M. Microalgae autoflocculation: An alternative to high-energy consuming harvesting methods. J. Appl. Phycol. 2013, 25, 991–999. [Google Scholar] [CrossRef]
  82. Jesus, C.S.d.; Jesus Assis de, D.; Rodriguez, M.B.; Menezes Filho, J.A.; Costa, J.A.V.; de Souza Ferreira, E.; Druzian, J.I. Pilot-scale isolation and characterization of extracellular polymeric substances (EPS) from cell-free medium of Spirulina sp. LEB-18 cultures under outdoor conditions. Int. J. Biol. Macromol. 2019, 124, 1106–1114. [Google Scholar] [CrossRef] [PubMed]
  83. Zhu, C.; Chen, C.; Zhao, L.; Zhang, Y.; Yang, J.; Song, L.; Yang, S. Bioflocculant produced by Chlamydomonas reinhardtii. J. Appl. Phycol. 2012, 24, 1245–1251. [Google Scholar] [CrossRef]
  84. Zhao, F.; Xiao, J.; Ding, W.; Cui, N.; Yu, X.; Xu, J.W.; Li, T.; Zhao, P. An effective method for harvesting of microalga: Coculture-induced self-flocculation. J. Taiwan Inst. Chem. Eng. 2019, 100, 117–126. [Google Scholar] [CrossRef]
  85. Gondi, R.; Kavitha, S.; Yukesh Kannah, R.; Parthiba Karthikeyan, O.; Kumar, G.; Kumar Tyagi, V.; Rajesh Banu, J. Algal-based system for removal of emerging pollutants from wastewater: A review. Bioresour. Technol. 2022, 344, 126245. [Google Scholar] [CrossRef] [PubMed]
  86. Wu, Y.; Rao, B.; Wu, P.; Liu, Y.; Li, G.; Li, D. Development of artificially induced biological soil crusts in fields and their effects on top soil. Plant Soil 2013, 370, 115–124. [Google Scholar] [CrossRef]
  87. Tiwari, O.N.; Bhunia, B.; Mondal, A.; Gopikrishna, K.; Indrama, T. System metabolic engineering of exopolysaccharide-producing cyanobacteria in soil rehabilitation by inducing the formation of biological soil crusts: A review. J. Clean. Prod. 2019, 211, 70–82. [Google Scholar] [CrossRef]
  88. Raposo, M.F.D.J.; De Morais, A.M.M.B.; De Morais, R.M.S.C. Influence of sulphate on the composition and antibacterial and antiviral properties of the exopolysaccharide from Porphyridium cruentum. Life Sci. 2014, 101, 56–63. [Google Scholar] [CrossRef]
  89. Tiwari, O.N.; Mondal, A.; Bhunia, B.; Bandyopadhyay, T.k.; Jaladi, P.; Oinam, G.; Indrama, T. Purification, characterization and biotechnological potential of new exopolysaccharide polymers produced by cyanobacterium Anabaena sp. CCC 745. Polymer 2019, 178, 121695. [Google Scholar] [CrossRef]
  90. Alvarez, X.; Alves, A.; Ribeiro, M.P.; Lazzari, M.; Coutinho, P.; Otero, A. Biochemical characterization of Nostoc sp. exopolysaccharides and evaluation of potential use in wound healing. Carbohydr. Polym. 2021, 254, 117303. [Google Scholar] [CrossRef]
  91. Uhliariková, I.; Šutovská, M.; Barboríková, J.; Molitorisová, M.; Kim, H.J.; Park, Y.I.; Matulová, M.; Lukavský, J.; Hromadková, Z.; Capek, P. Structural characteristics and biological effects of exopolysaccharide produced by cyanobacterium Nostoc sp. Int. J. Biol. Macromol. 2020, 160, 364–371. [Google Scholar] [CrossRef]
Figure 1. Interactions and structure of EPS with microalgae cultivation and their applications.
Figure 1. Interactions and structure of EPS with microalgae cultivation and their applications.
Polysaccharides 03 00015 g001
Table 1. Technoeconomic analysis of microalgae biomass recovery (adapted from Valdovinos-García et al. [47]).
Table 1. Technoeconomic analysis of microalgae biomass recovery (adapted from Valdovinos-García et al. [47]).
Drying ProcessHarvest MethodResponses
Electrical Power (kWh Year−1) *Unit Production Cost (US $ kg−1) *Operating Cost (US $ Year−1) *Biomass Production (kg Year−1)
Spray dryingAuto-flocculation followed by vacuum filtering13,9881.2561,00048,145.6–53,495.11
Auto-flocculation followed by plate press filter87381.1958,000
Drum dryerAuto-flocculation followed by plate press filter82970.8542,00048,145.6–53,495.11
Auto-flocculation followed by vacuum filtering13,5970.9145,000
Spray dryingAdding an iron salt followed by vacuum filtering13,9871.2159,00048,145.6–54,306.51
Adding an iron salt followed by plate press filter87371.1556,000
Drum dryerAdding an iron salt followed by plate press filter82960.8140,00048,145.6–54,306.51
Adding an iron salt followed by vacuum filtering13,5460.8743,000
Spray dryingFlocculation with chitosan followed by vacuum filtering13,9881.2662,00048,145.6–54,306.51
Flocculation with chitosan followed by plate press filter87281.2059,000
Drum dryerFlocculation with chitosan followed by plate press filter82970.8643,00048,145.6–54,306.51
Flocculation with chitosan followed by vacuum filtering25,5970.9246,000
* The values presented are the sum of respective data obtained from harvesting and drying processes.
Table 2. Comparative yield of microalgae flocculation/coagulation using different substances.
Table 2. Comparative yield of microalgae flocculation/coagulation using different substances.
MicroalgaeRecovery ProcessExperimental ConditionsSubstance Used in Biomass RecoveryProcess YieldReference
Chromochloris zofingiensisFlocculation and alkaline sedimentationBold’s Basal Medium, biomass concentration 0.5, 1.0, 1.5 g L−1, 200 L working volume, air sparged (~10 L min−1), centrifugation 12,000× g for 10 min, pH 7.0NaOH (4.6 and 8 mM) and MgSO4 (6, 8 and 10 mM)Sedimentation yield above 90%[52]
Chlorella vulgaris UTEX 395Flocculation with naturally available magnesium in brackish waterBG-11 medium, biomass concentration 0.3% v v−1, stirring at 700 rpm for 5 min, pH 9.0Mg2+ (0.3 mM) and MgCl2 (9.6 mM)Sedimentation yield of 100 cm h−1[53]
Chlorella marinha sp.Flocculation induced by NaOHF/2 medium Guillard, 5000× g for 5 minNaOH (5 and 7 mM)Flocculation yield of 90%[54]
Chlorella vulgarisReversible flocculationWright’s Cryptophyte medium, 10 L working volume, centrifugation 20 min of stirrung at 250 rpm, pH 8.5Mg (2.5 mM) and NaOH (4 mM)Maximum flocculation efficiency of 90%[55]
Phaeodactylum tricornutumMaximum flocculation efficiency of 73%
Chlorella salinaAlkaline autoflocculationGuillard’s F/2 media with artificial seawater media, optical density of 0.1, 11,500× g for 12 min, pH 8.0NaOH (4 mM)Biomass recoveries greater than 95% efficiency[56]
Scenedesmus sp., Kirchneriella sp., and Microcystis aeruginosaInduced flocculationMedium mixture, 50 L working volume, 200 rpm for 1 min, pH 7.7CaCl2 (20, 60, 120 and 180 mg L−1), NaC6H7O6 (10 and 20 mg L−1) and Tannin (10 and 20 mg L−1).Maximum flocculation efficiency for Tannin 95.35%, sodium alginate 90.49% and, calcium chloride 84.04% [57]
Chlorella vulgarisInduced natural flocculationN8 medium, 1 L working volume, 100 rpm for over 24 h, pH 6.8Chitosan (0.25 g L−1) and aluminum sulfate (2.5 g L−1)Flocculation yield of 90%[58]
Chlorella vulgarisInduced flocculationMLA medium, 350 L working volume, 100% CO2 for 1 min d−1, pH 9.0Cationic polyacrylamide polymer (2 g L−1)Flocculation yield of 97%[59]
Table 3. Techniques for recovering and identifying of microalgal EPS.
Table 3. Techniques for recovering and identifying of microalgal EPS.
TechniquesMicroalgaProcess ConditionsObjectiveResponsesReference
Membrane filtrationPorphyridium cruentumPermeate fluxes of 49.8, 68.9 and 81.9 L h−1 m−2 and 4 bar for, respectively, cross-flow velocities of 2.5, 3.3 and 4.2 m s−1; 49.7 L h−1 m−2.Influence of cross-flow velocities on filtration performances.EPS concentration at 6.3 to 10.4 times reaching from 1.74–2.26 g L−1 (80% (w w−1) recovery).[63]
Membrane filtrationBotryococcus braunii CCALA778Culture flow circulating in 110 cm2 area 0.2 μm microfiltration hollow fiber membrane (GE Healthcare®, CFP-2-E-35MA).Optimization and efficiency of extraction and recovery, ensuring high efficiency without compromising the viability of the culture.Increased EPS productivity by 25% (4 g m−2 d−1). Daily EPS extraction rate of 0.36 g m−2 d−1.[64]
Ultrafiltration (polymeric membrane)Porphyridium cruentumPES 50 kDa flat membrane in full recirculation mode, with permeate flow transmembrane pressure (TMP) curves (0.10–1.06 kg GlcEq m−3), tangential fluid velocity (0.3–1.2 m s−1), and temperature (20 and 40 °C).Parametric study of ultrafiltration of EPS solutions in organic membrane.The concentrated solution of 0.10 kg GlcEq m−3 (moderate fouling, portion of irreversible/reversible fouling was 88 and 12%).[65]
DiafiltrationFlintiella sanguinariaVivaflow ultrafiltration system (Sartorius) and 100 kDa NMWCO membranes.Native EPS extraction.EPS solution was concentrated (volume reduction factor of 5).[66]
High Pressure Anion Exchange Chromatography (HPAEC)Flintiella sanguinariaThe quantification of monosaccharides was achieved by injecting different concentrations of monosaccharides and plotting the response area as a function of concentration.Quantification and identification of native EPS extract.Identification of galactoxylan, with rhamnose and glucuronic acid, low content of sulfate groups (0.6%), and methylated and acetylated compounds (5.1 and 3.2%, w w−1).[66]
Molecular weight (Mw)Chlorella zofingiensisDetermination with gel permeation chromatography and refraction detector (RI), and at a flow rate of 0.5 mL min−1 using 0.2 M sodium nitrate as the mobile phase.Investigation of the physicochemical characteristics of EPS.EPS yields were 208.4 and 364.3 mg L−1 with average molecular weights of 2.66 × 104 and 1.88 × 104 Da, respectively.[20]
Table 4. Production and potential application of microalgal bioflocculants in microalgae harvesting.
Table 4. Production and potential application of microalgal bioflocculants in microalgae harvesting.
MicroalgaDescription of the Experimental SetChemical Composition and Identification of BioflocculantsProduced and Applied Concentration of Bioflocculants Bioflocculant
Application
Reference
Spirulina sp. LEB-18Outdoor cultivation, using a raceway (250 L), Zarrouk medium, under natural light for 30 d (probable stationary phase), and pH 9.8–10.5.Sugars composition: glucose, galactose, fructose, and organic acids were glucuronic, galacturonic and pyruvic.9.5 g L−1 was the highest production of extracellular polymeric substances.Possible application as a bioflocculant and/or other industrial applications.[82]
Scenedesmus obliquus AS-6-1Cultivation to stationary phase, 28 °C, 14/10 h light/dark cycle, 60 mol m−2 s−1.Cell wall-associated polysaccharides.
Monomers consist of glucose, mannose, galactose, rhamnose and fructose.
0.6 mg L−1 of bioflocculant was responsible for 88% of the flocculant activity of Scenedesmus obliquus FSP-3.Bioflocculation of Chlorella vulgaris CNW-11, Scenedesmus obliquus FSP-3 and Nannochloropsis oceanica DUT01.[7]
Chlamydomonas reinhardtiiCultivation at 5 to 25 °C, pH 6–10, 40–60 μmol photons m−2 s−1, 30 d of cultivation.Bioflocculant composition: proteins (42.1% w w−1), carbohydrates (48.3% w w−1), lipids (8.7% w w−1), and nucleic acid (0.01% w w−1).4 mg L−1 of bioflocculant was responsible for 96.6% of the flocculant activity of Chlamydomonas reinhardtii.Microalgae bioflocculation[83]
Ettlia texensisCultivation in a 4 L photobioreactor, batch mode, 24-h lighting, 300 rpm, 26 °C, pH 6.5, and 300 μmol m−2 s−1.Bioflocculants containing mainly glycoproteins patched to the cell surface._Microalgae autoflocculation and bioflocculation.[6]
Desmodesmus sp. ZFY and Monoraphidium sp. QLY-1Microalgae were used in co-culture, cultivated in mixotrophic medium BG-11 + ammonium nitrate and glucose, pH 6.8, 300 mL, 25 °C, 120 rpm, 3500 lux, and 7 d of cultivation (stationary phase).Bioflocculant consisted mainly of polysaccharides and proteins. The levels of polysaccharides in co-culture were 46.53% in substances loosely bound to cells (LB-EPS).Concentration of total extracellular polymeric substances was 368.40 mg L−1.Microalgae bioflocculation.[84]
Scenedesmus acuminatusCultivation performed in 15 L photobioreactors at 25 °C, modified BG-11 medium (NO3 reduction), 180 μmol m−2 s−1, light period 24 h d−1 and pH 6.5–7.0.High (>50 kDa; 35.1%) and low molecular weight (<3 kDa; 46.1%) polymeric substances were identified; being composed of galactose, glucosamine, mannose.3.2 mg g−1 of extracellular polymeric substances were added in the harvesting process together with Al3+ (4.5 mg g−1).Microalgae bioflocculation.[10]
Chlorella vulgaris JSC-7Modified Bold’s Basal Medium with nitrogen supplementation was used, pH 6.9, 28 °C, 13/11 h light/dark cycle and 25 μmol m−2 s−1.The bioflocculant is a cell wall polysaccharide; The monomers consist of glucose, mannose, and galactose.47 mg of the bioflocculant was extracted from 4 L of culture; Addition of 0.5 mg L−1 of the bioflocculant was responsible for >80% of the flocculation of the suspended microalgal cells.Bioflocculation of C. vulgaris CNW11 and Scenedesmus obliquus FSP.[9]
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Moreira, J.B.; Kuntzler, S.G.; Bezerra, P.Q.M.; Cassuriaga, A.P.A.; Zaparoli, M.; da Silva, J.L.V.; Costa, J.A.V.; de Morais, M.G. Recent Advances of Microalgae Exopolysaccharides for Application as Bioflocculants. Polysaccharides 2022, 3, 264-276. https://0-doi-org.brum.beds.ac.uk/10.3390/polysaccharides3010015

AMA Style

Moreira JB, Kuntzler SG, Bezerra PQM, Cassuriaga APA, Zaparoli M, da Silva JLV, Costa JAV, de Morais MG. Recent Advances of Microalgae Exopolysaccharides for Application as Bioflocculants. Polysaccharides. 2022; 3(1):264-276. https://0-doi-org.brum.beds.ac.uk/10.3390/polysaccharides3010015

Chicago/Turabian Style

Moreira, Juliana Botelho, Suelen Goettems Kuntzler, Priscilla Quenia Muniz Bezerra, Ana Paula Aguiar Cassuriaga, Munise Zaparoli, Jacinta Lutécia Vitorino da Silva, Jorge Alberto Vieira Costa, and Michele Greque de Morais. 2022. "Recent Advances of Microalgae Exopolysaccharides for Application as Bioflocculants" Polysaccharides 3, no. 1: 264-276. https://0-doi-org.brum.beds.ac.uk/10.3390/polysaccharides3010015

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