Polyelectrolyte flocculants in harvesting microalgal biomass for food and feed applications

Algal Research 24 (2017) 167–180

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Algal Research journal homepage: www.elsevier.com/locate/algal

Review article

Polyelectrolyte flocculants in harvesting microalgal biomass for food and feed applications Lucie Van Haver a,b, Sasi Nayar a,⁎ a b

Algal Production Group, South Australian Research and Development Institute – Aquatic Sciences, 2 Hamra Avenue, West Beach, SA 5024, Australia Ecole Supérieur de Chimie Organique et Minérale (ESCOM), 1 Allée du Réseau J-M Buckmaster, 60200 Compiègne, France

a r t i c l e

i n f o

Article history: Received 18 September 2016 Received in revised form 27 February 2017 Accepted 24 March 2017 Available online xxxx Keywords: Low toxicity flocculation Polyelectrolyte flocculant Anionic flocculant Cationic flocculant Non-ionic flocculant Microalgae harvesting

a b s t r a c t Harvesting microalgal biomass at industrial scale is a techno-economic bottleneck for the algal biomass industry, compounded by the small cell size of microalgae and dilute biomass concentrations in culture. As a result, large volumes of water need to be removed during harvesting, making the process energy and cost intensive, accounting for up to 30% of the total cost of biomass production. Among the various harvesting techniques adopted commercially, flocculation is convenient and cost effective. The choice of a flocculant depends on its effectiveness on multiple microalgal strains, efficiencies at low biomass concentrations, its environmental footprint, being inexpensive and non-toxic for end application of the recovered biomass. Of the various flocculants, polyelectrolyte flocculants are widely utilised for various industrial applications such as wastewater treatment and mining, but also for effective harvesting of mass cultures of microalgae. Polyelectrolyte flocculants are polymers that are either branched or linear, but carrying ionic charge along their chain. They are accordingly classified as cationic, anionic or non-ionic polymers. These flocculants neutralise surface charges on cells and bind particles together by physical or chemical forces. The efficiency of polyelectrolyte flocculants depend on the type of polymer used, its molecular weight and charge density, dosage concentrations, cell concentration in the medium, type of strain, ionic strength and pH of the medium, and other parameters. Bulk harvesting of toxicant free microalgal biomass by polyelectrolyte flocculants is regarded to be one of the most economically viable techniques, with the cost of flocculants ranging between US$1.50 and 7.50 kg−1, and requiring very low dosage for effective harvesting. This review focusses on polyelectrolyte flocculants to harvest cultivated microalgae for non-toxic residue free applications of the harvested biomass in the food and the feed industry and evaluates various commercial polyelectrolyte flocculants, their properties and application in harvesting microalgal biomass from high density cultures. © 2017 Elsevier B.V. All rights reserved.

Contents 1. 2. 3.

4.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Commercially adopted microalgal harvesting techniques . . . . . . . . . . . . . . . . . . . . . . Polyelectrolyte flocculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Types of polyelectrolyte flocculants . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1. Cationic polyelectrolyte flocculants . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Anionic polyelectrolyte flocculants. . . . . . . . . . . . . . . . . . . . . . . . . 3.1.3. Non-ionic polyelectrolyte flocculants . . . . . . . . . . . . . . . . . . . . . . . 3.2. Recent advances in polyelectrolyte flocculants . . . . . . . . . . . . . . . . . . . . . . . 3.3. Factors influencing the efficiency of polyelectrolyte flocculants in harvesting microalgal biomass. 3.3.1. Effect of molecular weight, charge density and ionic strength of the medium . . . . . 3.3.2. Properties of different microalgal strains . . . . . . . . . . . . . . . . . . . . . . 3.3.3. Properties of the culture media . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Applications of commercial polyelectrolyte flocculants in harvesting microalgal biomass . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

⁎ Corresponding author. E-mail address: [email protected] (S. Nayar).

http://dx.doi.org/10.1016/j.algal.2017.03.022 2211-9264/© 2017 Elsevier B.V. All rights reserved.

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Funding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction With an ever increasing population we face numerous challenges, in particular a significantly higher demand for feed, food and energy to meet the ever increasing global demand, predicted to increase in excess of 50% in the next decade [1]. Microalgae is touted to be a promising feedstock for the production of bioenergy [2], but hold much greater promise for food and feed applications as they are a rich source of proteins and lipids [3]. As microalgae belong to different evolutionary lineages than terrestrial plants, they are capable of producing unique metabolites such as polyunsaturated fatty acids (e.g., eicosapentaenoic acid or docosahexaenoic acid), natural photosynthetic and non-photosynthetic pigments (e.g., phycocyanin, different carotenoid pigments, etc.), antioxidants, anti-inflammatory and anti-tumour compounds [4]. A large number of polysaccharides and other metabolites that cannot be sourced from terrestrial plants [5] are known to be present in algae. Considering their rapid growth rate, ease of culture and the fact that they do not compete with traditional agriculture for arable land for cultivation [1], microalgae hold significant promise as a feedstock for new biotechnological products. As part of the production process, harvesting microalgal biomass at an industrial scale continues to be a techno-economic bottleneck for the algal biomass industry. This is largely attributed to the small cell size of microalgae (1–30 μm) and dilute biomass concentrations in the culture media (~ 1 g L−1). As a result, the harvesting process requires large volumes of water to be removed, making it energy and cost intensive. Increasing the efficiency of the process by adopting low-energy, cost effective and environmentally friendly harvesting techniques is a major challenge in microalgal biotechnology, especially when the cost of harvesting can account for N30% of the total cost of the biomass production [6]. Harvesting techniques such as centrifugation can be used efficiently to obtain a clean solid-liquid separation for high value applications, but is too energy-intensive and costly for large scale applications [7]. This study aims to review various commercially adopted microalgal harvesting techniques, with a specific focus on the utilisation of various commercially available polyelectrolyte flocculants to harvest residue free microalgal biomass for food and feed applications. The review also encompasses general properties and efficiency of different commercial polyelectrolyte flocculants, together with their advantages and limitations. From a review of the published literature, this study

178 178 178

evaluates various commercially available polyelectrolyte flocculants and their application in harvesting cultivated microalgal biomass. As far as we are aware, this comprehensive review is the first in the published literature that has undertaken an in-depth review of various commercially available polyelectrolyte flocculants for food and feed where the harvested biomass is expected to have no toxic chemical residues and is clean for human or animal consumption. From an environment and economic standpoint, it is also important that these flocculants are non-toxic to microalgae cultivation when the medium is recycled after the recovery of the flocculants for reuse.

2. Commercially adopted microalgal harvesting techniques Out of the several harvesting techniques commonly used in commercial scale algal production systems, microfiltration is not only simple, but is also cost-effective. However, the technique is limited in its efficiency to separate cells from high density microalgal cultures [8], leading to clogging of the filters that require periodic cleaning. Microfiltration is efficient in harvesting biomass of chain forming or larger cell size microalgae such as Arthospira. Smaller cells are recovered using microstrainers [9] or diatomaceous earth [6]. Flocculation is one of the most convenient and cost effective method to harvest microalgal biomass [10,11]. The technique has undergone considerable technological advancements over the years with various novel spinoff technologies developed and optimised. As the technique is cost effective and has a low energy demand, flocculation is regarded to be a promising choice for environmentally sustainable applications. Various techniques to flocculate microalgae are well documented in the published literature, accomplished by physical, chemical and biological means. Physico-chemical flocculation techniques involve the use of magnetic nanoparticles, differential pH gradient, mineral salts and polymers. Flocculation by biological means include autoflocculation and bioflocculation. The use of ultrasound in harvesting microalgae has been demonstrated to be satisfactory in the recovery of biomass [12]. The technique leaves no residue in the harvested biomass as the algal culture suspension does not come in contact with any chemical agents unlike other techniques. However, the cooling system required to control temperature gradient needed to homogenise the field, is energy intensive and therefore increased process costs especially at scale [12].

Fig. 1. A schematic illustrating the mechanism of action of polyelectrolyte flocculants in flocculating microalgal cells (modified from [115]).

L. Van Haver, S. Nayar / Algal Research 24 (2017) 167–180 Table 1 Classification of flocculants on the basis of their molecular weight and charge density [43,45]. Classification

Molecular weight (Da)

Charge density (%)

High Medium Low Anti-scalant/dispersant

N107 105–106 104–105 103–104

50–100 N25 N10 –

Commercial scale electrocoagulation-flocculation uses sacrificial aluminium electrodes. By applying an electric-chemical field between reactive electrodes, the charged microalgal cells are attracted to the anode, where upon contact they destabilised and form aggregates. The efficiency of the flocculation process can be further enhanced by increasing the charge through the media, which is energy intensive. Whilst the technique is effective, it is expensive and leaves metal traces of up to ~1% in the recovered biomass [13,14]. Magnetic separation using iron oxide (Fe2O3) is a promising method of harvesting microalgal biomass based on electro-attraction [15,16]. Magnetic nanoparticles bind to the surface of microalgal cells with flocculation induced in the presence of a strong magnetic field. Although the nanoparticles can be recovered and reused many times without any pH adjustment, it is worth a mention that the process of regenerating the nanoparticles is a challenge [16]. High cost of this technology and the requirement for specialised equipment required for the recycling of nanoparticles make it non-viable for large commercial scale systems. Flocculation of microalgal cells by altering pH, also known as alkaline flocculation, is a widely adopted technique accomplished by altering the ratio of hydrogen and hydroxyl ions (H+/OH−) and magnesium ions (Mg2+) [17]. The efficiency of this process is dependent on the biomass concentration in the medium, requiring it to be N108 cells mL−1, and is often problematic in naturally occurring low density cultures. The

169

medium can be reused after harvesting the biomass by re-adjusting the pH [18]. However, the residual magnesium hydroxide generated as result of high pH could accumulate in the harvested biomass which could be limiting when toxic residue-free biomass for food and feed is required [18]. Another popular chemical flocculation technique uses mineral salts of ferric chloride (FeCl3), aluminium sulphate (Al2(SO4)3), ferric sulphate (Fe2(SO4)3) or inorganic polymers such as poly-aluminium complex, that are regarded to be efficient flocculants [19,20]. The major drawback of chemical flocculants is the contamination of the harvested biomass by residual chemicals such as aluminium or iron salts that renders the biomass useless for wide range of applications that include food and feed. Chemical flocculants are often required in high doses, for e.g., the concentrations of FeCl3 and Al2(SO4)3 required for the flocculation to be effective are 383.5 μM and 438.1 μM, respectively [20]. At these operating concentrations used to achieve an effective large scale flocculation of Nanochloropsis salina, a residual aluminium concentration in the biomass of N 1 mg g−1 on a dry weight basis was recorded [21], a concern from a food safety perspective. A report from the European Food Safety Authority [22], highlights that the toxicity limits of aluminium for humans to be ~ 0.01 mg g− 1 of body weight. Exposure to higher levels of aluminium is known to alter iron homeostasis in patients with Alzheimer's disease [23–25]. Higher concentrations of aluminium have also been demonstrated to inhibit growth and affect gill performance in fish [26,27]. In waste water treatment plants, higher concentrations of aluminium can cause failure of anaerobic digestion systems [28]. Aluminium residues in harvested microalgal biomass are also known to affect the composition of fatty acids methyl esters (FAMEs) in extracted lipids [21], with implications in the production of biofuels such as biodiesel [29]. Bioflocculation is achieved by co-cultivating certain microalgal species with others without the addition of any chemical agents, for e.g., co-cultivation of Chlorella vulgaris JSC-7 and Ettlia texensis [30,31]. Some microalgal species flocculate more readily than others, making

Fig. 2. Chemical structures of cationic polyelectrolytes (i) polyDADMAC (PDADMAC), (ii) polymer from epichlorohydrin and dimethylamine (ECH/DMA), (iii) cationic Polyacrylamide (CPAM), and (iv) chitosan.

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Fig. 3. A schematic illustrating the mechanism of action of cationic flocculants in flocculating microalgal cells.

them a candidate species for co-cultivation to induce bioflocculation in the target species [32]. The process is achieved by the generation of surface charges due to the production of extracellular polymeric substances (EPS) or infochemicals during the co-cultivation process [33]. An infochemical isolated from a senescent and autoflocculating culture of Skeletonema sp. was observed to induce flocculation in another species of microalgae [34]. These EPS include sugars, polysaccharides and their derivatives such as cellulose, galactose, glucose, mannose, pectins, rhamnose, ulvans, uronic acid, xylose, and others [35]. The presence of functional groups such as carboxyl, sulphate, amino and other electronegative atoms in the matrix imparts a negative surface charge to microalgal cells. The presence of biopolymers with the negatively charged microalgal cells in the solution results in a reduction of electrolytic repulsion, reducing the zeta-potential and facilitating aggregation or flocculation [36]. Advances in genetic manipulation over the past few years have led to the development of microalgal strains that autoflocculate with relative ease when compared to wild strains [37– 39]. The stability of the floc formation and minimisation of the repulsive effect of microalgal cells due to differential surface charges can be addressed by manipulating the target genes of the cultivated microalgae to induce self-flocculation [37]. Specific consortia of bacteria and certain fungi have also been reported to induce bioflocculation in microalgal cultures [40]. However, the technique of bioflocculation requires further research to optimise the process for better efficiencies. Bioflocculation techniques are relatively energy efficient processes, but are very strain or species specific and may not work under all conditions. As the efficiency of flocculation of bioploymers is very pH dependant [41], they may not work effectively in flocculating marine microalgae from a seawater media. Despite these drawbacks, the technique is regarded to be safe, biodegradable and environmentally friendly, besides having the advantage of a short operation time [42].

mass cultures of microalgae [43,44]. These flocculants are polymers that can either be branched or linear but carry ionic charge along their chain. Depending on their ionic nature, these polymers can be either cationic (positively charged), anionic (negatively charged) or nonionic (b1% charged) [45]. These flocculants are either organic or inorganic. During the flocculation process, small stabilized particles are combined into larger aggregates or flocs due to a combination of charge neutralisation and particle bridging [46], the extent of each depending on the charge density and chain length of the polymer. The interaction of the polyelectrolyte to the algal cell surface is affected by charge interaction, dipole-dipole interaction, hydrogen bonding, and/or Van der Waals interaction [40] (Fig. 1). Polyelectrolyte flocculants can concentrate the suspension by up to 800-times [47], making it easier to dewater the flocculated microalgal biomass comprising of larger aggregates after sedimentation than individual cells. After the flocculation process, a low energy centrifugation or gravity filtration of larger flocs can lead to a further 10-fold higher concentration of the harvested biomass [48]. Evaluation of several harvesting methods have demonstrated that a combination of flocculation with flotation or sedimentation followed by dewatering by centrifugation or filtration, is quite effective besides being cost- and energy-efficient [32]. The choice of a flocculant will hinge on its small environmental footprint, being inexpensive, non-toxic for end application of the recovered biomass, and its effectiveness at low concentrations. Another important consideration is the broad spectrum efficiency of the selected flocculant across several strains and species. For a flocculant to be regarded as effective, it must be able to separate N90% of the biomass [11,49,50] from the culture medium in b 60 min. Most laboratory studies on flocculation efficiencies evaluate performance of the flocculants being studied for durations of 60 min or less. 3.1. Types of polyelectrolyte flocculants

3. Polyelectrolyte flocculation Polyelectrolyte flocculants such as chitosan, alginates, alum, acrylamides, etc. are widely used for many industrial applications such as wastewater treatment, paper making, mineral processing, oil recovery or colour removal, but can also be employed as a harvesting aide in

Polyelectrolytes are water soluble polymers and can be classed as organic and inorganic polymers. These polymers are effective at low concentrations, at a high concentration factor and strengthen the resultant flocs. Inorganic polyelectrolytes are salts of multivalent metals such as aluminium (Alum or Al2(SO4)3) or iron (Iron chloride or FeCl3). Organic

Fig. 4. Chemical structures of anionic polyelectrolytes (i) anionic polyacrylamide (APAM), polystyrene sulfonic acid (PSSA), and (iii) 2 acrylamido 2 methyl propane sulfonic acid (AAMPSA).

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Fig. 5. A schematic illustrating the mechanism of action of anionic or non-ionic flocculants in flocculating microalgal cells.

polymers include those that occur naturally (e.g., carrageenans, alginates, etc), those that are chemically modified (e.g., chitosan, starch, etc.) or artificially synthesized (e.g. acrylamides, acrylic acids, etc). Both organic and inorganic polyelectrolyte are characterized by their functional group along their chain which may be charged either positive (cationic), negative (anionic), both negative and positive (ampholyte) or not charged at all (non-ionic). All polyelectrolyte flocculants differ in how they function, which is largely dependent on their molecular structure and mode of operation [45]. More a polymer is substituted with active monomers, higher is its charge density. This results in a polyelectrolyte that is regarded to be “strong” [51]. This classification of polyelectrolytes based on their molecular weight and charge density is summarised in Table 1. Based on composition, polyelectrolytes can either be a homopolymer, i.e., those composed of one main monomer type, or co-polymers, i.e., those that involves two or more monomers. Typically, a polyelectrolyte flocculant works by neutralising the surface charge on particles suspended in a solution [9]. Microalgal cells possess a negative surface charge under culture conditions (up to pH of 5). The most effective flocculant that brings about a charge neutralisation followed by an effective flocculation is a cationic polyelectrolyte flocculant [52,53]. Acrylamide based polyelectrolyte flocculants are widely used in microalgal biomass harvesting commercially, however they are toxic to aquatic life [3]. As an example, 96 h acute toxicity (LC50) of acrylamide in water on the Mediterranean mussel (Mytilus galloprovincialis) was determined to be 411 mg L−1 [54] when compared to ~ 119.5 mg L−1 for goldfish (Carassius auratus) [55]. Aquatic toxicity studies of acrylamide flocculants conclude that the anionic class possess lower toxicity to aquatic organisms when compared to the cationic class which are ~100 times more toxic [56] due to their positive charge. Cationic polyelectrolytes affect the integrity of the cell membranes of the affected organisms, with the effect dependent on the charge density and hydrophobicity of the polymer [57]. Additionally polyacrylamide flocculants are non-biodegradable, possess neurotoxic characteristics [24] and have been shown to be a carcinogen [58], more so with polycations than polyanions or non-ionic polymers [59]. Because of the inherent risks identified above, their dosage is strictly limited to 0.5 μg L−1 for treatment of drinking water in most countries.

3.1.1. Cationic polyelectrolyte flocculants Polycations or polybases are composed of a positively charged group along their main frame rendering these flocculants a positive charge in solution. The substituted monomers are often amino groups (\\NH2), acrylamide (C3H5NO)n or co-polymer of amines and acrylamide, but could also be ethyleneimine, vinyleamine, diallyldimethylammonium chloride (DADMAC), chitosan, etc. Starch modified by the incorporation of quaternary ammonium groups and is used as an alternative to chitosan [60,61]. However, the positive charges of quaternary ammonium groups of starches are less sensitive to changes in pH unlike chitosan rendering these effective over a broader pH range. Cationic starches, a type of polyelectrolyte flocculant that possess a positively charged group such as amino, imino, ammonium, sulphonium or phosphonium groups. These flocculants are effective across a wide range of pH, readily biodegradable, and are therefore often used to flocculate organic and inorganic matter in waste water possessing negative charges [62]. Chemical structures of common cationic polymers are depicted in Fig. 2. Cationic polyelectrolytes are utilised mainly in waste water treatment for the removal of organic matter from water. The efficiency of these flocculants increases with their charge density and assist with the mechanical dewatering process resulting in a ‘cake’ containing 20% to 50% solids [45]. Flocculation by cationic polyelectrolytes occurs through a combination of charge neutralisation between the negatively surface charge of colloids and the positive active site of the polymer and polymer bridging [63]. Because organic particles are mostly negatively charged in solution (stable colloids), cationic polyelectrolytes work very effectively [49]. The colloids become destabilised due to electro-attraction of the cells to the polymer, followed by a charge-charge attraction resulting in a floc formation as depicted in Fig. 3. The choice of a cationic polyelectrolyte flocculant with the appropriate charge density depends on the mechanical dewatering process applied after flocculation. These mechanical dewatering processes include the drying belt, which is the most efficient of the lot, followed by vacuum filtration, belt press, filter press and lastly centrifugation. As the efficiency of the mechanical dewatering process increases, lower the charge density of the flocculant needed to flocculate the organic matter. Low to medium molecular weight and low charge density cationic polyelectrolytes are

Table 2 Evaluation of process conditions and efficiencies of techniques commonly to harvest microalgal biomass at industrial scale.

Source Recovery efficiency (%) Flocculant dosage (kg kg−1 dry weight basis) Flocculants cost (US$ ton−1) Process energy (kWh m−3) Energy cost (US$ KWh−1) Processing cost (US$ ton−1 dry weight)

Centrifuge Inorganic metal salts Organic synthetic polyelectrolyte flocculant

Chitosan

Tannin-based polyelectrolyte

[6,7,109] N95 – – 8–20 0.1 800–2000

[7,11,89,92,101] ~90 0.038–0.35 7000–100,000 – – 266–35,000

[11,109,110] N90 0.018–0.04 2000 – – 36–80

[11,49,89,92,95,101] 70–90 0.22–0.40 300–2000 – – 66–800

[11,49,89,95,101] N95 0.002–0.028 2000–8000 – – 4–224

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Table 3 Applications of commercially available cationic polymers to harvest cultivated marine and freshwater microalgae. Manufacturer

Technical specifications Commercial Price name (US$Kg-1) Polymer type [P]; Molecular weight [MW]; Charge [C]

Microalgal species

Flocculation condition Cell density [D] g L-1 / Medium cells mL-1 / OD750; Scale [S] mL; Flocculant Freshwater dose [FD] mg L-1; Settling time [T] min [Fresh]; Marine [Marine]

Efficiency Source (%)

Allied Chemicals Ltd, UK

Zetag 63

Chlorella stimatophora Chlorella stimatophora Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlorella vulgaris Nannochloropsis oculata Nannochloropsis salina Phaeodactylum tricornutum Neochloris oleoabundans Muriellopsis sp.

D: 106 mL-1; S: 1000; FD: 10; T: 30

Fresh

93

D:10 mL ; S: 100; FD: 5; T: 30

Fresh

93

D: 107 mL-1; S: 100; FD: 35; T: 30 D: 107 mL-1; S: 100; FD: 35; T: 30

Fresh Fresh

72 95

D:107 mL-1; S: 100; FD: 30; T: 30

Fresh

95

S: 1000; FD: 34; T: 60 D: 0.26; S: 3000; FD: 5; T: 30

Fresh Fresh

98 100

D: 0.29; S: 3000; FD: 0.55; T: 30

Marine

75

S: 2000; FD: 10

Marine

10

[111]

D: OD750-3.6; S: 5; FD: 0.01; T: 120

Marine

98

[112]

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

52

D: 2.0; S: 250; FD: 10; T: 15

Fresh

95

D: 2.0; S: 250; FD: 16; T: 15 D: 2.0; S: 250; FD: 25; T: 15

Fresh Fresh

95 95

D: 2.0; S: 250; FD: 12; T: 15 D: 2.0; S: 250; FD: 3; T: 15

Fresh Fresh

95 95

D: 2.0; S: 250; FD: 2; T: 15

Fresh

98

D: 2.0; S: 250; FD: 15; T: 15 D: 2.0; S: 250; FD: 8; T: 15

95 89

D: 0.15; S: 250; FD: 2.5; T: 60

Fresh Fresh Fresh Fresh

95

[52]

D: 107 mL-1; S: 100; FD: 70; T: 30 D: 107 mL-1; S: 100; FD: 70; T: 30

Fresh Fresh

48 90

[50]

D: 107 mL-1; S: 100; FD: 40; T: 30

Fresh

95

D: 0.6; S: 8000; FD: 20; T: 120

Fresh

98

D: 0.6; S: 8000; FD: 20; T: 120

Marine

91

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

93

[112]

D: OD750-0.8; S: 10; FD: 30

Marine

90

[85]

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

36

D: 107 mL-1; S: 100; FD: 30; T: 30 D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh Fresh

95 20

D: 107 mL-1; S: 100; FD: 70; T: 30

Fresh

70

D: 107 mL-1; S: 100; FD: 30; T: 30 D: 107 mL-1; S: 100; FD: 50; T: 30

Fresh Fresh

95 90

D: 107 mL-1; S: 100; FD: 60; T: 30

Fresh

95

D: 10 mL ; S: 100; FD: 30; T: 30 D: 107 mL-1; S: 100; FD: 80; T: 30

Fresh Fresh

65 80

D: 107 mL-1; S: 100; FD: 60; T: 30

Fresh

35

D: 10 mL ; S: 100; FD: 1.5; T: 30 D: 107 mL-1; S: 100; FD: 4; T: 30

Fresh Fresh

95 99

D: 107 mL-1; S: 100; FD: 2; T: 30

Fresh

98

D: 107 mL-1; S: 100; FD: 20; T: 30 D: 107 mL-1; S: 100; FD: 30; T: 30

Fresh Fresh

88 91

BASF

Zetag 92

NA

Magnafloc LT225

NA

Zetag 8819 Zetag 8185

Brenntag Quimica

Brenntag Quimica

NA

8 8

P: Polyacrylamide; MW: 1x107 P: Polyacrylamide; MW: 2x107 P: Polyacrylamide

P: Polyacrylamide; C: High P: Polyacrylamide; MW: High; C: High

Zetag 7570

8

P: Polyacrylamide

Zetag 7557

8

P: Polyacrylamide

EM16

1.5-6

EM22

1.5-6

FB1

1.5-6

P: Polyelectrolyte; MW: Medium; C: medium P: Polyelectrolyte; MW: Large; C: Medium P: Polyelectrolyte; MW: High; C: Medium

EM1

1.5-6

P: Polyelectrolyte; MW: High; C: High

DOW Chemical

C-31

NA

Emsland-Stärk GmbH

Emfloc KC750

1.4

P: Polyelectrolyte; MW: 5x106 P: Potato Starch

Rundo Biotech Japan Co.

Poly 4.5 (γ-glutamic acid)

Sachtleben Synthofloc Wasserchemie 5080H

Separ Chemi GmbH

POLY SEPAR® CFL25

POLY SEPAR® KW100

POLY SEPAR® KW45

POLY SEPAR® PK55H

POLY SEPAR® SK72

NA

2.2

2.7

2.7

3.6

3.37

Chitosan

P: Polyacrylamide

P: Tannin, quaternary ammonia salt; MW: Low; C: High

P: Quaternary ammonia compound, free of polyacrylamide; C: High

P: Quaternary ammonia compound, free of polyacrylamide; C: Low

P: Polyacrylamide; MW: High; C: High

P: Starch

Muriellopsis sp. Scenedesmus sp. Muriellopsis sp. Chlorella vulgaris Scenedesmus subspicatus Muriellopsis sp. Chlorella fusca Mixed Chlorophyta Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella protothecoides Chlorella vulgaris Phaeodactylum tricornutum Neochloris oleoabundans Neochloris oleoabundans Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii

6

7

7

-1

-1

-1

[53]

[50]

[95] [11]

[49]

[49]

[108]

[50]

L. Van Haver, S. Nayar / Algal Research 24 (2017) 167–180

173

Table 3 (continued) Manufacturer

Separ Chemi GmbH

Technical specifications Commercial Price name (US$Kg-1) Polymer type [P]; Molecular weight [MW]; Charge [C]

POLY SEPAR® SK72

POLY SEPAR® KW745 H

CFL 217

CFL 229

Sigma-Aldrich

Sigma-Aldrich

SNF-Floerger

Chitosan

Chitosan

FO4990

FO4800

FO4650

SNF-Floerger

TANAC (Brazil)

Nalco (Australia)

FO4550

3.37

2.7

2.5

2.5

90

90

7.9

3.37

7.9

7.9

Tannin

1.9

Tanfloc SL

2.25

71301

P: Starch

P: Polyacrylamide

P: Poly DADMAC; MW: Low; C: High

P: Poly DADMAC; MW: Low; C: High

P: Linked D-glucosamine; MW: Medium

MW: Low

P: Polyacrylamide; MW: 4.5-7.1x106; C: Very High

P: Polyacrylamide; MW: 4.9-7.1x106; C: High

P: Polyacrylamide; MW: 4.5-7.1x106; C: Medium

P: Polyacrylamide; MW: 4.1-7.1x106; C: Low

P: Natural polymer; MW: Low; C: Low-medium P: Natural polymer; MW: Low; C: Low-medium

P: Polyacrylamide; MW: Medium; C: Medium/High

Microalgal species

Flocculation condition Cell density [D] g L-1 / Medium cells mL-1 / OD750; Scale [S] mL; Flocculant Freshwater dose [FD] mg L-1; Settling time [T] min [Fresh]; Marine [Marine]

Efficiency Source (%)

Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlamydomonas reinhardtii Chlorella vulgaris Chlorella stimatophora Neochloris oleoabundans Nannochloropsis salina Nannochloropsis sp. Isochrysis galbana Chaetoceros calcitrans Chlorella vulgaris Nannochloropsis salina Nannochloropsis oculata Chlamydomonas reinhardtii Chlorella vulgaris Nannochloropsis salina Nannochloropsis oculata Chlorella vulgaris Nannochloropsis salina Nannochloropsis oculata Chlorella vulgaris Nannochloropsis salina Nannochloropsis oculata Microcystis aeruginosa Chlorella vulgaris Nannochloropsis oculata Chlorococcum sp.

D: 107 mL-1; S: 100; FD: 60; T: 30

Fresh

99

D: 10 mL ; S: 100; FD: 20; T: 30 D: 107 mL-1; S: 100; FD: 30; T: 30

Fresh Fresh

88 91

D: 107 mL-1; S: 100; FD: 60; T: 30

Fresh

99

D: 107 mL-1; S: 100; FD: 20; T: 30 D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh Fresh

90 90

D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh

90

D: 107 mL-1; S: 100; FD: 20; T: 30 D: 107 mL-1; S: 100; FD: 30; T: 30

Fresh Fresh

90 80

D: 107 mL-1; S: 100; FD: 50; T: 30

Fresh

95

D: 10 mL ; S: 100; FD: 40; T: 30 D: 107 mL-1; S: 100; FD: 40; T: 30

Fresh Fresh

76 87

D: 107 mLl-1; S: 100; FD: 50; T: 30

Fresh

96

D: 0.7; S: 500; FD: 25

Fresh

93

[101]

D: 0.25; S: 100; FD: 8; T: 30

Fresh

85

[14]

D: 106 mL-1; S: 1000; FD: 10; T: 30

Fresh

90

[53]

D: OD750-0.8; S: 10; FD: 90

Marine

66

[85]

D: 0.7; S: 1500; FD: 3; T: 60

Marine

98

[89]

FD: 100; T: 60

Marine

90

[113]

D: 106 mL-1; S: 1000; FD: 10; T: 30

Marine

90

[53]

S: 500; FD: 20; T: 240

Marine

83

[100]

D: 0.26; S: 3000; FD: 1.66; T: 30

Fresh

99

[11]

D: 0.7; S: 50; FD: 3; T: 60

Marine

94

[89]

D: 0.26; S: 3000; FD: 0.55; [T]: 30

Marine

90

[11]

D: 0.7; S: 500; FD: 13.5

Fresh

97

[101]

D: 0.26; S: 3000; FD: 1.66; T: 30

Fresh

99

[11]

D: 0.7; S: 50; FD: 3; T: 60

Marine

88

[89]

D: 0.26; S: 3000; FD: 0.55; T: 30

Marine

87

[11]

D: 0.26; S: 3000; FD: 1.66; T: 30

Fresh

100

D: 0.7; S: 50; FD: 3; T: 60

Marine

73

[89]

D: 0.26; S: 3000; FD: 0.55; T: 30

Marine

81

[11]

D: 0.26; S: 3000; FD: 1.66; T: 30

Fresh

99

[11]

D: 0.7; S: 50; FD: 3; T: 60

Marine

73

[89]

D: 0.26; S: 3000; FD: 0.55; T: 30

Marine

67

[11]

FD: 10; T: 30

Fresh

97

[110]

D: 0.26; S: 3000; FD: 5

Fresh

100

[11]

D: 0.29; S: 3000; FD: 5; T: 30

Marine

97

D: 0.6; S: 1000; FD: 3; T: 30

Marine

78

7

7

-1

-1

[50]

[114]

(continued on next page)

174

L. Van Haver, S. Nayar / Algal Research 24 (2017) 167–180

Table 3 (continued) Manufacturer

Technical specifications Commercial Price name (US$Kg-1) Polymer type [P]; Molecular weight [MW]; Charge [C]

Microalgal species

Flocculation condition Cell density [D] g L-1 / Medium cells mL-1 / OD750; Scale [S] mL; Flocculant Freshwater dose [FD] mg L-1; Settling time [T] min [Fresh]; Marine [Marine]

Efficiency Source (%)

71303

Chlorococcum sp. Chlorococcum sp.

D: 0.6; S: 1000; FD: 4; T: 30

Marine

90

D: 0.6; S: 1000; FD: 3; T: 30

Marine

85

P: Polyacrylamide; MW: Low/Medium; C: Medium P: Polyacrylamide; MW: Low; C: Medium/High

71305

often used in potable water treatment to avoid acrylamide contamination or in paper manufacturing process to improve the durability and tensile strength of the paper [43,45]. 3.1.2. Anionic polyelectrolyte flocculants Polyanions or polyacids are composed of a negatively charged group attached to the main structure rendering these flocculants negatively charged in solution. Anionic polyelectrolytes are mostly polyacrylamide or polymer containing weak acidic carboxylic acid group (\\COOH), whose efficiency is highly dependent on the pH of the medium. These flocculants also occur as polystyrene sulfonic acid (PSSA), -acrylamido 2-methyl propane sulfonic acids (AAMPSA), pectin, alginates etc. The chemical structures of the most common anionic polymers are shown in Fig. 4. Anionic polyelectrolytes, especially anionic polyacrylamide (APAM), are largely used by the mineral processing industry. Extraction of coal and inorganic minerals from waste material or coal washing requires a medium molecular weight and charge density flocculant, whilst a flocculant with high charge density is required to flocculate spent mud in drilling process and dewatering alkaline sludge from industrial effluents. Polyanionic flocculants of low or medium charge density are also

used in retention aids in paper manufacture or for clarification of sugar cane juice or wash-water. Anionic polyelectrolytes have been demonstrated to prevent scale formation in industrial pipes. Those possessing anti-scalant or dispersant properties include polymers of acrylic acid and its salts, co-polymers of acrylamide and acrylic acid or polymers containing sulfonic acid with a low molecular weight and charge density [45]. Anionic flocculation is governed by chemical interactions rather than electrostatic forces and flocculation is affected by the absorption of the polymer on to the microalgae cell by hydrogen bonding [64]. Tenney and co-workers observed no algal flocculation with anionic flocculants, concluding that although the polymer bound to the cell surface by chemical forces, it was unable to form bridges [52]. Fig. 5 depicts the mechanism of action of polyanionic flocculants. 3.1.3. Non-ionic polyelectrolyte flocculants Non-ionic polyelectrolytes are mainly polyacrylamide without any surface charge or those with 1–3% anionic sites. The mechanism of action of non-ionic polyelectrolyte flocculants in flocculating microalgae is depicted in Fig. 5. A natural non-ionic polymer utilised in water treatment is macerated seeds from the tree Moringa oleifera [65]. The flour from Moringa oleifera seeds possess high flocculation potential [66],

Table 4 Applications of commercially available anionic polymers to harvest cultivated marine and freshwater microalgae. Manufacturer Commercial Price name (US$ Kg-1)

Technical specifications Polymer type [P]; Molecular weight [MW]; Charge [C]

Microalgal species

Flocculation condition Cell density [D] g L-1 / cells mL-1 / OD750; Scale [S] mL; Flocculant dose [FD] mg L-1; Settling time [T] min

Medium Freshwater [Fresh]; Marine [Marine]

Efficiency Source (%)

BASF

Chlorella zofingiensis Muriellopsis sp.

D: 0.2; S: 7000; T: 60

Fresh

0

[95]

D: 2.0; S: 250; FD: 10; T: 15

Fresh

2

[49]

Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chaetoceros calcitrans Chlorococcum sp. Chlorococcum sp. Chlorophyta

D: 107 mL-1; S: 100; FD: 10; T: 30 D: 107 mL-1; S: 100; FD: 40; T: 30

Fresh Fresh

8 21

[50]

D: 107 mL-1; S: 100; FD: 10; T: 30

Fresh

8

D: 107 mL-1; S: 100; FD: 10; T: 30 D: 107 mL-1; S: 100; FD: 40; T: 30

Fresh Fresh

5 24

D: 107 mL-1; S: 100; FD: 10; T: 30

Fresh

4

S: 100; FD: 0.1; T: 240; pH = 10.2

Fresh

98

[100]

D: 0.6; S: 1000; FD: 2; T: 30

Marine

84

[114]

D: 0.6; S: 1000; FD: 3; T: 30

Marine

84

D: 0.15; S: 250; FD: 20; T] 60

Fresh

3

[52]

D: 10 mL ; S: 100; FD: 20; T: 30 D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh Fresh

10 20

[50]

D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh

10

D: 0.7; S: 500; FD: 40

Fresh

20

[101]

D: 0.6; S: 1000; FD: 5

Marine

84

[114]

Brenntag Quimica Ciba Specialty Chemicals

Magnafloc E-38 EM6

NA

P: Polyacrylamide

NA

Magnafloc LT27

NA

P: Polyelectrolyte; MW: Low to High P: Polyacrylamide

Magnafloc LT25

Magnafloc 155 Magnafloc 156 A-21

DOW Chemical Separ chemie POLY GmbH SEPAR® Ahrensburg AN10TW

SNF-Floerger Nalco (Australia)

Flopam AN910 82230

NA

NA NA NA NA

NA

P: Polyacrylamide

P: Polyelectrolyte; MW: High; C: Low/Medium P: Polyelectrolyte; MW: High; C: Medium P: Polystyrene; MW: High P: Polyacrylamide

P: Polyacrylamide P: Polyacrylamide; MW: High; C: Medium

Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlamydomonas reinhardtii Chlorococcum sp.

7

-1

L. Van Haver, S. Nayar / Algal Research 24 (2017) 167–180

low toxicity [67] and is less expensive [68]. Research has demonstrated Moringa seed flour to be effective in flocculation of freshwater microalgae such as Chlorella sp. and Scenedesmus sp. [10,68–70]. The flour from the seeds of Moringa contain peptides with an isoelectric pH value ranging between 9 and 10 or higher [71–73]. In a saline media this compound transformed to a polyelectrolyte with the ability to flocculate microalgae [74,75].

3.2. Recent advances in polyelectrolyte flocculants Recent advances in polyelectrolyte flocculant technology include the development of novel flocculants that are recoverable after harvesting microalgae. In doing so not only is the flocculant recovered, but also the supernatant, which in this case is the culture media. The recovered media could be recycled for microalgal cultivation without detrimental effects on cell growth [76], making this process clean and green. Besides reducing the environmental footprint, the use of these flocculants in industrial-scale microalgal cultivation could lead to significant reduction in the operating costs. One such flocculant is the crystalline nanocellulose (CNC), derived from the acid hydrolysis of cellulose fibres. The resulting crystals from hydrolysis are nano-sized rods with a size and aspect ratio dependent on the source of cellulose, hydrolysis time and the temperature of hydrolysis [77, 78]. These bio-based flocculants are biodegradable and possess unique optical properties besides high specific surface area and nanoscale dimensions [79,80]. Furthermore, by altering the hydroxyl group on the surface of the CNC, the properties of the flocculant can be manipulated [81]. CNC grafted with cationic pyridinium and imidazole functional groups was found to flocculate microalgal cells effectively, with the flocculation efficiency relatively unaffected by the presence of algogenic matter, when compared to conventional polymer flocculants [80,82].

175

The process of reversible flocculation and deflocculation is generally achieved by pH variation, most commonly by the use of bases and acids. The recurrent use of acids and bases, especially with recycling of the medium, leads to an increase in the ionic strength of the medium potentially inhibiting the cultures [80]. Formation and accumulation of salts also contribute to an increased ash content in the harvested biomass, another drawback of this process. Imidazole grafted CNC has been shown to have a pH dependent surface charge, positive below pH 6.2 and negative above pH 6.9. This narrow pH range can easily be accomplished in a culture system by sparging carbon dioxide or air to flocculate and deflocculate [80]. The authors demonstrated maximum flocculation efficiency of N 90% with Chlorella vulgaris at a flocculant dosage of 200 mg L−1 and pH of 3.5 achieved with carbon dioxide purging. This approach of carbon dioxide switchability of the flocculant allows for the reuse of the re-dispersed CNC for further flocculation, and the supernatant following sediment to be reused for cultivation [76]. Ge and coworkers also demonstrated growth of microalgal cultures in CNC-harvested recycled medium, potentially making the cultivation process at larger scale cost-effective. The flocculant is recovered from CNC-bound microalgal aggregates by carbon dioxide sparging after centrifugation. The recovered CNC could be reused by redispersing again into the new batch of algal suspension to be dewatered. Similarily, Morrissey and co-workers reported the use of charge tunable polyampholytes, viz., polymers of positively charged N,Ndimethylaminopropyl acrylamide (DMAPAA), neutrally charged methaacrylamide and negatively charged acrylic acid, as a reversible and recyclable flocculant to dewater cultures of Chlorella vulgaris [83]. Furthermore, they reported achieving N99% flocculation efficiencies with a 10 fold increase in concentration of the microalgal biomass. These authors further reported that the reversible and recyclable polymer flocculants could be recovered at N 98% yields and recycled through at least five flocculation processes, by adjusting the pH through the

Table 5 Application of commercially available non-ionic polymers to harvest cultivated marine and freshwater microalgae. Manufacturer

Commercial Price name (US$ Kg-1)

Technical specifications Microalgal Polymer type [P]; Molecular species weight [MW]; Charge [C]

Flocculation condition Cell density [D] g L-1 / cells mL-1 / OD750; Scale [S] mL; Flocculant dose [FD] mg L-1; Settling time [T] min

Medium Efficiency Source Freshwater (%) [Fresh]; Marine [Marine]

BASF

Magnafloc 351

P: Polyacrylamide

D: OD750-0.7; S: 5; FD: 0.01; T: 60

Marine

0

D: OD750-0.7; S: 5; FD: 0.01; T: 60

Marine

0

D: 107 mL-1; S: 100; FD: 10; T: 30 D: 107 mL-1; S: 100; FD: 10; T: 30

Fresh Fresh

3 12

D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh

10

D: 2.0; S: 250; FD: 10; T: 15 D: 106 mL-1; S: 1000; T: 30

Fresh Fresh

0 0

[49] [53]

D: 0.15; S: 250; FD: 20; T: 60

Fresh

0

[52]

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

0

[112]

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

0

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

0

D: OD750-0.7; S: 5; FD: 0.01; T: 120

Marine

0

D: 10 mL ; S: 100; FD: 10; T: 30 D: 107 mL-1; S: 100; FD: 10; T: 30

Fresh Fresh

5 3

D: 107 mL-1; S: 100; FD: 20; T: 30

Fresh

1

D: 0.6; S: 1000; FD: 10; T: 30

Marine

80

[114]

D: OD750-1; S: 500; FD: 1000; pH 9.2; T: 120

Fresh

88

[68]

Magnafloc LT20

NA

NA

P: Polyacrylamide

Brenntag Quimica Cyanamid

EM5 N-100

NA NA

P: Polyelectrolyte P: Polyacrylamide

DOW Chemical

N-670

NA

SNF-Floerger

22N

NA

P: Polyacrylamide; MW: Medium NA

H22N

Separ Chemie GmbH Ahrensburg

POLY SEPAR® AN20

Magnafloc 351 Federal University Moringa of Sergipe oleifera seed flour

NA

3.15

Ciba Chemicals

NA

NA

P: Polyacrylamide

P: Polyacrylamide; MW: High; P: Natural polymer

Phaeodactylum tricornutum Neochloris oleoabundans Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus. Muriellopsis sp. Chlorella stimatophora Chlorophyte Phaeodactylum tricornutum Neochloris oleoabundans Phaeodactylum tricornutum Neochloris oleoabundans Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorococcum sp. Chlorella vulgaris

7

-1

[112]

[50]

[50]

176

L. Van Haver, S. Nayar / Algal Research 24 (2017) 167–180

addition of an acid or base. Extension of this research involved the flocculation of Chlamydomonas reinhardtii, Synechococcus sp., Aulacoseira ambigua, Nannochloropsis gaditana and Chlorella vulgaris using polyampolyte flocculant developed from the random copolymerization of monomers of DMAPAA and acrylic acid [84]. These polyampholytic flocculants with reversible electrostatic interactions resulted in flocculation efficiencies of N 97%, recovery of N90% of the flocculant yield, and when recycled, retained flocculation efficiencies of N 95%. Techno-economic viability analysis of the dewatering process using recyclable polyampholytic flocculants showed a reduction in flocculation operating costs by 85% under standard conditions [84]. 3.3. Factors influencing the efficiency of polyelectrolyte flocculants in harvesting microalgal biomass Technological advancements have led to the development of effective commercial polyelectrolyte flocculants. These flocculants work by neutralising the surface charge on the cells and bind particles together by physical or chemical forces. Several researchers have evaluated polyelectrolyte flocculants for their efficacy on different strains and operating parameters [49,52,53] and concluded that the efficiency of polyelectrolyte flocculants are dependent on the type of polymer used, its molecular weight and charge density, in addition to operating conditions such as dosage concentrations, cell concentration in the medium, type of strains, ionic strength and pH of the medium, and other parameters as outlined below. 3.3.1. Effect of molecular weight, charge density and ionic strength of the medium The efficiency of a polyelectrolyte flocculant depends on its size and polarity [45,61]. Based on polarity, they can be categorised as cationic, anionic or non-ionic. Their charge depends on the amount of charged monomers bound to the polymer chain and can often vary between negligible levels to up to 100% [61]. As surface charges are negative under natural conditions, studies have demonstrated that cationic polymers are 90% more effective in flocculating the biomass than anionic or non-ionic polymers, whose efficacies range between 0 and 40% under the same set of conditions [11,49,85]. Bleeke and co-workers demonstrated that the flocculation efficiency (FE) varied considerably from N90% for a polymer chain completely substituted with a cationic group, to a reduced efficiency of ~50% for one substituted with 35% by the cationic group [50]. This result confirms the assumption that the degree of polymer ionization impacts on harvesting efficiency. Increasing the molecular weight (MW) and charge density (CD) of polymeric flocculants have reportedly led to better FE [50,86]. Low MW cationic polymer may not be efficient in flocculation or may be required at a much higher concentration to accomplish some level of flocculation. On the other hand, higher concentrations of high MW polymers can reverse surface charges on microalgal cells and stabilize the suspension [87, 88]. Garzon-Sanabria and co-workers reported that FE was only affected by MW and CD of polymeric flocculants [89]. These findings were based on a previous research where a variation in CD from 45 to 99% at a constant molecular weight of 106 Da, resulted in an increase in FE from 73% to 93%, respectively. However, a reduction in molecular weight to104 Da at a CD of 100% did not result in a substantial increase in the level of flocculation of Nannochloropsis salina, even at a concentration of 100 mg L−1 [89]. Roselet and co-workers studied the interaction of CD and MW in commercially available polymeric flocculants Flopam and Zetag in flocculating freshwater and marine microalgae [11]. These authors highlighted that FE increased with an increase in polyelectrolyte charge and that an increase in MW led to a decline in FE. It is worth noting that polymeric flocculants from the Flopam series exhibited lower CD with increasing MW. The size of polymers depends on the interaction between polymer segments. Therefore, with increasing CD, the polymer adopts an expendable configuration [90]. Polyacrylamide polymers, Flopam and Zetag were observed to be very effective in

flocculating freshwater microalga Chlorella vulgaris, but could only flocculate 75% of the marine microalga Nannochloropsis oculata from the media. The relatively poor performance of these flocculants with marine microalgae is not surprising as it is well acknowledged that polymers often undergo coiling due to the ionic strength of the medium [85,89]. Bilanovic and co-workers observed that reducing the salinity of the medium under 5 g L−1 improved polyelectrolyte flocculation of the marine alga Chlorella stigmatophora [53]. Natural polysaccharides such as chitosan as well as cationic starch have been shown to perform poorly in marine media [40,91]. Ions present in seawater media can also shield the charged sites on both the flocculants as well as the microalgae [92], causing ionic hindrance during flocculation [40]. Differential effects of ionic strengths on FE of two synthetic polymers (Zetag 63 and Zetag 92) and chitosan were attributed to their polymer size, molecular configuration and charge density [53]. However, the study could not conclusively differentiate between the effect of ionic strength and algogenic organic matter (AOM) on dosage demand, as the marine culture was diluted with fresh media leading to a proportionate dilution of the salts as well as AOM. Henderson and co-workers determined that AOM is basically constituted of hydrophilic polysaccharides and hydrophobic proteins, whose ratio and concentration varied with species and growth phase [93]. Other researchers established that AOM impacted on FE more significantly than salinity [14,18,89]. Flocculation efficiencies of 50% were achieved in pH-induced flocculation of Chlorella vulgaris at pH 10 and polysaccharide concentrations of 25 mg L−1. However, the efficiency with marine microalgae Nannochloropsis oculata reduced by 10% under the same set of conditions [18]. Vandamme and co-workers demonstrated that for different flocculation methods on C. vulgaris, all parameters will need to be proportionately increased in order to achieve 85% algal removal when the amount of AOM was increased [14]. However, Garzon-Sanabria and co-workers highlighted that an increase in salt concentration from 5 g L−1 to 35 g L−1 in the absence of AOM did not influence the efficiency of flocculation of Nannochloropsis salina, requiring only a small dose of 4 mg L−1 of the polyelectrolyte flocculant Flopam [89]. Whilst evaluating the removal efficiency in the presence and absence of AOM, these authors reported the need to increase the dosage of polyelectrolyte flocculants by about 7 times from 3 mg L−1 to 20 mg L−1 and by about 10 times from 5 mg L−1 to 50 mg L−1 for chitosan to achieve the same level of FE. Lam and co-workers also demonstrated the efficiency of cationic polyelectrolyte on marine species at very low dosages. From these studies it could be concluded that the carbohydrates present in the culture medium are more likely to inhibit flocculation or reduce FE than the salt concentration [85]. 3.3.2. Properties of different microalgal strains The degree of flocculation is dependent on the extent of polymer coverage of the active sites on the algal cell surface. If the coverage by the polymer is too low it will result in inadequate electrostatic bridging [52]. This goes to suggest that not all polymers are suitable for all strains of microalgae. Wu and co-workers demonstrated this hypothesis with high MW polyacrylamide (PAM) that showed different FE with two green microalgae of the same genera viz., Scenedesmus sp. and Scenedesmus obtiquus [94]. The only difference between the two species used in this study was the cell size, viz., ~8.3 μm for Scenedesmus sp. and ~ 12.5 μm for Scenedesmus obtiquus. This study further demonstrated that a flocculant dosage of 15 mg L−1 and 100 mg L−1 was required to achieve a FE of 95% respectively for Scenedesmus sp. and Scenedesmus obtiquus. Bleeke and co-workers also reported differences in FE attributed to cell size [50]. Cells of Chlorella sp. registered a FE of b 90%, whereas Chlamydomonas reinhardtii only registered an FE of 20% when a low MW and high cationic charge Polydiallyldimethylammonium chloride (polyDADMAC) derived from tannin (Poly Separ® CFL25) was used for flocculation. This could be explained by the smaller cell size of Chlorella sp. (2–10 μm diameter) and therefore smaller surface area when compared to the cells of Chlamydomonas reinhardtii (10–30 μm

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diameter). Polymers of low MW are entirely absorbed onto the surface of the cell forming regions of cationic charge. These regions can bind negatively charged regions of other algal cells. Larger cells may not bind effectively, potentially leading to a lower FE. Furthermore, this study demonstrated that species with smaller cell size required 30 mg L−1 of cationic flocculants (KW100) to achieve a FE of 90% in contrast to the larger cells that required 60 mg L−1. The degree of flocculation is dependent on the extent of polymer coverage of active sites on the surface of the algal cell, although this surface must be reduced to the minimal distance of separation imposed by electronic repulsion for the polymer to be able to bridge [52]. Tenney and co-workers reported an optimal dose of 2.5 mg L−1 for cationic polyelectrolyte flocculants used for harvesting chlorophytes [52]. Bilanovic and co-researchers demonstrated that cationic polyelectrolyte dosage required for flocculation range between 1 and 10 mg L−1 to flocculate freshwater strains [53]. It has already been shown in the literature that flocculant dosage depends on the proprieties of the flocculants and the culture medium. Roselet and co-workers highlighted that charge density of a polyelectrolyte flocculant is inversely related to the dosage needed for flocculation [11]. It was concluded that a dosage of 45 mg L−1 was required for polymers with lower charge density and 0.55 mg L−1 for ones with higher charge density. Flocculant dosage is also dependent on the cell concentration in algal cultures. Cationic polymer dose was reported to be linearly correlated with microalgal cell concentration, in the range of 100 to 350 mg DW L−1. At lower algal cell concentrations, the dosage of polymer required is reduced [19,52,95]. Granados and co-workers reported that for an optimal polyelectrolyte efficiency, the biomass concentration in the media need to be in the range of 0.5–2.5 g L−1 [49]. This works out well as the cell concentrations at the time of harvest in most algal mass culture systems are within this range. Gerde and co-workers studied flocculation in cultures of marine microalgae Schizochytrium limacinum (18 ppt seawater) and freshwater microalgae Chlamydomonas reinhardtii and Scenedesmus spp. at three different cell concentrations using aluminium sulphate (Al2(SO4)3) and two cationic starches with different charge densities, viz., DS02 and DS05 [61]. The authors reported that the cationic starch DS05 with the higher charge density was most effective for the three species requiring 10-fold less flocculants than Al2(SO4)3. The cationic starch DS05 was much more efficient at higher cell densities, probably attributed to higher efficiency in the interaction of the cells with the charges carried by the starch flocculant. It was concluded that at higher cell concentrations, the probability of the cationic group in the polymer to interact with more cells was reported to be greater. However, at higher flocculant concentrations the three species registered a decline in FE. This could be attributed to the flocculant concentration exceeding the optimum concentration resulting in an excess of positive charge that causes (i) stabilisation of the cell particles in suspension due to neutralisation of surface charges, (ii) floc instability due to repelling of like charges, and (iii) excess flocculants causing steric hindrance [14,96,97]. The effectiveness of different flocculant dosages for different strains depend on the zeta potential of cells of different microalgal strains. Zeta potential is the potential difference between the bulk fluid and the layer of counter ions that remain associated with the charged particle when the particle is moving through the solution. The zeta potential can be easily estimated from the mobility of the charged particles in an electric field, and is therefore a useful indicator of the degree of repulsion between charged particles in suspension [14]. A study by Henderson and co-workers observed that the flocculation of four algal species was dependent on their zeta potential, with maximum flocculation efficiencies at zeta potentials between −8 and +2 mV after the addition of the flocculants [98]. To achieve a FE of 90%, Gerde and co-workers concluded that the dosage of cationic starch required to flocculate Schizochytrium limacinum was 10 mg L−1 (zeta potential − 9.97 mV), 20 mg L−1 for Chlamydomonas reinhardtii (zeta potential − 19.95 mV) and 30 mg L−1 for Scenedesmus spp. (zeta potential −20.60 mV) [61].

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3.3.3. Properties of the culture media Many polymer flocculants are highly pH dependent. The effect of pH needs consideration as it not only influences the charge of the chemical flocculants but also the surface charges of an algal cell. Surface charges imparted on a cell surface due to pH is dependent on the type and number of charged groups such as amines, carboxylic acid and phosphates present on the cell surface, which would then interact with the flocculants [99]. Hence, pH of the culture medium impacts on the interaction of flocculants with these charged groups, thus influencing its flocculation efficiency [99]. Studies have shown that the efficiency of chitosan was optimal at a pH range of 8–10, as the microalgal cells were more negatively charged at this pH range, thereby inducing the strongest electro-attraction with a cationic polymer [15,92]. However, at higher pH levels of 10.2, surface charges on an algal cell is neutralised. Harith and co-researchers demonstrated a pH based strategy to obtain a FE of up to 90% for the anionic polymer LT25 for the recovery of Chaetoceros calcitrans [100]. Although an increase in pH before flocculation improves FE, it renders the media non-usable for recycling thereby impacting on the cost-effectiveness of the process. Tenney et al. demonstrated an increased efficiency for the cationic polymer C-31 from Dow at a pH between 2 and 4 [52].This has been attributed to the zeta potential, which is negligible at lower pH [101], leading to a reduction in the electrostatic repulsion between algal particles. This also brings about an increase in the rate of polymer bringing due to the expansion of the polymer chains. More recently, Morrissey and co-workers explored the potential of a new class of flocculants called polyamphoteric flocculants that comprise of both positively and negatively charged components, thus emancipating from the pH dependence of these flocculants [102]. In order to reduce their water footprint and to recycle residual nutrients, some commercial algal cultivation plants reuse the supernatant resulting from the flocculation process. For such processes, the nontoxic nature of the flocculants is of paramount importance, and especially so if the harvested biomass has to be utilised for high value and lowtoxicity applications such as food and feed. Studies on media re-use in batch culture systems after flocculation using polyelectrolyte flocculants followed by dewatering by centrifugation [50] or filtration [49] showed no inhibition in the growth of microalgae in the culture. These polyelectrolyte flocculants included Actipol range of flocculants (e.g., EM1, EM16, EM22, FB1, etc) and Poly Separ PK55H. In mass cultivation of microalgae where the supernatant media during the flocculation process is recycled, there will be a point in time when it will become necessary to replace the media with fresh media either due to depletion of nutrients in the media [101] or due to the excess build-up of metabolites that could potentially exert a feedback inhibition in microalgal cultures. 3.4. Applications of commercial polyelectrolyte flocculants in harvesting microalgal biomass In the last decade or so, with significant advancements in polymer engineering, several commercial polyelectrolytes with different molecular weights and charge densities have become available in the market. Polyelectrolyte flocculants are synthetised in water with oil emulsion at high temperature and are found to contain between 25% and 50% of the active polymer [43]. However, the drawbacks associated with increasing the percentage of active sites on the polymer is an increase in the viscosity of the polymer solution. Any increase in the viscosity could adversely affect the synthesis of high molecular weight polyelectrolytes commercially [45]. As the manufacturing process requires high temperature with the addition of surfactants to enable the water-in-oil emulsion to break and invert to oil-in-water emulsion, it is difficult to remove the oil needed for the emulsification process from the polyelectrolyte, especially when the end use application is for potable water treatment or for other clean flocculation processes. Significant improvements have been made to the photo-polymerisation process used in a range of cationic, anionic and non-ionic polyelectrolyte flocculants [103]. In the manufacturing process the photo-

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polymerisation occurs in a dedicated reactor with a monomer feed. Various types of monomers and chain terminators yield a gel sheet, which is further treated and dehydrated for a final granular product. This process is easy to manipulate and is precise in the manufacture of products with a specific molecular weight and charge density, accomplished by just changing the feed composition. The final product is non-toxic for end use application such as food and feed. Commercial organic polyelectrolyte flocculants have been demonstrated in the literature to be efficient for a range of applications. The process, though not widely adopted, is promising, efficient, economic and non-toxic for the harvest of microalgae. An analysis of the economics of the use of polyelectrolyte flocculants to harvest different microalgae is summarised in Table 2. If one were to factor in the recovery efficiency and the cost of recovery of algal biomass, organic polyelectrolyte and tannin-based flocculants stand out. Another important consideration is that the dosages required for effective flocculation for these flocculants are also low. An analysis of commercial flocculant characteristics with their efficiency to flocculate different microalgae species under different culture conditions is summarised in Table 3 for cationic flocculants, Table 4 for anionic and Table 5 for non-ionic polyelectrolyte flocculants. Some of the most popular and widely used commercial flocculants are Flopam series (SNFFloerger) and Magnafloc/Zetag (BASF). Whilst anionic polyelectrolyte flocculants have been predominantly used to flocculate fresh water microalgae, both cationic and non-ionic polyelectrolyte flocculants have been used to flocculate both fresh water and marine microalgae. Commercial polyelectrolyte flocculants are generally inexpensive and are easy to scale up. The efficiency of organic flocculation using polyelectrolyte polymers have been demonstrated by other researchers and have been proven to have many advantages including low-toxicity of residues to humans and animals and biodegradability [104]. The efficiency of these flocculants are linked to not only their own intrinsic proprieties [50] but also to proprieties of the microalgal cell [105] and composition of the medium [89]. Naturals polymers such as chitosan or cationic starches are effective in flocculating freshwater microalgae [14], but are pH dependent [92,106,107]. These flocculants are not very effective in harvesting microalgae cultivated in brackish or seawater [53]. Chitosan is also cost prohibitive. Poly (γ-glutamic acid) or γPGA is another natural microbial flocculant, but its efficacy is salinity dependent and require a longer flocculation time [108]. Commercial polyelectrolytes are therefore useful in the recovery of microalgae at concentrations ranging from as low as 2 to 25 mg g− 1 [49]. This is in contrast to other flocculants such as aluminium or iron salts and chitosan that are required at much higher dosages ranging from 200 to 270 mg g−1 [92]. There is a paucity of literature comparing efficacies of commercial flocculants under comparable conditions, with few studies focussing on small batch harvest with less emphasis on final product or biomass quality. However, harvesting microalgae at scale by low energy and low toxicity technique such as polyelectrolyte flocculation without contaminating the biomass is paramount in addressing to the global demand for food and feedstock. 4. Conclusion For any algal biomass harvesting technique, cost and efficiency are of paramount importance. Bulk harvesting of microalgal biomass by polyelectrolyte flocculants is regarded to be one of the most economically viable techniques, with residues that are not toxic to humans and animals making them suitable for various industrial processes and applications. Even though the cost of polyelectrolyte flocculants are variable, ranging between US $1.50 kg−1 and US $7.50 kg−1, it is important to take into consideration the dosage rate when evaluating a flocculant for a particular application such as utilisation of the harvested residue free clean microalgal biomass in nutraceuticals, cosmeceuticals, pharmaceuticals, food and feed. The cost of polyelectrolyte flocculants to harvest a metric ton of microalgal biomass is estimated to vary between US $35 to $40, in contrast to ~US $800 using a plate centrifuge.

Whilst most commercially available natural and synthetic flocculants have been evaluated for their performance at laboratory scale under controlled conditions, there is a paucity of literature on studies comparing the efficiencies of various commercial flocculants at scale, especially in outdoor mass culture systems. There is growing evidence that the efficiencies of flocculants determined in jar tests under laboratory conditions are often quite different to the efficiencies measured under outdoor mass culture conditions, attributed to changes in prevailing physico-chemical conditions in the media such as pH, dissolved oxygen, temperature, etc. To avoid discrepancies in performance, it is critical that flocculants are evaluated under the same conditions under which the cultures would be harvested. Conclusions can be drawn from this review that the high charge density possessed by cationic polyelectrolytes render them potentially promising in harvesting high density marine algal cultures at scale. Although chitosan is a natural flocculant that is biodegradable and nontoxic, it is expensive and is pH dependant, and therefore may not render itself to be a suitable choice for use at scale for bulk microalgal biomass harvesting. Anionic polyelectrolytes are generally a poor performer but can be efficient at higher dosages with or without pH adjustment. Nonionic polymers do not work with microalgal cultures. In summary, it is important to consider that the effectiveness of a flocculant will vary considerably with the species or strains in question, therefore needing strain specific testing. Modern day industrial scale microalgal cultures aim for a smaller environmental foot print and with that comes a significant emphasis on reuse of culture media. This review identified a gap in published literature on the reuse of the culture medium after flocculation with low toxicity flocculants. There was also a paucity of information on the ‘biochemical cleanliness’ of the biomass harvested using flocculants for high value applications such as food and feed that requires a clean biomass. Further research is needed to underpin the techno-economic viability of the use of commercial flocculants at scale for a bulk microalgal biomass production for low toxicity applications.

Funding This research did not receive any specific grant from funding agencies in the public, commercial or not-for-profit sectors.

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