Flocculation and electroflocculation for algal biomass recovery

CHAPTER

Flocculation and electroflocculation for algal biomass recovery

11

Tawan Chatsungnoen⁎, Yusuf Chisti† Department of Agro-industrial Biotechnology, Maejo University-Phrae Campus, Phrae, Thailand* School of Engineering, Massey University, Palmerston North, New Zealand†

1 ­INTRODUCTION This chapter focuses on flocculation and electroflocculation methods of separating the algal biomass from the culture medium. Separating the biomass from water contributes around 20%–30% to the total cost of producing the dry algal biomass [1–3]. Several species of microalgae and cyanobacteria are produced commercially mainly as feeds for aquaculture, human foods, and sources of high value products [4–7]. In nearly all these applications, the algal biomass is typically recovered from the culture medium by centrifugation and occasionally by filter screens. Biomass recovery by these methods is expensive to the point of being unfeasible if the product of interest has a low value. Algal oils for use as fuel feedstock are examples of such low-value products [8–10]. Algal oils, starch and other precursors for making biofuels must be produced in huge amounts to make any meaningful impact on fuel supply. Therefore, algal biomass must be produced in large quantities and a biomass recovery operation must be able to process large volumes of algal culture cheaply. At harvest, the biomass concentration in a photoautotrophic cultivation system is generally low because light limitation does not allow the culture to grow to a high density [8, 9]. In open pond types of culture systems commonly used in outdoor production of algae, the maximum dry biomass concentration is around 0.5 g L−1 [4, 9, 11]. The maximum dry biomass concentration may reach around 4–5 g L−1 in some of the closed commercial photobioreactors [8]. Both these concentrations are quite low: recovery of 1 kg of dry biomass requires processing of between 200 and 2000 L of algal culture. Other factors complicate biomass recovery. Individual cells of microalgae are small, with a diameter between 3 and 30 μm. In addition, the density of the cells is nearly the same as the density of water, therefore, gravity sedimentation of individual cells is difficult [1, 12–14]. The cells typically carry a net negative surface charge, and the consequent mutual repulsion prevents them from coagulating, or flocculating. Biomass, Biofuels and Biochemicals. https://doi.org/10.1016/B978-0-444-64192-2.00011-1 © 2019 Elsevier B.V. All rights reserved.

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FIG. 1 Dewatering by flocculation-sedimentation. A microalgal broth before (A) and after (B) flocculation-sedimentation. The entire treatment was completed in about an hour. Based on T. Chatsungnoen, Y. Chisti, Harvesting microalgae by flocculation–sedimentation, Algal Res. 13 (2016) 271–283.

Flocculation is the process of agglomerating the suspended cells by neutralizing the surface charge to minimize mutual electrical repulsion. Agglomeration increases the average size of a composite particle, or floc, to improve its speed of settling under gravity [1, 15]. Flocculation followed by gravity sedimentation is a relatively cheap method that is well established in large-scale water treatment processes for removing suspended solids from water. This proven method can be potentially extended for removing much of the water from an algal culture [13, 14, 16–20]. Efficacy of flocculation in settling cells out of suspension is demonstrated in Fig. 1. Although no single biomass recovery scheme can be prescribed as being universal [16, 21], based on efficiency and cost, flocculation followed by sedimentation is perhaps the most suitable primary concentration method for recovering microalgal biomass from the culture media [1, 13, 16–19] so long as the application at hand allows its use.

2 ­FLOCCULATION AND ITS VARIANTS Flocculation is achieved by adding chemicals, or flocculants, to reduce or neutralize the negative surface charge on cells [1, 22]. The flocculants used may also help to bridge cells loosely together, and also operate by other mechanisms. Flocculation works with a wide range of microalgal species and other suspended particles [14, 23]. Although flocculation has been widely used to remove microalgae mostly from water

3 ­ Flocculation and electroflocculation as primary dewatering methods

and wastewater [12, 24–33], it is rarely used in commercial production of algae. This is because contamination of aquaculture feeds, nutraceuticals, and human foods by a flocculant chemical is not wanted. While food-safe flocculants are available [34], they are generally expensive. Variants of flocculation include electroflocculation, microbial flocculation (bioflocculation), autoflocculation, and ultrasonic flocculation. Electroflocculation is also known as electrolytic flocculation or electro-coagulation-flocculation [19, 23, 35–41]. In electrolytic variants of flocculation, the surface charge on the suspended cells is neutralized by metal cations (e.g., Al3+, Fe3+) generated in situ from a sacrificial metal anode by application of an electric current in an electrolytic reactor. The cells lose their surface charge by complexing with the metal cations and agglomerate to form flocs [35, 37]. Electroflocculation is highly efficient [23, 36, 37, 42]. It has an electric power consumption of around 0.3 kWh kg−1 of dry algal biomass [35]. This is low compared to centrifugation, which requires almost 2 kWh kg−1 of the dry biomass recovered [16]. Many pilot-scale operations have used electroflocculation for harvesting microalgal biomass [16]. Electroflocculation has been demonstrated with many algae, including Chlorella vulgaris, Phaeodactylum tricornutum [37], Botryococcus braunii [42], Dunaliella salina [43], Scenedesmus acuminatus [40], and Tetraselmis sp. [38]. In microbial flocculation, or bioflocculation [3, 44, 45], the flocculating agent is produced by one or more microorganisms present inadvertently in the algal culture, or deliberately co-cultured. A bioflocculant may also be produced in a separate dedicated microbial process and then added to the algal culture [44, 46, 47]. Autoflocculation [13, 48] is a result of an alga producing a flocculating substance at some stage of its growth. Autoflocculation is probably specific to some algae and culture conditions. In some cases, this is due to the pH rise in cultures with an insufficient supply of carbon dioxide. It may be useful in specific cases only. Ultrasonic flocculation [49–51] is brought about by passing a suspension of cells through a standing wave ultrasound field. Because of the forces generated by the sonic field, the cells migrate and concentrate in certain regions (the pressure nodes) of flow and drop out of suspension by gravity. This method appears to be of some use in small-scale applications [49]. Of the possible flocculation variants, only the conventional flocculation, relying on added flocculants, and electroflocculation are broadly useful for large-scale ­applications, as discussed in this chapter.

3 ­FLOCCULATION AND ELECTROFLOCCULATION AS PRIMARY DEWATERING METHODS Flocculation increases effective particle size through forming large flocs from individual cells. It is essentially a pretreatment step to facilitate cell separation by other means such as gravity sedimentation, filtration, centrifugation, and flotation [1, 12, 21]. An ­algal culture typically contains 0.5–5 g L−1 suspended biomass (dry basis).

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Flocculation followed by gravity sedimentation thickens the algal slurry to a dry biomass concentration of 20–70 g L−1, or more. At this stage, the thickened slurry has 93%–98% water by weight. Further water removal is feasible through centrifugation or filtration. The wet biomass paste recovered from the centrifuge, or the filter, would have a dry biomass concentration of 150–250 g L−1 and a water content of 75%–85% by weight. Further moisture removal would require some form of thermal drying, including freeze-drying. Flocculation and gravity thickening prior to centrifugation or filtration greatly reduce the volume of the slurry that needs to be centrifuged or filtered. This reduces the capital and energy requirements of solids recovery, as both centrifugation and filtration tend to be expensive. Dehydration beyond the biomass paste stage (75%–85% moisture by weight) may not be necessary if the application is aquaculture feed, human food, or extraction of certain products such as oils [52]. Drying of the biomass to a powder (≤2% moisture by weight), improves keeping quality but is economically feasible only for exceptionally high value applications.

4 ­MICROALGAL CELL AND SUSPENSION PROPERTIES A suspension of microalgal cells may be thought of as a hydrophilic bio-colloid. The dispersed cells (i.e., the discontinuous phase, or dispersed phase) are distributed uniformly in an aqueous medium (i.e., the continuous phase) [17, 24]. The suspension is generally quite stable (Fig. 1A). That is, the cells typically do not settle out, or coagulate, if the suspension is left standing. The small size of the cells and their surface charge are the two factors contributing to stability of the suspension. Usually, the cells carry a net negative electric charge on the surface due to ionization or dissociation of the surface functional groups such as COOH. The negatively charged cells mutually repel due to electrostatic repulsion. This keeps the cells apart and prevents them from coalescing into larger aggregates. The small size of the cells and a density that is close to that of the suspending medium [12, 53] result in a low settling velocity under the normal gravitational field. In addition to electrostatic repulsion, a mutually attractive van der Waals force [24] also exists between cells, but this is insufficient to overcome electrostatic repulsion. A negatively charged cell suspended in a nutrient medium containing many cations (positive ions) pulls the cations from the surrounding medium toward itself and repels the negative ions (the anions) from the vicinity (Fig. 2). Thus, an electrical double layer exists at the surface of the cell [15, 53–55]: the surface bound negative charge surrounded by an adjacent layer of cations (Fig. 2). The layer of cations adjacent to the cell surface is also known as the Stern layer. This layer is static, or rigid. The Stern layer is surrounded by a diffuse layer of cations and anions (Fig. 2). The diffuse layer is rich in cations near the vicinity of the Stern layer, but at its outer limit the distribution of charges becomes equal [17, 56]. Each cell is surrounded by an attached Stern layer [12, 24, 56] that moves with the cell. The interface between the Stern layer and the surrounding diffuse layer is known

4 ­Microalgal cell and suspension properties

FIG. 2 A negatively charged algal cell surrounded by the rigid Stern layer of cations and the diffuse layer. Zeta potential is the electrical potential at the interface of the Stern layer and the diffuse layer. Based on J. Bratby, Coagulation and Flocculation in Water and Wastewater Treatment, 2nd ed., IWA Publishing, Seattle, 2006.

as electro-kinetic face, or the plane of shear (Fig. 2). The electrical potential at the plane of shear is known as the zeta potential [56, 57]. Zeta potential characterizes the electrical charge on a particle surrounded by its double layer and is a measure of the stability of suspension. A particle with a net negative charge has a negative zeta potential. Reducing the magnitude of zeta potential

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reduces electrostatic repulsion among particles. At some point the zeta potential may be reduced to a critical value at which the van der Waals attractive forces between particles overcome the electrostatic repulsion and particles agglomerate [53]. The zeta potential of microalgae is usually in the range of −10 to −35 mV [53]. The intensity of the surface charge, or the areal density of the charge, depends on the zeta potential and the cell surface area. For a given zeta potential, a cell with a larger surface area will have a lower surface charge density. Charge density depends on the microalgal species, the phase of growth, the ionic strength of the culture medium, the pH, and other environmental factors [2, 12, 58]. For example, the zeta potential of Tetraselmis suecica and Chlorococum sp. was around −43.2 mV in the exponential growth phase, but rose to −34.5 mV in the stationary phase [2]. Similarly, the zeta potential of Chlorella zofingiensis was found to increase as the cells transitioned from exponential growth to the stationary phase [58]. Changes in zeta potential during growth are at least partly linked to changes in the surface functional groups and the cell size during growth. The functional groups known to occur on cell walls of microalgae include the carboxyl (COOH), phosphate (PO4−2), amino (NH2), and hydroxyl groups (OH) [59]. Another factor affecting the stability of an algal suspension is its ionic strength [55, 60]. The ionic strength of a culture medium is a measure of the concentration of active ions in the medium. In a medium with a low ionic strength (i.e., a low salt concentration), the diffuse layer around the cell is thicker and prevents the cells from coming into contact (Fig. 3A). In a more saline medium, the diffuse layer thins and the cells can approach closer (Fig. 3B). This may allow the van der Waals attractive force to overcome the double layer repulsion. In a medium of high ionic strength, the zeta potential of a cell is typically lower than the zeta potential of the same cell in a medium of low ionic strength.

5 ­CONVENTIONAL FLOCCULATION Flocculation is a three-step process. First, the flocculant is rapidly dispersed in and mixed with the algal suspension. This destabilizes the suspension and flocs form. The next step requires an extended period of gentle mixing to allow the flocs to grow. Growth is a result of particle-particle interactions. Therefore, mixing must be sufficient to achieve this, but not so intense as to rupture the growing flocs. Once the flocs have attained a certain size, the mixing must cease to allow the flocs to settle under gravity. In a batch flocculation tank, the initial mixing, floc growth, and sedimentation occur in the same vessel and therefore the intensity of agitation varies at different stages of the process. In continuous flocculation-sedimentation, zones of intense mixing, floc growth and sedimentation are spatially separated, often within a single process unit. Flocculation of suspended cells with a flocculant involves at least two mechanisms that occur concurrently. First, the suspension is destabilized by a flocculantmediated change in the surface properties of the cells. The mutual repulsion among cells is reduced and they come together to form flocs. Second, the flocs grow in

5 ­ Conventional flocculation

FIG. 3 Interaction of negatively charged cells suspended in media of different ionic strengths: (A) a low ionic strength freshwater medium; and (B) a high ionic strength seawater medium. The diffuse layer of counterion around each cell extends to a greater distance from the cell surface in media with low ionic strengths (A) compared to media with a high ionic strength (B). The cells approaching each other begin to experience mutual repulsion when their diffuse layers overlap [60] and, thus, are kept further apart in media with low ionic strength (A) but can approach closer in media with a high ionic strength (B). Adapted from J. Gregory, Particles in Water: Properties and Processes, IWA Publishing, Seattle, 2006.

size by attaching more cells and are settled out by gravity [61]. A third mechanisms known as sweep flocculation operates under some conditions and is discussed separately (Section 5.1.4).

5.1 ­FLOCCULANTS Flocculants are the agents used to bring about flocculation. Chemical flocculants are highly effective and widely used. Inorganic flocculants or polymeric organic flocculants [1, 12, 17, 18, 24] may be used. For use in large-scale operations, as anticipated for production of microalgae biomass for fuels, a flocculant must meet certain key requirements: it must be effective, cheap, environmentally benign, and readily available. Only certain inorganic flocculants (e.g., aluminum sulfate, or alum, and ferric chloride) meet all these criteria. Many other effective organic flocculants have been developed [27, 28, 34, 62], but none is as cheap as the commonly used inorganic salts [16, 27, 63].

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5.1.1 ­Organic flocculants

Organic flocculants are mostly polymers. They may be polyelectrolytes, that is, polymers carrying anionic or cationic charge, or uncharged non-ionic polymers. They may be synthetic or natural [12]. Examples of natural polyelectrolytes include the polysaccharides cationic starch and chitosan (a cationic polymer) and the polypeptide poly-γ-glutamic acid (a cationic polymer) [30, 64]. Among synthetic polyelectrolytes, polyacrylamides (either cationic or anionic) are widely used. Polyelectrolytes are further reviewed by Haver and Nayar [34]. Polyelectrolytes act through a combination of cell surface charge neutralization and particle bridging to form flocs [24]. The effectiveness of polyelectrolyte flocculants is influenced by the following factors: the molecular mass or chain length of the polymer; the charge density on the molecule; the dose used; the biomass concentration; the ionic strength and pH of the broth; and the mixing in the fluid. High molecular weight polyelectrolytes (i.e., longer chain polymers) are better bridging agents. A high charge density tends to unfold the polymer molecule, which improves its ability to neutralize the surface charge on cells and its bridging performance. A high cell concentration in the broth helps flocculation by increasing the frequency of the cell-cell encounters. Cationic polyelectrolytes such as chitosan can induce efficient flocculation of freshwater microalgae at low dosages [65]. A chitosan dosage in the range of 1–10 mg L−1 may be quite effective with freshwater microalgae [65]. Flocculation efficiency declines in higher salinity media [13, 26, 66], because in such media the molecule of the polymer tends to fold onto itself [65], necessitating a higher dosage. For example, a 100% flocculation efficiency of marine microalgae required a chitosan flocculant dosage of above 40 mg L−1 [25]. The flocculation efficiency of cationic polymers such as chitosan is highly dependent on pH with a pH of 7 being optimal for flocculation in low salinity media [67]. Chitosan-based flocculants are further discussed by Yang et al. [68]. Flocculation with cationic starch is discussed by Vandamme et al. [27].

5.1.2 ­Inorganic flocculants

Inorganic salts of multivalent metals are effective flocculants. The multivalent metal cations in these salts neutralize the cell surface charge and bridge cells together to facilitate flocculation [15, 66]. Salts of aluminum and iron are the most widely used [15] because of their efficacy, availability, safety and relatively low cost. The aluminum-based flocculants include aluminum sulfate, aluminum chloride, sodium aluminate, aluminum chlorohydrate, and polyaluminum chloride. The iron-based flocculants include ferric chloride, ferric sulfate, ferrous sulfate, and ferric chloride sulfate [15, 69]. Aluminum sulfate or alum (Al2(SO4)3), ferric chloride (FeCl3) and ferric sulfate (Fe(SO4)3) are the most widely used flocculants [1, 70]. These flocculants have a long history of use in removing colloidal particles from water and wastewater [15, 17]. A higher dose of a flocculant salt is generally necessary for flocculation of cells suspended in a medium of high ionic strength (e.g., seawater) compared to the dose needed for flocculation from a freshwater medium [26]. Metal salts have proven effective for flocculation of microalgal biomass suspended in the culture medium (Table 1). Aluminum salts are generally more ­effective

Table 1  Some inorganic flocculants used for harvesting microalgal biomass from the culture broth

Flocculant

48 90–100 30–40 10.8 750 500 500 200 225 ± 21 140 ± 15 25 ± 4 120 ± 15 55 ± 9 11 ± 4

Microalga Nannochloris oculata Phaeodactylum tricornutum P. tricornutum Nannochloropsis salina Chlorella minutissima C. minutissima C. minutissima Botryococcus braunii Isochrysis galbana Chlorella stigmatophora Chlorella vulgaris I. galbana C. stigmatophora C. vulgaris

Biomass concentration 7

−1

3.0 × 10  cells mL 3.11 × 106 cells mL−1 3.11 × 106 cells mL−1 1.0 g L−1 2.2 × 108 cells mL−1 2.2 × 108 cells mL−1 2.2 × 108 cells mL−1 – 1.0 × 106 cells mL−1 1.0 × 106 cells mL−1 1.0 × 106 cells mL−1 1.0 × 106 cells mL−1 1.0 × 106 cells mL−1 1.0 × 106 cells mL−1

Flocculation efficiency (%)

Reference

97 70–80 >90 99 >90 >90 >90 90 >97 >97 >97 >97 >97 >97

[71] [29] [29] [72] [73] [73] [73] [74] [26] [26] [26] [26] [26] [26]

Source: Adapted from T. Chatsungnoen, An Assessment of Inexpensive Methods for Recovery of Microalgal Biomass and Oils (PhD thesis). Massey University, New Zealand, 2015.

5 ­ Conventional flocculation

Aluminum chloride Polyaluminum chloride Aluminum sulfate Aluminum nitrate sulfate Aluminum chloride Ferric sulfate Zinc chloride Aluminum sulfate Aluminum sulfate Aluminum sulfate Aluminum sulfate Ferric chloride Ferric chloride Ferric chloride

Optimal concentration (mg L−1)

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flocculants than ferric and zinc salts because the small ionic radius of aluminum results in a higher surface charge density [12, 73]. Ionic radii (nm) of some common metal cations of interest are as follows: 0.053 for Al3+; 0.100 for Ca2+; 0.077 for Fe2+; 0.069 for Fe3+; 0.072 for Mg2+; and 0.074 for Zn2+. A combination of flocculants is sometimes more effective than any one flocculant. For example, in flocculation of Isochrysis galbana, a combination of an organic polyelectrolyte (chitosan) and an inorganic salt (ferric chloride) was more effective than using any one of these flocculants [26]. The overall degree of flocculation was improved and the dosage of the inorganic flocculant was reduced by using the two flocculants together. Similarly, Danquah et  al. [75] used a combination of a high molecular weight synthetic cationic polyelectrolyte polymer (Zetag 7650) and an inorganic salt (aluminum sulfate). Some life-cycle assessment studies suggest that inorganic flocculants such as ferric chloride may have a lower overall environmental impact compared to organic polyelectrolytes [76].

5.1.3 ­Flocculant dose

The optimal dose of a flocculant depends on many factors including the alga, its concentration, the nature of the culture medium (ionic strength, pH, concentrations of specific dissolved ions, concentration and nature of any dissolved organics), and temperature. Data on optimal dosages of some flocculants have been published. For example, the equations in Table 2 can be used to calculate the dosage needed for harvesting the specified algae from suspensions of a given biomass concentration. Under otherwise fixed conditions, the minimum flocculant dose needed to flocculate and sediment an alga fully, depends on the biomass concentration in suspension (Fig. 4). The flocculant dose increases with increasing biomass concentration (Table  2). Optimal dosages cannot be predicted and must be determined experimentally. Batch flocculation-sedimentation tests carried out in a laboratory jar test apparatus (Fig. 5) are used to determine experimentally the optimal dose. Six small identical beakers are filled to the same level (e.g., 200 mL) with the same sample of the algal culture. Each beaker has its own agitator (diameter = 5 cm) (Fig. 5). The beakers are dosed with a flocculant solution to different concentrations in different beakers. All beakers are then subjected to the exact same 3-step flocculation-sedimentation treatment: (1) 2 min of rapid mixing at 80 rpm to disperse the flocculant; (2) 30 min of gentle mixing (20 rpm) to promote floc growth; and (3) 30 min of a quiescent (no mixing) sedimentation period. The concentration of the biomass remaining in the clear supernatant sampled a distance of 2 cm below the surface of the liquid, is determined. Flocculation efficiency is calculated as the fraction of the initial suspended biomass removed from the supernatant. See Chatsungnoen and Chisti [14] for further details.

5.1.4 ­Ionic strength and pH effects

As previously noted, ionic strength and the pH affect the stability of suspension and the dose of a flocculant required for complete flocculation. The pH is a measure of the concentration of H+ ions. The pH affects the zeta potential of the negatively

Table 2  The flocculant dose dependence on the microalgal biomass concentration (Cb) for 95% removal of the biomass from the culture broth Microalga and medium

Equation

Correlation coefficient

Aluminum sulfate  (mg L−1) = 318.980  Cb  (g L−1) Ferric chloride (mg L−1) = 257.815  Cb (g L−1) Aluminum sulfate  (mg L−1) = 11.796  Cb  (g L−1) Ferric chloride  (mg L−1) = 32.228  Cb (g L−1) Aluminum sulfate  (mg L−1) = 503.156  Cb  (g L−1) Ferric chloride (mg L−1) = 265.252  Cb  (g L−1)

0.995 0.991 0.999 0.970 0.980 0.905

Aluminum sulfate  (mg L−1) = 62.903  Cb  (g L−1) Ferric chloride  (mg L−1) = 67.742  Cb  (g L−1) Aluminum sulfate  (mg L−1) = 229.576  Cb  (g L−1) Ferric chloride  (mg L−1) = 290.060  Cb  (g L−1) Aluminum sulfate  (mg L−1) = 154.594  Cb  (g L−1) Ferric chloride  (mg L−1) = 132.690  Cb  (g L−1)

0.956 0.990 0.996 0.976 0.976 0.983

BG11 freshwater medium Choricystis minor Neochloris sp. Chlorella vulgaris BG11 seawater medium

Nannochloropsis salina Cylindrotheca fusiformis

Source: Adapted from Chatsungnoen and Chisti [14].

5 ­ Conventional flocculation

Chlorella vulgaris

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FIG. 4 Dose dependence of flocculation. Each beaker contained 200 mL of a broth of the microalga Chlorella vulgaris (dry biomass concentration = 3.5 g L−1) grown in a seawater medium (BG11 medium made using full strength artificial seawater). The beakers were dosed with different concentrations of the flocculant (0–1000 mg L−1 aluminum sulfate), and then treated the same way: (1) rapid mixing (80 rpm, or impeller tip speed = 0.209 m s−1) for 2 min; (2) gentle mixing (20 rpm, or impeller tip speed = 0.052 m s−1) for 30 min; and (3) no mixing for 30 min.

FIG. 5 A jar test system for determining optimal flocculation dosage.

charged cells in suspension and, therefore, the flocculation behavior. The pH also affects the chemical character of certain flocculants. For example, a pH close to neutral is optimal for cationic polymer flocculants such as chitosan, as discussed earlier. Flocculation processes using metal cation flocculants are particularly affected by pH. The optimal range of pH for flocculation with aluminum sulfate is 6–7. With iron salts, the optimal use pH is 5.5–6.5. Lowering the pH by adding acid reduces the metal cation dose required for flocculation from an alkaline medium. In the presence of a high concentration of certain metal cations (e.g., Al3+, Fe3+, Mg2+), gelatinous precipitates of metal hydroxides (e.g., Al(OH)3, Fe(OH)3) form through reaction of the cations with water. These gelatinous precipitates (also known as sweep flocs) entrap other suspended particles and sweep them down during settling [37, 55, 60].

5 ­ Conventional flocculation

This process is known as sweep flocculation. Sweep flocculation requires a metal cation dose greater than necessary for charge neutralization of suspended particles. Although it requires high dosages of metal salt flocculants, sweep flocculation is more effective compared with flocculation by charge neutralization alone. Sweep flocculation is used in treatment of drinking water.

5.1.5 ­Cost of flocculants

Fuels are low value products and, therefore, the algal biomass for their production must be harvested especially cheaply. Inorganic salts are probably the least expensive among flocculants. For example, the bulk prices of aluminum sulfate (Al2(SO4)3·14H2O) and ferric chloride (FeCl3) in 2013 were US $318.90 and US $354.33 per metric ton, respectively [77]. On an unhydrated basis (i.e., for Al2(SO4)3), the price of aluminum sulfate was US $440.39 per metric ton. The mass (F, mg) of these flocculants needed for harvesting a metric ton (dry basis) of microalgal biomass can be estimated using the optimal flocculant dosage data provided in Table 2; thus, F ( mg ) = constant ( from Table 2 ) ´ 106

(1)

Therefore, the cost of flocculant (Cf, US$/ton biomass) required to harvest one metric ton of dry microalgal biomass is: Cf =

F ´ 106 ´ Pf 10 9

(2)

where Pf (US$/ton salt) is the price of the flocculant (anhydrous basis). The relevant costs are shown in Table 3, based on the experimental data in Table 2 and the flocculant prices noted above. The cost of a flocculant depended on the dosage and varied between US $5.19 and $221.58 per metric ton of dry algal biomass (Table 3). Which flocculant was cheaper, depended on the alga. For most cases in Table 3, ferric chloride was cheaper, but for Neochloris sp. and Nannochloropsis salina, aluminum Table 3  Cost of flocculant for harvesting microalgal biomass from the culture broth US $ per metric ton of dried microalgal biomass Alga and culture medium

Aluminum sulfate (Al2(SO4)3)

Ferric chloride (FeCl3)

Grown in the BG11 freshwater medium Choricystis minor Neochloris sp. Chlorella vulgaris

140.44 5.19 221.58

91.35 11.42 93.99

Grown in the BG11 seawater medium Chlorella vulgaris Nannochloropsis salina Cylindrotheca fusiformis

27.70 101.10 68.08

24.00 102.78 47.02

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sulfate was cheaper. In view of Table 3, generalizations cannot be made about which flocculant may be cheaper and the cost needs to be evaluated on a case-by-case basis. The cost of harvesting algal biomass using organic polyelectrolytes is in the range of US$35 to US$40 per metric ton of dry biomass [34]. Polyelectrolytes range in price from US$1500 to US$7500 per metric ton [34].

5.2 ­CONTINUOUS FLOCCULATION-SEDIMENTATION PROCESSES Although batch flocculation of microalgae has been extensively reported [2, 3, 14, 18, 25–27, 29–33, 35, 46–48, 63, 71–73, 75], most large scale flocculation-­sedimentation processes are operated on a continuous flow basis [78–80]. In contrast with studies in water treatment, relatively few studies have discussed continuous flow flocculation for biomass recovery from actual algal broths [20, 78–82]. In a large-scale continuous flocculation-sedimentation operation, the various steps may be carried out in separate vessels. For rapid mixing of the flocculant solution with the algal suspension, a standard well-mixed stirred tank may be used. Design of the mixing stage is important for achieving a rapid and uniform distribution of the flocculant [83]. The mixing tank may be circular or square in cross-section [15]. The geometry of a circular tank is such that the ratio of the depth of liquid to the diameter of the tank is 1 [84–86]. Usually a single impeller is used. The impeller diameter ranges between 50% and 80% of the tank diameter. The impeller is placed one impeller diameter above the bottom of the tank. The recommended retention time in the mixing tank ranges from 9 s to 2.5 min for aluminum sulfate concentrations ranging from 10 to 100 mg L−1 [15, 69]. The flocculation stage is focused on growth of the flocs. This stage requires gentle mixing to favor floc-floc interactions and agglomeration but not disruption so that the floc size increases for ease of sedimentation in the next step [15, 69, 86]. The flocculation tank can be rectangular or circular. For circular tanks the ratio of depth of water to the diameter of the tank is 0.9–1.1. The position of the mixing impeller is 0.33–0.5 impeller diameters above the bottom of the tank. The impeller diameter usually ranges between 0.3 and 0.6 of the tank diameter, and the recommended flocculation time ranges between 10 and 30 min [87]. The sedimentation, or clarification tank, is where the flocs settle under gravity. This tank can be designed with rectangular, square or circular cross sections [88]. Generally, long rectangular tanks are more effective compared to the other configurations [89]. A rectangular continuous horizontal flow sedimentation tank consists of four zones according to function: [90] the inlet zone, the settling zone, the outlet zone and the sludge zone as shown in Fig. 6A. The inlet zone is designed to disperse the suspension over the cross section of the tank. The setting zone is where all settling occurs. The outlet zone in where clarified effluent is collected over the cross section of the tank. The sludge zone is the zone at the bottom of the tank where the solids accumulate [69, 80, 89, 90]. Generally, the rectangular sedimentation tank is designed to be narrow and long with a length-to-width ratio of between 4:1 and 6:1 [69, 80, 83]. A model tank being used to sediment flocs of the

5 ­ Conventional flocculation

FIG. 6 A rectangular continuous flow sedimentation tank for microalgal flocs: (A) the typical tank geometry and (B) a model tank in use. Adapted from T. Chatsungnoen, Y. Chisti, Continuous flocculation-sedimentation for harvesting Nannochloropsis salina biomass, J. Biotechnol. 222 (2016) 94–103.

marine microalga Nannochloropsis salina is shown in Fig. 6B. The sedimentation is incomplete because of a relatively short retention time [20]. Although continuous flocculation-sedimentation systems with physically distinct stages for mixing the flocculant, floc growth and sedimentation are used, other system configurations combine these different operations within a single piece of equipment. One such commonly used type of vertical flow mechanical flocculator is shown in Fig. 7. The algal slurry and the flocculant solution are mixed in an inline mixer as they enter the flocculation tank. The upper zone of the tank is where gentle mixing and floc growth occur. The lower part of the tank is quiescent and this is where the flocs sediment. The thickened slurry of settled flocs leaves the vessel at the

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FIG. 7 A continuous vertical flow industrial flocculation vessel combining flocculant dosing, floc formation and growth, and sedimentation in different zones of a single unit.

bottom. The biomass-free clarified effluent leaves the tank as overflow. The tapered design of the agitator assures regions of different mixing intensity in the top and bottom parts of the tank. For a given rotational speed of the agitator, the tip speed of the impeller is higher in the upper part of the tank compared the lower part. Continuous flocculation-sedimentation is further discussed by Chatsungnoen and Chisti [20]. Continuous flocculation-sedimentation is quite effective for dewatering. A N. salina biomass recovery of 86% was observed for an aluminum sulfate dose of 229 mg L−1 and a hydraulic retention time of 148 min in a rectangular channel flocculation system. In a batch flocculation-sedimentation operation, the flocculant dose requirement was 50% lower for the same algal broth. In the continuous flow system, the distribution of the floc sizes tended to be broader because of the nonuniform residence times of the different elements of fluid.

5.3 ­EFFECTS OF MIXING Floc growth and final size in a flocculation tank are affected by mixing and turbulence [91]. Gentle mixing is necessary for effective floc–floc contacting and growth. Shear rates, or turbulence levels, for rupturing flocs may be estimated using methods discussed elsewhere [91]. Methods for calculating the shear rate in different mixing systems and other process devices are also discussed in the literature [91–93]. The mixing intensity in a flocculation tank is typically characterized in terms of a velocity gradient (G, s−1). The tank is designed for a G-value in the range of

6 ­ Electroflocculation

20–75 s−1. The retention time t (s) in the tank should be such that the dimensionless factor G×t, or Camp number, is in the range of 10,000 to 150,000. G is related to the power input in the tank, as follows: æ P ö G =ç ÷ è Vm ø

0.5

where P (W) is the power input in the mixing tank, V (m3) is its volume, and μ (Pa s) is the viscosity of the algal suspension. The tip speed of the agitator in a flocculation tank is typically 0.4 m s−1, but may be in the range of 0.2–0.6 m s−1. Entry velocity in the tank may range from 0.45 to 0.9 m s−1. The superficial liquid velocity based on the surface area of the tank should be in the range of 0.2–0.8 m s−1. A retention time of 20 to 60 min is satisfactory.

6 ­ELECTROFLOCCULATION Electroflocculation or electrolytic flocculation was first developed in 1903 and has since been used in treating municipal and industrial wastewaters [94, 95]. The metal cations needed for flocculation are generated in situ from a sacrificial anode using electricity. An external source of flocculants is not required [35], therefore, anions such as chloride (Cl−) and sulfate (SO42−) do not contaminate the water [35, 37]. Electroflocculation is effective for recovering the biomass of both freshwater and marine microalgae [38, 40, 51, 96].

6.1 ­OPERATIONAL PRINCIPLE An electroflocculation reactor is shown in Fig. 8. It consists of two electrodes immersed in the algal suspension and connected to an external supply of direct current [97]. The positive electrode, or anode, is made of a sacrificial metal such as iron or aluminum. The negatively charged cathode is made of a relatively inert metal such as stainless steel. The current flow between the electrodes is adjusted by changing the voltage of the direct current (DC) power supply [37]. The electricity oxidizes the anode to release the metal cations responsible for flocculation. The following reactions occur at an aluminum anode [38]: Al ® Al3+ + 3e – 2H 2 O ® O2 + 4H + + 4e – 2OH – ® O2 + 2H + + 4e –

The reactions occurring at the cathode are as follows: O2 + 4H + + 4e – ® 2H 2 O 2H 2 O + 2e – ® H 2 + 2OH – 2H + + 2e – ® H 2

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FIG. 8 An electroflocculation reactor. Metal is oxidized at the anode to release cations that react with water and the OH− generated at the cathode, to produce hydrated metal cation complexes. Oxygen is evolved at the anode and hydrogen is released at the cathode (DC, direct current).

Thus, oxygen is evolved at the anode and hydrogen is generated at the cathode (Fig. 8). The cathode must be relatively inert compared to the anode [94]. The metal ions produced at the anode are immediately hydrolyzed to form various metal hydroxide complexes (Fig. 8). This hydrolysis results in neutralization of the charge on the metal cations [60]. For example, Al3+ reacts with a water molecule to produce H+ and Al(OH)2+. The latter reacts with another water molecule to produce H+ and Al(OH)2+. Further reaction with water [60] produces Al(OH)30. The bubbles of oxygen and hydrogen produced at the electrodes may be entrapped in the flocs causing them to float to the surface. Thus, electrolysis may cause both flocculation and separation of flocs by flotation [35, 37]. This type of electroflocculation is known as electroflotation. If the intention is to cause flocculation, but not flotation of the flocs, the current density between the electrodes, or the applied voltage, should be kept relatively low to minimize electrolysis of water and generation of the gases at the electrodes [51]. Generation of gases cannot be entirely prevented as electrolysis of water requires a potential difference of only around 1.2 V between the electrodes.

6 ­ Electroflocculation

6.2 ­ELECTROFLOCCULATION EQUIPMENT Electroflocculation equipment typically consists of a rectangular or cylindrical tank with electrodes connected to a power supply (Fig. 8). The tank has associated piping and pumps for feeding and removing the algal suspension. In addition, it may have a mixing mechanism and a cooling jacket. All electrochemical reactions occur in this tank. Large commercial electroflocculation units use multiple rectangular electrode plates arranged horizontally (Fig. 9A), or vertically (Fig. 9B), to from channels through which the algal suspension flows [98]. The electrodes in the reactor may be wired in various ways. A parallel connection of monopolar electrodes is shown in Fig.  10A [98a]. Each of the alternating anodes and cathodes is directly connected to the power supply. All the anodes are externally connected to each other and all the cathodes are externally linked together. Therefore, the electrical current is split among the electrodes. An in series arrangement of monopolar electrodes is shown in Fig. 10B. In this case, only the outer anode and cathode are directly connected to the power supply. The intervening anode and cathode pairs are externally linked, but the only connection between electrode pairs is through the electrolyte (i.e., the algal slurry). The in parallel electrical arrangement (Fig. 10A) requires a lower voltage for a given flow of current compared to the in series connection (Fig. 10B) because of the higher electrical resistance of the in series arrangement. A third possibility is shown in Fig. 10C. In this arrangement, the outer monopolar anode and cathode are connected to the power supply. The sacrificial electrodes (the bipolar electrodes) are placed between the monopolar electrodes without any external

FIG. 9 Continuous flow electroflocculation units with horizontal (A) and vertical (B) configurations of electrode plates. Adapted from G. Chen, Electrochemical technologies in wastewater treatment, Separ. Purif. Technol. 38 (2004) 11–41.

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FIG. 10 Batch electroflocculation tanks with various connecting arrangements of electrodes: (A) monopolar electrodes connected in parallel; (B) monopolar electrodes connected in series; and (C) bipolar electrodes connected in parallel (DC, direct current). Adapted from M.Y.A. Mollah, R. Schennach, J.R. Parga, D.L. Cocke, Electrocoagulation (EC)—science and applications, J. Hazard. Mater. 84 (2001) 29–41.

connections. Once the monopolar electrodes are connected to a power source, the two sides of a sacrificial electrode acquire a charge opposite to that of the adjacent electrodes [95]. This type of electrode arrangement is easier to implement and maintain compared to the other arrangements [98] shown in Fig. 10. Electroflocculation may be carried out as a batch process, or a continuous flow operation. Batch studies are often used for an initial feasibility evaluation in the laboratory [99], but continuous flow operation is the norm in industrial processing. In a continuous flow operation, one or two electroflocculation tanks in series are followed by a quiescent sedimentation tank where the flocculated biomass settles and the cell-free effluent is discharged. Such a multitank system is shown in Fig. 11. A full-scale electroflocculation system would not generally exceed 40 m3 in volume. Multiple such systems may have to be used if larger capacity is desired [100].

FIG. 11 A continuous electroflocculation system with multiple tanks in series.

6 ­ Electroflocculation

6.3 ­FACTORS AFFECTING ELECTROFLOCCULATION Electroflocculation is influenced by the design features of the electrolytic equipment, the nature of the algal suspension and the way the electroflocculation is carried out. Some of these aspects are discussed in the following sections.

6.3.1 ­Types of electrodes

The nature of the anode used impacts electroflocculation in multiple ways. Both the efficacy of the flocculation and its cost (power consumption, electrode consumption) are affected. In principle, anodes made of many different metals may be used. Anodes made of iron, aluminum, magnesium, copper, zinc, and brass may be used for harvesting microalgal biomass [40]. In practice, Al3+ and Fe3+ ions are most effective for flocculating the biomass, and therefore anodes made of aluminum and iron materials are the ones most commonly used [95]. The flocculation efficacy of aluminum(III) and iron(III) is associated with their high charge and surface charge density compared to the other metal ions [97]. A high charge density enhances their ability to adsorb to and neutralize the charge on negatively charged particles in suspension [94]. More than 90% of the freshwater alga Scenedesmus acuminatus could be recovered by electroflocculation using aluminum or magnesium anodes and an applied voltage of 40 V [40]. Aluminum is less electronegative than iron and, therefore, aluminum anodes are electrically more efficient in generating ions compared to iron anodes. As a result, for otherwise similar conditions, aluminum anodes result in a more rapid destabilization of an algal suspension [37]. Iron electrodes cause discoloration of the biomass and the internal surfaces of the electrolytic tank through formation of dark brown iron oxides. Discoloration does not occur with aluminum anodes [43]. A cathode needs to be inert relative to the anode. Cathodes made of stainless steel and lead can be used [35, 94], although any possibility of contamination of the residual medium or the algal biomass with lead should be avoided in view of its toxicity. Cathodes made of metal oxides (e.g., Ta2O5, IrO2, RuO2, Sb2O2, SnO2) or mixed metal oxides (e.g., IrO2/TiO2) may be used [37].

6.3.2 ­Spacing between electrodes

Distance between the electrodes in an electrolytic unit is an important design variable that affects the cost of operation [94]. In harvesting of Dunaliella salina in a batch flocculation process, the flocculation efficiency was reduced from 93% to 75% when the distance between electrodes increased from 1 to 4 cm. The electrical energy consumption for flocculation increased with increasing distance between electrodes: with a 1 cm spacing, the energy consumption was 0.13 kWh m−3 but rose to 0.23 kWh m−3 with a spacing of 4 cm [43]. An electrode spacing of 1 cm was recommended [43]. A spacing shorter than 1 cm affected the formation of gas bubbles around the electrodes and this, too, resulted in an increased power consumption [43]. For a continuous flow electroflocculation process with a residence time of <1 s in the electrolytic tank, Koren and Syversen [94] recommended a spacing of 3 mm between the aluminum anode and stainless steel cathode.

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6.3.3 ­Electrical energy input

Electroflocculation requires electrical power. One of the objectives of a large-scale flocculation operation is optimization of power consumption. An increasing current density (A m−2) results in increasing rate of oxidation of anode and release of the flocculant cations [95]. Increasing current density therefore increases the rate of flocculation. In an electrolytic reactor of a fixed geometry, the current density is increased by increasing the voltage. The suitable voltage likely depends to some degree on the concentration of the biomass in suspension, the algal species and the properties of the culture medium (e.g., ionic strength, pH). In electroflocculation of the freshwater alga Scenedesmus acuminatus, a 90% flocculation of the biomass was obtained in 43 min at a voltage of 10 V [40]. Increase in voltage to 20 V shortened the time for 90% flocculation to 18 min. At an operating voltage of 30 V, 90% flocculation was attained in 12 min and an operation voltage of 40 V further reduced the time for 90% flocculation to 7 min [40]. Similar results have been reported with the freshwater alga Chlorella vulgaris and the marine alga Phaeodactylum tricornutum [37]. Thus, an increase in the current density from 15 to 120 A m−2 reduced the time required for 95% flocculation of C. vulgaris from 50 to 10 min [37]. Seawater-based culture media are rich in NaCl and the use of a high current density in such media results in the formation of sodium hypochlorite (NaClO), or bleach [37]. This may limit acceptable current density in harvesting marine microalgae. If the production of toxic chemicals such as bleach is suppressed, the electroflocculation process does not seem to harm microalgae. The cells remain viable and can be used as inoculum for a fresh batch [43]. Flocculation efficiency of the marine alga Dunaliella salina increased from 63% to 92% when the applied current density was increased from 30 to 90 A m−2 [43]. Under optimal conditions, Vandamme et  al. [37] reported a power consumption of 2 kWh kg−1 of the freshwater C. vulgaris biomass harvested. For the marine diatom P. tricornutum the power requirement was ~0.3 kWh kg−1 [37]. The lower power requirement in the marine medium reflected its lower resistance to flow of electric current and the relative ease of flocculation in a medium with a high ionic strength.

6.3.4 ­The pH

The pH affects electroflocculation in many of the same ways as explained earlier in the context of conventional flocculation (Section 5.1.4). During electroflocculation, the pH of the algal suspension increases rapidly. A suspension with an initial pH of around 7.5 becomes highly alkaline (pH >11) within 10 min at an operating voltage of 40 V [40]. How much the pH rises and the rate of rise depend on the type of metal anode used [40]. In electroflocculation of both freshwater alga Chlorella vulgaris and the marine alga Phaeodactylum tricornutum, increasing the initial pH from 4 to 8 reduced the harvesting efficiency [37]. Increasing pH actually increases the cation dose required for flocculation.

7 ­ Concluding remarks

6.3.5 ­Mixing intensity

As in conventional flocculation (see Section 5.3), mixing affects the electroflocculation process. Effective mixing reduces flocculation time [37], but intense mixing disrupts flocs and prevents growth. An optimal impeller tip speed for mixing in an algal electroflocculation process was reported to be 150 rpm in a 1 L rectangular tank [37]. The diameter of the mixing impeller was not specified [37], hence the tip speed of the agitator could not be calculated.

6.4 ­COSTS Electrical power and the sacrificial anode are the main consumables in an electroflocculation process [94]. Power consumption depends substantially on the operating conditions, including the current density, the electrode type and number, the surface area of the electrodes, the electrode spacing and the duration of operation [95]. The relationship between electrical power consumption (Pe, W), the applied voltage (U, V) and current (I, A) is as follows: Pe = UI

(3)

In terms of the resistance R (ohm) to flow of the electrical current between electrodes, the power consumption can be shown to depend on the electrical current, as follows: Pe = RI 2

(4)

Thus, the power consumption depends strongly on the current density. For a fixed current density, power consumption can be reduced by reducing the electrical resistance R, for example, by reducing the distance between the electrodes or increasing the electrical conductivity of the culture medium [94]. The total cost of harvesting the marine microalga Tetraselmis sp. by electroflocculation has been estimated to be US$0.19 kg−1 of biomass, or $190 per metric ton [38]. This included the cost of electricity, the replacement of anodes, and a 10% annual depreciation for a facility with a potential biomass processing capacity of 57,480 metric tons per annum [38]. Electricity usage contributed 49.2% to the total operational cost, whereas the contributions of electrode replacement and capital depreciation were 37.3% and 13.5%, respectively. Table 4 compares the cost of harvesting of microalgae by some of the methods available. According to this table, harvesting by electroflocculation appears to be cheapest, but conventional flocculation combined with sedimentation is relatively cheap compared to the other methods.

7 ­CONCLUDING REMARKS Flocculation and electroflocculation followed by gravity sedimentation are economically viable methods for substantially concentrating the microalgal slurries prior to further dewatering by methods such as centrifugation and filtration. Both flocculation and

279

280

US $ per metric ton of dry biomass Harvesting method

Year

Original

2017a

Items included in costing

Reference

Centrifugation (selfcleaning disc separator) Flocculation with flotation

1995

1710

2755

[22]

1988

1390

1889

Flocculation with sedimentation Electroflocculation

1988

370

768

2012

190

203

Plant depreciation, maintenance and energy Plant depreciation, maintenance, flocculant and energy Plant depreciation, maintenance, flocculant and energy Plant depreciation, electrode consumption and energy

a

[22] [101] [38]

US inflation was calculated from http://www.usinflationcalculator.com. Source: Modified from A.K. Lee, D.M. Lewis, P.J. Ashman, Harvesting of marine microalgae by electroflocculation: the energetics, plant design, and economics, Appl. Energy 108 (2013) 45–53.

CHAPTER 11  Flocculation and electroflocculation

Table 4  Cost comparison of different microalgae harvesting methods

­References

electroflocculation are readily scaled up and have an established history of large-scale use in treatment of water and wastewater. These methods require the use of either an externally supplied flocculant, or a flocculant generated in situ by the use of electricity. Diverse factors influence flocculation and electroflocculation as discussed in this chapter. Both batch and continuous flocculation operations are effective for dewatering, but continuous flocculation/electroflocculation is clearly preferred for processing large volumes of algal suspensions.

­REFERENCES [1] E. Molina Grima, E.H. Belarbi, F.G. Acién Fernández, A. Robles Medina, Y. Chisti, Recovery of microalgal biomass and metabolites: process options and economics, Biotechnol. Adv. 20 (2003) 491–515. [2] M.K. Danquah, B. Gladman, N. Moheimani, G.M. Forde, Microalgal growth characteristics and subsequent influence on dewatering efficiency, Chem. Eng. J. 151 (2009) 73–78. [3] S.  Salim, R.  Bosma, M.  Vermuë, R.  Wijffels, Harvesting of microalgae by bio-­ flocculation, J. Appl. Phycol. 23 (2011) 849–855. [4] Y. Chisti, Biodiesel from microalgae, Biotechnol. Adv. 25 (2007) 294–306. [5] M.A. Borowitzka, High-value products from microalgae—their development and commercialisation, J. Appl. Phycol. 25 (2013) 743–756. [6] M.A. Borowitzka, N.R. Moheimani (Eds.), Algae for Biofuels and Energy, Springer, Dordrecht, 2013. [7] F. Bux, Y. Chisti (Eds.), Algae Biotechnology: Products and Processes, Springer, New York, 2016. [8] Y. Chisti, Fuels from microalgae, Biofuels 1 (2010) 233–235. [9] Y. Chisti, Raceways-based production of algal crude oil, in: C. Posten, C. Walter (Eds.), Microalgal Biotechnology: Potential and Production, de Gruyter, Berlin, 2012, pp. 113–146. [10] Y.  Chisti, Constraints to commercialization of algal fuels, J. Biotechnol. 167 (2013) 201–214. [11] Y. Chisti, Biodiesel from microalgae beats bioethanol, Trends Biotechnol. 26 (2008) 126–131. [12] G. Shelef, A. Sukenik, M. Green, Microalgae Harvesting and Processing: A Literature Review, U.S. Department of Energy, National Renewable Energy Laboratory Golden, Colorado, USA, 1984. [13] N. Uduman, Y. Qi, M.K. Danquah, G.M. Forde, A. Hoadley, Dewatering of microalgal cultures: a major bottleneck to algae-based fuels, J. Renew. Sustain. Energy 2 (2010) 1–15. [14] T.  Chatsungnoen, Y.  Chisti, Harvesting microalgae by flocculation–sedimentation, Algal Res. 13 (2016) 271–283. [15] J.  Bratby, Coagulation and Flocculation in Water and Wastewater Treatment, second ed., IWA Publishing, Seattle, 2006. [16] J. Milledge, S. Heaven, A review of the harvesting of micro-algae for biofuel production, Rev. Environ. Sci. Biotechnol. 12 (2013) 165–178. [17] S.  Pahl, A.  Lee, T.  Kalaitzidis, P.  Ashman, S.  Sathe, D.  Lewis, Harvesting, thickening and dewatering microalgae biomass, in: M.A. Borowitzka, N.R. Moheimani (Eds.), Algae for Biofuels and Energy, Vol. 5, Springer, Dordrecht, 2013, pp. 165–185.

281

282

CHAPTER 11  Flocculation and electroflocculation

[18] D. Vandamme, I. Foubert, K. Muylaert, Flocculation as a low-cost method for harvesting microalgae for bulk biomass production, Trends Biotechnol. 31 (2013) 233–239. [19] A.I. Barros, A.L. Gonçalves, M. Simões, J.C.M. Pires, Harvesting techniques applied to microalgae: a review, Renew. Sust. Energ. Rev. 41 (2015) 1489–1500. [20] T.  Chatsungnoen, Y.  Chisti, Continuous flocculation-sedimentation for harvesting Nannochloropsis salina biomass, J. Biotechnol. 222 (2016) 94–103. [21] L. Brennan, P. Owende, Biofuels from microalgae—a review of technologies for production, processing, and extractions of biofuels and co-products, Renew. Sust. Energ. Rev. 14 (2010) 557–577. [22] E.W.  Becker, Microalgae: Biotechnology and Microbiology, Cambridge University Press, Cambridge, 1994. [23] N.  Uduman, V.  Bourniquel, M.K.  Danquah, A.F.A.  Hoadley, A parametric study of electrocoagulation as a recovery process of marine microalgae for biodiesel production, Chem. Eng. J. 174 (2011) 249–257. [24] M.W.  Tenney, W.F.  Echelberger  Jr., R.G.  Schuessler, J.L.  Pavoni, Algal flocculation with synthetic organic polyelectrolytes, Appl. Environ. Microbiol. 18 (1969) 965–971. [25] J. Morales, J. de la Noüe, G. Picard, Harvesting marine microalgae species by chitosan flocculation, Aquac. Eng. 4 (1985) 257–270. [26] A. Sukenik, D. Bilanovic, G. Shelef, Flocculation of microalgae in brackish and sea waters, Biomass 15 (1988) 187–199. [27] D.  Vandamme, I.  Foubert, B.  Meesschaert, K.  Muylaert, Flocculation of microalgae using cationic starch, J. Appl. Phycol. 22 (2010) 525–530. [28] M.R.  Granados, F.G.  Acién, C.  Gómez, J.M.  Fernández-Sevilla, E.  Molina Grima, Evaluation of flocculants for the recovery of freshwater microalgae, Bioresour. Technol. 118 (2012) 102–110. [29] S. Şirin, R. Trobajo, C. Ibanez, J. Salvadó, Harvesting the microalgae Phaeodactylum tricornutum with polyaluminum chloride, aluminium sulphate, chitosan and alkalinityinduced flocculation, J. Appl. Phycol. 24 (2012) 1067–1080. [30] H. Zheng, Z. Gao, J. Yin, X. Tang, X. Ji, H. Huang, Harvesting of microalgae by flocculation with poly (γ-glutamic acid), Bioresour. Technol. 112 (2012) 212–220. [31] L. Chen, C. Wang, W. Wang, J. Wei, Optimal conditions of different flocculation methods for harvesting Scenedesmus sp. cultivated in an open-pond system, Bioresour. Technol. 133 (2013) 9–15. [32] M.S. Farid, A. Shariati, A. Badakhshan, B. Anvaripour, Using nano-chitosan for harvesting microalga Nannochloropsis sp, Bioresour. Technol. 131 (2013) 555–559. [33] N. Rashid, M.S.U. Rehman, J.-I. Han, Use of chitosan acid solutions to improve separation efficiency for harvesting of the microalga Chlorella vulgaris, Chem. Eng. J. 226 (2013) 238–242. [34] L.V. Haver, S. Nayar, Polyelectrolyte flocculants in harvesting microalgal biomass for food and feed applications, Algal Res. 24 (2017) 167–180. [35] E. Poelman, N. De Pauw, B. Jeurissen, Potential of electrolytic flocculation for recovery of micro-algae, Resour. Conserv. Recycl. 19 (1997) 1–10. [36] S. Gao, J. Yang, J. Tian, F. Ma, G. Tu, M. Du, Electro-coagulation–flotation process for algae removal, J. Hazard. Mater. 177 (2010) 336–343. [37] D.  Vandamme, S.C.V.  Pontes, K.  Goiris, I.  Foubert, L.J.J.  Pinoy, K.  Muylaert, Evaluation of electro-coagulation–flocculation for harvesting marine and freshwater microalgae, Biotechnol. Bioeng. 108 (2011) 2320–2329.

­References

[38] A.K.  Lee, D.M.  Lewis, P.J.  Ashman, Harvesting of marine microalgae by electroflocculation: the energetics, plant design, and economics, Appl. Energy 108 (2013) 45–53. [39] F.L. Souza, S. Cotillas, C. Saéz, P. Cañizares, M.R.V. Lanza, A. Seco, M.A. Rodrigo, Removal of algae from biological cultures: a challenge for electrocoagulation? J. Chem. Technol. Biotechnol. 91 (2014) 82–87. [40] F.  Bleeke, G.  Quante, D.  Winckelmann, G.  Klöck, Effect of voltage and electrode material on electroflocculation of Scenedesmus acuminatus, Bioresour. Bioprocess. 2 (2015) 36. [41] R. Misra, A. Guldhe, P. Singh, I. Rawat, T.A. Stenström, F. Bux, Evaluation of operating conditions for sustainable harvesting of microalgal biomass applying electrochemical method using non sacrificial electrodes, Bioresour. Technol. 176 (2015) 1–7. [42] L. Xu, F. Wang, H.-Z. Li, Z.-M. Hu, C. Guo, C.-Z. Liu, Development of an efficient electroflocculation technology integrated with dispersed-air flotation for harvesting microalgae, J. Chem. Technol. Biotechnol. 85 (2010) 1504–1507. [43] A.  Zenouzi, B.  Ghobadian, M.A.  Hejazi, P.  Rahnemoon, Harvesting of microalgae Dunaliella salina using electroflocculation, J. Agric. Sci. Technol. 15 (2013) 879–887. [44] A.K. Lee, D.M. Lewis, P.J. Ashman, Energy requirements and economic analysis of a full-scale microbial flocculation system for microalgal harvesting, Chem. Eng. Res. Des. 88 (2010) 988–996. [45] S.B. Ummalyma, E. Gnansounou, R.K. Sukumaran, R. Sindhu, A. Pandey, D. Sahoo, Bioflocculation: an alternative strategy for harvesting of microalgae—an overview, Bioresour. Technol. 242 (2017) 227–235. [46] H.-M. Oh, S. Lee, M.-H. Park, H.-S. Kim, H.-C. Kim, et al., Harvesting of Chlorella vulgaris using a bioflocculant from Paenibacillus sp. AM49, Biotechnol. Lett. 23 (2001) 1229–1234. [47] G. Prochazkova, P. Kastanek, T. Branyik, Harvesting freshwater Chlorella vulgaris with flocculant derived from spent brewer’s yeast, Bioresour. Technol. 177 (2015) 28–33. [48] Y. Shen, Z. Fan, C. Chen, X. Xu, An auto-flocculation strategy for Chlorella vulgaris, Biotechnol. Lett. 37 (2015) 75–80. [49] J.J. Hawkes, M.S. Limaye, W.T. Coakley, Filtration of bacteria and yeast by ultrasoundenhanced sedimentation, J. Appl. Microbiol. 82 (1997) 39–47. [50] R.  Bosma, W.  van Spronsen, J.  Tramper, R.  Wijffels, Ultrasound, a new separation technique to harvest microalgae, J. Appl. Phycol. 15 (2003) 143–153. [51] J.E.  Coons, D.M.  Kalb, T.  Dale, B.L.  Marrone, Getting to low-cost algal biofuels: a monograph on conventional and cutting-edge harvesting and extraction technologies, Algal Res. 6 (2014) 250–270. [52] T. Chatsungnoen, Y. Chisti, Optimization of oil extraction from Nannochloropsis salina biomass paste, Algal Res. 15 (2016) 100–109. [53] R.K. Henderson, S.A. Parsons, B. Jefferson, Successful removal of algae through the control of zeta potential, Sep. Sci. Technol. 43 (2008) 1653–1666. [54] R.K. Henderson, S.A. Parsons, B. Jefferson, The impact of algal properties and preoxidation on solid-liquid separation of algae, Water Res. 42 (2008) 1827–1845. [55] J. Gregory, Particles in Water: Properties and Processes, IWA Publishing, Seattle, 2006. [56] K.K.K. Kaisha, Kurita Handbook of Water Treatment, 2nd ed., Kurita Water Industries, Tokyo, Japan, 1999. [57] R.A. Speers, M.A. Tung, T.D. Durance, G.G. Stewart, Colloidal aspects of yeast flocculation: a review, J. Inst. Brew. 98 (6) (1992) 525–531.

283

284

CHAPTER 11  Flocculation and electroflocculation

[58] X.  Zhang, P.  Amendola, J.C.  Hewson, M.  Sommerfeld, Q.  Hu, Influence of growth phase on harvesting of Chlorella zofingiensis by dissolved air flotation, Bioresour. Technol. 116 (2012) 477–484. [59] S. Hadjoudja, V. Deluchat, M. Baudu, Cell surface characterisation of Microcystis aeruginosa and Chlorella vulgaris, J. Colloid Interface Sci. 342 (2010) 293–299. [60] J.  Gregory, Chapter  3 – Stability and flocculation of suspensions, in: P.A.  Shamlou (Ed.), Processing of Solid–Liquid Suspensions, Butterworth-Heinemann, Oxford, UK, 1993, pp. 59–92. [61] N. Tambo, Y. Watanabe, Physical aspect of flocculation process—I: fundamental treatise, Water Res. 13 (1979) 429–439. [62] N. Uduman, Y. Qi, M.K. Danquah, A.F.A. Hoadley, Marine microalgae flocculation and focused beam reflectance measurement, Chem. Eng. J. 162 (2010) 935–940. [63] A.  Schlesinger, D.  Eisenstadt, A.  Bar-Gil, H.  Carmely, S.  Einbinder, J.  Gressel, Inexpensive non-toxic flocculation of microalgae contradicts theories; overcoming a major hurdle to bulk algal production, Biotechnol. Adv. 30 (2012) 1023–1030. [64] S.  Sirisansaneeyakul, M.  Cao, N.  Kongklom, C.  Chuensangjun, Z.  Shi, Y.  Chisti, Microbial production of poly-γ-glutamic acid, World J. Microbiol. Biotechnol. 33 (2017). article 173. [65] D.  Bilanovic, G.  Shelef, A.  Sukenik, Flocculation of microalgae with cationic polymers—effects of medium salinity, Biomass 17 (1988) 65–76. [66] R.M. Knuckey, M.R. Brown, R. Robert, D.M.F. Frampton, Production of microalgal concentrates by flocculation and their assessment as aquaculture feeds, Aquac. Eng. 35 (2006) 300–313. [67] R.  Divakaran, V.N.  Sivasankara Pillai, Flocculation of algae using chitosan, J. Appl. Phycol. 14 (2002) 419–422. [68] R. Yang, H. Li, M. Huang, H. Yang, A. Li, A review on chitosan-based flocculants and their applications in water treatment, Water Res. 95 (2016) 59–89. [69] D.W. Hendricks, Water Treatment Unit Processes: Physical and Chemical, Taylor and Francis, Boca Raton, FL, USA, 2006. [70] J. Duan, J. Gregory, Coagulation by hydrolysing metal salts, Adv. Colloid Interf. Sci. 100–102 (2003) 475–502. [71] A.J. Garzon-Sanabria, R.T. Davis, Z.L. Nikolov, Harvesting Nannochloris oculata by inorganic electrolyte flocculation: effect of initial cell density, ionic strength, coagulant dosage, and media pH, Bioresour. Technol. 118 (2012) 418–424. [72] V.M.  Rwehumbiza, R.  Harrison, L.  Thomsen, Alum-induced flocculation of preconcentrated Nannochloropsis salina: residual aluminium in the biomass, FAMEs and its effects on microalgae growth upon media recycling, Chem. Eng. J. 200–202 (2012) 168–175. [73] A. Papazi, P. Makridis, P. Divanach, Harvesting Chlorella minutissima using cell coagulants, J. Appl. Phycol. 22 (2010) 349–355. [74] S.J. Lee, S.-B. Kim, J.-E. Kim, G.-S. Kwon, B.-D. Yoon, H.-M. Oh, Effects of harvesting method and growth stage on the flocculation of the green alga Botryococcus braunii, Lett. Appl. Microbiol. 27 (1998) 14–18. [75] M.K. Danquah, L. Ang, N. Uduman, N. Moheimani, G.M. Forde, Dewatering of microalgal culture for biodiesel production: exploring polymer flocculation and tangential flow filtration, J. Chem. Technol. Biotechnol. 84 (2009) 1078–1083. [76] I.  Udom, B.H.  Zaribaf, T.  Halfhide, B.  Gillie, O.  Dalrymple, Q.  Zhang, S.J.  Ergas, Harvesting microalgae grown on wastewater, Bioresour. Technol. 139 (2013) 101–106.

­References

[77] EPA, Wastewater Technology Fact Sheet: Chemical Precipitation (EPA 832-F-00-018), EPA, Washington, DC, USA, 2000. [78] R. Hogg, Flocculation and dewatering, Int. J. Miner. Process. 58 (2000) 223–236. [79] R. Hogg, C. Rattanakawin, The role of polymers in dispersion stability, in: D.R. Skuse (Ed.), Speciality Chemicals in Mineral Processing, The Royal Society of Chemistry, Cambridge, UK, 2002, pp. 3–20. [80] R. Riffat, Fundamentals of Wastewater Treatment and Engineering, CRC Press, Boca Raton, FL, USA, 2013. [81] R.  Hogg, Chapter  12: Flocculation and dewatering of fine-particle suspensions, in: D.  Bohuslav, S.  Hansjoachim (Eds.), Coagulation and Flocculation, second ed., Vol. 126, CRC Press, Boca Raton, FL, USA, 2005, pp. 805–850. [82] T. Chatsungnoen, An Assessment of Inexpensive Methods for Recovery of Microalgal Biomass and Oils (PhD thesis), Massey University, New Zealand, 2015. [83] J.K. Edzwald, Water Quality & Treatment: A Handbook on Drinking Water/American Water Works Association, sixth ed., McGraw-Hill, New York, 2011. [84] J.Y. Oldshue, Fluid Mixing Technology, McGraw-Hill, New York, USA, 1983. [85] E.L. Paul, Handbook of Industrial Mixing: Science and Practice, Wiley, Hoboken, NJ, USA, 2003. [86] F.R. Spellman, Handbook of Water and Wastewater Treatment Plant Operations, second ed., CRC Press, Boca Raton, FL, USA, 2009. [87] J.C. Crittenden, R.R. Trussell, D.W. Hand, K.J. Howe, G. Tchobanoglous, Coagulation and flocculation, in: J.C.  Crittenden, R.R.  Trussell, D.W.  Hand, K.J.  Howe, G. Tchobanoglous (Eds.), MWH’s Water Treatment: Principles and Design, third ed., Wiley, Hoboken, NJ, USA, 2012, pp. 541–639. [88] J.C. Crittenden, R.R. Trussell, D.W. Hand, K.J. Howe, G. Tchobanoglous, Gravity separation, in: J.C. Crittenden, R.R. Trussell, D.W. Hand, K.J. Howe, G. Tchobanoglous (Eds.), MWH’s Water Treatment: Principles and Design, third ed., Wiley, Hoboken, NJ, USA, 2012, pp. 641–725. [89] T.R. Camp, Sedimentation and the design of settling tanks, Trans. Am. Soc. Civ. Eng. 111 (1946) 895–936. [90] T.R. Camp, A.A. Estrada, Studies of sedimentation basin design, Sewage Ind. Wastes 25 (1) (1953) 1–14. [91] Y.  Chisti, Shear sensitivity, in: M.C.  Flickinger (Ed.), Encyclopedia of Industrial Biotechnology, Bioprocess, Bioseparation, and Cell Technology, Vol. 7, Wiley, New York, 2010, pp. 4360–4398. [92] J.A. Sánchez Pérez, E.M. Rodríguez Porcel, J.L. Casas López, J.M. Fernández Sevilla, Y. Chisti, Shear rate in stirred tank and bubble column bioreactors, Chem. Eng. J. 124 (2006) 1–5. [93] T. Serra, J. Colomer, B.E. Logan, Efficiency of different shear devices on flocculation, Water Res. 42 (2008) 1113–1121. [94] J.P.F. Koren, U. Syversen, State-of-the-art electroflocculation, Filtr. Separ. 32 (1995) 153–156. [95] I. Kabdaşlı, I. Arslan-Alaton, T. Ölmez-Hancı, O. Tünay, Electrocoagulation applications for industrial wastewaters: a critical review, Environ. Technol. Rev. 1 (2012) 2–45. [96] M.L. Gerardo, S. Van Den Hende, H. Vervaeren, T. Coward, S.C. Skill, Harvesting of microalgae within a biorefinery approach: a review of the developments and case studies from pilot-plants, Algal Res. 11 (2015) 248–262.

285

286

CHAPTER 11  Flocculation and electroflocculation

[97] T. Harif, M. Khai, A. Adin, Electrocoagulation versus chemical coagulation: coagulation/flocculation mechanisms and resulting floc characteristics, Water Res. 46 (2012) 3177–3188. [98] G. Chen, Electrochemical technologies in wastewater treatment, Sep. Purif. Technol. 38 (2004) 11–41. [98a] M.Y.A.  Mollah, R.  Schennach, J.R.  Parga, D.L.  Cocke, Electrocoagulation (EC)—­ science and applications, J. Hazard. Mater. 84 (2001) 29–41. [99] Y.M.  Vong, D.G.  Garey, Wastewater treatment by electrocoagulation, in: G.  Kreysa, K.-I.  Ota, R.F.  Savinell (Eds.), Encyclopedia of Applied Electrochemistry, Springer, New York, USA, 2014, pp. 2117–2122. [100] V.  Robinson, Wastewater treatment by electroflocculation, in: G.  Kreysa, K.-I.  Ota, R.F. Savinell (Eds.), Encyclopedia of Applied Electrochemistry, Springer, New York, USA, 2014, pp. 2122–2126. [101] J.R. Benemann, W.J. Oswald, Systems and Economic Analysis of Microalgae Ponds for Conversion of CO2 to Biomass, Final Report Submitted to Department of Energy, Pittsburgh Energy Technology Center, Pittsburgh, PA, USA, 1996, p. 201.