A comprehensive review on harvesting of microalgae for biodiesel – Key challenges and future directions

Renewable and Sustainable Energy Reviews 91 (2018) 1103–1120

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Renewable and Sustainable Energy Reviews journal homepage: www.elsevier.com/locate/rser

A comprehensive review on harvesting of microalgae for biodiesel – Key challenges and future directions Thangavel Mathimani, Nirupama Mallick

T

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Agricultural and Food Engineering Department, Indian Institute of Technology Kharagpur, Kharagpur 721302, West Bengal, India

A R T I C LE I N FO

A B S T R A C T

Keywords: Biodiesel Bioflocculation Flocculation Harvesting Magnetic separation Microalgae

Economically viable microalgal biodiesel production is unrealistic and unsustainable owing to expensive harvesting or dewatering techniques. Hence, immense and meticulous exploration of harvesting process is essential to identify knowledge leads by which suitable harvesting technique could be ascertained for lucrative biodiesel production. With this in view, this review aims to collate and highlight the spectrum of harvesting techniques applied to microalgae, i.e., conventional – modern, high cost- inexpensiveness, energy efficient- energy consuming process. At the outset, global energy outlook and demand had been critically addressed, and the scientific ways to tackle or satiate the fuel demand had also been highlighted in this reveiw. This review manuscript has thrown widespread light on the physical harvesting methods namely centrifugation, sedimentation, filtration, flotation and technical advantages thereof. Due to the energy-intensive and cost barrier of physical harvesting techniques, chemical methods entailing organic, inorganic, and electroflocculation have come to limelight and in this regard, microalgae used, floc recovery and the dose of flocculants have been compared and presented in detail. Further, state of the art harvesting techniques viz., bioflocculation by microalgae/bacteria, flocculation by pH adjustment, and magnetic nanocomposite based microalgal harvesting had been critically articulated. Besides discussing the several methods, this paper has summarized the key challenges in conventional and advanced harvesting techniques and also provided the scope thereof. Hence, the key suggestions and findings given in this manuscript would positively offer a well-defined roadmap in choosing foreseeable harvesting technology for cost-effective microalgal biofuel development.

1. Introduction

sources to encounter energy thirst. As a decisive consequence, biofuel (first generation) came to limelight along with certain shades of negative criticism of uprooting the food market by food vs fuel debate. In fast‐developing economies like India, China, Thailand and South Africa, increasing requirement for food and fuel has initiated an augmented race for already limited water resources [6]. Biofuel is a renewable and alternate fuel produced from the organic (biological) feedstocks which can readily be fueled to the existing transportation infrastructure without engine modification [7–10]. Many countries realized the importance of biofuel, and thus, they geared up for producing biofuel and blending it with current transportation fuel to reduce the dependency on fossil fuel. The order of top biofuel producing countries is the United States > Brazil > Germany > China > Argentina [11]. Indonesia set a probable target of replacing 15% gasoline by ethanol and 20% diesel by biodiesel in 2025, and in Thailand, twelve bioethanol plants are being constructed that will produce 2.6 million L ethanol per day [12]. Brazil set a 5% and 20% biodiesel blending target by 2013 and 2020

Globally, energy is the prime commodity for the development of any nation, and till date, the majority of energy necessities are satiated through fossil fuels namely petroleum, coal and natural gas [1,2]. Fossil fuel poses dual threats to the society, i.e., fuel demand and climatic change. Due to the depletion of fossil fuels, energy consumption is predicted to rise from 550 EJ (Exajoule) to 865 EJ by 2040 [3]. In this regard, India featured to 7th place in energy production, accounting 2.49% of the world's annual energy production and in parallel, ranked as world's 5th energy consumer by utilizing 4% world's total energy consumption [4]. Further, India has imported 163 million metric tonnes of crude oil during 2010–2011 by spending $100 billion due to prompt urbanization and oil demand [4]. On the other hand, the current CO2 level is 394.5 ppmv (parts per million volume), and it is projected to reach 500 ppmv in 2050 if emissions remain unrestricted [5]. In a view to culminate the hazardous diesel usage, in-depth R & D initiatives have been unleashed in wide dimension for exploring the renewable fuel

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Corresponding author. E-mail address: [email protected] (N. Mallick).

https://doi.org/10.1016/j.rser.2018.04.083 Received 3 August 2017; Received in revised form 8 April 2018; Accepted 14 April 2018 1364-0321/ © 2018 Elsevier Ltd. All rights reserved.

Renewable and Sustainable Energy Reviews 91 (2018) 1103–1120

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in biofuel include (i) imperiling food security due to tradeoff between food vs fuel through resource allocation, (ii) surplus land requirement and agricultural inputs, (iii) high capital cost and uncompetitive retail prices, (iv) low net energy returns, (v) higher claims over gaseous emission reductions (v) low productivity over seasons [10,24,25]. Among the biofuel feedstocks, algae (microalgae, cyanobacteria, diatoms) have secured rampant attention as a next-generation sustainable substitute to diesel fuel [26,27], in the burgeoning energy enterprises over the past decades as it holds dual potentials to abate climatic disaster, and to safeguard energy security. Algae are prokaryotic (cyanobacteria) or eukaryotic (green algae and diatoms) photoautotrophs, which may be either unicellular or multicellular or heterocystous or colonial in morphology [28], and algae are being explored as potential crops for biodiesel whose ancestral relationships is broader than terrestrial plants and rich in genetic diversity [26]. Utilizing algae for biodiesel application offer several pros as microalgae, do not compete with edible crops, do not affect food security, do not emit high gaseous pollutants, do not demand surplus fertile land and fertilizer supplements, do have high biomass density and sustainable lipid productivity over terrestrial crops [29–31], and can grow in various habitats like freshwater, seawater, wastewater, and brackish water [32]. Preferably, Microalga was able to produce 58,700 L oil hectare−1, which is two magnitudes higher compared to other bioenergy feedstock [33]. Although microalgae are considered as a viable alternate to offset fossil fuel, there are numerous obstacles need to be overcome for lucrative biofuel production. Using current production process, a barrel of algal biofuel cost is estimated to be US$300–2600, which is much higher than a barrel of petrol $40–80 [26]. To substitute petrol with microalgal biofuel cost-effectively, algal oil costing ~ US$ 1.619 L−1 is preferable [11]. US invested $800 million through American Recovery and Renewal Act for R & D on economic algal biofuel production [34]. Albeit many economical biodiesel production strategies are underway, it is yet in primitive stage demanding financial viability, and however, hitherto findings have portrayed microalgae as a positive candidate for biodiesel [28]. The bottlenecks or need of the hour in biodiesel production are resilient strain isolation, mass cultivation using low-cost nutrient inputs, ideal harvesting techniques, pertinent lipid extraction method, fuel production, coproduct development, residual biomass utilization [26]. In concern with strain selection, a microalgal monoculture that resistant to pathogens (algal pond sustainability) and capable accumulating high lipid content is a prerequisite for biodiesel since algal cells could be invaded by pests and pathogens [26,33]. Acclimatization of microalgae to an unconducive open environment, resistant to contamination and high biomass productivity in a low-cost medium is the favorable features in strain selection and outdoor cultivation. In low-cost medium formulation, inexpensive urea can be used for Chlorella sp., and Spirulina platensis culturing instead of expensive chemical nutrients to reduce the cost [35]. In this connection, marine or halophilic strain could be used as a potential feedstock since it requires only seawater with few nutrient supplements during large-scale cultivation. Another bottleneck in biodiesel production is lipid extraction, which is most commonly carried out by oil expeller, solvent extraction, and supercritical extraction [30]. These methods are expensive concerning energy consumption and device investment; however, are amenable to engineering improvements [26]. It is reported that drying of microalgal biomass and oil extraction occupies ~90% of overall biodiesel economy [36,37]. Among the challenges in microalgal biodiesel, the most pressing challenge lies in the harvesting [38], because harvesting costs individually occupy nearly 30% of the total capital investment for biodiesel [39,40]. Harvesting is an economical key for commercial biodiesel production and therefore, choosing a pertinent harvesting technique, which is able to dewater high voluminous culture medium inexpensively is essential to increase the scale of biomass yield and decrease the overall harvesting cost concurrently [41,42]. Considering the above issues, this review contextualizes various harvesting methods and their limitations and scope as listed below to

respectively and to achieve above goals, 2000 and 12,000 ML/year biodiesel is likely to be produced respectively [13]. With reference to biofuel policy of India, National Biodiesel Mission (NBM) was launched in 2003, and concurrently National policy on biofuels was enacted to regulate the biofuel usage and blending. The blending target set by India was initially 5% by 2012 and then, 10% by 2017 and 20% after 2017 [14]. In 2007, an amendment in blending mandate indicated 5% ethanol blending with petrol throughout India, except the North Eastern States, Jammu and Kashmir, and other Island territories [15]. Recent times, certain countries set 7% or less than 7% biofuel blending target. In Japan, bioethanol and biodiesel blending has been limited to 3% and 5% respectively to ensure safety and avert the degradation of engine components [16]. However, increasing the blending beyond 7% (B7) poses substantially two major concerns between light and heavyduty vehicles i.e., rapid lubricant deterioration during post injection and presence of undesired compounds or impurities in the blends and notably in heavy-duty trucks, 30% biodiesel-diesel blend can safely run 80% of heavy-duty trucks [17]. In a very recent policy draft, oil ministry of India has set a lower blending of 5% biodiesel in diesel by 2030 [18]. Besides the difficulty in speculating the rapid implementation of blending, National biofuel policy enacted by Malaysian government in 2006 stated 5% blending target nationwide and also it had considered the implementation of 7% biodiesel blend [19], and recently, Malaysian government decided to blend 7% B100 biodiesel in conventional diesel to ensure the availability of 7% biodiesel-diesel blend (B7) in 2018 [20]. Though biofuel blending target of 7% is set by various countries, certain countries like Brazil, European Union, and the United States of America set a target of 20%, 10%, and 25% biofuel blending respectively by 2020 [21]. In a line of above blending, 10% blend is considered to be technically feasible for spark-ignited gasoline engines [16]. As reported by Kampman [17], in European Union, all diesel locomotives are compatible with 7% biodiesel blend in diesel, and therefore, 12.8 Mtoe of fatty acid methyl ester (biodiesel) is predicted to be brought on the energy market for road transportation by 2020 considering 7% blend. Low biofuel blending with transportation fuel is quite desirable since the fueling of certain automotive engines by high biofuel blends demand technical modification [16]. In this regard, increasing the blending target from 7% to 10% or 15% might not be problematic considering the availability of suitable technologies [17]. It is interesting to note that, overproduction and oversupply of biodiesel were witnessed in two major biodiesel producing companies, which gives 8.5 million L day−1, and this caused issues with domestic producers since production competition was tightened up with more companies in terms of biodiesel production. Global Green Chemicals Plc and PTG Energy are producing B100 biodiesel and simultaneously using it for their own supply [20]. In fact, Thai Biodiesel Producers Association stated that 13 industries are currently producing 100% biodiesel (B100) at about 6.6–7 million L day−1, but present national demand is ~ 3.3–3.5 million L day−1 [20]. The present oversupply of biodiesel in Thailand affected the profit of numerous domestic biodiesel producing companies at < 50% of capacity and cutting the cost below 20 baht L−1. However, biodiesels stated above are produced from vegetable oil like corn, maize, palm or other sources, and microalgal biodiesel has not yet been commercialized. It is stated that, cultivation of corn, soybeans, and other edible crops for biofuel or bioethanol taking a substantial toll on the environment and also affecting the food market through elevated food prices [22]. In addition to energy and environmental benefits, biofuel industries play a key role in providing socioeconomic services to the rural peoples such as infrastructure development, poverty reduction by creating job opportunities, opening schools, hospitals especially in the countries Brazil, India, China. In Brazil, ethanol industry provides occupation to ~ 12% of the rural population either directly or indirectly and adding to that; sugarcane employs one million workers in Brazil at different levels [14]. Further, biodiesel can also be used a lubricant, which is 66% more competent than diesel [23]. However, the major limitations 1104

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T. Mathimani, N. Mallick

axis and lighter particles towards the axis [38]. Various types of centrifuges have been used at different efficiency for harvesting algal cells owing to their rapidity and reliability [38], either through a one-step process or two-step process involving preconcentration of biomass [51]. Tubular and multi-chamber centrifuges are used traditionally to harvest algal biomass. Though tubular centrifuge is most efficacious, it did not have provision for solid discharge, and thus it cannot be operated continuously (intermittent cleaning required). Hence, it is capable in bench scale harvesting and in predicting disc centrifuge performance [51]. In a case of multi-chamber centrifuge, it has closed bowl subdivided into concentric, vertical compartments. Biomass is fed through the zones of increasing acceleration, and cleaning needs to be done manually similar to a tubular centrifuge. Therefore, multi-chamber might not be considered as a proficient device for algae recovery due to its time-consuming cleaning process. The major issue over the cleaning process of the multi-chamber centrifuge can be averted by nozzle type disc centrifuges, which can be effortlessly cleaned and sterilized [52]. Based on algae removal efficacy, harvested biomass quantity, nozzle type disc centrifuge is appeared to be an ideal instrument compared to others although it is economically incompatible due to high capital investment and increased power consumption [52]. In concern with disc stack centrifuges, it is widely used for commercial applications like algal biofuel and products [41] in which centrifugal force applied to the feed is 4,000 to 14000 times higher than gravitational force and therefore, it reduces separation time [39]. As described by Milledge & Heaven [39] and Uduman et al. [42], disc stack centrifuges are suitable for separating 3–30 µm cells and 0.02–0.05% of cell density in a culture medium. It is stated that disc centrifuge has high energy consumption, for example, 1 kWh m−3 is required for the harvesting Scenedesmus sp. [41] while 1.4 kWh m−3 energy imposed for harvesting pig waste grown algae. In general, centrifugation requires ~ 1 MJ kg−1 energy for algal biomass, which is higher than the harvesting of wood ~ 0.7–0.9 MJ kg−1 [53]. The very recent work undertaken by Wang and Dandy [54] suggested a passive microfluidic approach for harvesting Synechocystis sp. PCC 6803, which does not requires extra reagents or external fields. This approach separates the cyanobacterial cells from suspension by introducing culture into a microchannel with set flow rate, and moving the cells laterally to the channel cross-section using the geometry of microfluidic network, and using this single microchannel device, 98.4% recovery was observed. Lipid yield and fatty acid profile of the Nannochloropsis oculata (N. oculata) was appraised with respect to centrifugation and pH-induced flocculation techniques. The pH increase by NaOH recovered > 90% cells with the low lipid content of 4.40% while centrifuged N. oculata biomass showed 45.4% lipid. Further, centrifuging the biomass washed with ammonium formate was identified to be the better method regarding high lipid yield and high eicosapentaenoic acid, eicosatetraenoic acid levels besides its cost [55]. As per the U.S. Department of Energy report, for large-scale application, the present stance of centrifugation process is “cost-prohibitive” (U.S. DOE, 2010) [38]. Economically successful harvesting of microalgae through centrifugation is hindered by certain limitations as detailed in Table 1.

grab economic discrepancy of burgeoning microalgal biodiesel production i. Forecasting of the global energy demand and exploring microalga as a paradigm towards next generation feedstock to satisfy the energy demand and identifying technicity hurdles and portraying microalgal energy sector in socioeconomic development. ii. Discussing the mechanism and comparing, evaluating the challenges, merits, and demerits of physical, chemical (organic, inorganic) and electroflocculation methods concerning flocculation efficiency and flocculant dose using plethora literature. iii. Presenting the mechanism and research work so far undertaken on advanced harvesting techniques viz., bioflocculation, autoflocculation, magnetic separation and advantages, disadvantages, future perspectives thereof. iv. Highlighting knowledge gaps in each harvesting process with an emphasis on lucrative biodiesel production and addressing the research needs to seal the gaps. 2. Harvesting Harvesting is a process by which algal biomass is separated from the medium using various techniques. Separation of cells from the dilute microalgal suspension makes harvesting more expensive than harvesting the terrestrial counterparts [43]. High harvesting costs are owing to the tiny veritable cell size (3–30 µm) in much-diluted cultures (inferior biomass concentration < 0.6 g L−1) which is near to water density [38,41]. In microalgal suspension, negatively charged cell surface (−7.5 to −40 mV) with algogenic organic material aids in the stability of dispersed cell [44,45]. It is assessed that 90% of instrument cost for outdoor microalgal biomass production originated from harvesting [46]. In spite of many harvesting methods are being implemented by the researchers worldwide, type, value, and properties of a product, sedimentation rate, cell viability, recycling of culture medium, cell density and size decide suitable harvesting method to be applied at large scale [47]. In this perspective, based on the hitherto reports none of the commonly used harvesting technologies are proven to be economical and best at scale till date [42,48]. Hence, reducing the harvesting costs is contemplated as a crucial factor for sustainable and inexpensive production of biofuel [38]. Till date, copious research efforts have been undertaken on improving lipid yield and composition of microalgae rather than on biomass dewatering or harvesting [49]. An ideal harvesting method must be suitable for most of the microalgal types and must achieve high biomass recovery, and must use minimal energy with nominal operative cost [44,50]. Various methods used for harvesting of microalgae are illustrated in Fig. 1. In general, physical, chemical, biological and, electrical based harvesting methods are used, and in some cases, two or more of the above methods were combined to obtain maximum biomass slurry. Presently, magnetic nanocomposite based separation is being practiced for efficient harvesting of microalgae. In the view of harvesting, there were numerous research works have been undertaken and published recently. Nonetheless, basic understanding and knowledge on harvesting generated by the publications are inadequate and fragmented to some degree. Hence, this review is attempted to pool and analyze all the available reports to provide overall forecasts of harvesting technique in microalgal biofuel enterprise.

2.1.2. Gravity sedimentation Gravity sedimentation is a solid-liquid separation method by which concentrated slurry is settled out under gravitation force and leaving above noticeably clear liquid supernatant; further settling characteristics or slurry formation is governed by sedimentation velocity [39,49,56]. Gravity sedimentation can be designated by Stokes’ law, which states that sedimentation velocity is linearly correlated to strain characteristics, i.e., radius and density of cells as given below [39].

2.1. Physical methods 2.1.1. Centrifugation Centrifugation is the process where gravitational acceleration is replaced by centrifugal force, which is employed to separate two miscible particles. Centrifugation is a type of conventional mechanical harvesting method in which substances are separated based on density variation as centrifugal force drives high dense particle away from the

Setting velocity = 1105

2 r2 g (ρs−ρl) 9 η

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Fig. 1. Schematic illustration of various methods used for harvesting of microalgae.

suspension is run through a filter or a porous membrane, which retains filtrate (algae slurry) and concentrate (remaining water) is passed through a filter by driving force [56]. Eventually, in most of the filtration process, the membrane is a “selective porous barrier” between retentate and permeate phases, when the suspension is fed, the membrane separates it into retentate (deposited on the face of a membrane) and permeate (which passes through the membrane) [66]. Filtration requires force drop across the system to accelerate the suspension through a membrane, and it can be pressure, temperature, concentration, etc. The movement of solute or solid through predefined porous filter relies on its size, shape, charge, and it is affected by viscosity and mixing rate of a suspension [67]. Membrane filtration is being used in various applications due to its advantages such as (i) devoid of chemical additives, (ii) low energy and cost (iii) low impact on slurry or feed quality (iv) feasibility to make hybrid harvesting process, (v) easy scale up as membranes can be assembled into modules [39,66–68]. Pressure drop forced membrane filtration modules typically has plate-and-frame, spiral wound, and tubular filtration configuration [67]. Considering its potential advantages, numerous membrane filtration techniques have been implemented with varying degree of success namely, microfiltration (0.1–10 µm), macrofiltration (10 µm), ultra-filtration (0.02–0.2 µm), reverse osmosis (0.001 µm), tangential flow filtration, dead-end filtration, vacuum filtration and pressure filtration [39,69]. Microfiltration is generally used for harvesting fragile smaller algal cells as it has the applicable pore size (0.1–10 mm) for many microalgal strains like Chlorella and Cyclotella, and microalgal cell size is typically > 30 µm [41,49,58]. Micro filters permit higher initial fluxes, but clogging or fouling occurs promptly and thus, requires repeated expensive filter replacement [70]. However, macro filtration is best suited for larger cell size and mostly for filtering flocs (biomass obtained through flocculation) [39], and recent past, extensive range of macrofiltration units were used for downstream processing of algae like vibrating macro-filters. Though vibrating screens can separate Coelastrum and Spirulina, these vibrating screens are not considered to be an effective process [71] due to energy intensiveness. For example, estimated energy to produce 6% dry microalgal slurry is 0.4 kWh m−3 [72]. Ultrafiltration can be an alternative to macrofiltration for recovery of very fragile cells, but it has not been extensively practiced for dewatering microalgae [28,41]. Further, ultrafiltration (pore size up to

where, r – radius, g – viscosity, ρs and ρl are densities of solid and liquid, respectively. According to Stokes equation, settling velocity of circular shaped (5 µm) microalga Chlorella (density of 1070 kg m−3) was 0.1 m day−1 whereas for Cyclotella a microalgae with a size similar to Chlorella, calculated sedimentation velocity was (0.04 m day−1) lower than the experimental settling velocity (0.16 m day−1) and thus, it is inferred that algal sedimentation could be driven by the slight change in cell density [57,58]. Low dense algal species are not successful in sinking rate while high-density cells settle out rapidly [50]. The cytoplasmic density of microalgae and cyanobacteria and marine diatom was commonly in the range between 1030 and 1100 kg m−3 [57], 1082 and 1104 kg m−3 [59], 1030 and 1230 kg m−3, respectively. Further, average sedimentation velocity of green algae, diatoms, and cyanobacteria was 0.1 m day−1, 0.2 m day−1 and 0.0–0.05 m day−1 respectively [60]. Therefore, Stokes’ law is mostly applicable to spherical shape algae, but all algal strains cannot be mostly spherical [61]. According to the study undertaken by Choi et al. [62] and Nurdogan and Oswald [63], of the 30 test microalgal strains, needle, long cylindrical strains and motile algal species like Euglena, Chlorogonium were found reluctant to settle out, while spiral shaped cyanobacteria Spirulina by virtue of large size, density and colonial forms Micractinim, Scenesdesmus were successfully gravity sedimented. As detailed in Table 1, gravity sedimentation is not a suitable technique for small size microalgal cells. In addition to cell density and size, gravity sedimentation process is also hampered by cell viability whereby sinking rate was maximum in senescent cells and spore-producing cells [39]. Despite the rudimental of this process, an addition of coagulant/flocculant to the suspension prior to gravity sedimentation fasten and improve the settling the tiny algal cells [41,56]. Furthermore, lamella separators (1.6% total suspended solids retrieval) and sedimentation tanks (3% total suspended solids retrieval) are also used to improve the sinking rate, which is attributable to autoflocculation process [42,64]. Though lamella separators and sedimentation tank processes are costeffective, reliability is uncertain owing to low slurry output and requisite of thickening [47,65]. 2.1.3. Filtration Filtration is a separation process in which culture medium or algal 1106

1107

Flotation

Filtration

Gravity sedimentation

Centrifugation

Harvesting method

of chemical • Devoid supplements energy consumption • Low impact on slurry or • Low biomass quality harvesting process is • Hybrid feasible scale up • Easy biomass recovery • High to recover shear sensitive • Able cells area requirement • Less time is short • Operation • Flexibility • Industrial scalability possible

• Cost-effective • Simple and effortless operation

reliable • Rapid, independent • Strain • High recovery of solid content

Advantages

Energy-intensive High capital investment Operational cost high Not pertinent to low-value product like biodiesel Cell damage due to shear force Long operation time for high voluminous culture Time-consuming due to slow sedimentation Settling of small size cells is probable. Not reliable and effective process Low slurry recovery Biomass quality is questionable High operational cost membrane replacement and pumping membrane life-span is short selectivity or permeance of membrane is low

➢ Investment high and energy intensive (except micro bubble flotation) ➢ Recovery can be hampered due to large bubbles ➢ Current consumption high ➢ Unsuitable for marine strains

➢ ➢ ➢ ➢ ➢ ➢ ➢ ➢ ➢

➢ ➢

➢ ➢ ➢ ➢

Disadvantages

Table 1 Comparison of various physical harvesting methods of microalgae [38,39,41,56].

<7

5–27

0.5–3

10–22

Solid Conc. (%)

50–90

70–90

10–90

> 90

Recovery (%)

Chlorella vulgaris (C. vulgaris) - ozone dispersed flotation, Dunaliella salina (D. salina)microflotation, Chlorella sp. (flotation) Scenedesmus quadricauda (S. quadricauda)-dispersed air flotation with Cetrimonium bromide), Microcystis aeruginosa (Electro-coagulation flotation)

Coelastrum proboscideum (Vacuum filtration), Phaeodactylum tricornutum (P. tricornutum)-Ultrafiltration, Spirulina (simple filtration)

Spirulina, Micractinim, Scenesdesmus sp. Cyclotella sp.

Nannochloropsis sp. (spiral plate centrifuge, continuous flow centrifuge), Scenedesmus sp. Coelastrum proboscideum (self-cleaning disc stack centrifuge, Nozzle discharge centrifuge, Decanter bowl centrifuge)

Strains used

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inappropriate method for filtering microalgae [56].

100 nm) is not seemingly relevant filtration method for recovering of 1–500 kDa molecular weight feed. The performance of ultrafiltration depends on hydrodynamics, concentration and culture characteristics [47], and its operating and maintenance cost is high, power consumption is intensive and frequent membrane replacement is required [28,70,73]. Vacuum filtration is also used for effective recovery of large size algae and particularly rotary vacuum filters a common vacuum design used for dewatering [74]. As reported by Goh [75], rotary vacuum filter with 12 µm pore size is able to filter filamentous cyanobacterium Spirulina and colonial Micractinium at the output slurry concentration of 1–3% dry weight−1, while unicellular, chlorophycean microalga Chlorella had not been recovered efficiently even with 5 µm vacuum filter. Mohn [71] recommended a suction filter, and belt filter for harvesting Coelastrum considering low power consumption, consistency, and biomass concentrating ability whereas precoated vacuum drum filter is suggested for small cell sized microalga like Scenedesmus. Vacuum filters need to be cleaned and replaced frequently owing to fouling and further, feed pumping makes it an energy-intensive process [70]. Belt type filtration design has been suggested for water treatment and separation of cyanobacteria like Spirulina [71]. Large size algae can be readily recovered at 18% slurry concentration if belt filter is run with 4% pre-concentrated algae with a power consumption of 0.5 kWh m−3 [41]. In concern with cost, recovering 80 m3 h−1 feed through three ‘‘Klampress’’ belt filter costs ~ £ 360,000 with estimated power utilization of 17–21 kW, which corresponds to 0.25 kWh m−3 energy input [39]. To reduce operational energy cost, filter aids and flocculants may be used, but accessory entities may escalate costs, and need to be cleaned from algal slurry and filtrate [39]. Other types of filtration proposed for harvesting algal cells are tangential flow filtration and dead-end filtration. Tangential filtration is known to be feasible high rate method than dead-end filtration for dewatering suspended smaller algal cells due to negligible fouling [38,44,76], whereas dead-end filtration is preferred for large microalgal cells (> 70 mm cell diameter) [41,70]. Tangential flow filtration (TFF) is capable of concentrating 70–89% freshwater algal cells, and notably, TFF retains intact structure and properties of cells in the harvested slurry [50,56]. TFF has anti-fouling property and is applicable for shear-sensitive algae [42] as the suspension is allowed to run tangentially at high velocity across a membrane and retentate can be recycled [77]. Both tangential and dead-end filtration techniques were found to unveil limited success in largescale application due to increased power requirements, often membrane substitute replacement and fluid pumping [44,56]. The performance of membrane is evaluated by two factors such as permeance and rejection. A typical membrane must uphold its permeance over prolonged operation without a considerable loss while sustaining an adequate rejection [67]. A research work compared hollow fiber filtration (tangential flow), indirect electrocoagulation and pH-induced flocculation of D. viridis cells at two phases i.e., 3–10 L and 30–150 L culture volume. Among the methods, hollow fiber filtration was identified to be best with 99% recovery compared to other harvesting techniques [78]. Besides their advantages, each filtration technique exhibits certain limitations in microalgal harvesting. The key shortcoming in filtration is fouling/clogging phenomenon, which boosts up operational cost [38]. Clogging of algal slurry on filter grow thicker with the progression of filtration, and thus, thickened slurry increases resistance and decreases filtration flux over persistent pressure drop [64]. In addition, exopolysaccharides (EPS) secreted by microalgae while exposed to a hostile environment is also affecting the filtration process. EPS forms a gel-like layer on the membrane and poses resistance to flow and hence, membranes should be cleaned to make sure sanitization and reusability [79]. Even though many types of filters were found to be effective in recovering relatively large, lengthy algal cells, and colonies [41], the performance of filter was impeded by minimum throughput and prompt clogging [71]. Operational hurdles like back mixing in dead-end filtration make it an

2.1.4. Flotation Flotation is an “inverted sedimentation process” (algal particles move upward in flotation whereas downwards in sedimentation) where air or gas bubbles are sparged to the suspension for attachment to solid or suspended particles, and then lifted to the liquid surface due to the low density for skimming off or separation [47,80,81]. Flotation has been recognized as a promising method for separation of freshwater microalga C. vulgaris [82], and requires low space, relatively short operation time and possesses better flexibility [80]. The success of flotation process is contingent on a phenomenon is known as bubbleparticle collision or adhesion, and further higher bubble - particles contact occur in the system with suspended particles of lower instability, and small particles are more likely to be risen up by bubbles [42,64]. Flotation is classified into various types according to the size of bubble (i) dissolved air flotation (ii) dispersed air flotation; (iii) electrolytic flotation; and (iv) ozonation-dispersed flotation (ODF) [50,56,64,70]. Dissolved air flotation is the flotation process where micro air bubbles in the range of 10–100 µm were generated by dissolving air at a very high pressure in water to levitate suspended solid to the surface for skimming [47,58]. It is efficient and preferred in conjunction with sedimentation or addition of chemical flocculants to the algal culture system, and however, flocculant/coagulant supplementation can be problematic in downstream processing [83,84]. In a recent work, Laamanen et al. [85] have used waste industrial heat aided flotation of Scenedesmus cells without addition of any chemicals. At 85 °C, maximal recovery and a concentration factor of 83% and 25.8 was observed respectively, which produces 2.78 g L−1 concentrated Scenedesmus sp. cells from 0.13 g L−1 initial cell density. Flotation technique is seemingly robust in the harvesting of microalgae cultivated in pig slurry, but a high dose of alum was required [75], and energy-intensive owing to high-pressure process [86]. In dispersed air flotation, bubbles (700–1500 µm diameter) were generated continuously through a porous material with a mechanical agitator. It is reported that microalga S. quadricauda was harvested competently with dispersed air flotation coupled with N-cetyl-N-N-N-trimethyl ammonium bromide or cetrimonium bromide [87]. Though this method is energy efficient, it entails costly instruments and high-pressure drop [47]. Electrolytic flotation works on the phenomenon known as electrolysis in which hydrogen bubbles (low solubility in water) generated at the cathode end by the electrolysis of water charges (adhesion) the algal floc and levitates them to the surface [64,88]. It is advocated to be an impending algal separation technique as it does not necessitate chemical flocculation and also holds several advantages such as ecocompatibility, versatility, selectivity [88]. However, electrolytic flotation is a high-energy intensive process than dissolved air flotation and is best suited for harvesting marine algal species rather than freshwater (due to potential conductivity variation) and suitable for bench scale experimentation than industrial scale [39]. In foam flotation process, the impact of bubble size and rise velocity on the harvesting of microalgae was also demonstrated using limewood sparger, and ceramic flat plate sparger with constant airflow and oscillating airflow. The biomass recovery and bubble rising velocity are high in ceramic sparger at both airflow compared to limewood spargers due to improved bubble-cells collision and attachment [89]. Efficacy of foam harvester (dispersed air flotation with foam fractionation) and its operative factors in microalgal biomass separation was also studied using polystyrene latex beads, and results showed that highest concentration factor was recorded at lower surfactant concentrations and high column heights by consuming 0.015 kWh/m3 energy [48]. In a biodiesel viewpoint, the influence of cetyl trimethylammonium bromide aided foam flotation on lipid recovery and fatty acid profile of microalgae was assessed. Lipid recovery from the cells harvested by cetyl trimethylammonium bromide-foam flotation was higher than centrifuged cells by assisting in situ cell lysis, and further, CTAB 1108

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efficient flocculant, Papazi et al. [95] have tested twelve inorganic salts in chlorophycean microalga C. minutissima, and data seemingly unveils Al2(SO4)3, AlCl3, Fe2(SO4)3, FeCl3, ZnSO4, ZnCl2 have flocculated C. minutissima cultures at the optimum concentration of 0.75 g L−1 (sulfates salts) and 0.5 g L−1 (chloride salts) within 24 h settling time, whereas other six salts CaSO4, CaCl2, MgSO4, MgCl2, (NH4)2SO4, and NH4Cl were not found to show a remarkable variation in flocculation efficiency relatively over control [95]. Aluminium salts were found to be the best flocculant for harvesting Chlorella and Scenedesmus cells [41,95]. In another study, it is stated that alum is proven to be an effective flocculant for a wide variety of microalgae viz., Scenedesmus and Chlorella [102]. Albeit aluminium salts ranked first in flocculation efficiency, it causes cell damage (cell viability in floc is low). Further, ferric and zinc salts hold second and third position respectively, however, change in color of cells were observed by using ferric salts at > 1 g L−1 dose, whereas zinc salts cause the adhering of floc to the wall of the container [95]. Among the salts, chlorides are highly efficacious than sulfate salts owing to the “solubility phenomenon.” Chloride has a high degree of solubility and wider concentration range, and thus enables chloride salts to achieve maximal flocculation efficiency than sulfate salts [95,103]. In addition to the solubility of chloride and sulfate, the flocculants CaSO4, CaCl2, MgSO4, MgCl2, (NH4)2SO4, and NH4Cl exhibited low efficiency than alum, iron and zinc salts even after 24 h due to its lower electronegativity. Higher the electronegativity, earlier the settling and greater the efficiency [95]. Of the aluminium, iron, zinc flocculants, aluminium was exceptionally competent flocculant in terms of settling time and floc recovery, which can be described by two characteristics, i.e., molecular weight and charge density. Aluminium cations have high charge density (+ 3), and low molecular weight and therefore high cationic charge (high electronegativity) leads to wider molecular conformation and bridging of cells and thus, results in rapid charge neutralization of culture suspension. In addition, the low molecular weight of aluminium relatively to ferric and zinc enables increased solubility in culture suspension and hence, better floc recovery [95]. Correspondingly, albeit ferric possess independent charge density equivalent aluminium; it has higher molecular weight, which leads to limited solubility and reduction in floc efficacy than aluminium flocculant. However, ferric cations are highly effective than zinc as the later one possess low charge (+ 2) and high molecular weight [95]. On the other hand, the ionic strength of the culture medium (either freshwater or marine) is also determining the flocculant concentration. Due to the conductivity difference between freshwater and seawater, the flocculation of marine/brackish microalgae necessitates 5–10 times additional flocculant concentration (seawater salts hamper flocculant activity) than freshwater microalgae [42]. As reported recently by Chatsung et al. [104], aluminium sulfate is better over ferric chloride in flocculating both freshwater and marine microalgae such as C. vulgaris, Choricystis minor, Cylindrotheca fusiformis, Neochloris sp., N. salina with 95% recovery at 62 min settling time. The effect of Al2(SO4)3, AlCl3, and FeCl3·6H2O on the flocculation of S. rubescens and marine D. tertiolecta was tested. Of the various chemical salts dosed, aluminium sulfate showed 99% turbidity removal in both the cultures whereas 20% and 93% was observed for S. rubescens and D. tertiolecta, respectively using FeCl3·6H2O [105]. In a study carried out by Şirin et al. [98], marine diatom P. tricornutum grown in a F/2 medium was flocculated with two inorganic flocculants, aluminium sulfate and polyaluminium chloride (a polymeric form of aluminium sulfate). Results revealed that aluminium sulfate has a high degree of flocculation efficiency at 82.6% with higher concentration factor whereas, 66.6% was observed with polyaluminium chloride at the dosage of 30 ppm. In concern with ferric salts mediated flocculation, 90% recovery of C. stigmatophora biomass was observed using 25 mg L−1 FeCl3 [98,106], whereas 58 mg L−1 FeCl3 was required to attain 63–74% recovery of Anabaena flosaquae and Asterionella formosa cells [107]. In addition to metal flocculants, the polymeric metal salt is

harvested cells possess high saturated and monounsaturated fatty acids suitable for biodiesel [90]. Ozonation dispersed flotation or Ozone flotation is also practiced by passing ozone gas to the suspension to produce charged bubbles, which further promote flotation of cells on the surface [47]. During this process, air bubbles were found to rupture the cells and releases biopolymers, which in turn act as a flocculant, and hence results in improved slurry separation and unlaborious lipid extraction [56]. Limited literature are available on the use of ozonation dispersed flotation for microalgal harvesting. According to Pragya et al. [56], chlorophycean C. vulgaris cells harvested by ozonation dispersed flotation unveiled high lipid content over control (increased from 31% to 55% dry weight). However, it is also expensive process and not appropriate for large-scale separation due to contamination issues [70]. Compared to other flotation methods, micro-bubble flotation is considered to be cost-effective method since small (micro) bubbles were generated with low power consumption. Despite its enhanced efficacy over recovery of algal slurry [86], large-scale energy efficient demonstration needs to be carried out for commercialization [39]. The merits and demerits of flotation method are broadly presented in Table 1. 2.2. Chemical methods 2.2.1. Inorganic flocculants Flocculation is the prominent algal harvesting process in which solute particles in the culture suspension are aggregate together and settled as floc or slurry or aggregate [42] by different types of flocculants or coagulants [56]. In general, microalgal cells possess negative charge owing to the ionization of functional groups on surface and adsorption of ions originated from organic matter [50,69], and therefore auto aggregation is averted due to the repulsion of negative charges of difference algal cells [91]. To enable microalgal aggregate formation, stable system of algae must be disrupted by neutralizing the charge on the algal surface through flocculant [50,91]. Flocculation method follows charge dispersion phenomenon [70] and particularly entails three-corner ionic interaction between particles and flocculants and salts present in the medium [56]. Flocculation occurs by any one of the following three modes, i.e., patching or bridging or sweeping. In patching, charged particle patched or adsorbed to the surface of algal cells and reversing the charge (negative) locally, which then attract opposite charged patches and results in settling of cells [91]. In bridging, flocculant bind to the surface of two or more algal cells and form a bridge between cells, and in the case of sweep flocculation, microalgal cells are entrapped in mineral precipitation [92]. In all the modes of flocculation, coalescence of small solutes into aggregates and agglomeration of aggregates into large flocs was occurred [47]. A wide range of approaches undertaken for flocculating microalgae entails conventional or traditional methods (by chemical flocculants) or ecofriendly methods using bioflocculants or natural flocculants or advanced methods using magnetic nanocomposites [92]. Chemicals flocculants are grouped into inorganic and organic flocculants depend on their nature (carbon chemistry) [93]. Multivalent or polyvalent metal salts, such as Al2(SO4)3, Fe2(SO4)3 and FeCl3, were used extensively as inorganic flocculants to separate microalgae from suspension [39,47,94]. By and large, the negative surface charge conferred by COOH terminal on the surface of the microalgal cells would be reduced or neutralized by adding aluminium or iron-based chemical flocculants [50]. More explicitly, dissolution of salts in the medium liberate its respective cations (Al3+, Fe3+ or Zn2+), which reduces electrostatic repulsive force between the cells, and enables large floc formation [70,95–97]. Efficiency of flocculants depends on their ionic strength, i.e., high ionic strength (high electronegativity), high flocculation efficiency and vice -versa [47]. Amongst flocculants, + 3 ionic charge of aluminium sulfate and ferric chloride make them widely used and best flocculants over other divalent cationic salts [38]. Efficacy of inorganic flocculants on microalgal harvesting studied by various researchers was tabulated in Table 2. With an aim to select 1109

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Table 2 Comparison of flocculation efficiency of different chemical flocculants in various microalgal strains [41,56,98–101]. Sl. No

Strain

Flocculant

Dose (mg L−1)

Recovery efficiency (%)

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23.

Chlorella consortium Chlorella sorokiniana Chlorella minutissima Scenedesmus obliquus Anabaena sp. Asterionella sp. Chlorococcum sp. Anabaena sp. Asterionella sp. Chlorococcum sp. Chlorella consortium Chlorella sorokiniana Scenedesmus obliquus Dunaliella salina Chlorella stigmatophora Muriellopsis sp. Anabaena sp. Asterionella sp. Phaeodactylum tricornutum Phaeodactylum tricornutum Chlorella minutissima Microcystis aeruginosa Chlorella vulgaris UTEX-265

Fe2(SO4)3 Fe2(SO4)3 Fe2(SO4)3 Fe2(SO4)3 Fe2(SO4)3 Fe2(SO4)3 Fe2(SO4)3 Polyferric sulfate Polyferric sulfate FeCl3 FeCl3 FeCl3 FeCl3 FeCl3 FeCl3 Al2(SO4)3 Aluminium sulfate Aluminium sulfate Al2(SO4)3 Polyaluminium Chloride AlCl3 AlCl3 + Chitosan Organoclays or nanoclay (Mg2+ or Fe3+ with 3- aminopropyl triethoxysilane (APTES)

250 250 750 100 0.25 mM L−1 × 100 0.25 mM L−1 × 100 150 0.25 mM L−1 × 100 0.25 mM L−1 × 100 150 250 250 100 8.0 × 10 −4 mol L−1 25 1.42 × 10 −4 mol L−1 0.25 mM L−1 × 100 0.25 mM L−1 × 100 0.27 kg/dcw algae 0.27 kg/dcw algae 500 15 + 7 1 gL−1

90 98 80 96 ≤ 78 ≤ 70 87 ≤ 95 ≤ 93 90 98 66 95 85 90 10 ≤ 95 ≤ 95 82.6 66.6 80 71.55 98

also used as a flocculant since it works in wide pH range than nonpolymerized metal salts. Polyferric sulfate is a type of prepolymerized salt that is identified to recover 95% of Anabaena and Asterionella cells than other sulfate salts of aluminium and iron, and further floc settled by polymeric metal salts can effortlessly be dewatered [41,107]. In order to effectively harvest D. salina cells, various inorganic salts like Al2(SO4)3, AlCl3, FeCl3, FeSO4, Fe2(SO4)3, and pH-induced flocculation by NaOH, and Ca(OH)2 was carried out. Among the flocculants studied, FeCl3 was identified as the potential flocculant for D. salina based on its flocculation efficiency, concentration factor, medium reuse, and downstream integration [108]. The diameter of the floc generated by aluminium salts is in the range between 30 and 400 µm [109], and however, removal of lowdensity flocs could be difficult by sedimentation [58]. Typical chemical or inorganic flocculant should: (i) not contaminate biomass; (ii) possesses maximal flocculation efficiency; (iii) aggregate the biomass in less settling time (iv) allow the reuse of spent medium; and (v) be costeffective and eco-friendly [41]. Despite the relatively improved efficiency of inorganic flocculants over organic counterpart, high dosage makes them expensive per unit of microalgae [71], and further, inorganic flocculants might obstruct the extraction of lipid from certain algae [70]. In addition, chemical flocculation might not be beneficial for sustainable harvesting of microalgae at large-scale since metal flocculant associated with floc need to be discarded (operational costs may increase) for further processing [110], and chemical flocculated biomass may be undesirable for aqua feed and other food applications [41].

minimizes the electronegativity of algae (freshwater) and act as a bridge between the cells [41,64]. However, polymeric flocculants were found to be incompetent for flocculating seawater/brackish watergrown algae since it does not work in wide dimension and fails to bridge algal cells if ionic strength of culture suspension is high [41,42,112]. Generally, reducing the salinity increases the efficiency of cationic polymers, and it is effective when the salinity value is beyond 5 g L−1 in culture suspension [42,47]. Chitosan – a polymer of enhance cationicity derived from crustacean shells (chitin – from shellfish production) is an effective organic flocculant at low pH for harvesting freshwater microalgal cultures and also used in wastewater treatment in food industry. Biodegradability is the major positive feature of chitosan, and thus, it does not contaminate the floc [49,92,112,113]. On the other hand, the vital demerit of chitosan is the requirement of high dosage (20–150 mg L−1) than other synthetic organic flocculants [41,113]. It is demonstrated that chitosan divulges low biomass recovery in green unicellular microalgae Muriellopsis and necessitates higher dosage than synthetic polymer [114]. In addition to the high dose, expensiveness of chitosan makes it inappropriate for large-scale processing [50]. Cationic starch is an alternative to chitosan, which works even at low pH; however, it demands higher concentration [115]. Another organic flocculant polyacrylamide might not be of interest for algal harvesting owing to its toxicity [47]. As shown in Table 3, cationic polymers are employed as a promising flocculant for the microalgal harvesting by many researchers [42]. Granados et al. [114] reported that researchers at University of Almeria had observed high algal biomass recovery using 2–25 mg L−1 of cationic polymers than inorganic flocculants. In the track of usage, Praestol, a cationic polyacrylamide has been found to recover 70% of Teraselmis, and Spirulina biomass at a dose of 1 mg L−1 and notably, the supernatant of the flocculated suspension was reused for subsequent culture scale-up [117]. Cationic starch (starch with quaternary ammonium groups) is also being applied as a substitute for inorganic flocculants and was found effective to harvest Scenedesmus and Parachlorella, at a concentration higher than chitosan [71,92,115]. However, the charge of ammonium groups is pH independent and therefore works in a broad range, and further modification of starch to improve microalgal settling in turn increase cost excessively [71,92,115]. Nonionic and anionic polyacrylamides, anionic polystyrene and cationic polyethyleneimine at concentrations of 0.01–1000 mg L−1 were studied for the flocculation of C. ellipsoidia by Tilton et al. [118]. The biomass

2.2.2. Organic flocculants Organic flocculants are natural or synthetic, or polyelectrolyte flocculants of either anionic, cationic, or non-ionic [111], and however, anionic or non-ionic flocculants did not hold much influence on flocculation as microalgal surface carries negative charge whereas cationic polyelectrolytes physically aggregate algal cells with greater efficacy [42]. Flocculation strength of organic flocculants depends on the algal surface charge, pH and cell density in suspension [50,69]. High cell density and slow mixing of suspension aid closer and frequent cell–cell contact and therefore increases flocculation efficiency. High-speed mixing, i.e., elevated shear forces might destabilize the floc [56]. Chitosan, polyacrylamides, cellulose, surfactants and certain synthetic fibers are the commonly used cationic organic polymers, which 1110

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Table 3 Utilization of different polymer flocculants for harvesting of microalgae [41,56,99–101,116]. Sl. No

Strain

Flocculant name

Flocculant type (technical specification)

Ionic charge of flocculant

Dose (mg L−1)

Floc recovery or removal efficiency (%)

1. 2. 3. 4. 5. 6.

Chlorella consortium Scenedesmus obliquus Chlorococcum sp. Chlorella sorokiniana Chlorella sorokiniana Phaeodactylum tricornutum

Chitosan Chitosan Chitosan Chitosan Chitosan Chitosan

– – – – – –

Cationic Cationic Cationic Cationic Cationic Cationic

58 20 38 30 99 91.80

7. 8. 9.

Phaeodactylum tricornutum Chlorella vulgaris Nannochloropsis sp.

Chitosan Chitosan Nanochitosan

Cationic Cationic Cationic

10. 11. 12. 13. 14. 15. 16. 17. 18.

Nannochloropsis sp. Phaeodactylum tricornutum Chlorella sp. Chlorella vulgaris Nannochloropsis oculata Phaeodactylum tricornutum Neochloris oleoabundans Chlorella sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Muriellopsis sp. Scenedesmus sp. Chlamydomonas reinhardtii Scenedesmus acuminatus Chlorella sp. Chlamydomonas reinhardtii Neochloris Oleoabundans Chlorophyta Muriellopsis sp. Chlorella vulgaris

Chitosan VAMfloc 704 Zetag 8819 Zetag 8185 Zetag 8185 Zetag 7557 Zetag 7557 Magnafloc LT225 Magnafloc LT225

– – (Chitosan -crosslinking with sodium tripolyphosphate) – – Polyacrylamide Polyacrylamide Polyacrylamide Polyacrylamide Polyacrylamide Polyacrylamide Polyacrylamide

25 25 25 25 10 mg g−1 0.18 kg/dcw algae < 40 ppm 30 60

Cationic – Cationic Cationic Cationic Cationic Cationic Cationic Cationic

100 mg 10 mg L−1 34 5 0.55 0.01 0.01 35 35

< 95 > 80 98 100 75 98 52 72 95

Magnafloc LT225 EM22 EM22 Emfloc KC750

Polyacrylamide Polyelectrolyte Polyelectrolyte Potato Starch

Cationic Cationic Cationic Cationic

30 16 25 70

95 95 95 90

Emfloc KC750 POLY SEPAR® CFL25 POLY SEPAR® CFL25

Potato Starch Tannin, quaternary ammonia salt Tannin, quaternary ammonia salt

Cationic Cationic Cationic

40 30 20

95 95 20

Chitosan A-21 EM5 Moringa oleifera seed flour

Linked D-glucosamine Polystyrene Polyelectrolyte Natural polymer

Cationic Anionic Non-ionic Non-ionic

90 20 10 1000

66 3 0 88

19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29.

concentration used was in the range 50–3000 mg L−1. Of the flocculants tested, cationic polyethyleneimine was found to be successful, and non-ionic, anionic polyacrylamides, anionic polystyrene did not show prominent floc even at 200 mg L−1 concentration, and noticeably, increasing molecular weight of cationic polyethyleneimine flocculants was positively correlated with flocculation efficiency or biomass recovery. Algal-bacterial consortia (axenic species of C. sorokiniana, S. obliquus, Chlorococcum sp.) grown in piggery wastewater were flocculated with two ferric salts (FeCl3 and Fe2(SO4)3) and five organic flocculants (Drewfloc 447, Flocudex CS/5000, Flocusol CM/78, Chemifloc CV/300 and Chitosan) to evaluate their flocculation efficiency or biomass removal efficiency. Biomass removal efficacy of 66–98% was recorded for ferric salts at 150–250 mg L−1 concentration while, polymer flocculants displayed similar efficiency even at a low dose of 25–50 mg L−1 [99]. As stated by Selesu et al. [119], the nontoxic Tanfloc SG at 210 mg L−1 dose was found to recover 96.7% microalgal cells at pH 7.8 based on central composite design based optimization. Unique polyelectrolyte “crystalline nanocellulose” synthesized from the acid hydrolysis of cellulose fibers has been found to show 90% floc recovery at 200 mg L−1 concentration in C. vulgaris culture with a pH of 3.5 adjusted through carbon dioxide purging [116]. Morrissey and coworkers [120], used recyclable various polyampholytes flocculant viz., cationic N,N-dimethylaminopropyl acrylamide, non-ionic methacrylamide and anionic acrylic, which was known to show 99% floc recovery in C. vulgaris culture suspension. Further, polyampholyte flocculant synthesized from N,N-dimethylaminopropyl acrylamide, and acrylic acid through copolymerization was able to flocculate Chlamydomonas reinhardtii, Synechococcus sp. Aulacoseira ambigua, N. gaditana cells [116]. On the other hand, Flopam and Zetag polyacrylamide polymers showed better flocculation efficiency in freshwater C. vulgaris culture,

> 95 92 98

but those polymers were incompetent to recover marine microalga N. oculata and showed only 75% recovery [116]. In the path of polyacrylamide flocculants, high-molecular-weight cum higher charge density (2–4 meq g–1) polyacrylamide at low dose of 2–5 mg L–1 was quite effective in recovering > 95% of log phase cells of S. obliquus, S. subspicatus, Synechococcus nidulans and C. sorokiniana [121]. Uduman et al. [122] documented 89.9%, 84.5% and 79.9% recovery for cationic polymer 71303, anionic polymer 82230 and nonionic polymer Magnafloc 351, respectively using marine microalga Chlorococcum sp. culture. Additionally, in situ focused beam reflectance measurements (FBRM) was adopted to monitor the kinetics of microalgal flocs formation during settling process by measuring particle counts, chord size and density of unflocculated cells in the supernatant. Furthermore, other biopolymers such as the poly-g glutamic acid (excreted by Bacillus subtilis) [123], and cationic polymers in Moringa oleifera seeds [92,124] were also used for microalgal harvesting. In a very recent study, several inorganic salts, polymer flocculant, Moringa seed powder and electroflocculation methods were investigated concerning maximal floc recovery, less solid co-precipitation with marine microalga C. vulgaris [125]. Of the flocculants, aluminium sulfate (70 mg L−1), cationic polymer flocculant (20 mg L−1) and Cu electrode mediated electroflocculation (50 V) unveiled ∼98% floc recovery at 120 min settling time whereas Moringa seed powder displayed poor flocculation efficiency. As a striking observation, polymer flocculant showed minimum solid co-precipitation relatively to other flocculants, and further, dye exclusion staining showed intact viable microalgal cells in flocs obtained through polymer flocculant. Gerchman et al. [126] have compared the flocculation potential of three polymers such as Polydiallyldimethylammonium chloride (PDADMAC), chitosan and Superfloc in C. vulgaris culture suspension, 1111

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and identified that PDADMAC was highly efficient (90%) in flocculating low cell density suspension of 5 mg L−1 in 60 min settling time at pH 10. Few researchers have demonstrated the flocculation of microalgae through “aminoclay,” which has a cationic metal center composed of Mg2+ or Fe3+ (through sol–gel reaction) and (CH2)3-NH2 organofunctional pendants (from 3-amino- propyltriethoxysilane precursor) covalently bonded to metal centers. Synthesized cationic aminoclay at 1 gL−1 dose was found to sediment 98% of microalgae swiftly within 5 min settling time through electrostatic interaction between aminoclay and algal cell surface [127]. Flocculation of Scenedesmus sp. and S. obliquus cultures was demonstrated by Wu et al. [128] using Al2(SO4)3, chitosan, polyacrylamide, and HNO3 and NaOH. Results revealed that all flocculants showed similar flocculation efficiency > 90% in both the cultures; however, dosages of chitosan and polyacrylamide are lower than other flocculants. In addition to pH increased flocculation and chemical flocculant ferric chloride, flocculation efficiency of the organic flocculants maize starch, rice starch on harvesting Botryococcus, Chlorella, Chlamydomonas, Chlorococcum was assessed. Based on the results, it is observed that pH induction showed the minimum efficacy of 72–76% in selected strains whereas ferric chloride and rice starch displayed similar efficiency ~ 87% in Botryococcus species [129]. A new harvesting method was designed using thermoresponsive and charged copolymers of N-isopropylacrylamide and allylamine for C. protothecoides cells in which, when culture and polymers (25–50 mg/mL concentration) are heated above the critical temperature of polymers (~32 °C), cells are separated from solution [130]. In concern with a natural flocculant, Moringa oleifera seed derivatives were used to evaluate its flocculation efficiency in freshwater Chlorella sp. [131]. Primary derivative, i.e., seed powder at 10 mg L−1 dose unveiled > 95% flocculation efficiency at 20 min settling time, whereas other primary and secondary derivatives such as cupule/ wings, seed coat, seed oil, and de-oiled seed were found to reveal < 10% flocculation efficiency. Further, plant-based natural flocculant offers advantages such as green harvesting technology with low environmental hazard and inexpensive microalgae harvesting Similarly, another study was carried out to examine the effect of settling time and pH on Moringa oleifera seed flour facilitated flocculation of C. vulgaris [124]. Data implied that Moringa oleifera seed flour at 0.6 g L−1 dose was capable of flocculating 80% C. vulgaris cells in 120 min time at pH 9.2. Borges et al. [132] stated anionic flocculants is the best in harvesting biomass for producing biodiesel with low unsaturated fatty acids. In this regard, microalgal cultures were flocculated with NaOH, NaOH with anionic Magnafloc®, NaOH with cationic Flopam®. Results depicted that N. oculata cells flocculated by anionic polyacrylamide flocculant Magnafloc® showed high C14:0, C16:0, C16:1 fatty acid contents, and low level of C20:5. Chemical flocculants exhibit following drawbacks:

technique in harvesting microalgae and it holds the advantage of reusing spent medium for scale-up [135]. Autoflocculation occurs either spontaneously when the cultures were cultivated prolonged tenure in sunny days with less CO2 supply or by induction through the addition of alkali to increase the medium pH [56]. In certain strains, e.g., P. tricornutum spontaneously reaches high pH by its regular photosynthesis process during log phase [98]. Autoflocculation of microalgal cells at basic pH attributed to precipitation of carbonate salts and co-precipitation of magnesium and calcium ions by means of reduced CO2 supply (CO2 consumption) [52,136]. In addition to the above, it is hyphothesized that calcium phosphate precipitate is positively charged due to the supersaturation of surplus calcium ions at elevated pH, and is solid supported by negatively charged algal cells in the medium for flocculation [137]. In general, 0.0031–0.0062 g L−1 phosphate and 0.06–0.1 g L−1 calcium is a prerequisite to ensure autoflocculation at 8.5–9 pH [136], and it can be overcome in the certain culture medium by lime addition to attain 80% floc recovery [138]. In addition to the spontaneous sedimentation of cells, autoflocculation is also induced by increasing culture suspension pH above neutral. Autoflocculation of C. vulgaris by increasing the medium pH to the basic range by different salts such as sodium hydroxide, sodium carbonate, magnesium hydroxide, potassium hydroxide, and calcium hydroxide has been demonstrated by Vandamme et al. [139]. Among the salts tested, sodium carbonate did not flocculate the cells, even raising the pH to 11 whereas sodium hydroxide (9 mg g−1 biomass), followed by calcium hydroxide (12 mg g−1 biomass) and potassium hydroxide (18 mg g−1 biomass) flocculated the cells with an efficiency > 95% at pH 11 [139]. Similarly, D. tertiolecta cells have been successfully flocculated at > 90% efficiency when the pH rose between 8.6 and 10.5 by NaOH addition [135]. In another study, flocculation efficiency of alum, FeCl3, chitosan, pH-induced flocculation, dissolved air flotation and electro-flotation on S. obliquus and C. vulgaris cultures were evaluated in lab and field-scale [140]. In lab-scale experimentations, flocculation efficiency of 83.2%, 80.2%, 95%, 91%, 99% and 91% was observed for pH increased flocculation (> 12), FeCl3 (200 mg L−1), alum (250 mg L−1), chitosan (20 mg L−1), electro-flotation (24 V) and dissolved air flotation (1 mg L−1 alum) respectively after 1 h settling time in both the cultures. Under field-scale studies with 1000 L culture, dissolved air flotation (10 mg L−1 alum) and electro-flotation (60 V) showed 90% flocculation efficiency after 24 and 7 h respectively, whereas pH-induced flocculation unveiled 60–75% after 1 h settling time. On an interesting note, pH increased flocculation can be economically feasible and environmentally benign method as the treated supernatant could be reused to scale up the culture with appropriate nutrient inputs. In a very recent research, Leticia Perez et al. [141] studied both pH increase and decrease flocculation methods in which biomass recovery of two different pH ranges such as 2–6 (acidic) and 8–12 (basic) was examined using marine Skeletonema Costatum and Chaetoceros Gracilis. Interestingly, total biomass recovery was noticed in both the culture suspension at basic pH, whereas only 60% recovery was seen in acidic pH experiments. Harvesting induced pH has been studied in various strains such as A. septentrionalis, N. oculata, Skeletonema sp. Tetraselmis suecica, Chaetoceros muelleri, Chaetoceros calcitrans, which exhibits beyond 85% biomass recovery [142]. In contrast to increasing pH for flocculation (alkali range), certain microalgae Chlorococcum nivale, Chlorococcum ellipsoideum and Scenedesmus sp. were harvested (> 90%) by reducing culture suspension pH to 4 (acidic range). The mechanism behind the acid-mediated flocculation was carboxylate ions on microalgal cell surface accept the protons and destabilize the network by neutralizing its negative charges, and eventually, cells agglomerate and settle down [143]. It is reported that marine Chlorella cells were successfully flocculated at 90% efficiency by increasing medium pH with 5–7 mM NaOH [144]. Similarly, high flocculation efficiency of 90% was noticed when pH was raised up to 10.5 in a marine diatom P. tricornutum culture [145].

➢ End product contaminated with metal salts produces surplus sludge, which imposes metal salts separation from algal biomass [50]. ➢ Metal salts associated with floc might deteriorate biomass quality. ➢ Metal salts flocculants are pH dependent. ➢ Optimal flocculant (either organic or inorganic) suitable for one algal strain, might be inappropriate for other microalgal strains. ➢ Inorganic flocculants do not perform well in the flocculation of seawater grown microalgae as they tend to coil at high ionic strength [92]. 2.3. Biological methods 2.3.1. Autoflocculation The word autoflocculation is coined by Golueke and Oswald [133], and later, the terminology autoflocculation and bioflocculation were used interchangeably in literature albeit their difference [134]. Autoflocculation or bioflocculation is low cost, ecofriendly, energy efficient 1112

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Table 4 Harvesting of microalgae by autoflocculation and bioflocculation methods [47,72,123,139,149–154]. Sl. No

Nonflocculating algae

Microorganism used as bioflocculant

Flocculant type (technical specification) or mechanism

Floc recovery or removal efficiency (%)

1.

Chlorella vulgaris

Paenibacillus sp.

83

2.

Desmodesmus brasiliensis

3. 4. 5.

Poly γ-glutamic acid Poly γ-glutamic acid Poly γ-glutamic acid

95 95 95

Bacillus subtilis Bacillus subtilis Ettlia texensis Ankistrodesmus falcatus Scenedesmus obliquus Tetraselmis suecica Bacteria Bacterial population

Poly γ-glutamic acid Poly γ-glutamic acid – – – – EPS –

> 90 > 90 40 36 31 50 97.5

14. 15.

Chlorella vulgaris Chlorella protothecoides Botryococcus braunii LICME 003 Phaeodactylum tricornutum Nannochloropsis oculata LICME 002 Chlorella vulgaris Chlorella vulgaris Chlorella vulgaris Neochloris oleoabundans Pleurochrysis carterae Consortium of Pediastrum sp., Phormidium sp. and Scenedesmus sp. Heterotrophic Chlorella vulgaris UMN235 Chlorella vulgaris UTEX 259

Bacillus licheniformis CGMCC 2876 Bacillus subtilis Bacillus subtilis Bacillus subtilis

Bioflocculant, and 6.8 mM CaCl2 as co flocculant Poly γ-glutamic acid

– –

63 99

16. 17. 18.

Selenastrum capricornutum Nannochloropsis oceanica Pediastrum sp.

– – –

< 95 88 98

19.

Scenedesmus sp. consortium

–

98

20.

Phormidium sp.

–

98

21. 22. 23. 24. 25. 26. 27. 28. 29. 30.

Pleurochrysis carterae Dunaliella tertiolecta Chlorella vulgaris Chlorella vulgaris Chlorella vulgaris Chlorella vulgaris Chlorococcum ellipsoideum Chlorococcum nivale, Scenedesmus sp. Chaetoceros muelleri Chaetoceros calcitrans Nitzschia closterium Tetraselmis suecica Thalassiosira pseudonana, Rhodomonas salina

Fungi Aspergillus oryzae Cunninghamella echinulate NRRL 3655 Paenibacillus sp. Solibacillus silvestris Bacterial consortium from waste water Bacterial consortium from waste water Bacterial consortium from waste water Tap water bacterial inoculum pH increase by NaOH Sodium carbonate Potassium hydroxide Calcium hydroxide Magnesium hydroxide 1 M HNO3 1 M HNO3 1 M HNO3 NaOH and non-ionic polymer

6. 7. 8. 9. 10. 11. 12. 13.

31. 32. 33. 34.

NaOH NaOH NaOH NaOH

and and and and

non-ionic non-ionic non-ionic non-ionic

polymer polymer polymer polymer

98

– pH pH pH pH pH pH pH pH pH

increase to 10.5 increase to 11 increase to 11 increase to 11 increase to 11 decrease to 3.5–4 decrease to 3.5–4 decrease to 3.5–4 increased between 10.0 and 10.6

< > 0 > > > > > > <

pH pH pH pH

increased increased increased increased

< 95 85–95 < 95 < 90

between between between between

10.0 10.0 10.0 10.0

and and and and

10.6 10.6 10.6 10.6

93 90 95 95 95 90 95 97 95

mediated microalgal flocculation by supplementing organic carbon sources to promote bacterial growth. Flocculation efficiency of 90% was observed in Pleurochrysis carterae by the EPS produced by the bacteria at 0.1 g L−1 acetate or glucose or glycerine and 24 h mixing [149]. The green microalgae C. vulgaris and Scenedesmus sp. was efficaciously flocculated using coagulant secreted by Paenibacillus polymyxa bacterium [41,146]. In another research, both the flocculant secreted by the bacterium Paenibacillus sp. AM49 and co-flocculant 6.8 mm CaCl2 were found to sediment 83% of C. vulgaris cells within 5–11 pH range, which was notably maximum than inorganic aluminium sulfate (72%) and organic polyacrylamide (78%) respectively [155]. Further, poly γ-glutamic acid produced by the bacterium Bacillus licheniformis CGMCC 2876 doubtlessly unveiled > 98% recovery in Scenedesmaceaen green alga Desmodesmus brasiliensis and notably, neither the lipids extracted from bioflocculated biomass nor from centrifuged biomass showed discrepancy [149]. The bacterial population is identified to settle down the consortium of microalgae Chlorella sp. Pediastrum sp. Phormidium sp. and Scenedesmus sp. from the vicinity of the power plant at 97.5% efficiency in 30 min incubation time [72]. Fungal based bioflocculation of microalgae is also carried out by Zhou et al. [153] with pellet forming Aspergillus oryzae for facultative heterotrophic algae C. vulgaris UMN235. The experimental set up was aimed to optimize spore density, glucose concentration and pH range for enhanced floc recovery and the data apparently revealed that

2.3.2. Bioflocculation Bioflocculation is slightly a different phenomenon from autoflocculation in which polymers specifically EPS secreted by the microorganisms tend to flocculate the target microalgae [38]. The rapid bioflocculation process depends on the quantity of EPS, capability rate of microalgal attachment to polymers and growth phase of bacteria (the deflocculating activity of enzymes was witnessed in the stationary phase bacterial cells) [47,146,147]. The most common EPS involved in cell adhesion during bioflocculation is uronic acids and pyruvic acids [148]. The harvesting of microalgae by autoflocculation and bioflocculation methods is described in Table 4. Recently, co-culturing of autoflocculating microalga with non-flocculating microalga is considered to be plausible microalgal harvesting method [91]. The research work carried out by Salim et al. [150] divulged that the addition of autoflocculating microalgae Ettlia texensis, Ankistrodesmus falcatus and S. obliquus to non-flocculating alga C. vulgaris increased the floc recovery from 25% to 40%, 36%, and 31% respectively and further, floc recovery of Neochloris oleoabundans was increased by 10% with autoflocculating Tetraselmis suecica. Co-culturing of autoflocculating alga Skeletonema sp. with non-flocculating oleaginous microalga Nannochloropsis was suggested to be an exceptionally potential harvesting method [110]. The EPS or polymer or bioflocculant secreted by the bacteria was evidently observed to flocculate microalgae extensively [52]. Lee et al. [149] carried out bacterial 1113

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Table 5 Harvesting of microalgae by electroflocculation and magnetic nanocomposite facilitated flocculation [152,164,165,169–177]. Sl. No

Strain

Flocculation type

Electrode material Flocculant type (technical specification)

Floc recovery or removal efficiency (%)

1. 2. 3. 4. 5. 6. 7.

Chlorella vulgaris Phaeodactylum tricornutum Phaeodactylum tricornutum Chlorella vulgaris Chlorella vulgaris Botryococcus braunii Botryococcus braunii

16.

Microcystis aeruginosa Microcystis aeruginosa Scenedesmus acuminatus Chlorococcum sp. Tetraselmis sp. Chlorella vulgaris Chlorella vulgaris Chlamydomonas reinhardtii, Chlorella vulgaris and Phaeodactylum tricornutum, Nannochloropsis salina Botryococcus braunii and Chlorella ellipsoidea

Aluminium at 3 V cm−1 Aluminium at 3 mA cm−2 current intensity Iron electrodes at 3 mA cm−2 current intensity Aluminium at 3 mA cm−2 current intensity Iron electrodes at 3 mA cm−2 current intensity Aluminium electrodes Aluminium plates and air flow rate at 0–60 mL min−1 Aluminium electrodes and chloride ions Iron electrodes Mg electrode Stainless steel Stainless steel Aluminium Iron Silica-coated magnetic particles called MagSilica at pH 8–12

90 ≤ 81 ≤ 78 ≤ 81 ≤ 91 93.6 99

8. 9. 10. 11. 12. 13. 14. 15.

Electroflocculation Electro-coagulation-flocculation Electro-coagulation-flocculation Electro-coagulation-flocculation Electro-coagulation-flocculation Electroflocculation Electroflocculation integrated with dispersed air flotation Electro-coagulation-flotation Electro-coagulation-flotation Electroflocculation Electrocoagulation Electrocoagulation Electro-coagulation- flocculation Electro-coagulation- flocculation Magnetic separation

Magnetic separation

17.

Chlorella sp.

Magnetic separation

18. 19. 20.

Nannochloropsis oceanica Chlorella vulgaris Chlorella ellipsoidea

Magnetic separation Magnetic separation Magnetic separation

21. 22.

Chlorella ellipsoidea Chlorella sp.

Magnetic separation Magnetic separation

23. 24. 25.

Nannochloropsis maritima Nannochloropsis maritima Chlorella vulgaris

Magnetic separation Magnetic separation Magnetic separation

Cationic polyacrylamide (0.1 mg/mL) coated iron oxide Magnetic iron oxide coated amino-riched polyamidoamine pH-8 Uncoated Iron oxide particles at pH ≥ 8 Iron oxide magnetic microparticles Naked Fe3O4 particles (chemical precipitation) pH 7 Fe3O4- Polyethylenimine nanocomposites, pH 9 Diallyldimethylammonium chloride) - Fe3O4 nanoparticles Silica-coated magnetic Particles, pH-8 Naked Fe3O4 particles pH-8 DEAE magnetic beads pH-4

1.2 × 104 fungal spores/mL, 20 gL−1 glucose, and 4.0–5.0 pH were able to recover 63% algal cells in heterotrophic mode whereas 10 gL−1 glucose was high enough to recover 100% autotrophically grown algal cells. Other fungi used to entrap algal cells are Rhizopus oryzae, Penicillium expansum and Mucor circinelloides [156]. Similarly, 95% recovery was attained for C. vulgaris, and C. protothecoides was by 19–22 gL−1 poly γ-glutamic acid obtained from Bacillus subtilis, and the same concentration showed > 90% flocculation efficiency in N. oculata LICME 002 and P. tricornutum (0.5–0.6 gL−1 biomass density) [123]. It is also described that poly γ-glutamic acid-mediated flocculation is a potential harvesting method in biodiesel perspective as it does not show the sign of lipid loss and cell damage [123]. Despite the better biomass removal efficiency observed in bioflocculation technique, the exact mechanism of bioflocculation process is not clearly understood, and it requires follow-up study [110,157]. The probable underlying mechanism is suggested by using Selenastraceaen algae Ankistrodesmus falcatus to flocculate non-selenastraceaen member C. vulgaris (Chlorellaceae). The part of EPS (excreted by autoflocculating strain) bind to the surface of Ankistrodesmus falcatus and another part possessing positively charged tails bridged with other Ankistrodesmus falcatus cells to form large network wherein Chlorella cells were entrapped [91]. Several researchers have cultured various bacterial strains to produce and purify bioflocculant polymer for microalgal harvesting viz., Alcaligenes cupidus KT201, Enterobacter sp., Bacillus coagulans, Pullularia pullulans and Rhodococcus erythropolis [149,158,159].

100 ≤ 79 90 98 99 99 64 > 90

> 95 > 95 99% 95 > 98 97 99 < 40 97.5 90

interfere with extraction of lipid from target microalgae into lowvalue biofuels product since added autoflocculating microalga comprising > 25% lipid could contribute a notable fraction in total lipid content [91,150,155]. 2.3.2.2. Limitations.

• Bacteria [149] and diatom [110] mediated flocculation of micro• • • •

algae demand different culturing conditions, which enables contamination risk and extra expensiveness of additional nutrients. Requires comparatively high organic carbon (approx. 20%) to culture autoflocculating microbes for microalgal harvesting [149]. Microalgal biomass obtained through flocculation by bacteria and fungi interferes with food or feed applications owing to contamination [92]. Time-consuming and inconsistent process over strains [110]. No hitherto literature on successful harvesting method at large scale [149].

2.4. Electrical based harvesting Electrical based microalgal dewatering is a rapid and strain independent method [47]. Different speculations on the underlying mechanism of electroflocculation have been reported since the definite and detailed mechanism of electroflocculation is not completely understood and disclosed. As reported by Richardson et al. [160], electrolytic flocculation entails nonreactive (non-sacrificial) anode and cathode. Negatively charged algal cells move towards positively charged anode due to electrophoretic motion where cells lose their charge, and form flocs or aggregates by the charge neutralization and may settle down at bottom or float on the surface based on the density [161]. On the contrary, electroflocculation entails reactive (sacrificial) electrodes to release metal flocculants, and settling occurs in

2.3.2.1. Advantages of Bioflocculation.

• Practically feasible and ecofriendly approach over chemical based flocculation [47]. • Power consumption is low and does not require chemicals [150]. • Bioflocculation using autoflocculating microalga does not appear to 1114

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efficiency in 1 h settling period with the addition of 5 mM chlorine ions to the freshwater microalga Microcystis aeruginosa, which is attributed to the release of Al3+ from the aluminium anode and increased positive zeta potential by active Cl− ions. Nonetheless, SEM-EDX analysis displayed formation of transient pores in the cell membrane attributed to the trans-membrane potential caused by electric field [171]. Similarly, 1 mA cm− 2 current density using aluminium anode, pH 4–7, 18–36 °C temperature, 0.55 × 109 –1.55 × 109 algal cells/L were found to be the optimal parameters to recover 100% Microcystis aeruginosa cells, which is confirmed by atomic force microscopy (AFM) analysis [167]. Lee et al. [178] used baffled hydraulic mixer and gravity clarifier in electroflocculation for energy effective mixing and settling of marine Tetraselmis sp. and the resulting cost including power consumption, dissolution of cations from an anode and capital depreciation is estimated to be $0.19 kg−1. Floc recovery of two marine microalgal species Chlorococcum sp. and Tetraselmis sp. with respect to temperature and salinity by stainless steel electrode mediated electrocoagulation were compared, and 98% floc recovery was observed from both marine strains at 60 °C temperature and 20% salinity [172]. Besides the use of sacrificial electrodes for electroflocculation, nonsacrificial electrodes are also for the electrochemical harvesting of microalgae. Tiny cell sized chlorophycean alga Ankistrodesmus falcatus was electroflocculated by non-sacrificial carbon electrodes and the efficiency and performance of electrochemical harvesting was compared to centrifugation, alum and chitosan flocculant. Maximum efficiency was in the order as centrifugation (93%) > electrochemical harvesting (91%) > alum (86%) > chitosan (55%) and nonetheless, the energy consumption of centrifugation (65.34 kWh kg−1) was much higher than electrochemical harvesting (1.76 kWh kg−1) [179]. Besides the widely used aluminium and iron electrodes for electroflocculation, other electrodes such as magnesium, copper, zinc, and brass were also tested in S. acuminatus culture at 10, 20, 30 and 40 V. The highest flocculation efficiency was achieved using Mg, Al, Zn and however Mg electrodes showed cost-effective flocculation in terms of power consumption than other electrodes. The settling time for Mg electrode to attain 90% of flocculation efficiency at 40, 30, 20, and 10 V was 7.3, 12.5, 18.5, and 43.2 min, respectively. Low toxicity or less hazardous property of Mg ions make them advantageous for flocculated biomass to be used for food or feed application [164]. Local sand enhanced electro-flocculation of D. Salina was also evaluated, as sand is effective in accelerating flocculation and settling of flocs. The maximal recovery of 98.09% was noticed in 4.5 min by sand enhanced electro-flocculation method, which consumes 51.03% less energy compared to traditional electroflocculation [180]. In recent times, the pair of aluminium and borondoped diamond electrodes was also used for the electrochemical harvesting of S. quadricauda cells [181].

consecutive stages: (i) electrolytic oxidation of reactive anode releases flocculants (metal ions) for dissolution to culture suspension (ii) destabilization of cell suspension network by compressing double layer around the cells by metal ions and (iii) strengthening of Van der Waals force by minimizing electrostatic repulsive forces between cells, (iv) aggregation (floc) of the destabilized particles in the suspension due to charge neutralization [42,88,162,163]. Aluminium and Iron electrodes are widely used in electrolytic flocculation [47], which release Al(aq)3+ and Fe(aq)3+ ions respectively in an electric field, and they are spontaneously hydrolysed to hydroxides and/or polyhydroxides [88]. Aluminium metal ions produce monomeric and polymeric species such as Al(H2O)63+, Al(H2O)5OH2+, Al(H2O)4(OH)2+, Al(OH)2+, Al(OH)2+, Al2(OH)24+, Al(OH)4−, Al6(OH)153+, Al7(OH)174+, Al8(OH)204+, Al13O4(OH)247+, All3(OH)345+ and similarly, ferric ions produce Fe(OH)3, Fe(H2O)63+, Fe(H2O)5(OH)2+, Fe(H2O)4(OH)2+, Fe2(H2O)8(OH)24+ and 4+ Fe2(H2O)6(OH)4 [88]. These positive metal hydroxides act as an active surface to adsorb negatively charged microalgal cells and therefore, algal cells were entrapped as flocs due to the neutralization of the surface charge [164]. As detailed in Table 5, better performance of aluminium electrode than iron electrode is reported by many authors [165–168]. Aluminium electrodes flocculation efficiency is about 100% than iron electrode (78%) as it possesses high electricity and thereby dissociates more Al3+ ions for rapid flocculation [47]. Electrical density and ionic strength (medium) are negatively correlated with settling time [167], release of metal ions (presence of maximal active binding sites on the surface) were increased at high current density and thus, fast sedimentation of microalgal cells was observed [47]. In concern with ionic strength, electroflocculation of freshwater microalgae consumes energy double the time than marine microalgae since freshwater medium has low ionic strength, which increases the settling period [163]. The impact of current density and pH of the medium on the flocculation efficiency of unicellular green alga C. vulgaris Beijerinck by electroflocculation was studied by Kumar et al. [169], and data divulged that 3 V cm−1 voltage density and pH 7.0, recovered 90% of C. vulgaris cells within 30 min settling time. Electro-coagulation-flocculation of freshwater (C. vulgaris) and marine (P. tricornutum) microalgae was demonstrated using a pair of aluminium or iron electrodes [165]. Results indicated that high power density of aluminium shows rapid flocculation efficiency than sacrificial iron electrode and further, aluminium content in the flocculated biomass and processed water was < 1% and 2 mg L−1 respectively. The power consumption of electrodes for marine P. tricornutum and freshwater C. vulgaris were determined to be 0.3 kWh kg−1 and 2 kWh kg−1 respectively, which make them foreseen to be energy efficient harvesting method for marine microalgae [165]. In certain cases, electroflocculation is integrated with flotation for effective microalgal harvesting. Xu et al. [170] reported 93.6% flocculation efficiency in 30 min settling time through electroflocculation of Botryococcus braunii (B. braunii) whereas the efficiency was increased to 99% in a shorter settling period of 14 min when electroflocculation is combined with dispersed-air flotation. The effect of several factors such as flocculation time, conductivity, current density, the distance between electrodes, and electrode type on harvesting efficiency of Dunaliella sp. was studied. Aluminium electrodes (1 cm distance), at 1.35 S m−1 conductivity, 90 A m−2 current density, 3 min settling time, showed 97.44% flocculation efficiency [163]. Continuous flow electro-coagulation and flotation involving three aluminium anodes were utilized to remove the algae from wastewater treatment plant. Total suspended solids (algal particles) removal efficiency was reached to 99.5% with 550 W current density in 15 min retention time, and further, the same efficiency was recorded in 30 min settling time when the current density was decreased [168]. An electro-coagulation-flotation process is also improved by the addition of chloride ions. Aluminium electrodes showed remarkably 100% removal

2.4.1. Advantages ✓ Eco-friendly, strain -independent, energy efficient, and inexpensive [42,88]. ✓ Reliable and not a laborious operation. Does not release undesirable sulfate or chloride ions to the biomass [165]. ✓ Simple instrumental set up with merely modest investment [178]. ✓ Does not demand chemical salts, which will contaminate the biomass [47]. ✓ Relatively efficient than polymer (chitosan) flocculants in harvesting dilute or low cell density suspensions [161]. 2.4.2. Disdvantages ➢ Increased power density may elicit unfavorable change in microalgal cells [166]. ➢ The current density of 0.5 mA cm−2 consumes 0.2 kWh m−3 energy, while 5.0 mA cm−2 current density shows the energy consumption of 2.28 kWh m−3 in microalgal harvesting [167]. 1115

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➢ Some electrode (ferric) is not beneficial, as it produces colored biomass after flocculation [47]. ➢ Cathode fouling, and harvesting system damage [87,161]. ➢ Reusability of electrodes is not much desirable due to internal resistance [165]. ➢ For large scale harvesting, electrophoretic equipment imposes high cost [38].

magnetic particles and were separated by a water-nonpolar solvent interface from hydrophilic microalgae. This method shows promise for high cell density microalgae ranging 1.5 gL−1 to 60 gL−1 and harvested biomass directed for nonpolar organic solvent assisted wet lipid extraction overcoming multiple downstream processing [189]. In situ magnetic particles facilitated method was also developed using uncoated iron oxide for B. braunii and C. ellipsoidea. At a stirring speed of 120 rpm/min, 98% algae were recovered within 1 min settling time. The adsorptivity of magnetic particles (electrostatic attraction) was 55.9 and 5.83 mg cells/mg particles for B. braunii and C. ellipsoidea respectively [185]. Coating of magnetite nanoparticles has the robust impact on zeta potential and load of magnetic nanoparticles. Polyethylenimine coating changes the zeta potential − 7.9 to + 39.0 mV at a pH of 7.0, and decrease the load requirement from 0.2 to 0.1 g g−1 S. dimorphus cells irrespective of separation efficiency (80%) [190]. In a recent work, long chain poly-arginine modified porous Fe3O4 microspheres were used to separate Chlorella cells from suspension at 10 mg L−1 dose and further, it has good recycling feature due to large microsphere size [191]. Polypyrrole/Fe3O4 magnetic nanocomposite was reported to be costeffective and energy intensive in biomass separation. The recovery efficiency of 99%, 92.4%, and 90.8% was observed from B. braunii, C. protothecoides, and C. vulgaris at pH 10.0 [192]. Effect of pH on harvesting N. oceanica using magnetic iron oxide particles was also studied. At basic pH and low particle concentration, 99% separation efficiency was obtained despite constant stirring speed and time. Further, adsorption characteristics of iron oxide particles to algal cell surface follows Freundlich model, i.e., adsorption process was multilayer onto heterogeneous situates [176]. Despite its rapidity in separation, postharvest separation, recycling of nanoparticles, practical applicability, and uncontaminated microalgal biomass production are still foremost bottlenecks [189]. In the line of key limitations, requirement of more magnetic nanoparticles may increase the overall process cost [174].

2.5. Magnetic particle facilitated separation Recent past, magnetic particles facilitated separation has been considered as an emerging technique for microalgal harvesting [182], in which suspended cells were tagged or adsorbed to micron-sized or nano-sized magnetic particles, and the tagged composites were separated using external magnet force owing to intrinsic paramagnetic movement [97,173]. It is most acclaimed modern harvesting method in terms of rapid, simple, cost-effective and energy efficient characteristics [174]. Various magnetic nanocomposites used for the separation of microalgae are tabulated in Table 5. Ideal magnetic particles should have the following the properties for efficient harvesting viz., reusability, stability, adsorptivity and low-cost bulk product generation [183]. In this connection, Iron oxides (Fe3O4) was used as a basis in the meadow of magnet based microalgal harvesting [184,185], wherein flame gas-phase processes were used to produce and coat the magnetite by iron salts precipitation [186]. Fe3O4 magnetic nanoparticles were widely used for harvesting various types of microalgal strains due to their specific surface area and super paramagnetism [152,185]. However, low harvesting efficiency of uncoated Fe3O4 magnetic nanoparticles necessitates modification [187]. In this juncture, polyethylenimine and cationic polyacrylamide were used to modify Fe3O4 magnetic particles to positive magnetite for enhancing its adsorptivity towards negatively charged algal cells [188]. Further, algal removal efficiency relies on magnetic particle load, pH, and composition of medium [173]. Silica-coated magnetic particles were used to separate fresh water algae Chlamydomonas reinhardtii, C. vulgaris and marine algae P. tricornutum, N. salina. High gradient magnetic filtration showed > 95% algal removal with maximum particle loads of 30–77 g g−1 at pH 8–12 within 5 min [173]. Cationic polyacrylamide (0.1 mg/mL) coated iron oxide showed recovery efficiency > 95% within 10 min at a load of 25 and 120 mg L−1 for B. braunii and C. ellipsoidea respectively, and their adsorption to magnetic particles was determined to be multilayer heterogeneous according to Freundlich isotherm. The highest adsorption was 114.8 and 21.4 mg cells/mg−1 for Botryococcus and Chlorella respectively at neutral pH, and the core electrostatic interactions between cells and particles were found to “bridging type flocculation” [174]. Magnetic iron oxide coated separately with dendrimer (amino-riched polyamidoamine), and amino acid was compared for efficient harvesting of Chlorella sp. The results showed that adsorption efficiency of magnetic iron oxide coated with highly positive branched dendrimer at 80 mg L−1 load had dramatically reached to 95% within 2 min at pH 8.0 compared to amino acid modified Fe3O4 and the efficiency further improved when pH brought down to 4.0 [188]. Phosphorus ions in the medium are identified as one of the vital factors interfering magnetic microalgal cell harvesting. Inexpensive iron oxide magnetic microparticles synthesized by microwave treatment showed phosphorus limited C. vulgaris cells were separated from its suspension at 95% efficiency than phosphorous repleted cells with 3:1 load of microparticles to cells. Ultrasound mediated demagnetization of separated cells were carried out at 40 °C assisted with 10% H2SO4 treatment [182]. To overcome energy consuming post-harvesting process, magnetic particles were designed with organosilane compounds namely triethoxysilane and octyltriethoxysilane, and were easily separated by water-nonpolar organic solvent using an external magnetic field. Triethoxysilane exhibits ~ 99% recovery at a dose of 1.6 g g−1 cells while octyltriethoxysilane confers lipophilicity to

3. Conclusions and way forward Broad marketing of microalgal biodiesel production stuck in a cost factor. For economic biodiesel production, inexpensive large-scale harvesting process yet to be developed through detailed research studies. With this in view, this review manuscript was undertaken to provide a comprehensive outlook of the harvesting techniques applied and being used in microalgae to date. Preliminary contents have dealt with the present global energy projections and its concerns with respect to the environment, and future energy security. The feasibility of microalgal biofuel in satiating energy demand and its challenges and scope have been widely analysed and discussed. The paramount challenge in microalgal biodiesel, i.e., “harvesting” has been taken as a core content of the review, and the various harvesting techniques and its limitations with an emphasis on cost factor were reviewed. The merits and demerits of conventional physical methods such as centrifugation, gravity sedimentation, filtration, flotation and organic, inorganic salts mediated flocculation, pH-induced flocculation, bioflocculation, electrical based harvesting, and advanced magnetic nanocomposites based separation have been enunciated. The key challenges of each harvesting methods were extensively signposted in this review to provide a holistic perspective about energy and cost-efficient separation of microalgae for biodiesel production. High energy requirement and time-consuming characteristics are the major issues in physical methods to dewater voluminous microalgal suspension, which paves the way for exploration of an effective harvesting technique. In concern with bioflocculation method, growth kinetics and contamination of flocculating microalgae have to be determined, and it also questions the dose of flocculating microalgae to be dispensed to target microalgae. Also cost of the separate pond for culturing flocculating algae has to be taken into account to enable its suitability in reducing cost obstacle. Traditional and advanced methods 1116

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have long been employed for the competent and low-cost harvesting of algae; however, lack of regimented algal removal systems is the prime impediment for separating algal cells from raceways. In a line of research activity requirements, electrical based methods eventually necessitate energetic optimization of large-scale trials such as the distance between the electrodes and corrosion or fouling of electrodes due to continuous operation. Intensive research efforts have to be undertaken to fill knowledge gap concerning the issues highlighted in this review for each harvesting process. Further, most of the harvesting methods are invariably confined to laboratory level and therefore, research studies must be carried out at large scale to check their outdoor compatibility. Therefore, major R and D initiatives are still needed to bypass its infancy in bench scale experimentations. Innate features, viability, and energetics of microalgal cell are the additional pivotal properties should be taken into account while reducing harvesting cost. Furthermore, harvesting method has to be selected based on the algal species and the product to be extracted and therefore, new experimental demonstration with various microalgae of different genera needs to be performed to understand the cell behavior on harvesting. Grounded in the pros and cons documented in this review, biodiesel from microalgae still has a long way to go for industrial scale trials to seal the knowledge gaps of expensive harvesting techniques.

[17]

[18] [19]

[20]

[21]

[22] [23]

[24] [25]

[26] [27]

Acknowledgement [28]

The authors thank Science and Engineering Research Board (SERB), Department of Science and Technology, Govt. of India (Grant No. PDF/ 2016/000541, SERB/F /5089/2016-17) for awarding National PostDoctoral Fellowship to Dr. T. Mathimani.

[29]

[30]

References [1] Behçet R. Performance and emission study of waste anchovy fish biodiesel in a diesel engine. Fuel Process Technol 2011;92:1187–94. [2] Mathimani T, Uma L, Prabaharan D. Homogeneous acid catalysed transesterification of marine microalga Chlorella sp. BDUG 91771 lipid–an efficient biodiesel yield and its characterization. Renew Energy 2015;81:523–33. [3] Creasey JJ, Chieregato A, Manayil JC, Parlett CMA, Wilson K, Lee AF. Alkali-and nitrate-free synthesis of highly active Mg–Al hydrotalcite-coated alumina for FAME production. Catal Sci Technol 2014;4:861–70. [4] Hemaiswarya S, Raja R, Carvalho IS, Ravikumar R, Zambare V, Barh D. An Indian scenario on renewable and sustainable energy sources with emphasis on algae. Appl Microbiol Biotechnol 2012;96:1125–35. [5] Caspeta L, Buijs NAA, Nielsen J. The role of biofuels in the future energy supply. Energy Environ Sci 2013;6:1077–82. [6] Hoogeveen J, Faurès J, Van de Giessen N. Increased biofuel production in the coming decade: to what extent will it affect global freshwater resources? Irrig Drain 2009:58. [7] Mathimani T, Uma L, Prabaharan D. Optimization of direct solvent lipid extraction kinetics on marine trebouxiophycean alga by central composite design–Bioenergy perspective. Energy Convers Manag 2017;142:334–46. [8] Bharathiraja B, Chakravarthy M, Ranjith Kumar R, Yogendran D, Yuvaraj D, Jayamuthunagai J, et al. Aquatic biomass (algae) as a future feed stock for biorefineries: a review on cultivation, processing and products. Renew Sustain Energy Rev 2015;47:635–53. http://dx.doi.org/10.1016/j.rser.2015.03.047. [9] Singh SP, Singh P. Effect of CO2 concentration on algal growth: a review. Renew Sustain Energy Rev 2014;38:172–9. http://dx.doi.org/10.1016/j.rser.2014.05. 043. [10] Doshi A, Pascoe S, Coglan L, Rainey TJ. Economic and policy issues in the production of algae-based biofuels: a review. Renew Sustain Energy Rev 2016;64:329–37. http://dx.doi.org/10.1016/j.rser.2016.06.027. [11] Abomohra AEF, Jin W, Tu R, Han SF, Eid M, Eladel H. Microalgal biomass production as a sustainable feedstock for biodiesel: current status and perspectives. Renew Sustain Energy Rev 2016;64:596–606. http://dx.doi.org/10.1016/j.rser. 2016.06.056. [12] Zhou A, Thomson E. The development of biofuels in Asia. Appl Energy 2009;86:S11–20. http://dx.doi.org/10.1016/j.apenergy.2009.04.028. [13] Volpi G. Sustainability and biofuels: lessons from Brazil, presentation for the Conference of the German network on renewable energy. North-South, Bonn; 2005. p. 20. [14] Chandel AK, Bhatia L, Garlapati VK, Roy L, Arora A. Biofuel policy in Indian perspective: socioeconomic indicators and sustainable rural development. Sustain Biofuels Dev India 2017:459–88. Springer. [15] Ray S, Miglani S, Goldar A. Ethanol blending policy in India: demand and supply issues. ICRIER Policy Ser 2011:9. [16] Kang S, Selosse S, Maïzi N. Strategy of bioenergy development in the largest

[31]

[32]

[33] [34]

[35]

[36] [37]

[38]

[39]

[40]

[41]

[42]

[43] [44]

[45]

1117

energy consumers of Asia (China, India, Japan and South Korea). Energy Strateg Rev 2015;8:56–65. Kampman BE, Verbeek R, Grinsven AH, van Mensch P, Croezen HJ, Patuleia A. Bringing biofuels on the market: options to increase EU biofuels volumes beyond the current blending limits. CE Delft; 2013. Economic times. Make ethanol from surplus food grains: draft policy. doi:available at 〈https://goo.gl/Af6zy7〉; 23 Nov 2017. Mofijur M, Masjuki HH, Kalam MA, Rahman SMA, Mahmudul HM. Energy scenario and biofuel policies and targets in ASEAN countries. Renew Sustain Energy Rev 2015;46:51–61. Bangkok post. Oversupply of biodiesel worries producers 〈https://www. bangkokpost.com/news/general/1386642/oversupply-of-biodiesel-worriesproducers〉; 27 December 2017. Mofijur M, Rasul MG, Hyde J, Azad AK, Mamat R, Bhuiya MMK. Role of biofuel and their binary (diesel–biodiesel) and ternary (ethanol–biodiesel–diesel) blends on internal combustion engines emission reduction. Renew Sustain Energy Rev 2016;53:265–78. Runge C. The case against biofuels: probing ethanol's hidden costs. Yale Environ 2010:360. Atabani AE, Silitonga AS, Badruddin IA, Mahlia TMI, Masjuki HH, Mekhilef S. A comprehensive review on biodiesel as an alternative energy resource and its characteristics. Renew Sustain Energy Rev 2012;16:2070–93. Demirbas A. Biofuels sources, biofuel policy, biofuel economy and global biofuel projections. Energy Convers Manag 2008;49:2106–16. Juneja A, Ceballos RM, Murthy GS. Effects of environmental factors and nutrient availability on the biochemical composition of algae for biofuels production: a review. Energies 2013;6:4607–38. http://dx.doi.org/10.3390/en6094607. Hannon M, Gimpel J, Tran M, Rasala B, Mayfield S. Biofuels from algae: challenges and potential. Biofuels 2010;1:763–84. http://dx.doi.org/10.4155/bfs.10.44. Singh J, Gu S. Commercialization potential of microalgae for biofuels production. Renew Sustain Energy Rev 2010;14:2596–610. http://dx.doi.org/10.1016/j.rser. 2010.06.014. Mata TM, Martins AA, Caetano NS. Microalgae for biodiesel production and other applications: a review. Renew Sustain Energy Rev 2010;14:217–32. http://dx.doi. org/10.1016/j.rser.2009.07.020. Mackenzie A. Synthetic biology and the technicity of biofuels. Stud Hist Philos Sci Part C Stud Hist Philos Biol Biomed Sci 2013;44:190–8. http://dx.doi.org/10. 1016/j.shpsc.2013.03.014. Mathimani T, Kumar TS, Chandrasekar M, Uma L, Prabaharan D. Assessment of fuel properties, engine performance and emission characteristics of outdoor grown marine Chlorella vulgaris BDUG 91771 biodiesel. Renew Energy 2017;105:637–46. Kumar K, Mishra SK, Shrivastav A, Park MS, Yang JW. Recent trends in the mass cultivation of algae in raceway ponds. Renew Sustain Energy Rev 2015;51:875–85. http://dx.doi.org/10.1016/j.rser.2015.06.033. Maity JP, Bundschuh J, Chen CY, Bhattacharya P. Microalgae for third generation biofuel production, mitigation ofgreenhouse gas emissions and wastewater treatment: present andfuture perspectives – a mini review. Energy 2014;78:104–13. http://dx.doi.org/10.1016/j.energy.2014.04.003. Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25:294–306. Ribeiro LA, Silva PP Da. Surveying techno-economic indicators of microalgae biofuel technologies. Renew Sustain Energy Rev 2013;25:89–96. http://dx.doi. org/10.1016/j.rser.2013.03.032. Hsieh CH, Wu WT. Cultivation of microalgae for oil production with a cultivation strategy of urea limitation. Bioresour Technol 2009;100:3921–6. http://dx.doi. org/10.1016/j.biortech.2009.03.019. Mubarak M, Shaija A, Suchithra TV. A review on the extraction of lipid from microalgae for biodiesel production. Algal Res 2015;7:117–23. Chaudry S, Bahri PA, Moheimani NR. Pathways of processing of wet microalgae for liquid fuel production: a critical review. Renew Sustain Energy Rev 2015;52:1240–50. http://dx.doi.org/10.1016/j.rser.2015.08.005. Christenson L, Sims R. Production and harvesting of microalgae for wastewater treatment, biofuels, and bioproducts. Biotechnol Adv 2011;29:686–702. http://dx. doi.org/10.1016/j.biotechadv.2011.05.015. Milledge JJ, Heaven S. A review of the harvesting of micro-algae for biofuel production. Rev Environ Sci Biotechnol 2013;12:165–78. http://dx.doi.org/10. 1007/s11157-012-9301-z. Jankowska E, Sahu AK, Oleskowicz-Popiel P. Biogas from microalgae: review on microalgae's cultivation, harvesting and pretreatment for anaerobic digestion. Renew Sustain Energy Rev 2017;75:692–709. http://dx.doi.org/10.1016/j.rser. 2016.11.045. Molina Grima E, Belarbi EH, Acién Fernández FG, Robles Medina A, Chisti Y. Recovery of microalgal biomass and metabolites: process options and economics. Biotechnol Adv 2003;20:491–515. http://dx.doi.org/10.1016/S0734-9750(02) 00050-2. Uduman N, Qi Y, Danquah MK, Forde GM, Hoadley A. Dewatering of microalgal cultures: a major bottleneck to algae-based fuels. J Renew Sustain Energy 2010:2. http://dx.doi.org/10.1063/1.3294480. Benemann JR, Weissman JC, Koopman BL, Oswald WJ. Energy production by microbial photosynthesis. Nature 1977;268:19–23. Danquah MK, Ang L, Uduman N, Moheimani N, Forde GM. Dewatering of microalgal culture for biodiesel production: exploring polymer flocculation and tangential flow filtration. J Chem Technol Biotechnol 2009;84:1078–83. Garzon-Sanabria AJ, Davis RT, Nikolov ZL. Harvesting Nannochloris oculata by inorganic electrolyte flocculation: effect of initial cell density, ionic strength, coagulant dosage, and media pH. Bioresour Technol 2012;118:418–24.

Renewable and Sustainable Energy Reviews 91 (2018) 1103–1120

T. Mathimani, N. Mallick [46] Amer L, Adhikari B, Pellegrino J. Technoeconomic analysis of five microalgae-tobiofuels processes of varying complexity. Bioresour Technol 2011;102:9350–9. [47] Barros AI, Gonçalves AL, Simões M, Pires JCM. Harvesting techniques applied to microalgae: a review. Renew Sustain Energy Rev 2015;41:1489–500. http://dx. doi.org/10.1016/j.rser.2014.09.037. [48] Coward T, Lee JGM, Caldwell GS. Development of a foam flotation system for harvesting microalgae biomass. Algal Res 2013;2:135–44. http://dx.doi.org/10. 1016/j.algal.2012.12.001. [49] Brennan L, Owende P. Biofuels from microalgae—a review of technologies for production, processing, and extractions of biofuels and co-products. Renew Sustain Energy Rev 2010;14:557–77. [50] Chen CY, Yeh KL, Aisyah R, Lee DJ, Chang JS. Cultivation, photobioreactor design and harvesting of microalgae for biodiesel production: a critical review. Bioresour Technol 2011;102:71–81. http://dx.doi.org/10.1016/j.biortech.2010.06.159. [51] Moraine RS, Sandbank E, Bar-Moshe Z, Shvartzburd L. Recovery of sewage-borne algae: flocculation, flotation, and centrifugation techniques. Algae Biomass Prod Use/[sponsored by Natl Counc Res Dev Isr Gesellschaft Fur Strahlen-Und Umweltforsch (GSF), Munich, Ger Ed Gedaliah Shelef, Carl J Soeder; 1980. [52] Shelef G, Sukenik A. Microalgae harvesting and processing: a literature review. Tech Res Dev Found Ltd 1984:65. http://dx.doi.org/10.2172/6204677. [53] Sawayama S, Minowa T, Yokoyama S-Y. Possibility of renewable energy production and CO 2 mitigation by thermochemical liquefaction of microalgae. Biomass Bioenergy 1999;17:33–9. [54] Wang L, Dandy DS. A microfluidic concentrator for cyanobacteria harvesting. Algal Res 2017;26:481–9. http://dx.doi.org/10.1016/j.algal.2017.03.018. [55] Borges L, Caldas S, Montes D’Oca MG, Abreu PC. Effect of harvesting processes on the lipid yield and fatty acid profile of the marine microalga Nannochloropsis oculata. Aquac Rep 2016;4:164–8. http://dx.doi.org/10.1016/j.aqrep.2016.10. 004. [56] Pragya N, Pandey KK, Sahoo PK. A review on harvesting, oil extraction and biofuels production technologies from microalgae. Renew Sustain Energy Rev 2013;24:159–71. http://dx.doi.org/10.1016/j.rser.2013.03.034. [57] SMAYDA TJ. The suspension and sinking of phytoplankton in the sea. Ocean Mar Biol Ann Rev 1970;8:353–414. [58] Edzwald JK. Algae, bubbles, coagulants, and dissolved air flotation. Water Sci Technol 1993;27:67–81. [59] Kromkamp J, Walsby AE. A computer model of buoyancy and vertical migration in cyanobacteria. J Plankton Res 1990;12:161–83. [60] Cole TM, Wells SA. CE-QUAL-W2: A two-dimensional, laterally averaged, hydrodynamic and water quality model, version 3.5; 2006. [61] Peperzak L, Colijn F, Koeman R, Gieskes WWC, Joordens JCA. Phytoplankton sinking rates in the Rhine region of freshwater influence. J Plankton Res 2003;25:365–83. [62] Choi SK, Lee JY, Kwon DY, Cho KJ. Settling characteristics of problem algae in the water treatment process. Water Sci Technol 2006;53:113–9. [63] Nurdogan Y, Oswald WJ. Tube settling of high-rate pond algae. Water Sci Technol 1996;33:229–41. [64] Show KY, Lee DJ. Algal biomass harvesting. Biofuels Algae Burlingt Elsevier 2014:85–110. [65] Shen Y, Yuan W, Pei ZJ, Wu Q, Mao E. Microalgae mass production methods. Trans ASABE 2009;52:1275–87. [66] Baker RW. Membrane technology and applications. Wiley Online Library; 2000. http://dx.doi.org/10.1002/0471238961.1305130202011105.a01. [67] Bilad MR, Arafat HA, Vankelecom IFJ. Membrane technology in microalgae cultivation and harvesting: a review. Biotechnol Adv 2014;32:1283–300. http://dx. doi.org/10.1016/j.biotechadv.2014.07.008. [68] Trivedi J, Aila M, Bangwal DP, Kaul S, Garg MO. Algae based biorefinery – how to make sense? Renew Sustain Energy Rev 2015;47:295–307. http://dx.doi.org/10. 1016/j.rser.2015.03.052. [69] Harun R, Singh M, Forde GM, Danquah MK. Bioprocess engineering of microalgae to produce a variety of consumer products. Renew Sustain Energy Rev 2010;14:1037–47. [70] Rawat I, Kumar RR, Mutanda T, Bux F. Dual role of microalgae: phycoremediation of domestic wastewater and biomass production for sustainable biofuels production. Appl Energy 2011;88:3411–24. [71] Mohn FH. Harvesting of micro-algal biomass. Micro-Algal Biotechnol 1988:395–414. [72] Van Den Hende S, Vervaeren H, Desmet S, Boon N. Bioflocculation of microalgae and bacteria combined with flue gas to improve sewage treatment. New Biotechnol 2011;29:23–31. [73] Purchas DB. Solid/liquid separation technology. Uplands Press; 1981. [74] Brennan JG, Butters JR, Cowell ND, Lilly AEV. Food engineering operations. Applied Science Publishers Ltd.; 1976. [75] Goh A. Production of microalgae using pig waste as a substrate. Berlin: Algal Biomass Technol Cramer; 1986. p. 235–44. [76] Petrusevski B, Bolier G, Van Breemen AN, Alaerts GJ. Tangential flow filtration: a method to concentrate freshwater algae. Water Res 1995;29:1419–24. [77] Borowitzka MA, Moheimani NR. Algae for biofuels and energy. Springer; 2013. [78] Mixson SM, Stikeleather LF, Simmons OD, Wilson CW, Burkholder JAM. pH-induced flocculation, indirect electrocoagulation, and hollow fiber filtration techniques for harvesting the saltwater microalga Dunaliella. J Appl Phycol 2014;26:1701–9. http://dx.doi.org/10.1007/s10811-013-0232-z. [79] Rossi N, Derouiniot-Chaplain M, Jaouen P, Legentilhomme P, Petit I. Arthrospira platensis harvesting with membranes: fouling phenomenon with limiting and critical flux. Bioresour Technol 2008;99:6162–7. [80] Rubio J, Souza ML, Smith RW. Overview of flotation as a wastewater treatment

technique. Miner Eng 2002;15:139–55. [81] Laamanen CA, Ross GM, Scott JA. Flotation harvesting of microalgae. Renew Sustain Energy Rev 2016;58:75–86. http://dx.doi.org/10.1016/j.rser.2015.12. 293. [82] Liu JC, Chen YM, Ju Y-H. Separation of algal cells from water by column flotation. Sep Sci Technol 1999;34:2259–72. [83] Greenwell HC, Laurens LML, Shields RJ, Lovitt RW, Flynn KJ. Placing microalgae on the biofuels priority list: a review of the technological challenges. J R Soc Interface 2009. [rsif20090322]. [84] Hoffmann JP. Wastewater treatment with suspended and nonsuspended algae. J Phycol 1998;34:757–63. [85] Laamanen CA, Scott JA. Development of heat-aided flocculation for flotation harvesting of microalgae. Biomass Bioenergy 2017;107:150–4. http://dx.doi.org/ 10.1016/j.biombioe.2017.09.020. [86] Hanotu J, Bandulasena HC, Zimmerman WB. Microflotation performance for algal separation. Biotechnol Bioeng 2012;109:1663–73. [87] Chen YM, Liu JC, Ju Y-H. Flotation removal of algae from water. Colloids Surf B Biointerfaces 1998;12:49–55. [88] Mollah MYA, Morkovsky P, Gomes JAG, Kesmez M, Parga J, Cocke DL. Fundamentals, present and future perspectives of electrocoagulation. J Hazard Mater 2004;114:199–210. http://dx.doi.org/10.1016/j.jhazmat.2004.08.009. [89] Coward T, Lee JGM, Caldwell GS. The effect of bubble size on the efficiency and economics of harvesting microalgae by foam flotation. J Appl Phycol 2015;27:733–42. http://dx.doi.org/10.1007/s10811-014-0384-5. [90] Coward T, Lee JGM, Caldwell GS. Harvesting microalgae by CTAB-aided foam flotation increases lipid recovery and improves fatty acid methyl ester characteristics. Biomass Bioenergy 2014;67:354–62. http://dx.doi.org/10.1016/j.biombioe. 2014.05.019. [91] Salim S, Bosma R, Vermuë MH, Wijffels RH. Harvesting of microalgae by bioflocculation. J Appl Phycol 2011;23:849–55. http://dx.doi.org/10.1007/s10811010-9591-x. [92] Vandamme D, Foubert I, Muylaert K. Flocculation as a low-cost method for harvesting microalgae for bulk biomass production. Trends Biotechnol 2013;31:233–9. http://dx.doi.org/10.1016/j.tibtech.2012.12.005. [93] Taher H, Al-Zuhair S, Al-Marzouqi AH, Haik Y, Farid MM. A review of enzymatic transesterification of microalgal oil-based biodiesel using supercritical technology. Enzym Res 2011;2011. [94] de la Noüe J, Laliberté G, Proulx D. Algae and waste water. J Appl Phycol 1992;4:247–54. [95] Papazi A, Makridis P, Divanach P. Harvesting Chlorella minutissima using cell coagulants. J Appl Phycol 2010;22:349–55. http://dx.doi.org/10.1007/s10811009-9465-2. [96] McGarry MG. Algal flocculation with aluminum sulfate and polyelectrolytes. J Water Pollut Control Fed 1970:R191–201. [97] Chalmers JJ, Zborowski M, Moore L, Mandal S, Fang B, Sun L. Theoretical analysis of cell separation based on cell surface marker density. Biotechnol Bioeng 1998;59:10–20. [98] Şirin S, Trobajo R, Ibanez C, Salvadó J. Harvesting the microalgae Phaeodactylum tricornutum with polyaluminum chloride, aluminium sulphate, chitosan and alkalinity-induced flocculation. J Appl Phycol 2012;24:1067–80. http://dx.doi.org/ 10.1007/s10811-011-9736-6. [99] de Godos I, Guzman HO, Soto R, García-Encina PA, Becares E, Muñoz R, et al. Coagulation/flocculation-based removal of algal-bacterial biomass from piggery wastewater treatment. Bioresour Technol 2011;102:923–7. http://dx.doi.org/10. 1016/j.biortech.2010.09.036. [100] Farid MS, Shariati A, Badakhshan A, Anvaripour B. Using nano-chitosan for harvesting microalga Nannochloropsis sp. Bioresour Technol 2013;131:555–9. http:// dx.doi.org/10.1016/j.biortech.2013.01.058. [101] Salama ES, Jeon BH, Kurade MB, Abou-Shanab RAI, Govindwar SP, Lee SH, et al. Harvesting of freshwater microalgae Scenedesmus obliquus and Chlorella vulgaris using acid mine drainage as a cost effective flocculant for biofuel production. Energy Convers Manag 2016;121:105–12. http://dx.doi.org/10.1016/j. enconman.2016.05.020. [102] Golueke CG, Oswald WJ. Harvesting and processing sewage-grown planktonic algae. J Water Pollut Control Fed 1965:471–98. [103] Maeda K, Kuramochi H, Shinkawa T, Fukui K. Solubility of two salts containing sulfate and chloride ions in water for ternary systems at 313 K. J Chem Eng Data 2002;47:1472–5. [104] Chatsungnoen T, Chisti Y. Harvesting microalgae by flocculation-sedimentation. Algal Res 2016;13:271–83. http://dx.doi.org/10.1016/j.algal.2015.12.009. [105] Vergini S, Aravantinou AF, Manariotis ID. Harvesting of freshwater and marine microalgae by common flocculants and magnetic microparticles. J Appl Phycol 2016;28:1041–9. http://dx.doi.org/10.1007/s10811-015-0662-x. [106] Sukenik A, Bilanovic D, Shelef G. Flocculation of microalgae in brackish and sea waters. Biomass 1988;15:187–99. [107] Jiang J-Q, Graham NJD, Harward C. Comparison of polyferric sulphate with other coagulants for the removal of algae and algae-derived organic matter. Water Sci Technol 1993;27:221–30. [108] Pirwitz K, Rihko-Struckmann L, Sundmacher K. Comparison of flocculation methods for harvesting Dunaliella. Bioresour Technol 2015;196:145–52. http:// dx.doi.org/10.1016/j.biortech.2015.07.032. [109] Hendricks D. Fundamentals of water treatment unit processes: physical, chemical, and biological. CRC Press; 2010. [110] Schenk PM, Thomas-Hall SR, Stephens E, Marx UC, Mussgnug JH, Posten C, et al. Second generation biofuels: high-efficiency microalgae for biodiesel production. Bioenergy Res 2008;1:20–43.

1118

Renewable and Sustainable Energy Reviews 91 (2018) 1103–1120

T. Mathimani, N. Mallick

[111] Rashid N, Ur Rehman MS, Sadiq M, Mahmood T, Han JI. Current status, issues and developments in microalgae derived biodiesel production. Renew Sustain Energy Rev 2014;40:760–78. http://dx.doi.org/10.1016/j.rser.2014.07.104. [112] Schlesinger A, Eisenstadt D, Bar-Gil A, Carmely H, Einbinder S, Gressel J. Inexpensive non-toxic flocculation of microalgae contradicts theories; overcoming a major hurdle to bulk algal production. Biotechnol Adv 2012;30:1023–30. [113] Harith ZT, Yusoff FM, Mohamed MS, Shariff M, Din M, Ariff AB. Effect of different flocculants on the flocculation performance of flocculation performance of microalgae, Chaetoceros calcitrans, cells. Afr J Biotechnol 2009:8. [114] Granados MR, Acién FG, Gomez C, Fernandez-Sevilla JM, Grima EM. Evaluation of flocculants for the recovery of freshwater microalgae. Bioresour Technol 2012;118:102–10. [115] Vandamme D, Foubert I, Meesschaert B, Muylaert K. Flocculation of microalgae using cationic starch. J Appl Phycol 2010;22:525–30. [116] Van Haver L, Nayar S. Polyelectrolyte flocculants in harvesting microalgal biomass for food and feed applications. Algal Res 2017;24:167–80. http://dx.doi.org/10. 1016/j.algal.2017.03.022. [117] Pushparaj B, Pelosi E, Torzillo G, Materassi R. Microbial biomass recovery using a synthetic cationic polymer. Bioresour Technol 1993;43:59–62. [118] Tilton RC, Murphy J, Dixon JK. The flocculation of algae with synthetic polymeric flocculants. Water Res 1972;6:155–64. [119] Selesu NFH, V de Oliveira T, Corrêa DO, Miyawaki B, Mariano AB, Vargas JVC, et al. Maximum microalgae biomass harvesting via flocculation in large scale photobioreactor cultivation. Can J Chem Eng 2016;94:304–9. [120] Morrissey KL, He C, Wong MH, Zhao X, Chapman RZ, Bender SL, et al. Chargetunable polymers as reversible and recyclable flocculants for the dewatering of microalgae. Biotechnol Bioeng 2015;112:74–83. [121] Mikulec J, Polakovičová G, Cvengroš J. Flocculation using polyacrylamide polymers for fresh microalgae. Chem Eng Technol 2015;38:595–601. [122] Uduman N, Qi Y, Danquah MK, Hoadley AFA. Marine microalgae flocculation and focused beam reflectance measurement. Chem Eng J 2010;162:935–40. http://dx. doi.org/10.1016/j.cej.2010.06.046. [123] Zheng H, Gao Z, Yin J, Tang X, Ji X, Huang H. Harvesting of microalgae by flocculation with poly (γ-glutamic acid). Bioresour Technol 2012;112:212–20. [124] Teixeira CMLL, Kirsten FV, Teixeira PCN. Evaluation of Moringa oleifera seed flour as a flocculating agent for potential biodiesel producer microalgae. J Appl Phycol 2012;24:557–63. [125] Mathimani T, Bhumathi D, Shan Ahamed T, Dineshbabu G, Deviram G, Uma L, et al. Semicontinuous outdoor cultivation and efficient harvesting of marine Chlorella vulgaris BDUG 91771 with minimum solid co-precipitation and high floc recovery for biodiesel. Energy Convers Manag 2017;149:13–25. [126] Gerchman Y, Vasker B, Tavasi M, Mishael Y, Kinel-Tahan Y, Yehoshua Y. Effective harvesting of microalgae: comparison of different polymeric flocculants. Bioresour Technol 2017;228:141–6. http://dx.doi.org/10.1016/j.biortech.2016.12.040. [127] Farooq W, Lee Y-C, Han J-I, Darpito CH, Choi M, Yang J-W. Efficient microalgae harvesting by organo-building blocks of nanoclays. Green Chem 2013;15:749. http://dx.doi.org/10.1039/c3gc36767c. [128] Wu J, Liu J, Lin L, Zhang C, Li A, Zhu Y, et al. Evaluation of several flocculants for flocculating microalgae. Bioresour Technol 2015;197:495–501. http://dx.doi.org/ 10.1016/j.biortech.2015.08.094. [129] Rakesh S, Saxena S, Dhar DW, Prasanna R, Saxena AK. Comparative evaluation of inorganic and organic amendments for their flocculation efficiency of selected microalgae. J Appl Phycol 2014;26:399–406. http://dx.doi.org/10.1007/s10811013-0114-4. [130] Zheng Y, Roberts M, Kelly J, Zhang N, Walker T. Harvesting microalgae using the temperature-activated phase transition of thermoresponsive polymers. Algal Res 2015;11:90–4. http://dx.doi.org/10.1016/j.algal.2015.06.006. [131] Abdul Hamid SH, Lananan F, Din WNS, Lam SS, Khatoon H, Endut A, et al. Harvesting microalgae, Chlorella sp. by bio-flocculation of Moringa oleifera seed derivatives from aquaculture wastewater phytoremediation. Int Biodeterior Biodegrad 2014;95:270–5. http://dx.doi.org/10.1016/j.ibiod.2014.06.021. [132] Borges L, Morón-Villarreyes JA, D’Oca MGM, Abreu PC. Effects of flocculants on lipid extraction and fatty acid composition of the microalgae Nannochloropsis oculata and Thalassiosira weissflogii. Biomass Bioenergy 2011;35:4449–54. http://dx.doi.org/10.1016/j.biombioe.2011.09.003. [133] Golueke Jr CG, Oswald WJ. Surface properties and ion exchange in algae removal. J Water Pollut Control Fed 1970:R304–14. [134] González-Fernández C, Ballesteros M. Microalgae autoflocculation: an alternative to high-energy consuming harvesting methods. J Appl Phycol 2013;25:991–9. [135] Horiuchi J-I, Ohba I, Tada K, Kobayashi M, Kanno T, Kishimoto M. Effective cell harvesting of the halotolerant microalga Dunaliella tertiolecta with pH control. J Biosci Bioeng 2003;95:412–5. [136] Sukenik A, Shelef G. Algal autoflocculation—verification and proposed mechanism. Biotechnol Bioeng 1984;26:142–7. [137] Lavoie A, De la Noüe J. Harvesting of Scenedesmus obliquus in wastewaters: auto‐or bioflocculation? Biotechnol Bioeng 1987;30:852–9. [138] Nurdogan Y, Oswald WJ. Enhanced nutrient removal in high-rate ponds. Water Sci Technol 1995;31:33–43. [139] Vandamme D, Foubert I, Fraeye I, Meesschaert B, Muylaert K. Flocculation of Chlorella vulgaris induced by high pH: role of magnesium and calcium and practical implications. Bioresour Technol 2012;105:114–9. http://dx.doi.org/10. 1016/j.biortech.2011.11.105. [140] Koley S, Prasad S, Bagchi SK, Mallick N. Development of a harvesting technique for large-scale microalgal harvesting for biodiesel production. RSC Adv 2017;7:7227–37. [141] Pérez L, Salgueiro JL, Maceiras R, Cancela Á, Sánchez Á. An effective method for

[142]

[143]

[144]

[145]

[146]

[147] [148]

[149]

[150]

[151]

[152]

[153]

[154]

[155]

[156] [157]

[158] [159] [160]

[161]

[162]

[163] [164]

[165]

[166]

[167] [168]

[169] [170]

[171]

1119

harvesting of marine microalgae: pH induced flocculation. Biomass Bioenergy 2017;97:20–6. http://dx.doi.org/10.1016/j.biombioe.2016.12.010. Knuckey RM, Brown MR, Robert R, Frampton DMF. Production of microalgal concentrates by flocculation and their assessment as aquaculture feeds. Aquac Eng 2006;35:300–13. http://dx.doi.org/10.1016/j.aquaeng.2006.04.001. Liu J, Zhu Y, Tao Y, Zhang Y, Li A, Li T, et al. Freshwater microalgae harvested via flocculation induced by pH decrease. Biotechnol Biofuels 2013;6:98. http://dx. doi.org/10.1186/1754-6834-6-98. Yang F, Xiang W, Fan J, Wu H, Li T, Long L. High pH-induced flocculation of marine Chlorella sp. for biofuel production. J Appl Phycol 2016;28:747–56. http://dx.doi.org/10.1007/s10811-015-0576-7. Vandamme D, Pohl PI, Beuckels A, Foubert I, Brady PV, Hewson JC, et al. Alkaline flocculation of Phaeodactylum tricornutum induced by brucite and calcite. Bioresour Technol 2015;196:656–61. http://dx.doi.org/10.1016/j.biortech.2015. 08.042. Kim D-G, La H-J, Ahn C-Y, Park Y-H, Oh H-M. Harvest of Scenedesmus sp. with bioflocculant and reuse of culture medium for subsequent high-density cultures. Bioresour Technol 2011;102:3163–8. Kurane R, Nohata Y. Microbial flocculation of waste liquids and oil emulsion by a bioflocculant from Alcaligenes latus. Agric Biol Chem 1991;55:1127–9. Frølund B, Palmgren R, Keiding K, Nielsen PH. Extraction of extracellular polymers from activated sludge using a cation exchange resin. Water Res 1996;30:1749–58. Lee AK, Lewis DM, Ashman PJ. Microbial flocculation, a potentially low-cost harvesting technique for marine microalgae for the production of biodiesel. J Appl Phycol 2009;21:559–67. http://dx.doi.org/10.1007/s10811-008-9391-8. Salim S, Vermuë MH, Wijffels RH. Ratio between autoflocculating and target microalgae affects the energy-efficient harvesting by bio-flocculation. Bioresour Technol 2012;118:49–55. Vandamme D, Foubert I, Fraeye I, Muylaert K. Influence of organic matter generated by chlorella vulgaris on five different modes of flocculation. Bioresour Technol 2012;124:508–11. http://dx.doi.org/10.1016/j.biortech.2012.08.121. Hu Y-R, Wang F, Wang S-K, Liu C-Z, Guo C. Efficient harvesting of marine microalgae Nannochloropsis maritima using magnetic nanoparticles. Bioresour Technol 2013;138:387–90. Zhou W, Min M, Hu B, Ma X, Liu Y, Wang Q, et al. Filamentous fungi assisted bioflocculation: a novel alternative technique for harvesting heterotrophic and autotrophic microalgal cells. Sep Purif Technol 2013;107:158–65. Ndikubwimana T, Zeng X, Murwanashyaka T, Manirafasha E, He N, Shao W, et al. Harvesting of freshwater microalgae with microbial bioflocculant: a pilot-scale study. Biotechnol Biofuels 2016;9:47. http://dx.doi.org/10.1186/s13068-0160458-5. Oh H-M, Lee SJ, Park M-H, Kim H-S, Kim H-C, Yoon J-H, et al. Harvesting of Chlorella vulgaris using a bioflocculant from Paenibacillus sp. AM49. Biotechnol Lett 2001;23:1229–34. Gultom SO, Hu B. Review of microalgae harvesting via co-pelletization with filamentous fungus. Energies 2013;6:5921–39. Taylor RL, Rand JD, Caldwell GS. Treatment with algae extracts promotes flocculation, and enhances growth and neutral lipid content in Nannochloropsis oculata—a candidate for biofuel production. Mar Biotechnol 2012;14:774–81. Yokoi H, Yoshida T, Mori S, Hirose J, Hayashi S, Takasaki Y. Biopolymer flocculant produced by an Enterobacter sp. Biotechnol Lett 1997;19:569–73. Salehizadeh H, Vossoughi M, Alemzadeh I. Some investigations on bioflocculant producing bacteria. Biochem Eng J 2000;5:39–44. Richardson JW, Johnson MD, Lacey R, Oyler J, Capareda S. Harvesting and extraction technology contributions to algae biofuels economic viability. Algal Res 2014;5:70–8. http://dx.doi.org/10.1016/j.algal.2014.05.007. Poelman E, De Pauw N, Jeurissen B. Potential of electrolytic flocculation for recovery of micro-algae. Resour Conserv Recycl 1997;19:1–10. http://dx.doi.org/ 10.1016/S0921-3449(96)01156-1. Sasson M Ben, Calmano W, Adin A. Iron-oxidation processes in an electroflocculation (electrocoagulation) cell. J Hazard Mater 2009;171:704–9. http://dx. doi.org/10.1016/j.jhazmat.2009.06.057. Zenouzi A, Ghobadian B, Hejazi MA, Rahnemoon P. Harvesting of microalgae Dunaliella salina using electroflocculation. J Agric Sci Technol 2013;15:879–87. Bleeke F, Quante G, Winckelmann D, Klöck G. Effect of voltage and electrode material on electroflocculation of Scenedesmus acuminatus. Bioresour Bioprocess 2015;2:36. http://dx.doi.org/10.1186/s40643-015-0064-6. Vandamme D, Pontes SCV, Goiris K, Foubert I, Pinoy LJJ, Muylaert K. Evaluation of electro-coagulation-flocculation for harvesting marine and freshwater microalgae. Biotechnol Bioeng 2011;108:2320–9. http://dx.doi.org/10.1002/bit.23199. González-Fernández C, Ballesteros M. Microalgae autoflocculation: an alternative to high-energy consuming harvesting methods. J Appl Phycol 2013;25:991–9. http://dx.doi.org/10.1007/s10811-012-9957-3. Gao S, Yang J, Tian J, Ma F, Tu G, Du M. Electro-coagulation–flotation process for algae removal. J Hazard Mater 2010;177:336–43. Azarian GH, Mesdaghinia AR, Vaezi F, Nabizadeh R, Nematollahi D. Algae removal by electro-coagulation process, application for treatment of the effluent from an industrial wastewater treatment plant. Iran J Public Health 2007;36:57–64. Kumar HD, Yadava PK, Gaur JP. Electrical flocculation of the unicellular green alga Chlorella vulgaris Beijerinck. Aquat Bot 1981;11:187–95. Xu L, Wang F, Li H, Hu Z, Guo C, Liu C. Development of an efficient electroflocculation technology integrated with dispersed‐air flotation for harvesting microalgae. J Chem Technol Biotechnol 2010;85:1504–7. Gao S, Du M, Tian J, Yang J, Yang J, Ma F, et al. Effects of chloride ions on electro-

Renewable and Sustainable Energy Reviews 91 (2018) 1103–1120

T. Mathimani, N. Mallick

[172]

[173]

[174]

[175]

[176]

[177]

[178]

[179]

[180]

[181]

06.012. [182] Prochazkova G, Safarik I, Branyik T. Harvesting microalgae with microwave synthesized magnetic microparticles. Bioresour Technol 2013;130:472–7. [183] Franzreb M, Siemann-Herzberg M, Hobley TJ, Thomas ORT. Protein purification using magnetic adsorbent particles. Appl Microbiol Biotechnol 2006;70:505–16. [184] Bitton G, Fox JL, Strickland HG. Removal of algae from Florida lakes by magnetic filtration. Appl Microbiol 1975;30:905–8. [185] Xu L, Guo C, Wang F, Zheng S, Liu C-Z. A simple and rapid harvesting method for microalgae by in situ magnetic separation. Bioresour Technol 2011;102:10047–51. [186] Kroell M, Pridoehl M, Zimmermann G, Pop L, Odenbach S, Hartwig A. Magnetic and rheological characterization of novel ferrofluids. J Magn Magn Mater 2005;289:21–4. [187] Hu Y-R, Guo C, Wang F, Wang S-K, Pan F, Liu C-Z. Improvement of microalgae harvesting by magnetic nanocomposites coated with polyethylenimine. Chem Eng J 2014;242:341–7. [188] Wang T, Yang W-L, Hong Y, Hou Y-L. Magnetic nanoparticles grafted with aminoriched dendrimer as magnetic flocculant for efficient harvesting of oleaginous microalgae. Chem Eng J 2016;297:304–14. [189] Lee K, Na J-G, Seo JY, Shim TS, Kim B, Praveenkumar R, et al. Magnetic-nanoflocculant-assisted water–nonpolar solvent interface sieve for microalgae harvesting. ACS Appl Mater Interfaces 2015;7:18336–43. [190] Ge S, Agbakpe M, Wu Z, Kuang L, Zhang W, Wang X. Influences of surface coating, UV irradiation and magnetic field on the algae removal using magnetite nanoparticles. Environ Sci Technol 2015;49:1190–6. [191] Liu P, Wang T, Yang Z, Hong Y, Hou Y. Long-chain poly-arginine functionalized porous Fe 3 O 4 microspheres as magnetic fl occulant for e ffi cient harvesting of oleaginous microalgae. Algal Res 2017;27:99–108. http://dx.doi.org/10.1016/j. algal.2017.08.025. [192] Hena S, Fatihah N, Tabassum S, Lalung J, Jing SY. Magnetophoretic harvesting of freshwater microalgae using polypyrrole/Fe3O4 nanocomposite and its reusability. J Appl Phycol 2016;28:1597–609. http://dx.doi.org/10.1007/s10811-0150719-x.

coagulation-flotation process with aluminum electrodes for algae removal. J Hazard Mater 2010;182:827–34. Uduman N, Bourniquel V, Danquah MK, Hoadley AFA. A parametric study of electrocoagulation as a recovery process of marine microalgae for biodiesel production. Chem Eng J 2011;174:249–57. Cerff M, Morweiser M, Dillschneider R, Michel A, Menzel K, Posten C. Harvesting fresh water and marine algae by magnetic separation: screening of separation parameters and high gradient magnetic filtration. Bioresour Technol 2012;118:289–95. Wang S-K, Wang F, Hu Y-R, Stiles AR, Guo C, Liu C-Z. Magnetic flocculant for high efficiency harvesting of microalgal cells. ACS Appl Mater Interfaces 2013;6:109–15. Fayad N, Yehya T, Audonnet F, Vial C. Harvesting of microalgae Chlorella vulgaris using electro-coagulation-flocculation in the batch mode. Algal Res 2017;25:1–11. http://dx.doi.org/10.1016/j.algal.2017.03.015. Boli E, Savvidou M, Logothetis D, Louli V, Pappa G, Voutsas E, et al. Magnetic harvesting of marine algae Nannochloropsis oceanica. Sep Sci Technol 2017;6395:1–8. http://dx.doi.org/10.1080/01496395.2017.1296463. Wang SK, Stiles AR, Guo C, Liu CZ. Harvesting microalgae by magnetic separation: a review. Algal Res 2015;9:178–85. http://dx.doi.org/10.1016/j.algal.2015.03. 005. Lee AK, Lewis DM, Ashman PJ. Harvesting of marine microalgae by electroflocculation: the energetics, plant design, and economics. Appl Energy 2013;108:45–53. http://dx.doi.org/10.1016/j.apenergy.2013.03.003. Guldhe A, Misra R, Singh P, Rawat I, Bux F. An innovative electrochemical process to alleviate the challenges for harvesting of small size microalgae by using nonsacrificial carbon electrodes. Algal Res 2016;19:292–8. Xiong Q, Pang Q, Pan X, Chika AO, Wang L, Shi J, et al. Facile sand enhanced electro-flocculation for cost-efficient harvesting of dunaliella salina. Bioresour Technol 2015;187:326–30. http://dx.doi.org/10.1016/j.biortech.2015.03.135. Ryu B-G, Kim J, Han J-I, Kim K, Kim D, Seo B-K, et al. Evaluation of an electroflotation-oxidation process for harvesting bio-flocculated algal biomass and simultaneous treatment of residual pollutants in coke wastewater following an algal-bacterial process. Algal Res 2017. http://dx.doi.org/10.1016/j.algal.2017.

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