Effect of Mechanical Pretreatment on Hydrocarbon Extraction from

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Effect of mechanical pretreatment on hydrocarbon extraction from concentrated wet hydrocarbon-rich microalga, Botryococcus braunii Shun Tsutsumi, Yasuhiro Saito, Yohsuke Matsushita, and Hideyuki Aoki Energy Fuels, Just Accepted Manuscript • DOI: 10.1021/acs.energyfuels.7b02753 • Publication Date (Web): 05 Jan 2018 Downloaded from http://pubs.acs.org on January 5, 2018

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Effect of mechanical pretreatment on hydrocarbon extraction from concentrated wet hydrocarbon-rich microalga, Botryococcus braunii Shun Tsutsumi*, Yasuhiro Saito, Yohsuke Matsushita, Hideyuki Aoki Department of Chemical Engineering, Graduate School of Engineering, Tohoku University, Sendai, Miyagi, Japan *Corresponding author

KEYWORDS Botryococcus braunii; Hydrocarbon extraction; Mechanical pretreatment; Membrane filtration; Intracellular substance

ABSTRACT In the present study, we investigated the effects of concentrating a cell suspension of a hydrocarbon-rich microalga, Botryococcus braunii, by membrane filtration on hydrocarbon recovery efficiency. B. braunii suspensions before and after filtration were mechanically pretreated with a high-pressure homogenizer or a JET PASTER high-speed mixer. Concentrating the suspension increased the apparent viscosity of the sample and altered the particle size and shape distributions. The increase of viscosity was derived from the existence probability that was caused by shear forces in the pump that introduced the suspension into the membrane filter and/or between the membrane wall and

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algae. Homogenizer treatment decreased the sample viscosity from 20 to 5 mPa⋅s due to the collapse of the bridge between the algae and polysaccharides. The treatment also decreased the hydrocarbon recovery efficiency from 60% to 15% because of the release of intracellular substances that prevented hydrocarbon extraction. In contrast, JET PASTER treatment increased the hydrocarbon recovery efficiency by removing polysaccharides surrounding the colonies and disrupting colonies, without disrupting the cells. Adding oleic acid as a model intracellular substance to the concentrated sample before extraction decreased the amount of extracted hydrocarbon. These results demonstrate that concentrating the sample by filtration combined with JET PASTER treatment can improve the hydrocarbon recovery efficiency of B. braunii. In addition, an energy analysis was performed in the present study. The energy consumption of the JET PASTER treatment combined with filtration was 7.6 times as high as the energy produced.

INTRODUCTION In recent years, there has been increasing interest in the utilization of biofuels, especially in liquid form, due to concerns regarding fossil fuel depletion, the rapid global growth of the transport sector, and provisions for global warming.1-3 Microalgae contain high levels (20–50 wt% in general) of lipids, mainly triacylglycerols, that can be used as biofuels1, 4 and do not compete with food demand, which is a problem with materials currently used as biofuels. Moreover, microalgae have a high CO2

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fixation capacity as compared to conventional energy crops, and thus hold promise as next-generation biofuels.2, 3, 5, 6 In general, the conversion of microalgae into biofuels such as biodiesel involves cultivation, harvesting, and extraction. Organic solvents such as n-hexane are frequently used for lipid extraction.2, 5, 7-10

This requires drying of the microalgal suspension, which consumes 80–90% of the total energy

required for the entire conversion process.11-14 As such, the amount of energy needed for biofuel production is greater than that produced.15 Hydrocarbon or lipid extraction from wet microalgae has been proposed to overcome this problem.6-12, 15-35 Because common microalgae store lipids inside their cells, disrupting the cell wall prior to the extraction of lipids or other intracellular substances can increase the efficiency of this process. To this end, various methods such as application of mechanical force,7, 10, 11, 16-24 novel extraction solvents,20, 26, 30, 31 and high temperature and water pressure have been investigated.27, 32-35 However, because the influence of pretreatment methods on the amount of extractable lipid varies due to several factors such as algal species,16, 18, 36, 37 organic solvent used for extraction,18, 20, 38 sample preparation method,20, 38 and moisture content (i.e., dry cell weight),38 it is difficult to determine the best pretreatment methods for microalgal lipid extraction. Thus, the accumulation of knowledge regarding several pretreatment methods performed under the several algal conditions is necessary for future process design. The green microalga Botryococcus braunii is approximately 10 µm in size and forms colonies that are tens of microns in size.21, 39, 40 B. braunii has 3 races, namely race A, race B, and race L.39, 41 In

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particular, race B algae produce the largest amount of C30–C37 polymethylated, unsaturated triterpenes (27–86% dry cell weight),34, 39, 42 over 90% of which are stored within the colony matrix, in contrast to other algae.20, 43 Therefore, extracting hydrocarbons using an organic solvent should be straightforward even under wet conditions. However, when employing widely used organic solvents such as n-hexane,5 this has not been possible without pretreatment16,

32

due to an amphiphilic

polysaccharide envelope around the colonies that blocks solvent penetration.33,

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Kanda et al.31

showed that using dimethyl ether did not require any pretreatment. Despite this advantage, their method requires relatively intensive dewatering (up to 74 wt%), and dimethyl ether extracts higher amounts of polar components, which can be contaminants in biofuel production, than n-hexane does. In contrast, n-hexane extraction shows selectivity towards neutral lipids or hydrocarbons and avoids co-extraction of the polar lipid fractions.10, 22 Thus, extraction with an organic solvent such as n-hexane would be effective for selectively extracting hydrocarbons from B. braunii. Dispersal and removal of these polysaccharides by heating the cell suspension33 or disrupting colonies by intense agitation in the presence of beads21 was shown to increase the amount of extractable hydrocarbon, presumably because removing the polysaccharides allowed the organic solvent to access the location in the colony where hydrocarbons were stored. In general, microalgae cultures are dilute suspensions44,

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; the cell suspension must be

concentrated by aggregation, membrane filtration, or centrifugation to minimize the energy required to extract hydrocarbons using an organic solvent. B. braunii secretes mucilaginous polysaccharides

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around the colony or in cell suspension,46 thereby increasing the sample viscosity and altering the flow properties, which can in turn affect the amount of hydrocarbon extractable by wet extraction. Membrane filtration is one of the most common methods for concentrating microalgae, and is an energy-saving and relatively simple procedure.47, 48 Filtration using a cross-flow (e.g., hollow fiber) membrane prevents fouling of the membrane surface. As the sample flows tangentially along the membrane wall, a shear force is generated that prevents sample deposition on the wall.49 This force also develops between the colony surface and membrane wall during cross-flow filtration. In membrane filtration, a pump is used to circulate and concentrate the cell culture; however, shear stress in the pump causes damage to microbial cells.50 Thus, concentrating B. braunii by membrane filtration can potentially disrupt cells and colonies, and remove surrounding polysaccharides to increase the amount of extractable hydrocarbon. In earlier studies, microalgae other than B. braunii were concentrated by flocculation or centrifugation, and their lipids were extracted.22, 23 However, the concentration of B. braunii was achieved by simple membrane filtration without a pump31, 33 or by resuspending lyophilized algal cells,16 which is impractical and consumes a large amount of energy. For other species such as Chlorella sp., the amount of extractable lipids from wet microalgae increased according to the degree of cell disruption.10 While B. braunii stores almost all hydrocarbons in the colony matrix, cell disruption seems to be effective for this alga, similar to other microalgae. Experimentally, disruption of B. braunii cells increased the amount of extractable hydrocarbon for non-filtered cell suspensions.16 However, no previous reports have described the

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influence of filtration on cell disruption and the amount of extractable hydrocarbon from B. braunii. We previously reported an improvement in the amount of extractable hydrocarbon by mechanical pretreatments using unconcentrated B. braunii suspensions24, but did not examine the effect of mechanical pretreatments on the concentration of algal suspensions. To address this issue, in the present study, cell suspensions of B. braunii with or without concentration by membrane filtration were mechanically treated, and their hydrocarbons were extracted using n-hexane. A high-pressure homogenizer and a JET PASTER high-speed mixer were used to apply external forces to the microalga. The cell size and shape were measured to evaluate the effects of the concentration process and/or application of external forces on B. braunii cells and/or colonies. We also analyzed the effect of the concentration process and/or mechanical pretreatments on the polysaccharides surrounding colonies by negative staining using India ink.

MATERIALS AND METHODS Microalgae. B. braunii BOT-22 (race B)51-53 was used in the present study. The algal sample was provided and cultivated at the Algae Biomass and Energy System R&D Center (University of Tsukuba). Details of the culture medium and cultivation method have been previous reported.53 Dry cell weight was measured by filtering a cell suspension through pre-weighed glass fiber filters and drying at 30°C under atmospheric pressure for 24 h. The dry cell weight was determined as approximately 0.72 g/L.

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Filtration and mechanical treatments. The B. braunii cell suspension was passed through a ceramic cross-flow membrane filter (Noritake Co., Tokyo, Japan) and concentrated before mechanical treatment. The nominal pore size and membrane area were 0.8 µm and 0.24 m2, respectively. Twenty-four filters were used in parallel. The membrane inlet pressure was set at 0.2 MPa of absolute pressure for filtration. The initial flow rate in the filter was set to 530 ± 15 L/min, and the initial linear velocity in the filter was 1.5 ± 0.04 m/s. Algae deposited on the membrane were cleaned by backwashing the membrane for 1 min at 4-min intervals. An algal suspension of approximately 2000 L was introduced in the filter used by the centrifugal pump (SE76-150, rated power outlet: 15 kW; OSAKA SANITARY Co., Ltd, Japan). The dry cell weight of the filtered sample was approximately 6.82 g/L, which was determined by the same procedure used for the unfiltered sample. The apparent viscosity of 1 ml of sample before and after filtration was measured at 25ºC with a rotatory viscometer (TV-20; Toki Sangyo Co., Tokyo, Japan) circulating in water at 100 rpm. Cells and colonies were mechanically treated and polysaccharides were removed using a high-pressure homogenizer (LAB2000, rated power output: 3 kW; APV, Delavan, WI, USA) or a JET PASTER high-speed mixer (JP-SS, rated power output: 5.5 kW; Nihon Spindle Manufacturing Co., Amagasaki, Japan). The high-pressure homogenizer, which was of the Manton–Gaulin type, uses a tubular flow system that can disrupt or disperse materials in a fluid by compressing the liquid; it produces shear force, impingement of cells on the channel wall in the device, and cavitation

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impulsive force.54 The sample was subjected to 30 MPa of pressure in the high-pressure homogenizer. The number of passes through the homogenizer was one. The JET PASTER comprised a casing surrounding a rotating impeller (located similarly to a centrifugal pump) and a closed-loop circulation pipe. The casing outlet was connected to the casing inlet via the circulation pipe. A schematic representation of the JET PASTER is shown in Figure 1. The impulsive force caused by hydrodynamic cavitation and the shear force caused by rotation of the impeller drew the microalgae into the device.24 The rotational speed of the internal impeller was 4800 rpm, and algal samples were treated for 3 min. Approximately 900 mL of algal sample was introduced into the device, and 200 mL of treated sample was used for analyses. A chiller (MTC-3000; AS-ONE, Osaka, Japan) was used to maintain the temperature in the sample chamber at 20°C.

Determination of particle size and shape distributions. To investigate the effects of membrane filtration and mechanical treatments on the size and shape of microalga, particle size and shape distributions were determined using a fully automated dynamic flow particle-imaging instrument (Sysmex FPIA-3000; Malvern Instruments, Malvern, UK). The number of particles was 10,000; measurements were performed in triplicate. The high particle concentration after filtration made the measurement of particle properties difficult. Hence, the filtered samples were diluted by approximately three fold before the particle size measurement.

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Negative staining with India ink. To visualize the structures surrounding microalgae, cell suspensions were negatively stained with India ink after each treatment. Soot particles in India ink cannot penetrate a fibrillar sheath with a cross-linked structure, thereby enabling examination of the microalgae surface.40 A 5-µL volume of sample and India ink was placed on a glass slide and covered with a coverslip. Samples were observed with an optical microscope connected to a digital camera (IX73PI-33FL/DIC-TR and DP73-SET-C-2; Olympus, Tokyo, Japan).

Hydrocarbon extraction. To extract hydrocarbons from the samples, 20 mL of algal sample and 50 mL of n-hexane were introduced into separation funnels, and the mixture was shaken at 318 rpm for 3 min on a universal shaker (AW-2; AS-ONE). Twenty milliliters of the organic phase were collected from the upper layer and used as the extracted sample from B. braunii. The extracted hydrocarbon was quantified using a gas chromatography-flame ionization detector (GC-FID,

Shimadzu GC-2014) with a UA-1 (MS/HT)-30M-0.25F column (Frontier Laboratories Ltd., Japan). The GC column oven program was as follows: the column temperature was raised from 130 to 270°C at 20°C/min, and then from 270 to 300°C at 2°C/min, after which it was held for 8 min. The injection temperature was set at 320°C, and the detector temperature was set at 350°C. The linear velocity of the carrier gas (He) was set at 51.9 cm/s. n-Triacontane (C30, Purity: 99%, GL Sciences Inc., Japan) was used as an internal standard. The hydrocarbons detected by GC-FID using this column were reported as highly purified botryococcenes (C34H58, >95% purity).52 Thus, all of the

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hydrocarbons detected and quantified in the present study were defined as botryococcenes with a molecular formula of C34H58. To obtain the maximum hydrocarbon recovery efficiency, the B. braunii cell suspension was dried in the same manner as that for the measurement of dry cell weight. A 50-mL volume of n-hexane was used as the extractant, and the mixture consisting of the dried sample and extractant was shaken for 3 h. The hydrocarbon concentration was calculated from the amount of hydrocarbon in the recovered n-hexane layer and the concentration of hydrocarbon recovered from wet samples was normalized to that of hydrocarbon recovered from the dried sample, according to the following equation: % =

mg/mL × 100 … (1) mg/mL

where RHC is the hydrocarbon recovery efficiency, and CHC-wet sample and CHC-dried sample [mg/mL] are the hydrocarbon concentrations in the recovered n-hexane layers of the wet and dried samples, respectively. The hydrocarbon recovery efficiency of the dried sample was 10–11% of the dry cell weight. The extraction was performed in triplicate for each sample condition, and the average value was reported as the hydrocarbon recovery efficiency.

Energy analysis. To investigate the efficiency of the methods used in this study, an energy analysis was performed and compared with that in a reported in a previous study32, 33, 55, 56 that used a thermal heating method and a method based on drying of the algal suspension. The analytical conditions are summarized in Figure 2. The JET PASTER treatment was considered as the

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mechanical pretreatment. The energy for filtration and the JET PASTER treatment were measured with an electrical energy meter (POWER ANALYZER 3390, HIOKI E. E Co., Japan). Among the n-hexane-extraction methods, thermal pretreatment methods performed by Kita et al.32, Magota et al.55, Saga et al.56, and Atobe et al.33, in which B. braunii culture broth was heated to 90°C and held at that temperature for 20 min, were regarded as comparative pretreatment methods because of their simplicity and high extraction yields. The authors of these studies reported that almost all hydrocarbons were extracted via their methods. The consumption energy for the thermal heating method was assumed to be the heating energy of water from 20°C to 90°C, using the heat capacity of water at 20°C, and the extractable hydrocarbon was assumed to 90% of the hydrocarbon content. The drying energy was the vaporization energy of water calculated using the latent heat of water at 20°C, and extractable hydrocarbon was assumed to 100% of the hydrocarbon content. The energies for filtration and pretreatment were considered, although other processes such as cultivation and the hydrocarbon-extraction process were not considered in order to focus on the influence of pretreatment methods on the energy balance. The production energy, higher heating value of oil, and extracted residue were calculated using the Dulong' formula 57: Q = 33.83C + 144.2H − 18.03O where Q is higher heating value [MJ], and C, H, and O are the mass fraction of carbon, hydrogen, and oxygen, respectively. The oil structure was assumed to be C34H58, and the ultimate composition of the extracted residue was used based on the study by Watanabe et al.58. The extracted residues

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shown in Figure 2 contained unextracted hydrocarbons and algal components. In addition, the energy ratio between the extracted hydrocarbon energy and consumption energy was evaluated. The energy ratio was defined as follows: Energy ratio % =

Extracted hydrocarbon energy MJ × 100. Consumption energy MJ

RESULTS AND DISCUSSION Microalgae filtration. The B. braunii cell suspension was concentrated by membrane filtration. The apparent viscosity of samples before and after concentration is shown in Figure 3. The concentration process increased the apparent viscosity of the samples irrespective of mechanical treatments, likely due to the condensation of polysaccharides secreted by B. braunii during filtration, resulting in artificial cell aggregation. Treatment of filtered samples with the high-pressure homogenizer significantly decreased the viscosity as compared to filtered samples without mechanical treatment. This suggests that the bridges between polysaccharides and algae were disrupted, thereby increasing sample fluidity. When the concentrated sample was treated with the JET PASTER, there was no decrease in apparent viscosity. This suggests that the JET PASTER treatment did not generate external forces that can destroy the polysaccharide bridges connecting cells and colonies.

Cell and colony size and shape distributions. Samples were treated with the high-pressure homogenizer or JET PASTER before and after membrane filtration, and the size and shape of B.

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braunii cells, i.e., the circle-equivalent diameter and roundness of algae, were examined (Figs. 4 and 5). Particles in the unfiltered sample without mechanical treatments had a diameter of 5–90 µm and a roundness value of 0.4–1 (Fig. 4 (a-1)). Colonies formed by agglomeration of single cells contained particles with various sizes and shapes. In the concentrated sample without mechanical treatment, most particles had a size of 10–20 µm, whereas a small fraction had a size greater than 20 µm, in contrast to the filtered sample without mechanical treatment (Fig. 4 (a-2)). Furthermore, the roundness distribution of concentrated algal samples without mechanical treatment showed a sharper peak than that of the unfiltered sample without mechanical treatment. This is because the shear force generated in the pump and/or between the membrane and cells broke up the colonies into smaller ones during membrane filtration. Treatment of non-concentrated samples with the high-pressure homogenizer decreased the percentage of particles with a size of 10 µm, while increasing the fraction of particles with a size less than 5 µm (Fig. 4 (b-1)). High-pressure homogenizer treatment destroyed B. braunii cells and colonies, leaving cellular debris. The concentrated sample treated with the high-pressure homogenizer had more particles smaller than 5 µm and fewer that were larger than 10 µm, as compared to the non-concentrated sample (Fig. 4 (b-2)). This was due to its higher apparent viscosity, which would tend to reinforce the shear force on microalgae. Additionally, the roundness distribution was broader after the concentration process (Fig. 5 (b-1), (b-2)), likely due to greater disruption by the homogenizer

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of the concentrated sample as compared to the non-concentrated sample, resulting in particles with a range of roundness values. JET PASTER treatment of B. braunii increased the percentage of particles with a diameter of 10 µm, while decreasing the fraction of particles larger than 20 µm (Fig. 4 (c-1)). In contrast to high-pressure homogenization, there was no increase in the percentage of 5-µm particles, indicating that the JET PASTER perturbed B. braunii colonies without rupturing the cells. Thus, JET PASTER treatment had a weaker effect on microalgae than the high-pressure homogenizer, in agreement with our previous observations.24 The particle and roundness distributions of concentrated and non-concentrated samples treated with the JET PASTER were similar (Fig. 5 (c-1), (c-2)); thus, the increase in viscosity due to the concentration of the B. braunii cell suspension likely did not contribute to the effects of JET PASTER treatment.

Observation of extracellular polysaccharides around B. braunii colonies. Samples before and after treatment were examined by negative staining using India ink (Fig. 6). B. braunii colonies were observed within the unstained parts of the unfiltered sample without mechanical treatments (Fig. 6 (a-1)). These white-colored parts were the fibrillar polysaccharides or fibrillar sheath reported in previous studies.42 Polysaccharides surrounding the colonies prevent the penetration of soot particles in Indian ink, leaving the colonies unstained. It is also known that polysaccharides trap moisture59 and thus exclude n-hexane, resulting in a low hydrocarbon recovery efficiency from unfiltered samples

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without mechanical treatments. The polysaccharide area decreased after filtration, likely because the shear force between the membrane wall and algal surface and/or that in the pump used to deposit the sample onto the membrane removed polysaccharides from the colony surface. In samples mechanically treated with the homogenizer, no polysaccharides were observed around B. braunii, irrespective of membrane filtration (Fig. 6 (b-1), (b-2)), in contrast to the non-homogenized samples. Polysaccharides in the sample were removed by the external force originating from the homogenizer. Moreover, because polysaccharide removal did not significantly differ between samples with and without filtration, the effect of polysaccharide removal was likely not influenced by sample concentration. Examination of negatively stained images of samples treated with the JET PASTER revealed that polysaccharides were partly or almost completely removed from the colonies, in contrast to membrane-filtered samples without mechanical treatments (Fig. 6 (c-1), (c-2)). External forces used in the JET PASTER treatment that disrupted the colonies of B. braunii included the shear force caused by the rotation of the impeller and the impulsive force derived from cavitation caused by the local pressure depression in the device. Little difference was observed between micrographs before and after membrane filtration. Thus, sample concentration did not influence the removal of polysaccharides with the JET PASTER treatment, as was the case with high-pressure homogenization.

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Hydrocarbon extraction. Hydrocarbon was extracted from samples using n-hexane. The hydrocarbon recovery efficiency of the unfiltered sample without mechanical treatments was very low (approximately 10%) (Fig. 7), likely because polysaccharides surrounding the B. braunii colonies prevented the penetration of n-hexane. Concentrating the cell suspension greatly increased the hydrocarbon recovery efficiency; membrane filtration disrupted the colonies and removed the polysaccharides through the shear force between the membrane wall and colonies and/or in the pump used to deposit the cell suspension onto the membrane filter, thereby increasing access by n-hexane to the hydrocarbon in the colonies. The hydrocarbon recovery efficiency of the non-concentrated sample was higher with homogenization as compared to without homogenizer treatment. This is because disrupting the colonies and cells enhanced the mixing of n-hexane and hydrocarbons in B. braunii. Mechanical treatment of the concentrated sample reduced the hydrocarbon recovery efficiency relative to the non-concentrated sample with the same treatment, down to a value similar to that of the unfiltered sample without mechanical treatment. We speculate that homogenization released intracellular substances such as fatty acids into n-hexane, preventing the dissolution of hydrocarbons into the solvent and thereby decreasing their hydrocarbon recovery efficiency. Irrespective of concentration, the hydrocarbon recovery efficiency of samples treated with the JET PASTER was much higher than that of samples without mechanical treatments; that is, the particle size and roundness distributions of the concentrated sample treated with the JET PASTER

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were similar to those of the filtered sample without mechanical treatment. In contrast, their hydrocarbon recovery efficiencies differed markedly. This is because colony disruption and polysaccharide removal were achieved by the JET PASTER, in addition to the effects of shear force caused by membrane filtration. Furthermore, although JET PASTER treatment of the condensed sample decreased the hydrocarbon recovery efficiency, the degree of decrease was small compared to that in the condensed sample treated by the homogenizer. According to the particle size distributions, the cells themselves were not disrupted by the JET PASTER, and therefore hydrocarbon extraction would not be prevented, in contrast to the case with homogenization. Homogenizer treatment of the filtered sample resulted in a greater decrease in the hydrocarbon recovery efficiency than identical treatment of the non-concentrated sample. As stated above, it is possible that homogenization released hydrophobic intracellular substances that prevented hydrocarbon extraction. To test this hypothesis, we examined the effect of hydrophobic intracellular substances on hydrocarbon recovery efficiency from concentrated samples. Oleic acid, which is one of the main fatty acids in B. braunii race B60, 61 and a precursor of the biopolymer used to synthesize the colony matrix62, 63, was used as a model hydrophobic intracellular compound. The concentrated sample filtered under the same conditions was used (dry cell weight ~3 g/L). The hydrocarbon recovery efficiency of the dried sample was 26% of the dry cell weight. The model compound was introduced into algal samples for hydrocarbon extraction by several methods (Fig. 8). Under Condition 1, the filtered sample (20 ml) and n-hexane (50 ml) were mixed and shaken for 3 min; under Condition

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2, the filtered sample (20 ml) and n-hexane (50 ml) were mixed and shaken for 3 min before adding oleic acid (1 ml) and shaking again for 3 min; and under Condition 3, the filtered sample (20 ml) and oleic acid (1 ml) were mixed and shaken for 3 min before adding n-hexane (50 ml) and shaking again for 3 min. All of the hexane layer was recovered and its hydrocarbon content was determined by GC-FID. The extraction was performed in duplicate, and the average value is presented (Fig. 9). The hydrocarbon recovery efficiency of the sample to which oleic acid was added after hydrocarbon extraction (Condition 2) was comparable to that of the sample without oleic acid (Condition 1). Thus, adding oleic acid to the hexane layer after extraction did not affect the hydrocarbon recovery efficiency. However, the hydrocarbon recovery efficiency was markedly decreased by adding oleic acid to the sample before hydrocarbon extraction (Condition 3). Under Condition 3, hydrocarbons were dissolved in oleic acid, and then the oleic acid-containing hydrocarbons could interact with the water-soluble materials including algal amphiphilic polysaccharides. Amphiphilic materials such as proteins previously showed strong adsorption at the oil–water interface and stabilized oil-in-water emulsions.64 The adsorption occurred through hydrophobic groups present within the structure of the amphiphilic materials. Because algal polysaccharides also have amphiphilic characteristics,65 polysaccharides were adsorbed on the surface of oleic acid droplets, which contained hydrocarbons. Hence, hydrocarbon recovery became low under Condition 3. In the other cases, because hydrocarbons were adsorbed to the colony matrix,42 they would not disperse into the liquid, and oil-in-water emulsions did not form. Thus, hydrocarbon extraction was not prevented by the

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formation of emulsions in the filtered sample and the JET PASTER-treated sample, in which the cells showed little to no disruption. This supports our hypothesis that intracellular substances such as oleic acid can prevent hydrocarbon extraction and decrease the hydrocarbon recovery efficiency from samples treated by high-pressure homogenization. The hydrocarbon recovery efficiency was enhanced by concentrating B. braunii cell suspensions by cross-flow filtration. In contrast, disrupting cells in the concentrated sample by homogenization decreased the hydrocarbon recovery efficiency. Using the JET PASTER for mild mechanical treatment of concentrated samples should thus result in additional improvements in hydrocarbon recovery efficiency.

Energy analysis. The energy required for filtration and pretreatment methods was measured or calculated, and the ratios of the consumption energy to the production energy were analyzed according to the analytical conditions shown in Fig. 2. The analytical results for the consumption and production energies determined via five methods are shown in Figure 10. The production energies were smaller than the consumption energies in all cases. The consumption energy was 6.7 to 100 times as high as the production energy. However, because the membrane-filtration process is expected to be scaled up66-69, the relative consumption energy required for filtration might be reduced on a commercial scale. In particular, Gerardo et al.68 reported that the operating energy was decreased by half when a filter with twice the membrane area was used. Figure 11 represents the energy ratio between the extracted hydrocarbon energy and the consumption energy. Figure 11

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shows that thermal treatment combined with filtration resulted in the highest energy ratio among the five methods tested. Figure 12 shows the input energy per sample weight for each pretreatment method and that the input energy for thermal treatment was higher than for mechanical treatments. These findings mean that filtration reduced the large amount of consumption energy required for downstream processes, even if the energy per processing weight was higher than with the mechanical treatment. The difference in hydrocarbon recovery efficiency resulted in a high energy ratio for thermal treatment combined with filtration, compared to mechanical treatment combined with filtration. However, the consumption energy per sample weight with mechanical treatment was approximately two-thirds of that observed with thermal treatment, indicating that mechanical treatments may be superior to thermal treatments in terms of the processing sample weight (i.e., the water content of algae).

CONCLUSION In the present study, we investigated the effects of concentrating microalgal cell suspensions on mechanical treatment and the hydrocarbon recovery efficiency. B. braunii cell suspensions were concentrated by cross-flow filtration and treated with a high-pressure homogenizer or JET PASTER before extracting hydrocarbons. Concentrating the cell suspensions increased the apparent viscosity and altered the particle size and shape distributions. The condensation of polysaccharides by the concentration process likely accounts for the increase in viscosity, whereas the shear force in the pump

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used to introduce the suspension onto the membrane and/or between the algae and membrane wall disrupted colonies and removed the surrounding polysaccharide matrix. Mechanical treatment of the concentrated sample by high-pressure homogenization decreased the viscosity and hydrocarbon recovery efficiency. The former effect may have been caused by destruction of the polysaccharide bridges and the latter effect by the release of intracellular substances that prevented hydrocarbon extraction. Irrespective of membrane filtration, JET PASTER treatment increased hydrocarbon recovery efficiency as compared to samples without any mechanical treatment. In contrast to the homogenizer, the JET PASTER disrupted B. braunii colonies without destroying the cells, and removed the surrounding polysaccharides. Furthermore, cell disruption released intracellular substances that decreased the hydrocarbon recovery efficiency. These results indicate that the hydrocarbon recovery efficiency of a B. braunii cell suspension is increased by JET PASTER treatment as compared to membrane filtration alone, providing a basis for the use of B. braunii as a raw material for biofuel production. In addition, an energy analysis was performed. The consumption energy of the JET PASTER treatment combined with filtration was 7.6 times higher than the production energy. However, because the consumption energy per processing weight of the JET PASTER treatment was lower than those of a previous study and the conventional drying method, mechanical treatments may be superior to other treatments in terms of the processing sample weight.

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FIGURES

Figure 1. Schematic diagram of the JET PASTER high-speed mixer.

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Figure 2. Analytical conditions of the energy analysis.

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Figure 3. Apparent viscosities of the samples. (a) Sample without mechanical treatment. (b) Sample mechanically treated with the high-pressure homogenizer. (c) Sample mechanically treated with the JET PASTER. Error bars represent the standard error (n = 3).

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Figure 4. Particle size distributions of samples. (a-1) Unfiltered sample without mechanical treatment. (a-2) Filtered sample without mechanical treatment. (b-1) Unfiltered sample mechanically treated with the high-pressure homogenizer. (b-2) Filtered sample mechanically treated with the high-pressure homogenizer. (c-1) Unfiltered sample mechanically treated with the JET PASTER. (c-2) Filtered sample mechanically treated with the JET PASTER. Error bars represent the standard error (n = 3).

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Figure 5. Roundness distributions of samples. (a-1) Unfiltered sample without mechanical treatments. (a-2) Filtered sample without mechanical treatment. (b-1) Unfiltered sample mechanically treated with the high-pressure homogenizer. (b-2) Filtered sample mechanically treated with the high-pressure homogenizer. (c-1) Unfiltered sample mechanically treated with the JET PASTER. (c-2) Filtered sample mechanically treated with the JET PASTER. Error bars represent the standard error (n = 3).

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Figure 6. Micrographs of cells. Upper figures: micrographs without negative staining by India ink. Lower figures: micrographs with negative staining by India ink. (a-1) Unfiltered sample without mechanical treatments. (a-2) Filtered sample without mechanical treatment. (b-1) Unfiltered sample mechanically treated with the high-pressure homogenizer. (b-2) Filtered sample mechanically treated with the high-pressure homogenizer. (c-1) Unfiltered sample mechanically treated with the JET PASTER. (c-2) Filtered sample mechanically treated with the JET PASTER.

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Figure 7. Hydrocarbon recovery efficiency (RHC) from each sample. (a) Sample without mechanical treatments. (b) Sample mechanically treated with the high-pressure homogenizer. (c) Sample mechanically treated with the JET PASTER. Error bars represent the standard error (n = 3).

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Condition 1: Shaken for 3 min n-Hexane

Oleic acid

Condition 2:

Shaken for 3 min

Shaken for 3 min n-Hexane

Microalgal sample

n-Hexane Collect the organic layer

Condition 3:

Oleic acid Shaken for 3 min

Shaken for 3 min

Microalgae + oleic acid

Figure 8. Schematic illustration of the experimental procedure used to investigate the effect of oleic acid on the hydrocarbon recovery efficiency.

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Figure 9. Hydrocarbon recovery efficiency (RHC) from the mixture of samples and oleic acid. Extraction conditions are shown in Figure 8. Error bars represent the standard error (n = 2).

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Figure 10. Energy consumption and production from B. braunii via the filtration and pretreatment methods. Thermal treatment was performed considered the heating energy of water from 20°C to 90°C, using the heat capacity of water at 20°C. EC represents the consumption energy, and EP represents the production energy. (i) Filtration without any pretreatment. (ii) Mechanical treatment combined with filtration. (iii) Thermal treatment without filtration. (iv) Thermal treatment combined with filtration. (v) Drying method combined with filtration.

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Figure 11. Energy ratio between the extracted hydrocarbon energy and consumption energy. (i) Filtration without any pretreatment. (ii) Mechanical treatment combined with filtration. (iii) Thermal treatment without filtration. (iv) Thermal treatment combined with filtration. (v) Drying method combined with filtration.

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Figure 12. Input energy per sample weight. Input energy for the thermal* treatment is the calculated value, and others are the measured values. Thermal: energy required for the thermal treatment calculated using the heat capacity of water, HPH: energy required for the high-pressure homogenizer treatment, JP: energy required for the JET PASTER treatment, and FLT: energy required for filtration.

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AUTHOR INFORMATION *Corresponding Author Shun Tsutsumi. Department of Chemical Engineering, Graduate School of Engineering, Tohoku University, Aoba, Aramaki, Aoba-ku, Sendai, Miyagi 980-8579 Japan. Phone: +81-22-795-7251, e-mail: [email protected].

Author Contributions The manuscript was written through the contributions of all authors. All authors have approved of the final version of the manuscript. S. Tsutsumi contributed to the conception and design of the present study, analysis and interpretation of the data, drafting of the manuscript, and collection and assembly of data. Y. Saito contributed to the analysis and interpretation of the data, critical revision of the manuscript for important intellectual content, and logistic support for the present study. Y. Matsushita contributed to the analysis and interpretation of the data, critical revision of the article for important intellectual content, and logistic support for the present study. H. Aoki contributed critical revision of the article for important intellectual content, final approval of the manuscript, and obtaining funding and administrative and logistic support for the present study.

Funding Sources Ministry of Education, Culture, Sports, Science, and Technology (MEXT), Japan.

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Conflict of interest statement The authors declare no competing financial interest. ACKNOWLEDGMENT This work was supported by the Next-generation Energies for Tohoku Recovery (NET) project of the Ministry of Education, Culture, Sports, Science, and Technology (MEXT), Japan. The non-concentrated B. braunii cell suspension was kindly provided by Prof. Makoto M. Watanabe of the University of Tsukuba. The authors thank Dr. Demura of the University of Tsukuba and Mr. Michikawa of Sobio Technologies, Inc. for performing filtration of B. braunii.

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