A new method to produce cellulose nanofibrils from microalgae and the measurement of their mechanical strength

Accepted Manuscript Title: A new method to produce cellulose nanofibrils from microalgae and the measurement of their mechanical strength Authors: Hyun-Ro Lee, KyuHan Kim, Sung Cik Mun, Yong Keun Chang, Siyoung Q. Choi PII: DOI: Reference:

S0144-8617(17)31146-3 https://doi.org/10.1016/j.carbpol.2017.09.104 CARP 12849

To appear in: Received date: Revised date: Accepted date:

18-7-2017 29-8-2017 30-9-2017

Please cite this article as: Lee, Hyun-Ro., Kim, KyuHan., Mun, Sung Cik., Chang, Yong Keun., & Choi, Siyoung Q., A new method to produce cellulose nanofibrils from microalgae and the measurement of their mechanical strength.Carbohydrate Polymers https://doi.org/10.1016/j.carbpol.2017.09.104 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

A new method to produce cellulose nanofibrils from microalgae and the measurement of their mechanical strength

Hyun-Ro Lee, KyuHan Kim, Sung Cik Mun, Yong Keun Chang, Siyoung Q. Choi*

Department of Chemical and Biomolecular engineering, Korea Advanced Institute of Science and Technology (KAIST), Daejeon 34141, Korea

*Corresponding author, E-mail: [email protected]

Revised submission date: August 29, 2017

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Highlights  This work provides a new route to produce cellulose nanofibrils (CNFs) from microalgae.  To measure the tensile strength precisely, we develop the method to distinguish between straight and buckled fibrils.  The CNFs from N. oceanica show ~ 9 nm in diameter and 3-4 GPa of the tensile strength. 

The tensile strength of N. oceanica CNF is similar and even higher than that of other CNFs and general reinforcements.

Abstract Despite the enormous potential of cellulose nanofibrils (CNFs) as a reinforcing filler in various fields, the use of them has been limited by high-energy mechanical treatments that require a lot of energy and time consumption. To reduce the demands of energy and time required for mechanical treatments, microalgae, in particular, Nannochloropsis oceanica, which has small size, rapid growth rate, and high productivity was used as a CNFs source. This study obtains the CNFs by lipid/protein extraction, purification, and TEMPO-mediated oxidation processes under gentle mixing without high-energy mechanical treatments. Furthermore, to evaluate the applicability of microalgal CNFs as a reinforcing filler, this study estimated the mechanical strength of the fibrils by the sonication-induced scission method. To achieve a precise estimation, an effective method to distinguish straight fibrils from buckled fibrils was also developed, and subsequently, only straight fibrils were used to calculate the mechanical strength in the sonication-induced scission method. Consequently, the tensile strength of the N. oceanica CNFs is around 3-4 GPa on average which is comparable with the mechanical strength of general reinforcing fillers and even higher than that of wood CNFs. Thus, this study has shown that the newly proposed simplified method using N. oceanica is very successful in

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producing CNFs with great mechanical strength which could be used in various reinforcement fields.

Keywords: Microalgae; cellulose nanofibrils; tensile strength; sonication-induced scission

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1.

Introduction Cellulose nanoparticles are rod-like particles that have a diameter in the nanoscale and

consist of multiple cellulose chains combined by intra- and inter-molecular hydrogen bonds (Moon, Martini, Nairn, Simonsen, & Youngblood, 2011). Since they were pioneered by Turbak and Herrick in 1983 (Herrick, Casebier, Hamilton, & Sandberg, 1983; Turbak, Snyder, & Sandberg, 1983), they have been classified into several types depending on their morphologies, size, crystallinity, extraction method by International Nanocellulose Standards Coordination Committee at 2011: cellulose microfibrils (Dufresne, 2013; Shinichiro Iwamoto, Abe, & Yano, 2008; Siaueira, Bras, & Dufresne, 2009), cellulose nanofibrils (CNFs) (Miao et al., 2016; Saito, Kuramae, Wohlert, Berglund, & Isogai, 2013), and cellulose nanocrystals (Elazzouzi-Hafraoui et al., 2008; Lin & Dufresne, 2014). Among them, CNFs, which are sometimes called nanofibrillated celluloses, have been referred to separated elemental fibrils with smaller diameters extracted from the cellulose-based materials by mechanical treatment (Moon et al., 2011; Siró & Plackett, 2010). They generally have generally narrow diameter distribution (420 nm) even though their diameter varies depending on the species and production routes (Dufresne, 2013; Moon et al., 2011). Because of their high tensile modulus, light weight, surface activity, and biocompatibility, they have recently attracted much attention for use in a variety of applications: for example, CNFs have been used as a filler in the composites field (packaging and paper industry) (Dufresne, 2013; Lavoine, Desloges, Dufresne, & Bras, 2012; Lee, Heo, Lee, Kim, & Shin, 2017), a stabilizer in emulsions (Blaker, Lee, Li, Menner, & Bismarck, 2009), a conducting sheet in electronics (Shah & Brown, 2005), or a biomaterial in a medical application (Klemm, Schumann, Udhardt, & Marsch, 2001). Despite their versatility, the production of CNFs from wood in an industrial scale is quite difficult. Typically, high-energy mechanical treatments such as grinding, high-shear 4

homogenization, high-intensity ultrasonication, and cryocrushing are required for pulverization of wood pulp or sawdust into micrometer-scale particles to increase the efficiency of swelling and purification (e.g., pulping) as well as separation of cellulose fibers into the individual fibrils (Abdul Khalil et al., 2014; Chen et al., 2011; Dufresne, 2013; Shinichiro Iwamoto et al., 2008; Moon et al., 2011; Siró & Plackett, 2010). However, mechanical treatments require high-energy input (5,000-70,000 kWh/ton) (Eriksen, Syverud, & Gregersen, 2008; Wang et al., 2012) and time-consuming process performed with repeated cycles (Hassan, Mathew, Hassan, El-Wakil, & Oksman, 2012; Stelte & Sanadi, 2009), thus hindering the industrial use of CNFs. Although chemical and enzymatic methods performed before the mechanical treatment could help reduce the energy consumption (Pääkko et al., 2007; Siró & Plackett, 2010), combination of mechanical and chemical treatment could not completely resolve high-energy and time consuming problems, and they could damage the crystalline phase of the fibrils, thus deteriorating their mechanical properties (Felix & Thomas, 2004; S. Iwamoto, Nakagaito, & Yano, 2007; Moon et al., 2011). Non-timber plants and agricultural byproducts also have been used as sources of CNFs, such as cotton (Miao et al., 2016), wheat straw (Singh, Kaushik, & Ahuja, 2016), palm (Benhamou, Dufresne, Magnin, Mortha, & Kaddami, 2014), sisal (Siaueira et al., 2009), and kenaf (Karimi, Tahir, Karimi, Dufresne, & Abdulkhani, 2014), etc. Because they have less hydrophobic matrix contents (e.g., lignin) and less tightly-coiled fibrils than wood (Dinand, Chanzy, & Vignon, 1996; Dufresne, 2013; Siró & Plackett, 2010), they could retain more water, resulting in effective mechanical fibrillation and simplified purification process (Dufresne, 2013; Spence, Venditti, Habibi, Rojas, & Pawlak, 2010). However, high-energy mechanical treatments including retting, pulping, and mechanical fibrillation are still required to produce the CNFs, and requirements of vast land for massive harvests also limit their bulk production. 5

Meanwhile, some researchers produced MFCs or CNFs from tunicate (Shinichiro Iwamoto, Kai, Isogai, & Iwata, 2009; Saito et al., 2013) and macroalgae, such as Valonia (Hanley, Giasson, Revol, & Gray, 1992; Imai & Sugiyama, 1998), Cladophora (Imai & Sugiyama, 1998; Mihranyan, 2011; Mihranyan, Nyholm, Bennett, & Strømme, 2008; Saito et al., 2013), and Boergesenia (Imai & Sugiyama, 1998), but commercial production is also limited to mechanical fibrillation and additional mechanical/chemical process to isolate mantles of the tunicate and to remove large portion of non-cellulose matrix substances (Shinichiro Iwamoto et al., 2009; Saito et al., 2013; Zhao & Li, 2014). To overcome the limitations of the current CNFs production methods, this study used microalgae as a source. Like macroalgae, microalgae are one of the photosynthetic organisms found in bodies of fresh and salt water, but microalgae greatly differ from macroalgae in their size and cell structure. Compared to macroalgae, multicellular organisms that has similar structures with higher plants, microalgae are composed of only one cell and has far simpler microscopic structure (John, Anisha, Nampoothiri, & Pandey, 2011). It is advantageous that microalgae can be massively cultivated by photosynthesis throughout the year (Jorquera, Kiperstok, Sales, Embiruçu, & Ghirardi, 2010; Radakovits et al., 2012). They have fast growth rate (microalgal biomass typically doubles in a day) (Ho et al., 2013; K. W. M. Tan & Lee, 2016; Yusuf, 2007), short harvesting cycle (~1-10 days) (Harun, Danquah, & Forde, 2010), and higher biomass areal productivity (100-260 ton/ha∙year) (Rodolfi et al., 2009; Yusuf, 2007) than woody biomass (8-22 ton/ha∙year) (Heaton, 2004; Ragauskas, 2006), non-woody plants (7-87 ton/ha∙year) (Huber, Iborra, & Corma, 2006), and macroalgae (19-61 ton/ha∙year) (De Paula Silva, Paul, De Nys, & Mata, 2013). In particular, the Nannochloropsis species are a few micrometers in size, and their cell wall mainly consist of cellulose without hemicellulose and

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lignin (Scholz et al., 2014), thus large amounts of CNFs could be easily isolated by simplified purification process. Additionally, microalgal CNFs can be obtained from the abundant waste from the microalgae industries producing biofuels and co-products. Microalgae are typically used to produce useful materials such as lipids (e.g., fatty acid methyl esters and eicosapentaenoic acid), proteins, and pigments for biofuel, food and feed, cosmetics, and pharmaceutical drugs (Chiu et al., 2009; Gerde et al., 2013; Grimi et al., 2014; Jeon, Jeong, & Chang, 2016; Wen & Chen, 2003). After extracting the useful materials from microalgae, abundant amounts of solid waste including cellulose are left and used to produce low-value products such as animal feed (Z. Yang, Guo, Xu, Fan, & Luo, 2011a) and biogas (Stephens et al., 2010; Williams & Laurens, 2010; Z. Yang et al., 2011a; Z. Yang, Guo, Xu, Fan, & Luo, 2011b), but they also could be reused to produce a high-value product such as CNFs, thereby improving the economic feasibility of NFC production and even the sustainability of microalgal industries. Thus, microalgae can be a potentially superb candidate for massive and inexpensive CNFs production; however, experimental studies on the CNFs production from microalgae have not yet been reported. To understand the underlying characteristics of CNFs that are produced by a variety of methods and to utilize them as reinforcing fillers for various applications, it is very important to measure the mechanical properties of CNFs, especially a single fibril of CNFs. Several groups have measured the tensile strength of a single fibril using various techniques including a nano tensile tester (E. P. S. Tan, Ng, & Lim, 2005) and tensile and bending test by atomic force microscope (AFM) cantilevers (Shinichiro Iwamoto et al., 2009; Yu, 2000). However, these techniques have several disadvantages when measuring tensile properties of CNFs: first, it is hard to grip and apply a force delicately due to the very small diameter of CNFs which is 7

generally below 10 nm; second, these techniques are not adequate for statistically estimating the mechanical properties of the whole fibrils. Instead of directly applying the mechanical deformation of a single fibril, some researchers have calculated the elastic modulus indirectly from the crystallite extension of a bundle of fibrils with an X-ray diffractometer (Sakurada, Nukushina, & Ito, 1962) and from the shift of a specific Raman band for cellulose Ⅰ(1095 cm1

) under deformation of NFC-reinforced epoxy composites by the four-point bending test

(Eichhorn & Young, 2001; Rusli & Eichhorn, 2008; Šturcová, Davies, & Eichhorn, 2005). However, it is still challenging and elaborate to grip the bundle of fibrils during the determination of the elastic modulus by X-ray diffraction (Šturcová et al., 2005), and the elastic modulus estimation of a single fibril could be inaccurate by Raman spectroscopy because the bending test is conducted with an epoxy resin that is reinforced by CNFs, not with only NFC fibrils (Moon et al., 2011). Recently, sonication-induced scission was suggested for measuring the tensile strength of a single fibril indirectly (Huang, Knowles, & Terentjev, 2009; Saito et al., 2013). This technique has been developed mainly for investigating the cavitation kinetics of carbon nanotubes (Hennrich et al., 2007; Lucas et al., 2009; Pagani, Green, Poulin, & Pasquali, 2012; Stegen, 2014); however, it was demonstrated that it could be also used for other types of fibril materials including amyloid fibrils, silver nanowires, and cellulose fibrils (Huang et al., 2009; Saito et al., 2013). This technique is based on the fact that the elongation force generated by the bubble burst can be balanced by the mechanical strength of a fibril. Then, the tensile strength of a fibril can be calculated by measuring the dimensions of the fibril as a result of their breaking by sonication. However, this technique could not be directly applied when buckling occurs in many fibrils (e.g., kinks) after sonication in contrast to rod-like fibrils that are expected to be produced by the stretching mechanism introduced in a previous study (Pagani et al., 2012). 8

Because fibrils with buckling possess a longer contour length than that of broken ones, the tensile strength might be overestimated. To adapt the sonication induced method to measure the tensile strength by eliminating the buckled fibrils, it is thus necessary to develop a precise method to determine whether the fibril is buckled. Here, this study has two major purposes. First, we introduce a method to produce CNFs from microalgae without high-energy mechanical treatments. Second, to check whether the microalgal CNFs could be a potential candidate for use in various applications, the mechanical strength of the microalgal CNFs was measured by the sonication-induced scission method with some modifications. To precisely measure the mechanical strength of the CNFs using this method, this study adapted the ratio of the persistence length to the contour length, which is generally used to assess the flexibility of linear polymers in the polymer field (Käs et al., 1996), and calculated the mechanical strength of the CNFs whose ratios were larger than 5. As a result, the tensile strength of the microalgal CNFs is higher than that of wood CNFs reported in previous studies (Saito et al., 2013). This strength difference is explained by the crystallinity and the cellulose Iα fraction of the microalgal CNFs. Finally, this study considered the applicability of the mciroalgal CNFs as general reinforcing fibrils.

2.

Experimental

2.1.

Materials Microalgae strains, Nannochloropsis oceanica and Nannochloropsis salina, were

supplied by Chloland Inc. (Republic of Korea). Solid waste was obtained through the extraction of lipids and proteins from raw microalgae (Fig. 1A). Lipids were extracted by the Bligh-Dyer method, which is a well-known protocol for lipid extraction of a biomass (Bligh & Dyer, 1959). Proteins were extracted by the alkaline extraction method using 55 mM NaOH solution (Sari, 9

Syafitri, Sanders, & Bruins, 2015). The remaining solid was separated by centrifugation, and then the solid waste in slurry form was stored in a refrigerator at 3-5oC to prevent its decomposition with time. All chemicals including chloroform, hexane, methanol, ethanol, sodium hydroxide, hydrochloric acid, sodium chlorite, sodium acetate solution, 2,2,6,6tetramethyl-1-piperidinyloxy (TEMPO), sodium bromide, and sodium hypochlorite solution were purchased from Sigma-Aldrich.

2.2.

Composition analysis of the solid waste For the composition analysis of the solid waste, the freeze-dried sample was used. The

lipid content in the dried solid waste was determined by a previously reported method with a few modifications (F. Yang et al., 2014). The dry solid waste (200 g) was mixed with hexane (800 ml) and ethanol (800 ml) in a shaking incubator at 30oC to extract the lipids. After 24 h, the suspension was centrifuged at 9000 rpm for 10 min, and the lipids in the liquid phase were collected and gravimetrically measured. The protein content was calculated by multiplying the weight percent of nitrogen by the nitrogen-to-protein conversion factor for marine microalgae, 6.28 (Safi et al., 2013). The weight percent of nitrogen was measured by an element analyzer (FLASH 2000 series, Thermo Scientific, USA). Inorganic material in the dried solid waste was analyzed with a NREL Laboratory Analytical Procedure for microalgae (Laurens et al., 2012): the organic compounds of the dried solid waste (~100 mg) were completely oxidized in an electric furnace (JSMF-45T, Laftech, Australia) by placing the samples at 105oC for 24 h at 250oC for 30 min. and at 575oC for 3 h, sequentially, and the inorganic compounds were then gravimetrically determined. Because microalgae is mostly composed of lipids, proteins, inorganic materials, and carbohydrates (Yoo, Park, Yang, & Choi, 2015), the dried solid waste mainly possesses these four components. Therefore, the total carbohydrate content in the dried 10

solid waste was calculated by subtracting the amount of the lipids, proteins, and ash in the dried solid waste from the total weight of the dried solid waste.

2.3.

Preparation of microalgal cellulose nanofibrils To obtain CNFs with an extremely high carbohydrate content, further purification was

required to remove the remaining lipids, proteins, and inorganic materials in the solid waste based on previously reported methods (Dinand et al., 1996; Siaueira et al., 2009; F. Yang et al., 2014). The solid waste in slurry form prepared in section 2.1 was used to go through the purification process. Remaining lipids was removed by mixing the solid waste (50 g) with hexane (200 ml) and ethanol (200 ml) in the shaking incubator at 30oC, and then the liquid phase including lipids were removed by centrifugation. The lipid-extracted solid waste were mixed with 2 wt% NaOH solution for 2 h at 80oC and washed with distilled water three times each. Subsequently, it was bleached by mixing 1.7 wt% of sodium chlorite in 0.2 M sodium acetate buffer solution for 4 h at pH 4.9 and 70oC and washed three times each followed by the collection of fibrils after centrifugation. After the purification, TEMPO-mediated oxidation was used to facilitate the delamination of the cellulose fiber and dispersion of the fibrils in water by changing the hydroxyl groups on the fibril surface to carboxylic groups (Okita, Saito, & Isogai, 2010). This treatment was done by mixing the fibrils in slurry form (1 g of the fibrils content) with 0.1 mmol of TEMPO, 1 mmol of sodium bromide, and 5 mmol of sodium hypochlorite solution for 2 h, while the pH was maintained at 10 using 0.5 M NaOH solution. Oxidized fibrils were washed with distilled water and collected by centrifugation three times, and then oxidized fibrils in slurry form were stored at 3-5oC.

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2.4.

Crystallinity and the fraction of cellulose Iα phase of microalgal cellulose nanofibrils The crystallinity and the fraction of cellulose Iα phase of the CNFs were measured by

X-ray diffraction. The freeze-dried CNFs were prepared in a powder form to measure the Xray data with a high resolution powder X-ray diffractometer (SmartLab, Rigaku) operated at 40 kV and 250 mA with a Cu-Kα1 optics monochromator. The sample was scanned at a speed of 4°/min in the range of 2θ from 10° to 40° with a step interval of 0.01°. The crystallinity index (𝐶𝑟𝐼) was calculated with the Segal method (Segal, Creely, Martin, & Conrad, 1959), 𝐶𝑟𝐼 (%) =

𝐼200 −𝐼𝑎 𝐼200

× 100 ....................................................................................................... (1)

where 𝐼𝑎 is the intensity of amorphous phase of the cellulose at the lowest value around 18°, and 𝐼200 is the intensity of both the crystalline and amorphous phase at the highest value around 22.7°. The intensity of the samples was measured by excluding the background intensity. The fraction of the cellulose Iα phase was calculated depending on the shift of the d1 and d2 spacings whose details are well introduced in a previous study (Masahisa Wada, Okano, & Sugiyama, 2001).

2.5.

Determination of the dimensions and persistence length of single fibril The contour length and diameter of the CNFs were identified by AFM images with a

10 μm × 10 μm (5,100 pixels × 5,100 pixels) size (INNOVA-LABRAM HR800, Bruker). Thus, a droplet of the fibril suspension at a concentration of ~0.01 wt% was first placed on a mica substrate which was pretreated with plasma (myPL-150, Atmospheric process plasma Inc.) for 1 min to increase the polarity of the mica surface, and excessive liquid on the surface was then removed by spin coating for 15 min. Using the tapping mode of the AFM whose silicon cantilever had a resonant frequency of 70 kHz and a spring constant of 2 N/m (OLTESPA-R3, 12

Bruker), the dimensions of the fibrils were accurately investigated When the CNFs were overlapped each other and entire image of a fibril was not visualized, they were not counted as experimental samples because their lengths were unmeasurable. The diameter of a fibril was considered to be the average height of a fibril in the AFM images, which was calculated by averaging the heights of six different spots selected randomly in the fibril using the section mode of the Nanoscope Analysis software. The contour length of a fibril was measured with the Fiberapp software, which could analyze and track the fibril contour quickly (Usov & Mezzenga, 2015). The persistence length of a fibril was estimated by the mean squared end-toend distance method from the contour line of a fibril previously described elsewhere (Usov et al., 2015; Usov & Mezzenga, 2015). Fibrils smaller than 100 nm in contour length or 1 nm in diameter were excluded for measuring the dimensions because their contour was poorly captured by the tracking program.

2.6.

Tensile strength of a single fibril The tensile strength of a NFC can be calculated by a sonication-induced scission

method. This method is based on the fact that fibrils arranged axially with bubbles are broken by a stretching force (Hennrich et al., 2007). Because the stretching force at the center of the fibril can be expressed as 𝐹𝐶 =

1 2

𝜋𝜂𝜖̇𝐿2 , where 𝜂 is the viscosity of the liquid; 𝜖̇ is the shear

rate, and 𝐿 is the contour length of fibril, it decreases continuously as the contour length decreases until the stretching force is balanced with the breaking force which is proportional to the tensile strength. Assuming the NFC follows a continuum cylindrical model with the diameter (𝐷), the tensile strength (𝜎) is given by 𝜎 = 2𝜂𝜖̇

𝐿2 𝐷2

,............................................................................................................................. (2) 13

Here, we use 10-3 Pa·s and 109 s-1 for the water viscosity and shear rate, respectively. Although it is hard to calculate the exact value of the shear rate due to the significant variation of the shear rate at each experimental condition (Kuijpers, van Eck, Kemmere, & Keurentjes, 2002), 109 s-1 was considered here as a reasonable value for the shear rate with a sonication frequency and sound pressure of 20 kHz and 10 bar, respectively, (Hennrich et al., 2007; Pagani et al., 2012). The fibril suspension of ~0.01 wt% was sonicated for 10-420 minutes with a pulse mode of 5s-on/5s-off. An ultrasonic processor (VCX 750, SONIC & MATERIALS Inc.) with tapered microtips (630-0420, SONIC & MATERIALS Inc.) was used for the sonication. It was operated at a vibration amplitude of 30%. The temperature of the suspension was maintained at 20oC with a customized temperature controller.

3.

Results and discussion

3.1.

Production and characterization of the microalgal CNFs After extracting useful materials including lipids and proteins by the Bligh-Dyer

method (Bligh & Dyer, 1959) and alkaline extraction method from Nannochloropsis oceanica (N. oceanica) and Nannochloropsis salina (N. salina), the solid wastes of each species were collected, and their composition was analyzed before the producing the microalgal CNFs. From the composition analysis (Section 2.2) of the solid waste which has carbohydrates, proteins, lipids and inorganic matter, it is clearly seen that the carbohydrates comprise the largest proportion of the waste from the N. oceanica and N. salina (Fig. 1C), and this is caused by high cellulose contents in the cell wall of the Nannochloropsis species (~75 wt%) (Scholz et al., 2014). The cellulose content was calculated from the ratio of dried cellulose weight to dried 14

solid waste weight. Because other components except for cellulose could be intensively removed through sequential cellulose isolation processes (lipids and proteins extraction, purification, bleaching) which are introduced in previous researches (Bligh & Dyer, 1959; Dinand et al., 1996; Sari et al., 2015; Siaueira et al., 2009; F. Yang et al., 2014), the remaining material could be ascribed exclusively to cellulose. The initial cellulose content was 34 wt% of the solid waste, which means the total carbohydrate content in solid waste consists of 47 wt% of the cellulose contents and 53 wt% of other carbohydrates including sugar and starch. However, compared to other cellulose sources, it is cautious to present the exact amount of cellulose in solid waste due to significant variation of cellulose content in microalgae depending on culture conditions. For example, previous research reported nitrogen deprivation in microalgae cultivation could induce the thickening of the cell wall and 1.5-fold enhancement of the cellulose content in Nannochloropsis sp. (Jeong et al., 2017). Likewise, the deficiency of salt, phosphate, and sulfur in culture medium was reported to increase the width of the cell wall (Beacham, Bradley, White, Bond, & Ali, 2014; Jeong et al., 2017). Therefore, if the culture conditions of microalgae are optimized for CNFs production by further researches, the cellulose content could occupy more than 34 wt% of the solid waste of microalgae, and then the productivity of CNFs from microalgae could be improved. The 14-22 wt% inorganic compounds of the dry solid waste were mainly from the sea salt and mineral accumulation in a cell body. Using the solid waste of N. oceanica whose carbohydrate contents are far larger than that of N. salina, we obtained the fibrils by the purification process and the TEMPO-mediated oxidation process (Section 2.3) without any further mechanical treatments required for pulping and fibrillation. We visualized them using the tapping mode of AFM (Fig. 1D) and estimated their diameter by measuring the height of a fibril whose log-normal average value was 9.0±2.4 15

nm. To check whether these fibrils are individually separated fibrils or multiple ones, we performed the ultrasonication on the fibrils suspension. Because the average diameter of the fibrils was reduced from 9.0±2.4 nm to 6.7±2.0 nm after sonication for 420 min (Fig. 2F), we conclude that they are not aggregates, but mainly elemental fibrils. The CNFs production without high-energy mechanical treatments is worth noting because the CNFs from other species were generally produced by the high-energy mechanical treatments (Eriksen et al., 2008; Lavoine et al., 2012; Siró & Plackett, 2010). This result might lead to less demands of energy and time of mechanical fibrillation for the CNFs production, except for the production of short CNFs (a few micrometers in length). Therefore, microalgae have a potential to produce the CNFs with less energy and time. We suggest that the fibrillation of the cellulose fiber into the nanometer sized fibrils without the mechanical treatments is because our production process does not have any “drying” process, which could induce an aggregation of the fibrils. If raw materials containing cellulose fibrils is dried, the fibrils were tightly agglomerated by hydrogen bonding between each other (Shinichiro Iwamoto et al., 2008; Moon et al., 2011). Even though dried raw materials go through the swelling process with water, disintegration of agglomerated fibrils is limited because water could not permeate into all the space between the fibrils. However, microalgae are dispersed well in the water, so they go through the CNFs production route without any drying processes. In addition, it is reported that Nannochloropsis species have delicate cellulose fibrils without tightly entangled fibrous structure of secondary cell wall and rosettes complexes (Giddings & Staehelin, 1988; Scholz et al., 2014). Thus, untangled fibrous structure of Nannochloropsis species is easier to be separated than higher plants and macroalgae.

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Fig. 1. (A) Schematic illustration of production of cellulose nanofibrils (CNFs) from microalgae which contains high-value products including lipids, proteins, pigments, and CNFs. (B) Optical microscopic image of the Nannochloropsis species, which generally has a spherical shape that is a few micrometers in diameter. (C) Composition of the dry solid waste from N. oceanica and N. salina. (D) Atomic force microscopic images of N. oceanica CNFs.

3.2. The dimensions of N. oceanica cellulose fibrils with sonication time To measure the tensile strength of a single N. oceanica NFC by sonication-induced scission method, various sizes of N. oceanica CNFs was produced depending on the sonication 17

time. Aqueous solutions of cellulose fibrils with 0.01 wt% were sonicated for 10, 60, 240, and 420 min, and the dimensions of the fibrils at each condition were estimated by AFM shown in Fig. 2A-D. It is clearly shown that the populations of the contour length at each sonication time fit quite well with a log-normal distribution, as seen in Fig 2E, and the average contour length at each sonication time became slightly smaller as the sonication time was increased while its standard deviation was reduced drastically. In addition, the diameter of the fibrils at each sonication time was also distributed log-normally; however, no significant dependence of the diameter distribution and log-normal average diameter on the sonication time was observed here: average diameter of ~7 nm was observed at 60 and 420 min, shown in Fig 2F. Because the average diameters were maintained around 7 nm despite of performing sonication for 420 min, we believe that most of the fibrils are individual fibrils. Compared with the CNFs from other species, the diameter of the CNFs from N. oceanica is larger than the diameter of CNFs from plants, such as wood and cotton (3-5 nm) (Saito et al., 2013; Saito, Nishiyama, Putaux, Vignon, & Isogai, 2006) but less than the diameters of tunicate (10-20 nm) and macroalgae, such as valonia (~20 nm), micrasterias (2060 nm), and cladophora (20-30 nm) (Mihranyan, 2011; Moon et al., 2011).

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Fig. 2. Atomic force microscopic images of the N. oceanica cellulose nanofibrils (CNFs) sonicated for 10 (A), 60 (B), 240 (C), and 420 min (D), respectively. The distributions of the contour length (E) and diameter (F) of the N. oceanica CNFs with varying sonication times. The solid lines correspond to the best-fitted curves for the log-normal distribution.

3.3.

Applying the sonication-induced method even in the presence of buckled fibrils. During the sonication, a cavitation force is applied to the CNFs, and this could stretch out

the CNFs in the direction of the fibril axis thereby producing a rod-like structure. This stretching mechanism has been reported frequently in previous studies on carbon nanotubes (Hennrich et al., 2007; Lucas et al., 2009) (Fig. 3A). However, it was clearly observed that a large fraction of CNFs from N. oceanica were not straightened out by the sonication process in contrast to carbon nanotubes (Fig. 2A-D), suggesting that the CNFs presumably follow another mechanism, namely a buckling mechanism (Pagani et al., 2012). Pagani et al. suggested that fibrils with a 19

large aspect ratio could be arranged tangentially with the direction of the cavitation force first followed by the center of the fibril being pulled toward the center of the bubble (Fig. 3B) resulting in the buckling with irreversible deformation.

Fig. 3. Illustration of the sonication-induced scission mechanism of cellulose nanofibrils. (A) Stretching mechanism. (B) Buckling mechanism. (C) The average contour length of fibrils that follow a power-law dependence on the sonication time. In the arithmetic mean value of the contour length, its power is -0.25, and the log-normal mean value of the contour length has a power of -0.14. The two solid black lines whose powers are -0.5 and -0.2, respectively, correspond to the two different mechanisms, stretching and buckling, induced by the sonication (Hennrich et al., 2007; Pagani et al., 2012).

Indeed, the mechanism for changing the dimensions of the CNFs during the sonication could be specified by identifying the relation between the average length of the fibrils and the sonication time (Hennrich et al., 2007; Lucas et al., 2009; Pagani et al., 2012). It has been reported that the average length (L) of fibrils follows a power-law dependence on the sonication 20

time (t), L ∝ 𝑡 𝑛 , and the fibrils undergoing the stretching and buckling mechanisms have the power of n ~ -0.5 (Hennrich et al., 2007) and n ~ -0.2 (Lucas et al., 2009; Pagani et al., 2012), respectively. Our results clearly show that the average contour length of the CNFs decreased with the power law dependence whose n was ~ -0.25 (arithmetically averaged) and -0.14 (lognormally averaged) shown in Fig. 3C. Regardless of the averaging method, the powers are close to -0.2, and this strongly suggests that the CNFs of N. oceanica are affected significantly by the buckling mechanism during the sonication induced scission process. Unfortunately, the current theoretical model for tensile strength estimation (equation (2)) considers only straight fibrils and could not explain the bulked ones. Therefore, the values of the contour length and the diameter for only the straight fibrils, excluding the buckled ones, are required to estimate precisely the tensile strength of the N. oceanica CNFs. However, it is very difficult to identify whether fibrils are straightened or buckled, and there are no quantitative standards yet. Thus, we used the ratio of a persistence length (𝐿𝑝 ) to a contour length (𝐿𝑐 ) for distinguishing the two different shapes, a straightened one and a buckled one. 𝐿𝑝 is defined as the characteristic length from the point on a fibril to another one where the tangent vectors along the contour line of the fibril are no longer correlated. In the polymer field, the flexibility of linear polymers has been assessed by the relationship between 𝐿𝑝 and 𝐿𝑐 (Käs et al., 1996). If the 𝐿𝑝 of a linear polymer is longer than its 𝐿𝑐 , it is considered to be a rod-like polymer, and in the opposite case, it is considered to be a flexible one. When the 𝐿𝑝 is comparable with the 𝐿𝑐 , the polymer is referred to as a semi-flexible polymer. To describe the straightness of fibrils by the 𝐿𝑝 ⁄𝐿𝑐 ratio, we used the end-to-end distance, 𝑅⃗, which indicates the vector from one end of a fibril to the other end. Assuming a

21

linear polymer follows a worm-like chain model, the mean squared end-to-end distance, ⟨𝑅2 ⟩, can be expressed by 𝐿𝑝 and 𝐿𝑐 (Grebikova et al., 2016). ⟨𝑅2 ⟩

=

𝐿 2𝐿𝑝 2 ( 𝑐 𝐿𝑝

−1+𝑒

𝐿 − 𝑐

𝐿𝑝

) ................................................................................................. (3)

When both sides in equation (3) are divided by 𝐿𝑐 2 , 〈𝑅2 〉⁄𝐿𝑐 2 can be written as the function of the 𝐿𝑝 ⁄𝐿𝑐 . ⟨𝑅2 ⟩ 𝐿𝑐 2

𝐿𝑝 2

𝐿𝑝 −1

𝐿𝑐

𝐿𝑐

= 2 ( ) {( )

−1+𝑒

𝐿𝑝 −1

−( 𝐿 ) 𝑐

} ................................................................................ (4)

Here, as a fibril becomes straighter, |𝑅⃗| approaches the 𝐿𝑐 , and as a result, 〈𝑅2 〉⁄𝐿𝑐 2 is close to 1. As shown in the graph of 〈𝑅2 〉⁄𝐿𝑐 2 against 𝐿𝑝 ⁄𝐿𝑐 (Fig. 4), 〈𝑅2 〉⁄𝐿𝑐 2 increases rapidly when 0 < 𝐿𝑝 ⁄𝐿𝑐 < 1 while the slope of the graph decreases significantly when 𝐿𝑝 ⁄𝐿𝑐 > 1. This clearly indicates that a higher 𝐿𝑝 ⁄𝐿𝑐 value of a fibril corresponds to that fibril being stiffer.

Fig. 4. A graph showing the correlation between the ratio of the mean squared end-to-end distance and the squared contour length, 〈𝑅2 〉⁄𝐿𝑐 2 , and the ratio of the persistence length to 22

the contour length, 𝐿𝑝 ⁄𝐿𝑐 , described in equation (4). As the 𝐿𝑝 ⁄𝐿𝑐 increases, 〈𝑅2 〉⁄𝐿𝑐 2 increases rapidly at the beginning; however, it increases slowly when the 𝐿𝑝 ⁄𝐿𝑐 > 1. Because the minimum value of the persistence length is the distance between the carbon atoms, the 𝐿𝑝 ⁄𝐿𝑐 and 〈𝑅2 〉⁄𝐿𝑐 2 cannot be zero.

3.4.

Estimation of the tensile strength for N. oceanica CNFs After a sonication of 240 min, straightened fibrils began to be observed frequently in the

AFM images (Fig. 2), and this is in good agreement with previous research which reported that CNFs from wood and tunicate have an equilibrium size after a sonication of 200 min (Saito et al., 2013). To get a more precise tensile strength, we picked the fibrils after a sonication of 420 min because fibrils with a longer sonication time could have dimensions that are closer to the equilibrium size although the contour length distribution of the fibrils did not change significantly between 240 and 420 min. We first disregarded the buckled CNFs which were determined by measuring the 𝐿𝑝 and 𝐿𝑐 of the fibrils. To do so, we established a specific value for the 𝐿𝑝 ⁄𝐿𝑐 from 1 to 10 as a baseline to divide all the fibrils into two different groups: buckled ones and straightened ones. The group that possessed a lower value than the baseline of the 𝐿𝑝 ⁄𝐿𝑐 was considered to be buckled fibrils, and the others were considered as straightened fibrils. We then picked straighten fibrils whose 𝐿𝑝 ⁄𝐿𝑐 ratio was bigger than the baseline. Thereafter, the tensile strength was statistically analyzed with equation (2). As well as the arithmetic mean, we calculated the geometric mean value using the log-normal distribution fit with the parameters of μ and σ, which are the mean and standard deviation (STD), respectively (Usov et al., 2015).

23

As indicated in Table 1, when two different cases, no baseline and a 𝐿𝑝 ⁄𝐿𝑐 of 5 were compared to each other, the arithmetic mean value was reduced by about half while the geometric mean decreased slightly. The reason for the lower mean value with the higher baseline might be because fibrils with a longer 𝐿𝑐 could be more easily buckled and excluded from the samples for calculating the tensile strength, and this is well observed in Fig. 5. Meanwhile, when the baseline of the 𝐿𝑝 ⁄𝐿𝑐 is larger than 5, the 〈𝑅2 〉⁄𝐿𝑐 2 values are already very close to 1, and the two different mean values remain almost constant even with the increasing the baseline. Therefore, we expect that if we choose fibrils whose 𝐿𝑝 ⁄𝐿𝑐 value is 5, we can readily and reasonably estimate the tensile strength of CNFs, and this procedure strongly suggests that the tensile strength of N. oceanica CNFs was 3-4 GPa on average.

Fig. 5. The tensile strength distribution of the N. oceanica cellulose nanofibrils (CNFs) depending on the ratio of the persistence length to the contour length ( 𝐿𝑝 ⁄𝐿𝑐 ). (A) Approximately 200 fibrils from among all the fibrils that included straight fibrils and bucked fibrils sonicated for 420 min were used to estimate the tensile strength. The fibrils for which the 𝐿𝑝 ⁄𝐿𝑐 was bigger than 2 (B) and 5 (C) were then picked from among all the fibrils to 24

estimate the tensile strength. The lines represent best-fitted curves for the log-normal distribution.

Table 1. The tensile strength of N. oceanica CNFs depending on the pretreatment and baseline of the 𝐿𝑝 ⁄𝐿𝑐 with other species Baseline Species

Tensile strength (GPa)

𝐿𝑝 ⁄𝐿𝑐

〈𝑅2 〉⁄𝐿𝑐 2

-

Arithmetic mean Geometric mean (STD.)

(STD.)

-

6.7 (7.2)

3.6 (2.8)

1

0.74

5.4 (5.3)

3.5 (2.5)

2

0.85

4.5 (5.0)

3.2 (2.0)

3

0.90

4.3 (5.3)

3.1 (1.9)

5

0.94

3.7 (3.0)

3.0 (2.1)

8

0.96

3.7 (3.2)

2.8 (2.0)

10

0.97

3.8 (3.4)

2.7 (1.9)

Wood*

-

-

2.6-3.6 (-)

0.8-1.5 (3.5)

Tunicate*

-

-

3.0-6.4 (-)

1.5 (3.9)

N. oceanica

*Data are from previous research (Saito et al., 2013).

3.5.

Comparison of N. oceanica CNFs with CNFs of other species and general

reinforcements Compared to CNFs of other species (Saito et al., 2013), the average tensile strength of N. oceanica CNFs is higher than wood CNFs and comparable with tunicate CNFs (Table 1). A previous studies reported that the crystallinity of the cellulose significantly influences the elastic modulus of the fibrils (Dufresne, 2013; Eichhorn & Young, 2001; Guhados, Wan, & 25

Hutter, 2005; Saito et al., 2013), and the elastic modulus of the crystalline phase is ~20 times larger than that of the amorphous phase (Eichhorn & Young, 2001). Because the tensile strength generally increases as the elastic modulus increases, fibrils with a higher crystallinity could be expected to have a higher tensile strength. Furthermore, the tensile strength could also be influenced by the fraction of the cellulose Iα phase in the crystalline phase of the cellulose because the cellulose Iα phase might have a higher elastic modulus than that of the cellulose Iβ phase based on theoretical modeling approaches (Eichhorn & Davies, 2006; Neyertz et al., 2000). Using the X-ray diffraction method (Experimental 2.4) (Segal et al., 1959; Masahida Wada, Heux, & Sugiyama, 2004), we measured the crystallinity of the N. oceanica cellulose and the fraction of the cellulose Iα phase, respectively. As a result, we showed that the cellulose of the N. oceanica CNFs possesses a very high crystallinity at 91.7%±0.4% and a high fraction of the cellulose Iα phase at 69%±3% which are considerably higher than that of wood and plants (Dufresne, 2013; Gröbe, 1989; Thygesen, Oddershede, Lilholt, Thomsen, & Ståhl, 2005; Masahida Wada et al., 2004), bacteria (Watanabe, Tabuchi, Morinaga, & Yoshinaga, 1998), and tunicate (Kono et al., 2002; Šturcová et al., 2005; Zhao & Li, 2014) and comparable to that of cladophora (Carlsson, Lindh, Strømme, & Mihranyan, 2015; Mihranyan, Llagostera, Karmhag, Strømme, & Ek, 2004; Masahida Wada, Heux, & Sugiyama, 2004; Masahisa Wada et al., 2001), as expected (Table 2).

Table 2. The crystallinity and the fraction of cellulose Iα phase of the microalgal CNFs and other species Species

N. oceanica

Crystallinity

Cellulose Iα

(%)*

fraction (%)**

91.7±0.4

69±3

References

This work 26

Wood and plants Bacteria

47-86

10-36

63-71

61-73

(Dufresne, 2013; Gröbe, 1989; Thygesen et al., 2005; Masahida Wada et al., 2004) (Watanabe et al., 1998) (Carlsson, Lindh, Strømme, & Mihranyan, 2015; Mihranyan, Llagostera,

Cladophora

92-95

54-80

Karmhag, Strømme, & Ek, 2004; Masahida Wada, Heux, & Sugiyama, 2004; Masahisa Wada et al., 2001)

Tunicate

70-95

(Kono et al., 2002; Šturcová et al., 2005;

8

Zhao & Li, 2014)

*Crystallinity index was measured by Segal method (Segal et al., 1959) with X-ray diffraction. **The fraction of cellulose Iα phase was measured by X-ray diffraction or CP/MAS 13C NMR spectroscopy.

When N. oceanica CNFs were also compared with other reinforcements, such as Eglass, aramid, carbon fiber, and steel alloy (Fig. 6), the comparison showed that the mechanical strength of N. oceanica CNFs is still comparable with other reinforcements (Callister & Rethwisch, 2006; Chiao & Chiao, 1982; Dufresne, 2013; Fu, Lauke, Mäder, Yue, & Hu, 2000; Khot et al., 2001; Petrović, Guo, Javni, & Zhang, 2003; Vallittu, 1998) while the density of the N. oceanica CNFs might be comparatively lower than that of the others (Abe & Yano, 2009; Callister & Rethwisch, 2006). Therefore, oceanica CNFs could be adequate for strengthening composites while maintaining their light weight.

27

Fig. 6. The tensile strength of the N. oceanica cellulose nanofibrils (CNFs) and other reinforcements. The red box indicates the tensile strength of the N. oceanica CNFs that was estimated in this work. Black boxes describe the tensile strength of general reinforcements measured by different methods, which were reported previously (Callister & Rethwisch, 2006; Chiao & Chiao, 1982; Dufresne, 2013; Fu et al., 2000; Khot et al., 2001; Petrović et al., 2003; Vallittu, 1998). The three different boundaries, the bottom boundary of a box, the line in a box, and top boundary of a box, correspond to the 25th percentile, median, and 75th percentile, respectively.

4.

Conclusions In summary, our study demonstrates two notable improvements. First, we produced

microalgal CNFs through lipid/protein extraction, purification, and TEMPO-mediated oxidation under gentle mixing without mechanical treatment. The cellulose fibrils obtained from Nannochloropsis oceanica has a few nm in diameter, so they are likely to be a single nanofibril. This result shows that the using microalgae as a NFC source has a potential to reduce 28

the consumption of energy and time. Second, after successfully producing the CNFs from microalgae, we estimated the tensile strength of the NFC fibrils with the sonication-induced scission method to evaluate the applicability of N. oceanica CNFs as a reinforcing filler. However, it was found that there are many buckled fibrils that prevent an accurate measurement of the tensile strength by the sonication-induced scission; thus, we suggested a new approach adapting the ratio of the persistence length to the contour length to choose only straight fibrils for more accurate estimation of the tensile strength. As a result, we choose more straight fibrils with a persistence length at least 5 times larger than that of the contour length, and the tensile strength of the N. oceanica CNFs was then estimated with these straight fibrils. Based on this procedure, we confirmed that the average tensile strength of the N. oceanica CNFs is 3-4 GPa, and this is considerably higher than that of wood CNFs due to their higher crystallinity and higher fraction of cellulose Ⅰα phase. Furthermore, the N. oceanica CNFs have a comparable tensile strength and even a lower density compared with other general reinforcing fillers. Therefore, we expect that the N. oceanica CNFs could be used in various applications that require a high stiffness, lightweight, and biocompatibility.

Acknowledgment This research was supported by the Advanced Biomass R&D Center (ABC) of Global Frontier Project funded by the Ministry of Science, ICT, and Future Planning (ABC-20100029728).

29

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