Bioresource Technology 173 (2014) 415–421

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Enzymatic and acid hydrolysis of Tetraselmis suecica for polysaccharide characterization Azadeh Kermanshahi-pour a,b,1, Toby J. Sommer a, Paul T. Anastas a,c, Julie B. Zimmerman a,b,c,⇑ a

Center for Green Chemistry and Green Engineering at Yale, 225 Prospect Street, New Haven, CT 06520, United States Department of Chemical and Environmental Engineering, Yale University, 9 Hillhouse Avenue, New Haven, CT 06511, United States c School of Forestry and Environmental Studies, Yale University, 195 Prospect Street, New Haven, CT 06511, United States b

h i g h l i g h t s  Cell wall and intracellular polysaccharides of T. suecica are characterized.  Kdo, a sugar with applications in medical research is present in T. suecica.  Cell wall polysaccharide composition is constant at varying nitrate concentrations.  Enzymatic processing of T. suecica for fermentable sugars production is feasible.  T. suecica consist of up to 45% starch and 5% Kdo on dry weight basis.

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Article history: Received 22 July 2014 Received in revised form 8 September 2014 Accepted 11 September 2014 Available online 19 September 2014 Keywords: Microalgae Tetraselmis suecica Pectinase 3-deoxy-D-manno-oct-2-ulosonic acid (Kdo) Polysaccharides

a b s t r a c t Carbohydrate composition of the marine microalgae, Tetraselmis suecica was characterized following acidic and enzymatic hydrolysis. Monitoring intracellular starch as a function of cultivation time at varying nitrate concentrations showed a maximum cellular starch content of 45% of dry biomass when grown under nitrate depleted conditions. Characterization of the cell wall methanolysates using GC/ MS showed that the monosaccharide composition did not change in response to the nitrate concentration and consisted of 54% 3-deoxy-D-manno-oct-2-ulosonic acid (Kdo), 17% 3-deoxy-lyxo-2-heptulosaric acid (Dha), 21% galacturonic acid and 6% galactose. Presence of up to 5% Kdo in the dry weight of T. suecica established in this study demonstrates the potential of the cell wall of this species as a feedstock for Kdo, a sugar that is difficult to obtain by chemical synthesis and that has applications in medicinal chemistry. Ó 2014 Elsevier Ltd. All rights reserved.

1. Introduction Use of microalgal polysaccharides for energy production has been proposed as early as the late 1950s in the process of anaerobic digestion for methane production (Meier, 1955). Conversion of microalgal intracellular and cell wall polysaccharides into fermentable sugars for bioethanol production has been the focus of several works in recent years (Johna et al., 2011). Some species of green microalgae possess a cellulosic cell wall (e.g., Chlorella sp.) (Northcote et al., 1958) and some can accumulate intracellular carbohydrates (e.g., Chlorella vulgaris) (Hirano et al., 1997); therefore, microalgae ⇑ Corresponding author at: School of Forestry and Environmental Studies, Yale University, 195 Prospect Street, New Haven, CT 06511, United States. Tel.: +1 203 432 5215, +1 203 432 9703. E-mail address: [email protected] (J.B. Zimmerman). 1 Current address: Department of Process Engineering and Applied Science, Dalhousie University, Halifax, Nova Scotia B3H 4R2, Canada. http://dx.doi.org/10.1016/j.biortech.2014.09.048 0960-8524/Ó 2014 Elsevier Ltd. All rights reserved.

have been employed as a feedstock for fermentable sugars production via saccharification (Fu et al., 2010; Harun et al., 2010; Hirano et al., 1997) or used in self-fermentation to produce ethanol (Hirano et al., 1997). The enormous diversity of microalgal and plant polysaccharides and their potential industrial applications have motivated research on their compositional characterization (Popper and Tuohy, 2010; U.S. DOE, 2010; Williams and Laurens, 2010). The characterization of polysaccharides often relies on their depolymerization into simpler oligomers using enzymatic or acid hydrolysis, followed by acetylation or silylation for further structure elucidation of the monosaccharides by mass spectrometry techniques (Becker et al., 1995; Blakeney et al., 1983). Development of novel polysaccharide-degrading enzymes (Bauer et al., 2006) and solutionstate 2D NMR spectroscopy for whole plant cell wall analysis (Mansfield et al., 2012) are some of the new advances in polysaccharide characterization.

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Despite identification of thousands of algal species and their recognition as sources of broad spectrum of chemicals including polysaccharides, proteins and triacylglycerols (TAGs), commercial application of microalgae is confined to a few species and products due to low biomass and product yield in microalgal cultivation systems (Raja et al., 2008). A potential route toward broadening the commercial value of microalgal bio-products is to start with the characterization of the biochemical composition of the species that are produced in large scale, aiming to identify new value-added chemicals that can be co-produced with the existing commercial products (Kermanshahi-pour et al., 2013; Yeh, 2011). Tetraselmis suecica (Prasinophyceae, Chlorophyta) is one of the few species of algae that is currently produced for aquaculture feed due to the large amount and high quality of its intracellular protein content. Therefore, T. suecica was chosen for the present study to further explore this species as a source of polysaccharides for fermentation and specialty chemicals. To evaluate, this potential both intracellular and cell wall polysaccharides should be characterized. The main focus was to investigate the possible presence of a valuable monosaccharide, 3-deoxy-D-manno-oct-2-ulosonic acid (Kdo), in the cell wall of T. suecica with applications for antibiotic development research (Camci-Unal et al., 2012). This sugar has been identified in other species of Tetraselmis such as T. striata and T. tetrathele (Becker et al., 1991). However, to the knowledge of the authors, the cell wall of T. suecica has not been previously characterized. Similarly, the effect of nitrate concentration on cell wall polysaccharide composition/content has not yet been investigated. Additionally, T. suecica is known for its rich intracellular polysaccharide content and it is desirable to fully utilize it. The effect of nitrate concentration in cultivation media has been shown to be an important factor that affects accumulation of starch in several microalgae species including T. suecica (Myklestad, 1974; Myklestad et al., 1972; Thomas et al., 1984). This work also extends the knowledge on the profile of cellular starch content and biomass concentration in response to varying the initial nitrate concentrations. 2. Methods 2.1. Cultivation of T. suecica T. suecica UTEX LB 2286 was obtained from UTEX, The Culture Collection of Algae at the University of Austin at Texas. Seawater, used for microalgal growth was collected from Hammonasset beach (New Haven County, Connecticut, United States), filtered through 0.45 and 0.2 micron Millipore sterile filters and pasteurized overnight in an incubator at 73 °C. Microalgae cultivation broth was prepared by dissolving various volumes of Enrichment Solution in 0.75 L of pasteurized seawater to target a final nitrate concentration as N of 3–90 ppm. Enrichment solution was prepared according to the recommended UTEX recipe for enrichment solution for seawater medium. The stock solution was stored in the refrigerator at 4 °C. T. suecica were grown autotrophically in light with an approximate light irradiance of 250 lmol m2 s1 and light/dark cycle of 14/10 h under air and CO2 flow rates of 0.6 and 0.015 mL/min, respectively. Microalgae was recovered from the cultivation media in pellet form by centrifugation at 4000g for 15 min and re-suspension in sodium acetate buffer at the desired pH (ranging from 4 to 5.2) and enzyme load (ranging from 0.005 to 0.01 protein/ microalgae dry weight, g/g) for enzymatic hydrolysis. 2.2. Nitrate measurement Nitrate concentration in cultivation broth was measured using the Manual Nitrate Reductase (Na-R) Nitrate Analysis method

(Nitrate Elimination Company, Inc.). Nitrate standard solutions were prepared in seawater collected and treated the same as samples. 2.3. Microalgal biomass dry weight measurement Triplicate samples of 10 mL were taken aseptically from the photobioreactor during the cultivation. The samples were centrifuged at 10,000 rpm at 4 °C for 10 min. The supernatant was discarded and the pellet was resuspended in 10 mL of DI water and centrifuged again. The rinsing and centrifugation steps were repeated three times to remove the salt and then the final pellet was re-suspended in deionized water and placed in a pre-weighed aluminum dish. The dishes were placed in an oven 70 °C for 48 h. The dishes were cooled and the dry weight was obtained using an XP-26 micro balance (Mettler-Toledo XP26). 2.4. Measurement of protein content of pectinase The protein content of pectinase (p 4716, Sigma Aldrich), which was used for hydrolysis of the whole cells as well as the cell walls, was estimated using the Bradford method with the Bio-Rad Protein Assay. This method relies on a dye reagent concentrate in which a differential color change of a dye occurs in response to various concentrations of protein. The blue–black color generated in the reaction between Coomassie Brilliant Blue G-250 in the dye reagent concentrate and protein was measured by a spectrophotometer (Varian Cary 3E UV–Vis Spectrophotometer using Cary WinUV software) at the wavelength of 595 nm. Bovine serum albumin (BSA) was used as the standard solution. The amount of protein in pectinase was found to be 14.7 g protein/L pectinase with a standard deviation of less than 9% from preparation of various pectinase solutions in deionized water. 2.5. Enzymatic hydrolysis of whole cells and determination of the amount and type of reducing sugar in the enzymatic hydrolysates 50 mM sodium acetate buffer was used as enzymatic buffer solution. The pH of the buffer solution was adjusted to the desired value, ranging from 4 to 5.2 using acetic acid. Pectinase load of 0.005, 0.001 and 0.0075 protein/microalgae (g/g) was used to determine the optimum hydrolysis rate and efficiency. The total concentration of reducing sugars in the enzymatic hydrolysates over the course of the reaction was measured using a standard dinitrosalicylic (DNS) colorimetric method (Miller, 1959). Reducing sugars (e.g., glucose) either have an aldehyde group or are transformable to an isomer that has an aldehyde group. The aldehyde groups reduce the yellow 3,5-dinitrosalicylic acid (DNS) to orange 3-amino-5-nitrosalicylic acid. During the course of hydrolysis reaction, 100 lL of sample was taken from the reaction mixture, placed in 1.5 mL microcentrifuge tubes and was centrifuged (Thermo Scientific™ Sorvall™) at 17000 g and 4 °C. For analysis, 60 lL of the supernatant was mixed with 180 lL of DNS solution in a 1.5 mL microcentrifuge tubes and was heated in boiling water for 10 min. After the samples were cooled to room temperature, 960 lL of DI water was added to each sample. Optical density was measured spectrophotometrically (Varian Cary 3E UV–Vis Spectrophotometer using Cary WinUV software) at 540 nm. Characterization of monosaccharides was carried out by reduction and acetylation of enzymatic hydrolysates according to the procedure of Blakeney et al. (1983) and analyzed using GC/MS. For starch determination, freeze dried T. suecica was hydrolyzed in sodium acetate buffer at pH of 6 using a combination of aamylase (A3403, Sigma Aldrich) and amyloglucosidase (A7420, Sigma Aldrich).

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2.6. GC/MS analysis of acetylated enzymatic hydrolysates of whole cells Aliquots of 1 lL of acetylated enzymatic hydrolysates were analyzed by Agilent GCD G1800C GC–MS with a 6890 Series injector. An HP-5MS capillary column (30 m  0.25 mm I.D. and film thickness of 0.25 lm) was used with helium as the carrier gas at a constant flow rate of 1 mL min1. The injector and detector temperatures were maintained at 250 °C and 280 °C, respectively. The temperature profile consisted of initial column temperature of 130 °C followed by a temperature increase to 220 °C at a rate of 4 °C min1 and another increase to 240 °C at a rate of 20 °C min1. The MS was operated in the electron impact mode with ionization energy of 70 eV. The scan range was set from 45 to 450 m/z. Data were acquired and processed with the Agilent GC ChemStation software. Monosaccharides were identified by comparison of their chromatographic retention times and mass spectra to those of authentic standards. 2.7. Starch assay Portions of 5–6 mg of freeze dried microalgae were hydrolyzed by pectinase until the concentration of reducing sugar produced by the enzymatic process reached a constant value. Microalgae were subsequently recovered from the enzymatic reaction mixture via centrifugation (Heraeus Fresco 17 centrifuge). The starch content of the microalgae paste was determined using an enzymatic assay kit (STA20, Sigma–Aldrich). 2.8. Isolation and purification of the cell wall of T. suecica Cell walls of microalgae grown at initial nitrate concentrations of 68 ppm and 90 ppm were isolated at the stationary phase of growth on day 12 of cultivation when biomass researched its maximum concentration. The cells were centrifuged at 5000g for 10 min at 4 °C and the pellet was further washed with deionized water and stored at 15 °C until the cell wall isolation procedure was performed. Thawed pellets were diluted in deionized water then sonicated with a Branson 250 Digital Sonifier with a flat tip probe at 30% power for 3 min (1 s on, 2 s off). The procedure of cell wall isolation was adapted from Russell (1995) to remove intracellular starch, lipid and soluble proteins and was further modified to remove the protein. To remove the intracellular protein contamination, the cell wall pellet was suspended in 1 mL of high salt buffer solution consisting of 0.5 mM NaCl, 20 mM HEPES buffer, and 0.05% Triton X-100 and then centrifuged at 13,000g for 10 min. This rinsing step was repeated 3 times, followed by frequent rinsing with 6 M guanidine hydrochloride (Sigma, 99%) until the amount of protein in the supernatant, measured at 280 nm (Shimadzu UV-1800 UV/Visible Scanning Spectrophotometer) was negligible. Cell wall samples were frozen at 80 °C and dried in a vacuum concentrator (Savant SpeedVac Plus SC110A). 2.9. Enzymatic hydrolysis of cell wall and determination of total sugar in enzymatic hydrolysates A 3–5 mg portion of the purified cell wall was exposed to pectinase in 4 mL sodium acetate buffer solution at pH of 4 while stirring at the constant temperature of 35 °C. Pectinase load was 0.02 protein/cell wall (g/g). The concentration of total sugar released in this process was measured using the Phenol–Sulfuric Acid colorimetric method (Chow and Landhäusser, 2004; DuBois et al., 1956). A 0.1 mL aliquot of enzymatic hydrolysates was mixed with 200 lL of 4% (w/v) phenol, followed by rapid addition of 0.8 mL concentrated

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sulfuric acid. After 30 min of color development in the dark at room temperature, absorbance was measured at 490 nm (Varian Cary 3E UV–Vis Spectrophotometer using Cary WinUV software). Dextrose (Sigma–Aldrich) was used as a standard to generate the calibration curve. 2.10. Methanolysis of cell wall and whole cells of T. suecica and trimethylsilylation of methanolysates for GC/MS analysis Cell wall (0.9–3.4 mg, weighed using METTLER TOLEDO XP26 micro balance) isolated from whole cells of microalgae, grown at nitrate concentrations of 62 or 90 ppm and whole cells of T. suecica (0.7–2 mg weighed using METTLER TOLEDO XP26 micro balance) grown at nitrate concentration of 90 ppm were subjected to methanolysis using 3 mL 1 M methanolic HCl at 85 °C for 24 h. 1 M methanolic HCl was prepared by dissolving 3 mmol acetyl chloride (99%, Acros Organics) in 3 mL methanol (99.9%, EMD Chemicals). The methanolysis reaction was carried out in 12 mL screw capped vials (LABCO limited, UK) in a reaction block at 85 °C for a period of 24 h. Then the reaction mixture was neutralized with 3 mmol silver carbonate (99.6%, Baker) and incubated with 50 microliters (0.53 mmol) of acetic anhydride (99%, Sigma–Aldrich) for 18 h. Then, the mixture was centrifuged and the supernatant was collected. The pellet was rinsed with methanol and centrifuged. The washing steps were repeated three times and the supernatant of all the steps were combined and evaporated by rotatory evaporation at 35 °C, and then the residue was dried in a dessicator over P2O5 overnight. Derivatization of sugars generated in methanolysis of cell wall (0.9–3.4 mg) and whole cells (0.7–2 mg) was performed using 100 lL (0.38 mmol) N,O-bis-(trimethylsilyl)trifluoroacetamide (BSTFA) (99%, Sigma–Aldrich), 1 lL (0.008 mmol) trimethylchlorosilane (TMCS) (99%, Sigma–Aldrich) and 100 lL (1.24 mmol) anhydrous pyridine (99.8%, Sigma–Aldrich) in screw capped vials for 3 h at 70 °C. Immediately preceding GC–MS analysis, derivatized extracts were evaporated to dryness using nitrogen gas and redissolved in 1 mL (7.6 mmol) of hexane for injection. 2.11. GC/ MS analysis of TMS derivatives of methanolysates of cell wall and whole cells Aliquots of 2 lL of derivatized samples and standards were analyzed using an Agilent 7890A gas chromatograph system and 5975C TAD Series Gas Chromatograph/Mass Selective Detector System with the Triple-Axis Detector. An HP Fast Residual Solvent chromatography column (30 m  0.53 mm I.D. and film thickness of 1 lm) was used with helium (Airgas) as the carrier at a constant flow rate of 1.3 mL min1. The injector and MS source temperatures were maintained at 250 and 230 °C, respectively. The temperature program began with a 2 min hold at 65 °C (2 min), then rising to 300 °C at 6 °C min1, followed by an isothermal hold at 300 °C for 15 min. The MS was operated in the electron impact mode with ionization energy of 70 eV. The scan range was set from 33 to 700 Da. Data were acquired and processed with the Agilent GC ChemStation software. Monosaccharide compounds were identified by comparing their chromatographic retention time and mass spectra with those of authentic standards as well as the reported mass spectra. Compounds were quantified using the total ion current (TIC) peak area, and converted to mass using calibration curves of standards. 2.12. Optical microscope T. suecica in seawater, in diluted seawater and in sodium acetate buffer were visualized using a Nikon Eclipse TE2000 microscope equipped with 40 and 100, 1.4 NA objective lens. Images were

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captured with a Cool Snap HQ CCD camera (Photometrics) using Metamorph software (Molecular Devices). 3. Results and discussion 3.1. Enzymatic hydrolysis of T. suecica Enzymatic hydrolysis has been widely studied as a process for the production of fermentable sugars from biomass waste (Harun and Danquah, 2011; Harun et al., 2010; Johna et al., 2011) and, to a lesser extent, as an analytical method for the depolymerization of polysaccharides for further characterization by GC/MS and LC/ MS (Bauer et al., 2006). Enzymatic treatment of microalgae has mainly been explored from two perspectives: enzymatic hydrolysis of the cell wall to enhance lipid extraction (Fu et al., 2010) and enzymatic hydrolysis of the whole cell for fermentable sugar production (Harun and Danquah, 2011). In this work, enzymatic hydrolysis was used as an analytical procedure for characterization of carbohydrate composition. However, the information obtained here also provides insights for the development of enzymatic process for production of fermentable sugars from T. suecica. Selection of enzymes capable of depolymerization of the polysaccharides of interest and optimization of the enzymatic activity to maximize the hydrolysis efficiency and rate are the primary steps in enzymatic process development (Bauer et al., 2006). The next step involves characterization of the sugars generated to determine the corresponding polysaccharide of origin. Pectinase is known to hydrolyze pectin, a complex polysaccharide present in plant cell walls. The cell walls of several species of T suecica have been characterized and found to consist of complex carbohydrates (Becker et al., 1991, 1998). Therefore, pectinase was selected for hydrolysis of whole cells of T. suecica. Temperature, pH and enzyme load significantly influence enzyme activity and, thus, the rate and efficiency of enzymatic hydrolysis (Fu et al., 2010). An optimization was performed using a two-level factorial design of experiments, shown in Table 1, to determine the effect of temperature, pH and enzyme load on the production of reducing sugar as a proportion of microalgae dry weight (g/g). As shown in Fig. 1, increasing the temperature from 25 °C to 35 °C resulted in an increase in the amount of sugar/microalgae (g/g) released in the hydrolysis step. Changing the pH and enzyme load within the examined range did not affect the amount of maximum sugar/microalgae dry weight (g/g) produced, but hydrolysis was faster at lower pH. Therefore the conditions for the enzymatic hydrolysis of microalgae were selected to be pH 4, 35 °C and a pectinase load of 0.005 protein/microalgae dry weight (g/g). These conditions led to the production of approximately 0.4 reducing sugar/microalgae dry weight (g/g). 3.2. Identification of the origin of the reducing sugar generated in enzymatic hydrolysis A set of experiments was designed to identify the separate contributions of the cell wall and intracellular polysaccharides to the 0.4 reducing sugar/microalgae dry weight (g/g) generated by pectinase hydrolysis. The mass spectra and retention times of the Table 1 Factorial experimental design to determine the optimum enzymatic hydrolysis of T. suecica. Experimental condition

Lower level

Higher level

Center point

Temperature (°C) pH Enzyme load (protein/microalgae, g/g)

25 4 0.005

35 5.2 0.01

30 4.7 0.0075

Fig. 1. Optimization of enzymatic hydrolysis of polysaccharides of T. suecica.

compounds identified in the acetylated enzymatic hydrolysates were consistent with those of glucose (retention time: 19.58 min and characteristics ions: m/z of 115, 145, 187, 217, 259, 289, 361), which suggested that the reducing sugar originated from intracellular starch. Pectinase hydrolysis of starch resulted in 1 g reducing sugar/g starch within 40 h of hydrolysis, indicating that pectinase was capable of complete hydrolysis of starch. Additionally, no starch was detected in pectinase treated microalgae paste using starch assay kit (Sigma SA20). Freeze dried T. suecica, which was treated by a-amylase and amyloglucosidase, a suite of enzymes that are exclusively used for starch determination, led to 0.4 reducing sugar/microalgae dry weight (g/g). All of the above results indicated that the reducing sugar generated in pectinase hydrolysis of T. suecica originated from intracellular starch and that the cell wall, in the process of enzymatic treatment, has been disrupted, which subsequently led to the exposure of intracellular starch to pectinase. 3.3. Insight into cell wall disruption in enzymatic hydrolysis Cell walls are known to act as barriers for access to the intracellular starch. Several studies have shown that disruption of the cell wall using enzymes or mechanical means (e.g., freeze drying, bead beating) enhances extraction of intracellular products from microalgae (Fu et al., 2010; Mendes et al., 2003). In this work, intracellular starch was depolymerized in the enzymatic process indicating that the cell wall was disrupted. To develop an understanding of the underlying reasons for cell wall disruption, the effect of physical properties of the enzymatic buffer solution, such as salt concentration and pH, on T. suecica cell integrity was investigated. The approximate concentration of salt in seawater in which marine microalgae such as T. suecica are cultivated is typically 30–40 g/L. Salt concentration in enzymatic buffer solutions is approximately 4 g/L. To explore the effect of the low osmotic pressure of buffer solution, seawater containing live cells was diluted to reach the salt concentration approximately equal to that of buffer solution and the mixture was stirred using a magnetic stir bar. Comparison of the microscopic image of the live cells in seawater and the cells in diluted sea water revealed that microalgae cells remained intact under the low osmotic condition of diluted seawater as swelling or bursting of the cells was not observed. Gentle magnetic stirring did not mechanically disrupt the cells, either. To explore the effect of

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the mildly acidic buffer solution on cell integrity, pH of the seawater containing live cells was dropped to pH 4 using acetic acid and the solution was mixed, as above, using a magnetic stir bar. Microscopic imaging of the cells under these conditions indicated that the cell wall was completely disrupted, which resulted in the exposure of intracellular starch to enzyme and further hydrolysis. Interaction of the isolated cell wall with sodium acetate buffer at pH 4, without the presence of pectinase, resulted in the production of approximately 0.14 reducing sugar/microalgae (g/g) in approximately 40 h, indicating that the cell wall was partially hydrolyzed due to the mild acidity of the buffer solution. Enzymatic hydrolysis of the cell wall using pectinase led to the production of the same amount of sugar. These results imply that pectinase does not have an observable effect on the hydrolysis of the cell wall of T. suecica as the cell wall appears to be resistant to enzymatic hydrolysis. Rather, it is the effect of pH that leads to the partial hydrolysis of the cell wall. Recalcitrance of the cell wall of some microalgal species (e.g., Chlorella sorokiniana) to enzymatic hydrolysis has previously been observed and was attributed to the high concentration of rhamnose containing polysaccharide (Russell, 1995). 3.4. Effect of freeze drying and mixing intensity on enzymatic hydrolysis of T. suecica To investigate the effect of pretreatment and the intensity of mixing on the hydrolysis of T. suecica, freeze dried and wet microalgae, which were directly recovered from cultivation media were enzymatically processed under vigorous and mild mixing. As seen in Fig. 2, enzymatic hydrolysis led to a similar profile of sugar production under these conditions indicating that no pretreatment or intense mechanical mixing is required to enhance the process of sugar production. These results show the feasibility of the enzymatic processing of T. suecica for production of fermentable sugar without requiring pretreatment prior to enzymatic hydrolysis. In general, renewable feedstocks that do not require any energyintensive mechanical pretreatment prior to enzymatic processing are more preferable due to the lower energy input. 3.5. Effect of nitrate concentration in cultivation media on accumulation of reducing sugar The effect of nitrate concentrations in the cultivation media, ranging from 3 ppm to 90 ppm, on the biomass concentration and the profile of the reducing sugars was investigated as shown in Fig. 3. As nitrate at concentrations of 30–90 ppm was assimilated by T. suecica (Fig. 3a) and biomass concentration increased

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(Fig. 3b), gradual increase in intracellular reducing sugar content (starch) (Fig. 3c) was observed. On the other hand, at 3 ppm nitrate concentration, the amount of reducing sugar in T. suecica was overproduced within only two days. Fig. 3a–d, not only clearly show that cellular starch content was overproduced under nitrate-deplete condition but also demonstrates the correlation of starch with biomass and nitrate concentrations during the batch cultivation. 3.6. Methanolysis of the cell wall and whole cell of T. suecica for polysaccharide characterization Microalgae, which were grown at initial nitrate concentrations of 68 ppm and 90 ppm were harvested on day 12 of cultivation when cells were at the stationary phase of growth (Fig. 3) and their cell wall were isolated and purified according to the method described in Section 2.10 and characterized using GC/MS, described in Section 2.8. As shown in Table 2, cell wall polysaccharide consisted of isomers of Kdo, 3-deoxy-lyxo-2-heptulosaric acid (Dha), galacturonic acid and galactose. These compounds were identified by comparing their fragmentation pattern and retention time with those of standards. Due to the lack of a commercial standard for Dha, the identification of this compound was based on the comparison of its mass spectrometric fragmentation pattern with the one which was previously reported for this compound (Becker et al., 1989). As illustrated in Fig. 4a, cell wall isolated from microalgae grown at initial nitrate concentration of 62 and 90 ppm, respectively found to have similar monosaccharide composition consisting of 54% Kdo, 17% Dha, 21% galacturonic acid and 6% galactose. Characterization of the cell wall of T. striata shed to the media during cell division showed that 42% of the polysaccharides consisted of Kdo (Becker et al., 1989). GC/MS on whole cell methanolysates of T. suecica, grown at 90 ppm initial nitrate concentration and harvested at stationary phase on day 12 of cultivation showed presence of approximately 42% glucose (Fig. 4b). The glucose of the isolated cell wall was below the detection limit (Fig. 4a). Therefore, all the glucose detected in whole cell methanolysates originated from the intracellular starch. As shown in Fig. 4b, the same quantity of glucose was obtained in pectinase hydrolysis of T. suecica, cultivated and harvested under the same condition, which confirms the complete hydrolysis of intracellular starch by pectinase and the accuracy of the compositional analysis. More importantly, Fig. 4b shows that this species consists of approximately 42 wt% glucose as starch and 5 wt% Kdo on dry weight basis of microalgae when grown under the conditions specified earlier. Although, the glucose content is much higher, Kdo is currently over 2 million times more expensive than glucose per mole (Camci-Unal et al., 2012). Due to its difficult chemical synthesis, development of biological system for production of Kdo appears favorable. Biosynthesis of Kdo via a glucose fermentation pathway in engineered Escherichia coli presents such an effort (Camci-Unal et al., 2012). Cultivation strategies to overproduce the cellular content of Kdo and technology development for its economical recovery from microalgal biomass remain to be further explored. 4. Conclusions

Fig. 2. Enzymatic hydrolysis of (a) (h) freeze dried T. suecica mixed using a stir bar and wet T. suecica (b) (4) mixed using a stir bar (c) (s) shaken in an incubatorshaker and (d) (d) while no mixing was provided. Temperature was set to 37 °C under all the above conditions.

Characterization of feedstock to identify potential products of commercial value allows the determination of their most appropriate applications. This work represents an effort to evaluate T. suecica as feedstock for a biorefinery through a comprehensive characterization of the polysaccharide content. Given that T. suecica already has commercial applications in aquaculture industries, utilization of the intracellular starch and cell wall sugars will likely

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Fig. 3. (a) Assimilation of nitrate, (b) biomass concentration profile, (c) reducing sugar content and (d) volumetric reducing sugar during the cultivation of T. suecica at nitrate concentrations as N of 3 (}), 30 ( ), 62 (N) and 90 ( ) ppm.

Table 2 Monosaccharides components of the cell wall and whole cells of T. suecica as TMS (methyl ester) methyl glycoside derivatives. Compound

Molecular mass-TMS (methyl ester) methyl glycoside derivatives

m/z

Retention time (min, a-,b-)

Galacturonic acid

438

Galactose Kdo Dha Glucose

482 554 408 482

Furano-ring: 73,133,217, 438 Pyrano-ring: 73,147,204,217, 438 73, 133, 147, 204, 217 73, 147, 204, 217, 539 73, 204, 217, 259, 349, 361, 393 73, 133, 147, 482

31.82, 34.97 34.98 33.26, 34.19 34.4 38.41,38.75, 39.22, 39.36 33.64,34.52, 35.41, 35.45, 35.51 35.07, 35.7

Fig. 4. (a) Profile of monosaccharide composition of T. suecica cell wall versus the total monosaccharides at 62 and 90 ppm nitrate concentrations, (b) monosaccharide composition of whole cells of T. suecica, grown at 90 ppm initial nitrate concentration.

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Enzymatic and acid hydrolysis of Tetraselmis suecica for polysaccharide characterization.

Carbohydrate composition of the marine microalgae, Tetraselmis suecica was characterized following acidic and enzymatic hydrolysis. Monitoring intrace...
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