Optimization of a wet microalgal lipid extraction procedure for improved lipid recovery for biofuel and bioproduct production

Accepted Manuscript Optimization of a Wet Microalgal Lipid Extraction Procedure for Improved Lipid Recovery for Biofuel and Bioproduct Production Ashik Sathish, Tyler Marlar, Ronald C. Sims PII: DOI: Reference:

S0960-8524(15)00843-3 http://dx.doi.org/10.1016/j.biortech.2015.06.052 BITE 15132

To appear in:

Bioresource Technology

Received Date: Revised Date: Accepted Date:

6 April 2015 10 June 2015 11 June 2015

Please cite this article as: Sathish, A., Marlar, T., Sims, R.C., Optimization of a Wet Microalgal Lipid Extraction Procedure for Improved Lipid Recovery for Biofuel and Bioproduct Production, Bioresource Technology (2015), doi: http://dx.doi.org/10.1016/j.biortech.2015.06.052

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.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Optimization of a Wet Microalgal Lipid Extraction Procedure for Improved Lipid Recovery for Biofuel and Bioproduct Production Ashik Sathish, Tyler Marlar, and Ronald C. Sims a

Department of Biological Engineering, Utah State University, 4105 Old Main Hill, Logan, UT 84322, United States

a

Corresponding author. Tel.: + 1 435 797 2576; Fax: + 1 435 797 1248

Biological Engineering Department, Utah State University, 4105 Old Main Hill, Logan UT 84322-4105 Email address: [email protected]

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Abstract Methods to convert microalgal biomass to bio based fuels and chemicals are limited by several processing and economic hurdles. Research conducted in this study modified/optimized a previously published procedure capable of extracting transesterifiable lipids from wet algal biomass. This optimization resulted in the extraction of 77% of the total transesterifiable lipids, while reducing the amount of materials and temperature required in the procedure. In addition, characterization of side streams generated demonstrated that: (1) the C/N ratio of the residual biomass or lipid extracted (LE) biomass increased to 54.6 versus 10.1 for the original biomass, (2) the aqueous phase generated contains nitrogen, phosphorous, and carbon, and (3) the solid precipitate phase was composed of up to 11.2 wt% nitrogen (70% protein). The ability to isolate algal lipids and the possibility of utilizing generated side streams as products and/or feedstock material for downstream processes helps promote the algal biorefinery concept.

Keywords: Microalgae, Wet Lipid Extraction, Lipid Extracted Algae (LEA), Protein, Bio refinery

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

1. Introduction Addressing the need to reduce global dependence on fossil fuel and crude oil based products has motivated the development of technologies for the production of renewable fuels, chemicals, and bioproducts (Huang et al., 2010; Mata et al., 2010; Wijffels et al., 2010). Several generations of biofuel producing feedstock materials have been researched and developed, with the latest being microalgae due to several advantages microalgae possess as a source of renewable biomass and lipids (Chisti, 2007; Demirbas and Fatih Demirbas, 2011). Current hurdles to processing and transforming algal biomass into fuels and chemicals have included challenges related to the extraction of algal lipids for conversion to biodiesel, dewatering of microbial biomass, and development and isolation of co-products (Halim et al., 2012; Harun et al., 2010; Huang et al., 2010). Several methods exist for the extraction and conversion of algal lipids to biodiesel. However, traditional methods such as solvent based extractions followed by transesterification or direct transesterification methods suffer from their lowered efficiency when performed in the presence of water, and attempting to dry the moisture remaining in the biomass after harvesting requires a significant amount of energy (Collet et al., 2011; Halim et al., 2012). Therefore, additional methods have been researched and developed to extract algal lipids from wet biomass (Halim et al., 2012; Kanda and Li, 2011; Sathish and Sims, 2012). Several methods were developed and successfully demonstrated, however the feasibility of successfully scaling up many of these methods has not been demonstrated at a commercial scale for producing biodiesel alone from algal lipids. Production of biodiesel alone from algal biomass has been considered too costly without the production of additional bioproducts or co-products as additional sources of revenue. For example, interest has increased in the utilization of algal biomass, or lipid extracted algal biomass (LEA biomass), as a protein source for agricultural applications. Other co-products such as high value chemicals and commodities are also being investigated including plastics, nutraceutical compounds, and pre-cursor compounds for the production of other products (Mata et al., 2010; Sun et al., 2011). These co-products can supplement costs of processing and producing fuels from algal biomass.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Thermo-chemical processing of algal biomass is an additional pathway for producing fuels and chemicals. Hydrothermal liquefaction (HTL) has received considerable attention due to its ability to directly process wet algal biomass and convert non-lipid fractions of the biomass into a liquid bio crude oil, via thermal decomposition in an oxygen free environment. However, it is in the early stages of development for application to algal biomass. Developmental issues related to fundamental understanding of the biomass feedstock, thermo-chemical processing, and bio crude oil refining and quality (high feedstock nitrogen content) are challenges that require addressing prior to scale up and commercialization (López Barreiro et al., 2013; Tian et al., 2014). The Wet Lipid Extraction Procedure (WLEP), as previously described by Sathish et al. (Sathish and Sims, 2012), provides a means for the isolation and extraction of lipids from wet algal biomass. In addition to isolating the algal lipids, the WLEP is also capable of fractionating the algal biomass into several streams that can be utilized as bioproducts or as feedstock materials for the production of additional bioproducts or bioprocesses. For this study, the wet lipid extraction procedure (WLEP) was modified with the objectives of (1) maximizing the lipid fraction recovery (extraction and isolation) of algal lipids from the biomass while minimizing consumption of materials and energy and (2) determining the fundamental properties and characteristics of the solid and aqueous fractions generated that can be utilized as is or further processed into valuable chemicals. 2. Materials and Methods 2.1. Chemicals and Reagents Reagents used in this study include ACS grade sulfuric acid and sodium hydroxide obtained from Fisher Scientific (Fair Lawn, NJ) and methanol obtained from Pharmco-AAPER (Brookfield, CT). HPLC grade hexanes was obtained from Fisher Chemicals (Pittsburgh, PA). Fatty acid methyl ester (FAME) standards were obtained from Supelco Analytical (Bellefonte, PA). All macronutrients used for algal growth media were laboratory or ACS grade, while micronutrients were technical or laboratory grade. 2.2. Algae Growth and Collection

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Algal biomass used in this study was obtained from two sources. The first source was an indoor 100 L laboratory open raceway system, with a single paddlewheel to maintain suspension of the algal cells. Lighting was provided by GE plant and aquarium bulbs and one high pressure sodium bulb. The average photosynthetic photon flux density was measured as 297 µmol m-2 s-1 (from various positions within the raceway) (Young, 2011). The initial inoculum of algae was obtained from the City of Logan Environmental Department’s Logan Lagoons Municipal wastewater treatment facility and grown in a modified SE medium (Li et al., 2008) and was composed mainly of Chlorella and Scenedesmus species of algae (Griffiths, 2009). The medium was prepared with the following components per liter: 0.85 g NaNO3, 0.35 g KH2PO4, 0.15 g MgSO4∙7H2O, 0.15 g K2HPO4, 50 mg CaCl2∙2H2O, 50 mg NaCl, 15 mg Ammonium Ferric Citrate, and 1 mL of a trace solution containing the following per liter: 2.86 g H3BO3, 1.81 g MnCl2∙4H2O, 0.22 g ZnSO4∙7H2O, 79 mg CuSO4∙5H2O, and 39 mg (NH4)6Mo7O24∙4H2O. Algal biomass was harvested using a Cepa Z-41 continuous bowl centrifuge when the culture reached stationary phase. The harvested algal biomass was immediately frozen at -80oC and lyophilized. The second source was algal biomass harvested by centrifugation directly from the Logan Lagoons Wastewater treatment plant using an Alpha Laval Clara 80 clarifier. The collected biomass was oven dried and ground using a Strand Lab Mill Grinder S102DS grinder. After grinding, the algal biomass was stored at -20oC. 2.3. Modification of the Wet Lipid Extraction Procedure (WLEP) The first objective of the study focused on modifying the WLEP procedure, previously described by Sathish et al. (Sathish and Sims, 2012), to maximize the isolation of algal lipids as a separate lipid phase. The WLEP is capable of fractionating the algal biomass into several streams through acid and base hydrolysis steps followed by several phase separation steps, as illustrated in Figure 1 (Sathish and Sims, 2012). Through these steps the algal lipids are extracted from the biomass and isolated as a separate phase. For this study lyophilized algal biomass was utilized to better control the solid to liquid ratio during the procedures to ensure comparable and consistent results. A previous study has established application of this procedure to wet algal biomass (Sathish and Sims, 2012).

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

2.3.1. Acid and Base Hydrolysis Steps Development of the modified WLEP involved reducing the amount of materials used for the acid and base hydrolysis of the algal biomass, which utilizes sulfuric acid and sodium hydroxide solutions. Temperature and time of the hydrolysis steps were also varied with the intent of reducing energy demand. Table 1 identifies the conditions tested. For the different concentrations of the acidic and basic solutions used, the levels are referred to as full (F), half (H), or quarter (Q), corresponding to the strongest to weakest concentrations respectively. For each condition tested three test tubes were prepared with 100 mg of dry algal biomass added to each. One mL of sulfuric acid solution of specified concentration was added to the biomass in each test tube and the test tubes were sealed using PTFE lined screw caps, mixed, and placed in a Hach DRB-200 heat block set to the specified temperature and time. Upon completion of the acid hydrolysis step, the test tubes were removed from the heat block and 0.75 mL of basic solution of specified concentration was added directly to the reaction mixture. The test tubes were again sealed, mixed, and placed in the heat block for the predetermined amount of time. For both the acid and base steps, the tubes were gently mixed every 15 minutes to keep the biomass suspended during the reaction. After completion of both the acid and base hydrolysis steps, the test tubes were removed from the heat block and allowed to cool. Once cooled, the test tubes were centrifuged using a Thermo-Sorvall RC 6 Plus centrifuge at 6,500 RPM for 10 minutes. The supernatant from each test tube was collected in a separate tube and the pelleted biomass was washed with 1 mL of DDI (Distilled Deionized) water using a vortex mixer for 10 seconds. The washed biomass was re-centrifuged and the supernatant collected and added to the original supernatant. The test tubes containing the residual hydrolyzed biomass were immediately frozen at -80oC and subsequently lyophilized. To the supernatant collected, 1.0 mL of a 1 M sulfuric acid solution was added to form a solid precipitate. This suspension was centrifuged to pellet the precipitated solid phase and collect the supernatant separately. Both phases were immediately frozen at -80oC for lyophilization and analysis. This procedure was repeated for all of the conditions stated in Table 1 and all three phases from each

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

sample of biomass were collected and immediately frozen, freeze dried, and analyzed for lipid content (as described in Section 2.6.1). 2.3.2. Washing of the Residual Hydrolyzed Algal Biomass Results from a previous study indicated that after the base hydrolysis step and a single wash of the residual biomass with DDI water, lipids remained in the residual hydrolyzed biomass phase (Sathish and Sims, 2012). To improve the recovery of transesterifiable lipids from the hydrolyzed biomass phase, variations of the wash procedure were evaluated. For each wash 1 mL of liquid was used, DDI water or a 0.1 M sodium hydroxide solution, and the pellet was washed a total of 4 separate times and centrifuged as described in the previous section. The supernatant from each wash was collected in a separate test tube and the sample frozen at -80oC, freeze dried, and analyzed for transesterifiable lipid content. The hydrolyzed biomass was also collected for analysis as well. 2.3.3. Separation of Lipids from the Precipitated Protein Phase The isolation of transesterifiable lipids as a separate phase, using the WLEP is accomplished by the separation of lipids, as free fatty acids, from the solid precipitate that is formed after the acid and base hydrolysis steps (Sathish and Sims, 2012). In this study the procedure was varied to improve the separation of lipids from the precipitate phase into the solvent phase by altering the separation procedure. This involved changing the temperature, time, and the number of times the separation was repeated (number of cycles) to evaluate changes in the efficiency of lipid isolation. These variations are described in further detail below. For this evaluation, the lyophilized precipitated solid phase generated from 100 mg of dry algal biomass, using a single condition (Full, 60 min, 75oC from Table 1) was contacted with 2 mL of fresh hexanes. Contact time was set at 15 minutes with 10 seconds of mixing by a vortex mixer every 5 minutes at the maximum RPM. After 15 minutes the suspension was cooled and centrifuged to collect the lipid containing solvent phase in a separate test tube using a glass gas tight glass syringe. This procedure was repeated a total of 5 times using 2 mL of fresh hexanes for each of the five cycles. The collected solvent phases from each extraction cycle was evaporated under a filtered air stream and the residual lipid

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

material analyzed for transesterifiable lipid content. This procedure was repeated using three different separation temperatures 75, 90, and 105oC. For each temperature three samples of algal biomass was processed using conditions of (F, 60, 75oC) to generate the solid precipitate phase, which was subsequently used for this portion of the experiment. 2.4. Generation of Side Streams for Characterization Algal biomass harvested directly from the Logan City Wastewater Treatment Lagoons system was processed using the WLEP with the goal of generating larger quantities of the side streams for analysis and characterization. In this case 50 g of dried algal biomass harvested from the wastewater treatment facility was processed as described previously (Anthony et al., 2013). Conditions used were the (F, 30, 90oC) and the resulting aqueous phase and solid precipitate phases were collected for analysis (Figure 1). Side streams generated were treated as follows: (1) the residual hydrolyzed biomass was immediately frozen at -80oC and lyophilized, (2) the aqueous phase was stored at 4oC and filtered using a Whatman GF/B filters prior to analysis, and (3) the solid precipitate phase was collected and immediately frozen at -80oC and lyophilized. For analysis of the protein content of the solid precipitated phase the material was treated further: After lyophilization, the dried powder was washed with DDI water three times to remove soluble salts (mainly sodium sulfate) and the powder was collected and re-lyophilized. Once dried the lipids associated with the solid precipitate phase was separated by boiling the solid phase in hexanes for approximately 1 hour with a reflux condenser and filtered to separate the solid phase from the solvent. The solids were allowed to air dry to remove any residual hexanes before being stored in sealed containers for analysis. 2.5. In situ Transesterification Procedure In situ transesterification of lyophilized algal biomass was performed: (1) to determine the maximum biodiesel (Fatty Acid Methyl Ester or FAME) yield obtainable from the harvested algal biomass as a control value and (2) to quantify the mass of transesterifiable lipids in the various intermediate phases generated during the WLEP. The procedure has been previously described by Sathish

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

et al. 2012 (Sathish and Sims, 2012). Fatty acid methyl esters (FAMEs), generated from the algal biomass or intermediate phases via the reaction with methanol and sulfuric acid, were extracted into hexanes for analysis by gas chromatography (GC). 2.6. Transesterifiable Lipid Analysis by Gas Chromatography (GC) The solvent (hexanes) phases containing FAMEs described in section 2.5 were analyzed using an Agilent 7890B equipped with a FID detector to quantify the mass of FAMEs. An HP-INNOWax column (30 m x 0.25 mm ID x 0.25 µm film thickness) was used to separate the individual FAMEs using helium as the carrier gas flowing at a constant 4 mL/min. The front inlet was operated in split less mode with an initial temperature of 60oC for 0.1 min and heated at a rate of 720oC/min to 250oC for 5 minutes with the injection volume set to 1µL. Initial temperature of the oven was held at 60oC for 4 min then heated at a rate of 15oC/min to 250oC and held for 5 min. The FID detector was maintained at 250oC for the duration of the analysis. Calibration of the GC system for FAME quantification and identification was performed as described previously using a Supelco Analytical FAME standard (C8 to C24) (Sathish and Sims, 2012). 2.7. Analytical Methods Associated with WLEP Stream Characterization 2.7.1. Chemical Characterization of the Aqueous Phase Analysis of the aqueous phase was performed by the Utah State University Analytical Laboratory (USUAL). The aqueous phase was analyzed for both total nitrogen content (ammonium nitrogen and nitrate nitrogen) and total phosphorous. Nitrogen analysis was performed via Lachet while phosphorous was measured using elemental analysis by ICP. Measurement of the total carbon content of the aqueous phase was also performed by the USUAL by combustion using a Leco instrument. All values are reported as averages of duplicate measurements. 2.7.2. Analysis of the Precipitated Solid Phase Analysis of the precipitate solid phase was performed by the Utah State University Analytical Laboratory. Prepared precipitate material was provided the USUAL, where it was analyzed for total nitrogen content via combustion using a Leco instrument.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

2.7.3. Elemental Analysis of Hydrolyzed Biomass Phase Chemical analysis of the harvested algal biomass from the wastewater treatment plant and the residual hydrolyzed algal biomass phase generated from that biomass was performed using a Thermo Scientific FLASH 2000 Organic Elemental Analyzer. The furnace temperature was set to 950⁰C with the oven temperature maintained at 75⁰C. Helium was utilized as a carrier gas at a flow rate of 140 mL/min as well as the reference gas at a constant 100 ml/min. Oxygen was supplied at a flow rate of 250 mL/min. 2.8. Statistical Analysis of Data Data generated from Section 2.3.1 of this study was analyzed using SAS statistical software version 9.4 (Cary, NC). Section 2.3.1 was designed as a 3x3x4 completely randomized factorial experiment with acid and base concentration (F, H, and Q), temperature (50, 75, and 90 oC), and time (15, 30, 45, and 60 minutes) as the 3 factors with triplicate measurements for each factor combination. A three way ANOVA analysis of this data was performed after checking for model assumptions of normal residual distribution and constant variance. An alpha value of 0.05 was used to determine significant differences between all the factor combinations tested using the generated Tukey adjusted p-values. 3. Results and Discussion 3.1. Variation of Acid and Base Hydrolysis Steps of the WLEP The first objective of this study was to improve the extraction, or recovery, of transesterifiable lipids from algal biomass by modifying the original WLEP. Modifications made to the acid and base hydrolysis steps are detailed in Table 1. The recovery of algal lipids from the biomass was determined as the mass of FAMEs measured by GC in the residual hydrolyzed biomass phase, and this value was compared to the total FAMEs measured from the control biomass (lyophilized algal biomass). This ratio is defined as the extraction efficiency, where a lower percentage of FAMEs remaining in the biomass corresponds to a higher percentage extraction from the biomass. The measured transesterifiable lipid content of the control algal biomass was 13.68 ± 0.07 wt%, based on six replicates. Figure 2 presents the measured extraction of lipids from the algal biomass using the hydrolysis conditions presented in Table 1.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

The most significant trend observed from the data in Figure 2 is the increase in the percentage of transesterifiable lipids that were extracted as the temperature of the hydrolysis reaction increases, i.e., a decrease in the percentage of lipids that remained associated with the algal biomass. In addition, for most of the conditions shown in Figure 2, as the hydrolysis reaction time increased the amount of transesterifiable lipids remaining in the residual biomass phase decreased, i.e., the fraction extracted increased. Only two of the conditions resulted in the opposite trend. Based on the slopes from the linear regression of each of the curves in Figure 2 with respect to time versus percentage of lipid remaining in the residual biomass, the two conditions that resulted in a positive slope were the 90oC Full and 90oC Half series. These can be considered the most aggressive conditions, due to the high concentrations of acid and base used and the high temperature. At those conditions, increasing hydrolysis reaction time led to a detrimental effect on the extraction of the algal lipids. However, the slope become negative when the temperature was decreased to 75oC and the acid and base concentrations were between the Full and Half concentrations, indicating a possible optimum between these two sets of conditions. The correlation between reaction time and lipid extraction is supported in Figure 3, which shows that the four most effective conditions used were from the 75oC Half and Full acid and base conditions. When the data presented in Figure 2 is arranged from lowest to highest measured FAMEs remaining in the hydrolyzed biomass across all conditions, the effect of the different conditions can be ranked from most effective to least effective as shown in Figure 3. The most effective condition tested in extracting algal lipids from the biomass during the hydrolysis steps was F, 60 min, 75oC. Using these conditions up to 77% of the algal lipids were extracted from the algal biomass. Statistical analysis of this data using SAS version 9.4 indicated that there were five conditions that were statistically equivalent. These five conditions are the first five conditions on the x-axis in Figure 3 from left to right. Data generated from this stage of the study illustrates the ability to modify the WLEP and achieve high rates of lipid extraction from the algal biomass. The five most effective conditions, as concluded by statistical analysis of the data presented in Figures 2 and 3, illustrates the possibility of reducing both

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

material and energy requirements of the WLEP. The original WLEP required conditions of (F, 30, 90oC), while the top four conditions tested in this study required a temperature of 75oC. This represents a reduction in the thermal energy required. In addition, one of the conditions evaluated (H, 30, 75oC) uses half of the acid and base as the original WLEP. A 50% reduction in the materials consumed in the form of acid and base could lead to substantial reduction in the cost of the procedure. Just outside of the top five, is a condition that requires a quarter of the acid and base (Q, 60, 90oC). Although this condition was not in the top five, it was able to extract approximately 70% of the transesterifiable lipids from the algal biomass, while reducing the acid and base consumption by 75%. 3.2. Washing of the Residual Hydrolyzed Algal Biomass for Additional Lipid Recovery Following the acid and base hydrolysis steps of the procedure, the residual hydrolyzed algal biomass was collected as Stream 1 (Figure 1). This phase contained residual lipids that remained associated with the hydrolyzed biomass. This study utilized two approaches to attempt the recovery of the residual transesterifiable lipids, by washing with DDI water or a dilute solution of sodium hydroxide. The use of dilute sodium hydroxide was motivated by the need to maintain a basic environment to ensure lipids remained in their saponified form. This would allow them to remain more soluble in an aqueous environment over their free fatty acid form. Analysis of the liquid supernatant phase after each washing step for transesterifiable lipids resulted in the data presented in Figure 4A, after comparing to the mass of transesterifiable lipids in the hydrolyzed biomass. After the first three washes, the use of DDI water versus the basic 0.1M sodium hydroxide solution resulted in statistically the same mass of lipids recovered from the residual biomass based on standard deviation. The first wash in both cases resulted in the highest recovery of lipids from the residual biomass. However, after the third wash the DDI water outperformed the basic solution used. After four washes using DDI water, recovery of additional lipids was not achieved. Without any washing of the residual biomass nearly 35% of the total transesterifiable lipids would remain un-extracted in the hydrolyzed biomass. Therefore, washing is necessary to recover a

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

significant portion of the transesterifiable lipids. These results are consistent with previous results after one DDI wash (Sathish and Sims, 2012). Attempts to recover additional lipids from the residual biomass may require an organic solvent or mixture of water and a solvent, such as ethanol. The addition of a solvent such as ethanol would also necessitate an additional separation stage to recover solvent used from the solvent and water mixture. The effects of utilizing solvents or solvent mixtures is an area of future study in recovering additional lipids from this stream and was not evaluated in this study. 3.3. Separation of Lipids from the Precipitated Protein Phase Isolation of the algal lipids for conversion to FAMEs is the final step of the procedure and is accomplished by the separation of the lipids from the solid precipitate phase (Stream 3 Figure 1) using hexanes. The objective of this portion of the study was to observe a reduction in the mass of transesterifiable lipids remaining within Stream 3 as the separation parameters were changed. These parameters were the temperature and the number of separation cycles. Figure 4B illustrates the reduction of transesterifiable lipids in the precipitate phase as the number of solvent separation cycles increase at various temperatures. Previous results had demonstrated that one separation cycle using 5 mL of hexanes, 15 minute cycle time, and 90oC resulted in approximately 17% of the total transesterifiable lipids in the algal biomass remaining unseparated from the solid precipitate phase. Separation of the lipids from the precipitated fraction in this study was accomplished by contacting the protein phase with 2 mL of hexanes for the specified time, temperature, and number of cycles. After five separation cycles the transesterifiable lipids remaining in the protein phase was significantly lower than if only one cycle was used. However, performing more than three separation cycles did not result in significant recovery of additional transesterifiable lipids for all three temperatures evaluated. This modification to the WLEP improved separation of lipids from the precipitated solid phase from approximately 78% with the original WLEP (Sathish and Sims, 2012) to 99% with the modified WLEP, with respect to the total transesterifiable lipids in the precipitated solids phase prior to separation. 3.4. Modified Wet Lipid Extraction Procedure (MWLEP)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Variations of the wet lipid extraction procedure described in the previous sections were combined into one procedure to evaluate the effectiveness of the improved parameters. The modified procedure (MWLEP) included the best determined conditions for the hydrolysis reactions, as stated in Section 3.1 (F, 60 min, 75oC), the increased number of DDI water washes of the residual biomass (Section 3.2), and the procedure for separating the lipids from the protein phase three separate times using a volume of 2 mL of hexanes each time at a temperature of 75oC (Section 3.3). The results from the MWLEP are compared to results generated from the original WLEP conditions as presented in a previous study (Sathish and Sims, 2012), in Figure 4C. Most notable is the improvement in the percentage of lipids extracted from the precipitate phase, i.e., reduction in the mass of transesterifiable lipids remaining in the precipitate phase. This contributed significantly in the procedure’s improvement in isolating lipids. However, the transesterifiable lipid content of the residual biomass remained unchanged between the MWLEP and WLEP. Solvents may be able to recover additional lipids, but this was not evaluated. Although these lipids remain in the residual biomass, they can be utilized in downstream processes by other microbial processes for example. The aqueous phase retained approximately 5% of the transesterifiable lipids in the algal biomass. Previous results indicated approximately 1% remained in the aqueous phase and this reduced the mass of lipids in the final lipid phase. Application of this procedure to wet algal biomass requires adjusting the amount of sulfuric acid and sodium hydroxide added to account for the water present in the biomass since these solutions are water based. This was verified by comparing the results of processing lyophilized biomass versus rehydrated biomass using the (F, 30 min, 90oC) condition. Rehydration of dried algal biomass was achieved by the addition of DDI water to lyophilized biomass to recreate an algal slurry with 80% moisture. This mixture was allowed to set for 30 minutes prior to further processing using the stated condition. Measured transesterifiable lipids remaining in the residual biomass phase was on average (of triplicates) 29.43 ± 1.34% for the dry algal biomass and 29.98 ± 0.73% for the rehydrated algal biomass.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

The final result of the MWLEP is an isolation of up to 71.8 ± 1.44% of the total transesterifiable algal lipids as a separate lipid fraction. This is a significant improvement over the previous procedure capable of isolating 60.55 ± 7.80% (Sathish and Sims, 2012), relative to the total lipids accounted for in all four streams generated. In addition, 77% of the transesterifiable lipids were extracted from the algal biomass after the acid and base hydrolysis steps. This improvement in the isolation of the algal lipids over the original procedure was possible with a reduction in the temperature of the procedure and equivalent results can be achieved with reduced acid and base usage as well. This can lead to potential cost benefits in the processing of algal biomass. 3.5. Characterization of Solid and Aqueous Streams Generated from the WLEP 3.5.1. Residual Hydrolyzed Biomass (Stream 1) The algal biomass that is processed is initially subjected to acid and base hydrolysis steps at elevated temperatures, providing conditions for the hydrolysis of various components of the algal biomass. This can be advantageous in downstream processes where biomass is utilized as a substrate for the production of other bio-products. An example would the production of acetone, butanol, and ethanol (ABE) via fermentation of algal biomass as a substrate by Clostridium sp. Initial pre-treatment of the algal biomass resulted in increased yields of ABE when a similar pre-treatment is applied to the algal biomass (Ellis et al., 2012). Processes such as anaerobic digestion are also enhanced with some form of pre-treatment of the algal biomass to degrade its structure, which is possible after the hydrolysis of the biomass using both an acid and base (González-Fernández et al., 2012; Passos et al., 2013). Algae typically contain a high proportion of protein leading to a carbon to nitrogen ratio (C/N) ratio between 3 and 9, depending on the species and culture conditions (Prajapati et al., 2013). Through the WLEP the nitrogen content of the algal biomass is reduced resulting in an increase in the C/N ratio from a value of 10.1 (original algal biomass) to 54.6 after the acid and base hydrolysis steps. Hydrolysis of the algal biomass solubilizes nitrogen into the generated aqueous phase and separates protein from the biomass (Section 3.5.3), which reduces the nitrogen content of the residual hydrolyzed biomass. Figure 5

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

illustrates this reduction based on the CHNS analysis of both the unprocessed algal biomass and the residual hydrolyzed algal biomass (Stream 1). The C/N ratio is critical in many processes that utilizes biomass for the production of fuels and chemicals. In anaerobic digestion for example, low (C/N) ratios can limit methane production and may require co-digestion of the algal biomass with a carbon rich stream to help increase the C/N ratio (Angelidaki and Ahring, 1993; Chen et al., 2008; Prajapati et al., 2013). The production of bio crude oil from thermo-chemical processes such as pyrolysis and hydrothermal liquefaction (HTL) are also affected negatively by the presence of high concentrations of nitrogen in the processed feedstock material (Du et al., 2013; Kim et al., 2014; López Barreiro et al., 2013; Tian et al., 2014). Nitrogen in the feedstock biomass leads to the formation of nitrogenous compounds in the bio crude oil resulting in instability of the oil, degradation of performance when utilized as a fuel, and higher emissions of nitrous oxides when combusted (Du et al., 2013; Kim et al., 2014). Removal of nitrogen from the algal biomass provides a residual biomass phase that is significantly lower in nitrogen content, which could be beneficial for producing a higher quality bio crude oil. 3.5.2. Aqueous Phase (Stream 2) The aqueous phase generated from this procedure was characterized to determine the concentration of nitrogen, phosphorous, carbon within the liquid phase. Analysis performed by the USUAL demonstrated that the aqueous phase contained 0.23% (m/v) nitrogen or 2.3 g/L N, 0.02% (m/v) phosphorous or 200 mg/L P, and 0.27% (m/v) of carbon or 2.7 g/L C. These reported values are averages of duplicate measurements. The SE medium used for the growth of algae in this study contains 141 mg/L N and 106 mg/L P, when compared, the aqueous phase contains a greater concentration of both nutrients. Previous research in utilizing aqueous co-products from algal biofuels production has centered on the aqueous solution produced via the hydrothermal liquefaction (HTL) of algal biomass (López Barreiro et al., 2013; Tian et al., 2014). Several studies have investigated the capability of HTL derived aqueous phase to support the growth of E.coli (as well as Pseudomonas putida and Saccharomyces cerevisiae Nelson et al.) (Nelson et al., 2013) and microalgae (Biller et al., 2012) as a method of nutrient recovery

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

and to increase carbon utilization efficiency of the whole process. However, the major limitation to this approach is the presence of toxic compounds such as phenols present in the HTL aqueous phase, which can inhibit microbial growth. Studies utilizing the aqueous fraction from HTL processing required significant dilution of the liquid prior to growth of microorganism to reduce the concentration of these inhibitory compounds (Biller et al., 2012; Nelson et al., 2013). Previous studies have established that the aqueous phase generated in this study is capable of supporting the growth of E.coli for the production of Polyhydroxybutyrate (PHB) (Sathish et al., 2014) and has also been observed to support the growth of algal biomass (unpublished results). Both were achieved without modification other than pH adjustment. Results also indicated that growth of E.coli in the aqueous phase was limited by dissolved carbon, rather than any inhibiting substance within the medium and therefore no dilution of the aqueous phase was necessary (Sathish et al., 2014). Use of this aqueous phase as a growth medium for the production of bacterial biomass or microalgae provides an opportunity to generate additional bioproducts, and/or, recycle nutrients (Anthony et al., 2013). 3.5.3. Solid Precipitate Phase (Stream 3): High Nitrogen/Protein Stream Evaluation of the precipitated solid phase determined that the solids consisted of 11.2% nitrogen by mass (corresponding to a 70% crude protein content based on 6.25 factor) (Du et al., 2013). Due to the high estimated protein content of this solid phase, this protein fraction has potential as a feed material for aquaculture and for agricultural applications. A clear need is present for alternative protein sources in aquaculture as fish meal production is declining and prices are increasing while fish stocks used for fish meal production are also decreasing (Gatlin et al., 2007; Li et al., 2014; Lum et al., 2013). Interest in alternative protein sources for animal feeds is also growing due to the increasing cost of traditional protein sources such as soybean meal (Campos et al., 2014). Several studies have been performed that illustrate the ability to utilize defatted or lipid extracted algal (or cyanobacterial) biomass as a protein source for animal feed (Becker, 2007; Lodge-Ivey et al., 2014; Lum et al., 2013).The ability to produce a concentrated protein material from algal biomass, while generating feedstock material for additional

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

bioproducts would be an additional advantage rather than using the whole biomass or lipid extracted algal biomass as a protein source. Additionally, if the lipids are not separated from the protein phase, approximately 70% of the total transesterifiable lipids in the algal biomass would be associated with the protein phase (Figure 6). Animal and fish feeds require a certain proportion of lipid or oil content and the presence of these lipids as free fatty acids in the protein precipitate fraction could be advantageous in these applications as well as potentially increase the value of the protein phase as well (Gatlin et al., 2007; Li et al., 2014; Lum et al., 2013). 3.6. Techno-Economic Considerations Although a detailed techno-economic analysis of the presented procedure is outside the scope of this study, a brief discussion is provided in terms of materials and energy required in this process. Sulfuric acid and sodium hydroxide are raw materials consumed at the highest rate in this procedure and is utilized in the hydrolysis and protein phase precipitation steps. For the different concentrations used in this study, the amount of acid and base utilized per kg of dry (or dry equivalent) algal biomass for the Full, Half, and Quarter conditions is 1.96 kg H2SO4 and 1.50 kg NaOH, 0.98 kg H2SO4 and 0.75 kg NaOH, and 0.49 kg H2SO4 and 0.37 kg NaOH respectively for each condition. The five most effective conditions tested were all statistically equivalent and utilized both the Full and Half conditions, while the sixth most effective condition (second most effective group) used the Quarter concentration (Figure 3). Based on the bulk cost associated with these materials the Full, Half, or Quarter conditions can be chosen based on the desired result or product and cost. Hexanes usage was 60 L per kg of dry algal biomass for the described procedure (three separation stages using 20 L each). However, hexanes can be recovered by distillation effectively to minimize the overall consumption of the solvent. Additionally, the volume of hexanes used can be reduced to a volumetric ratio of 0.5 (solvent to solid phase) (Stephenson et al., 2010), which would reduce the hexanes required significantly. This ratio was not evaluated for in this study and can be evaluated as the procedure is scaled.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Highest energy demands in this procedure can be expected during the separation steps, heating for the hydrolysis of the algal biomass, lipid separation stage using hexane, and the recovery of hexanes. The method of separation presented in this study was centrifugation, which was used due to its ease of use in the laboratory. Centrifugation, which is an energy intensive process can be replaced by several other methods of separation. For example, it was observed that the protein phase (Stream 3) when precipitated is capable of settling quickly, effectively separating itself from the liquid phase. Therefore, sedimentation could be a feasible alternative to centrifugation, or the two processes can be used simultaneously to help reduce the volumetric flow through a centrifuge and thus minimize energy demand. A preliminary calculation of the energy consumption was performed based on just the thermal energy required for the algae hydrolysis and lipid separation stages of the procedure per kg of dry algal biomass (separation steps were not included) for the best case conditions from this study. For the hydrolysis stage, calculations were based on the density of the corresponding solution, specific heat capacity, and temperature of the hydrolysis (heated from 20oC). Energy required per kg of dry algal biomass was calculated as 3.7 MJ for the hydrolysis step for this specific condition. Heat required for the hexane lipid separation stage of the procedure was calculated based on the volume of 60 L per kg of dry algal biomass processed as well as the volumetric 0.5 solvent to solids ratio proposed by Stephenson et al. (Stephenson et al., 2010), where the density of the algal biomass slurry was assumed to be one just for this evaluation. The calculation was based on heating the hexane from 30oC to the extraction temperature of 75oC, followed by cooling to 30oC (during the protein phase solid separation), and final vaporization of the hexane to recover the solvent. The specific heat capacity and latent heat of vaporization of hexane were utilized for the calculation. Utilization of 60 L of hexane per kg of dry algal biomass required 21.7 MJ per kg of dry algae processed, while the Stephenson et al. solvent ratio would require 0.18 MJ (this value could be lower due to the solvent contacting the fractionated protein phase and not the whole algal biomass in this procedure). This decrease can be attributed to the difference in mass of hexane used in each case.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

The total energy required for both the hydrolysis and hexane lipid separation steps would require 3.9 MJ using the 0.5 volumetric hexanes solvent to solid ratio (Stephenson et al., 2010) or 25.4 MJ when using the higher hexane ratio used in this study per kg of dry algal biomass. In comparison, one kg of dry equivalent algal biomass with 20% solids content would require 10.4 MJ of heat to just completely dry the algal biomass, if a traditional solvent based extraction was utilized. A complete mass and energy balance would need to be developed for a detailed analysis and coupled with the associated costs to begin developing a techno-economic analysis. In addition, the ability to produce several bioproducts from this process provides additional flexibility that can be evaluated in a detailed techno-economic analysis. 4. Conclusion Results obtained demonstrated recovery of 71.8 ± 1.44% of the transesterifiable lipids in the algal biomass with a 77% extraction rate from the algal biomass via modification of the WLEP. Characterization determined that (1) the residual hydrolyzed biomass has a higher C/N ratio after processing, (2) the aqueous phase generated contains 2.3 g/L N, 300 mg/L P, and 2.7 g/L C, and (3) the precipitated solid phase contains an estimated 70% protein by mass. Isolation of transesterifiable algal lipids for fuel and/or product production while generating additional material as bioproducts or feedstock material aids in developing the algal biorefinery concept. Acknowledgements The authors would like to acknowledge the help of Brian Smith, Jordan Morley, Charles Roberts, Chris Peterson, Justin Jones, and Guevara Che Nyendu. As well as the Utah Water Research Laboratory (Tessa Guy and Joan McLean), the Utah State University Analytical Laboratory, and Utah State University’s Synthetic Biomanufacturing Institute.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

References: 1. Angelidaki, I., Ahring, B.K., 1993. Thermophilic anaerobic digestion of livestock waste: the effect of ammonia. Appl. Microbiol. Biotechnol. 38, 560–564. 2. Anthony, R.J., Ellis, J.T., Sathish, A., Rahman, A., Miller, C.D., Sims, R.C., 2013. Effect of coagulant/flocculants on bioproducts from microalgae. Bioresour. Technol. 149, 65–70. 3. Becker, E.W., 2007. Micro-algae as a source of protein. Biotechnol. Adv. 25, 207–210. 4. Biller, P., Ross, A.B., Skill, S.C., Lea-Langton, A., Balasundaram, B., Hall, C., Riley, R., Llewellyn, C.A., 2012. Nutrient recycling of aqueous phase for microalgae cultivation from the hydrothermal liquefaction process. Algal Res. 1, 70–76. 5. Campos, A.F., Pereira, O.G., Ribeiro, K.G., Santos, S.A., Valadares Filho, S. de C., 2014. Impact of replacing soybean meal in beef cattle diets with inactive dry yeast, a sugarcane by-product of ethanol distilleries and sugar mills. Anim. Feed Sci. Technol. 190, 38–46. 6. Chen, Y., Cheng, J.J., Creamer, K.S., 2008. Inhibition of anaerobic digestion process: A review. Bioresour. Technol. 99, 4044–4064. 7. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. 8. Collet, P., Hélias, A., Lardon, L., Ras, M., Goy, R.-A., Steyer, J.-P., 2011. Life-cycle assessment of microalgae culture coupled to biogas production. Bioresour. Technol. 102, 207–214. 9. Demirbas, A., Fatih Demirbas, M., 2011. Importance of algae oil as a source of biodiesel. Energy Convers. Manag. 52, 163–170. 10. Du, Z., Hu, B., Ma, X., Cheng, Y., Liu, Y., Lin, X., Wan, Y., Lei, H., Chen, P., Ruan, R., 2013. Catalytic pyrolysis of microalgae and their three major components: Carbohydrates, proteins, and lipids. Bioresour. Technol. 130, 777–782. 11. Ellis, J.T., Hengge, N.N., Sims, R.C., Miller, C.D., 2012. Acetone, butanol, and ethanol production from wastewater algae. Bioresour. Technol. 111, 491–495. 12. Gatlin, D.M., Barrows, F.T., Brown, P., Dabrowski, K., Gaylord, T.G., Hardy, R.W., Herman, E., Hu, G., Krogdahl, Å., Nelson, R., Overturf, K., Rust, M., Sealey, W., Skonberg, D., J Souza, E., Stone, D., Wilson, R., Wurtele, E., 2007. Expanding the utilization of sustainable plant products in aquafeeds: a review. Aquac. Res. 38, 551–579. 13. González-Fernández, C., Sialve, B., Bernet, N., Steyer, J.P., 2012. Comparison of ultrasound and thermal pretreatment of Scenedesmus biomass on methane production. Bioresour. Technol. 110, 610–616. 14. Griffiths, E.W., 2009. Removal and utilization of wastewater nutrients for algae biomass and biofuels. Grad. Theses Diss. Paper 631. http://digitalcommons.usu.edu/etd/631. 15. Halim, R., Danquah, M.K., Webley, P.A., 2012. Extraction of oil from microalgae for biodiesel production: A review. Biotechnol. Adv. 30, 709–732. 16. Harun, R., Singh, M., Forde, G.M., Danquah, M.K., 2010. Bioprocess engineering of microalgae to produce a variety of consumer products. Renew. Sustain. Energy Rev. 14, 1037–1047. 17. Huang, G., Chen, F., Wei, D., Zhang, X., Chen, G., 2010. Biodiesel production by microalgal biotechnology. Appl. Energy 87, 38–46. 18. Kanda, H., Li, P., 2011. Simple extraction method of green crude from natural blue-green microalgae by dimethyl ether. Fuel 90, 1264–1266. 19. Kim, S.W., Koo, B.S., Lee, D.H., 2014. A comparative study of bio-oils from pyrolysis of microalgae and oil seed waste in a fluidized bed. Bioresour. Technol. 162, 96–102. 20. Li, S., Xu, J., Chen, J., Chen, J., Zhou, C., Yan, X., 2014. The major lipid changes of some important diet microalgae during the entire growth phase. Aquaculture 428, 104–110.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

21. Li, Y., Horsman, M., Wang, B., Wu, N., Lan, C.Q., 2008. Effects of nitrogen sources on cell growth and lipid accumulation of green alga Neochloris oleoabundans. Appl. Microbiol. Biotechnol. 81, 629– 636. 22. Lodge-Ivey, S.L., Tracey, L.N., Salazar, A., 2014. RUMINANT NUTRITION SYMPOSIUM: The utility of lipid extracted algae as a protein source in forage or starch-based ruminant diets. J. Anim. Sci. 92, 1331–1342. 23. López Barreiro, D., Prins, W., Ronsse, F., Brilman, W., 2013. Hydrothermal liquefaction (HTL) of microalgae for biofuel production: State of the art review and future prospects. Biomass Bioenergy, 20th European Biomass Conference 53, 113–127. 24. Lum, K.K., Kim, J., Lei, X.G., others, 2013. Dual potential of microalgae as a sustainable biofuel feedstock and animal feed. J. Anim. Sci. Biotechnol. 4, 53. 25. Mata, T.M., Martins, A.A., Caetano, N.S., 2010. Microalgae for biodiesel production and other applications: A review. Renew. Sustain. Energy Rev. 14, 217–232. 26. Nelson, M., Zhu, L., Thiel, A., Wu, Y., Guan, M., Minty, J., Wang, H.Y., Lin, X.N., 2013. Microbial utilization of aqueous co-products from hydrothermal liquefaction of microalgae Nannochloropsis oculata. Bioresour. Technol. 136, 522–528. 27. Passos, F., García, J., Ferrer, I., 2013. Impact of low temperature pretreatment on the anaerobic digestion of microalgal biomass. Bioresour. Technol. 138, 79–86. 28. Prajapati, S.K., Kaushik, P., Malik, A., Vijay, V.K., 2013. Phycoremediation coupled production of algal biomass, harvesting and anaerobic digestion: Possibilities and challenges. Biotechnol. Adv. 31, 1408–1425. 29. Sathish, A., Glaittli, K., Sims, R.C., Miller, C.D., 2014. Algae Biomass Based Media for Poly(3hydroxybutyrate) (PHB) Production by Escherichia coli. J. Polym. Environ. 22, 272–277. 30. Sathish, A., Sims, R.C., 2012. Biodiesel from mixed culture algae via a wet lipid extraction procedure. Bioresour. Technol. 118, 643–647. 31. Stephenson, A.L., Kazamia, E., Dennis, J.S., Howe, C.J., Scott, S.A., Smith, A.G., 2010. Life-Cycle Assessment of Potential Algal Biodiesel Production in the United Kingdom: A Comparison of Raceways and Air-Lift Tubular Bioreactors. Energy Fuels 24, 4062–4077. 32. Sun, A., Davis, R., Starbuck, M., Ben-Amotz, A., Pate, R., Pienkos, P.T., 2011. Comparative cost analysis of algal oil production for biofuels. Energy 36, 5169–5179. 33. Tian, C., Li, B., Liu, Z., Zhang, Y., Lu, H., 2014. Hydrothermal liquefaction for algal biorefinery: A critical review. Renew. Sustain. Energy Rev. 38, 933–950. 34. Wijffels, R.H., Barbosa, M.J., Eppink, M.H.M., 2010. Microalgae for the production of bulk chemicals and biofuels. Biofuels Bioprod. Biorefining 4, 287–295. 35. Young, A., 2011. Zeolite‐Based Algae Biofilm Rotating Photobioreactor for Algae and Biomass Production. Grad. Theses Diss. Paper 986. http://digitalcommons.usu.edu/etd/986

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

Figure Legends Figure 1. Illustration of the algae fractionation procedure for isolation of algal lipids (WLEP). Figure 2. The three charts presented above show the percentage of total FAMEs remaining in the residual algae biomass using different hydrolysis conditions. The efficiency of lipid extraction is determined by subtracting the values shown from 100%. The y-axis represents the relative amount of transesterifiable lipids remaining within the biomass after the hydrolysis step compared to the total transesterifiable lipids present in the algal biomass used. The x-axis in each plot is the hydrolysis reaction time in minutes, Full (F), Half (H), and Quarter (Q) refer to the concentration of acid and base (Table 1), and (A), (B), and (C) refer to the hydrolysis reaction temperatures used of 90, 75, and 50oC respectively. Error bars represent one standard deviation of triplicate measurements. Figure 3. Illustrates the percentage of the total transesterifiable lipids of the algal biomass that remain in the residual algal biomass after the acid and base hydrolysis steps and DDI water wash. Data are arranged from most effective condition (left side) to least effective (right). Error bars represent one standard deviation of triplicate measurements. Figure 4. (A) Reduction of residual lipids present in the residual hydrolyzed biomass fraction (Stream 1) of the WLEP with increasing number of washes using both DDI water and a 0.1 M solution of sodium hydroxide. Error bars represent one standard deviation of triplicate measurements. (B) Reduction in transesterifiable lipids remaining in the precipitated solid phase as the number of hexanes separation cycles increases. Error bars represent one standard deviation of triplicate measurements. (C) Distribution of transesterifiable lipids in the four side streams generated in this study using the modified WLEP compared to previous results (Sathish and Sims, 2012) (error bars represent one standard deviation of six replicates) Figure 5. Chemical analysis of the algal biomass pre-processing and the residual biomass after acid and base hydrolysis and washing. Error bars represent one standard deviation of triplicate measurements.

Figure 1

Figure 2

Figure 3

Figure 4

Figure 5

Table 1. Conditions tested to evaluate lipid extraction efficiency. Time corresponds to the length of time used for acid and base hydrolysis steps individually. Conditions were evaluated as a 3x3x4 factorial in which the factor levels were acid and base concentration (F, H, and Q), temperature (50, 75, and 90oC), and Time (15, 30, 45, and 60 minutes). Acid and Base Designation “Full” (F) “Half” (H) “Quarter” (Q)

Acid Solution Concentration 1M 0.5 M 0.25 M

Base Solution Concentration 5M 2.5 M 1.25 M

Temperature (C)

Time (Minutes)

50, 75, and 90 50, 75, and 90 50, 75, and 90

15, 30, 45, and 60 15, 30, 45, and 60 15, 30, 45, and 60

Highlights 

Modified wet lipid extraction procedure extracts 77% of transesterifiable lipids



Lipids isolated account for 72% of transesterifiable lipids in the algae biomass



Side streams can be used as bioproducts or as feedstock for bioproduct production



Solid precipitate phase formed contains 11.2 wt% nitrogen (70% protein by mass)



LEA biomass has higher C/N ratio after procedure and aqueous phase rich in nutrients