Fuel 95 (2012) 63–70

Contents lists available at SciVerse ScienceDirect

Fuel

journal homepage: www.elsevier .com/locate / fuel

Concomitant extraction of bio-oil and value added polysaccharides from Chlorellasorokiniana using a unique sequential hydrothermal extraction technology

Moumita Chakraborty a, Chao Miao a, Armando McDonald b, Shulin Chen a,⇑a Department of Biological Systems Engineering, Washington State University, Pullman, WA 99164-6120, United Statesb Forests Products Program, University of Idaho, Moscow, ID 83844-1132, United States

a r t i c l e i n f o a b s t r a c t

Article history:Received 4 March 2011Received in revised form 3 October 2011Accepted 20 October 2011Available online 28 December 2011

Keywords:Bio-oilChlorella sorokinianaHydrothermal liquefactionPolysaccharidesSequential subcritical hydrothermalliquefaction

0016-2361/$ - see front matter � 2011 Elsevier Ltd. Adoi:10.1016/j.fuel.2011.10.055

⇑ Corresponding author. Tel.: +1 509 335 3743; faxE-mail address: chens@wsu.edu (S. Chen).

To make algal bio-fuel economically viable, extraction of value added co-products along with oil, appearsabsolutely necessary. The major barrier in algal co-product development is the lack of an efficient sepa-ration technology. To address this issue a unique two-step sequential hydrothermal liquefaction(SEQHTL) technology for the simultaneous production of value-added polysaccharides and bio-oil fromalgal-biomass was developed. The first step involves the subcritical water extraction of valuable algal(Chlorella sorokiniana) polysaccharides at 160 �C. The polysaccharide rich water extract was removedand precipitated with ethanol. In the next step, the extracted biomass was liquefied to bio-oil at300 �C. The yield of bio-oil by SEQHTL was 24% of the dry weight. In addition, this method also extracted26% of the polysaccharides present whereas; direct hydrothermal liquefaction (DIRHTL) only generated28% bio oil. In the SEQHTL method bio-char formation was remarkably low, and as such, SEQHTL produce63% less bio-char than DIRHTL. The yield of bio-char production is negatively correlated to polysaccharidecontent (p > 0.98) suggesting majority of carbohydrates present in algal biomass were converted into bio-char. This conversion did not significantly influence the bio-oil production. Comparative GC-FID, GC–MS,NMR, FT-IR analysis and ESI-MS of the bio-oil extracted by SEQHTL with DIRHTL showed no significantdifferences. Elemental analysis of the SEQHTL bio-oil demonstrated that it contained 70% carbon, 0.8%nitrogen, and 11% oxygen. This method demonstrates an important proof-of-concept step towards theconcomitant production of value added co-products, along with bio-oil, from algal biomass.

� 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Algae are considered to be a potential biomass for the produc-tion of biofuels. One of algae’s outstanding features is its potentialfor high productivity [1], requiring much less acreage than tradi-tional feedstocks. Algae can be grown on non-arable land andwaste water. Despite these advantages, commercialization of algalderived fuel still remains challenging chiefly because of varioustechno-economic constraints. Nonetheless, one important ap-proach to making algal biofuel cost effective is the production ofco-products, which in turn augment the cost of biofuel production.Two major aspects need to be addressed in order to generate co-products from the algal biomass: 1. Identify compounds that canbe developed into value added co-products and 2. Enhance algalbio-fuel production by novel extraction methods so as to producea portfolio of oil and co-products.

The extraction of oil from algae by hydrothermal liquefaction(HTL) is advantageous over other methods. Some of the benefitsof HTL are: This process does not require drying of the biomass,

ll rights reserved.

: +1 509 335 2722.

nor high energy or high separation efficiency (since water remainsin liquid phase, the phase change is avoided). Additionally, highthroughputs, versatility of the chemistry process (solid, liquidand gaseous fuels), reduced mass transfer resistance in the hydro-thermal conditions, and improved selectivity for the desired en-ergy products (methane, hydrogen, liquid fuel) are characteristicsof this production process. Since the HTL extraction solvent iswater, which is a benign chemical, HTL is also favorable for extract-ing co-products targeted to the food, cosmetic, or nutraceuticalindustries. HTL has been demonstrated to be effective for produc-ing bio-oil using several species of micro-algae [2–4]. Recently,several literatures have reported the hydrothermal liquefactionof different micro algal species with different biochemical compo-sition [5–8]. Savage’s group from Michigan State University haveeven gained some success in further up gradation of the hydrother-mally liquefied algal bio oil by using heterogenous catalyst insupercritical water [9,10]. Ross’s group [11] has also reportedabout the catalytic up gradation of hydrothermally processed algalbio oil.

HTL converts, in addition to lipid, other organic molecules to fuelcomponents; thus the total bio-oil yield is greater than the lipidcontent. For example, Dote et al. [2] liquefied a strain of micro-algae

64 M. Chakraborty et al. / Fuel 95 (2012) 63–70

that contained 50% natural oils at 300 �C with Na2CO3 as catalyst,and produced 64% oil (on a mass basis) from this feedstock showingthat not only lipid, but also other organic compounds like protein,fiber, and carbohydrate are converted into an oil. Influence of alkaliand organic acid catalyst on the yield of bio oil from algae and fromdifferent model organic compounds like carbohydrate, protein andlipid are also elaborately studied by Ross and Biller group [7,12].During HTL, the solvent properties of water change significantlynear its critical point (Tc) [13]. In fact, the dielectric constant ofwater decreases considerably near the Tc, which causes a changein viscosity and an increase in its diffusion coefficient [14]. Underhydrothermal treatment, organic compounds are liquefied througha series of reactions such as solvolysis, depolymerization, dehydra-tion, and decarboxylation. Solvolysis reactions results in micellar-like substructures in the feedstock. Depolymerization reactionslead to smaller molecules. Decarboxylation and dehydration leadsto degradation products and the evolution of CO2 through elimina-tion of carboxyl groups [15]. Thus, the substantial changes that oc-cur in the physical and chemical properties of water in the vicinityof its Tc can be utilized advantageously for converting algal biomassto desired biofuels. In fact, reactions in subcritical water also pro-vide a novel medium to conduct tunable reactions for the extractionand synthesis of valuable chemicals from biomass [16]. Based onthe literature the two expected major co-products from HTL arehydrolyzed sugars and bio-char.

Algae are immensely diversified cells. Algal polysaccharidespossess a unique combination of functional properties. As such,they are renewable materials, generally non-toxic, and biodegrad-able. Due to their physical and chemical properties, namely theirwater retention ability, film-forming capacity, and rheology, poly-saccharides are used in a wide variety of industrial applications,including food, textiles, painting, cosmetics, paper, and pharma-ceuticals as emulsifiers, stabilizers, or thickening agents [17,18]Other uses of algal polysaccharides have also been explored[19,20]. For instance, polysaccharides from Chlorella sp. have beenshown to exhibit strong anti-inflammatory and immunomodula-tory properties [21,22]. Therefore, algal polysaccharides, in nativeform, are more valuable than simply being used as sugar sourcesor bio-char.

This study develops a process for tuning the hydrothermal reac-tion temperature so that it can be utilized to isolate algal polysaccha-ride without degradation. In so doing we develop a temperaturegradient hydrothermal extraction method to isolate polysaccha-rides at low temperature followed by hydrothermal liquefaction ofthe extracted biomass into bio-oil at higher temperature.

2. Experimental

2.1. Raw materials

The green alga Chlorella sorokiniana (UTEX 1602) was obtainedfrom the Culture Collection of Alga at the University of Texas (Aus-tin, TX, USA). Heterotrophic culturing was supplemented with var-ious concentrations of glucose according to specific experimentaldesigns. All media were autoclaved at 121 �C for 20 min before use.

2.2. Algae sample preparation

The culture medium recipe used was that of Kuhl and Lorenzen[23]. Algal culture development involved three steps. Each flask of250 ml medium was inoculated with 1 ml of algal biomass and al-lowed to grow for 3 days. In the second step, 25 ml of algal richmedium from each starter flask was subsequently inoculated intotwo flasks each containing 225 ml medium and cultured for3 days. In the third step, both of the 250 ml algal biomass cultureswere introduced into a 5 l fermentor. After 13 days of culture algae

were harvested and centrifuged at 20 �C and 5000 rpm for 5 min.The algal paste was collected and 10 g algae paste was placed intoa 120 �C oven for 12 h in order to dry it and measure pastemoisture.

2.3. Sequential subcritical hydrothermal liquefaction (SEQHTL)

Prior to the sequential hydrothermal extraction, to identify thetemperature require for optimum yield of polysaccharide extrac-tion of polysaccharide was conducted under four different temper-atures (140 �C, 160 �C, 180 �C, 200 �C) and autogenous pressure. Onthe basis of the optimum polysaccharide extraction temperatureSEQHTL experiments were conducted in three modes under differ-ent temperature conditions; two-step SEQHTL (160 �C, 300 �C),three step SEQHTL (160 �C, 180 �C, 300 �C), and DIRHTL (300 �C)respectively to further evaluate the influences of different sequen-tial steps on the yield of polysaccharides and bio oil. These pro-cesses were developed using a PARR 4522 1 l bench top reactor.A two-step SEQHTL and a three-step SEQHTL were developed.The three-step SEQHTL is comprised of three sequential tempera-ture sets: 160 �C, 30 min; 180 �C, 30 min; and 300 �C, 1 h. In thefirst step, algal paste (14 g dry algae) and distilled water weremixed thoroughly (at a 1:9 biomass to water ratio) and introducedinto the reactor. Nitrogen was bubbled for 5 min to purge oxygenfrom the system. The temperature of the reactor was ramped to160 �C at a rate of 7 �C/min and held for 30 min. The reactor wasthen cooled with cold water to room temperature. The reactionmixture was subsequently collected and filtered with 90 lm filterpaper. The filtered algal residue was returned back to the reactorfor the next step while the filtrate was treated with ethanol to pre-cipitate polysaccharides. To maintain the water to biomass ratio,moisture content and weight of the filtered biomass were mea-sured and water was added as necessary, and the mixture was re-turned to the reactor to continue the second step of SEQHTL, whichwas heating to 180 �C and holding at this temperature for 30 min.The subsequent reaction steps were the same as the first step. Forthe third step reaction moisture content and weight of the twotime treated algal residue were again measured, accordingly waterwas added to the biomass and the reactor temperature was raisedto 300 �C. At 300 �C reactor temperature was keep on hold for 1 h.After treatment, solids were found attached to the cooling pipewall of the reactor. Methylene chloride (CH2Cl2, 150 ml) was addedto the reactor to solubilize these solids. The two-step SEQHTL wasconducted in the same way except the 180 �C for 30 min step wasnot conducted. In the case of direct hydrothermal liquefaction(DIRHTL), both low temperature sequential extractions were omit-ted and instead the temperature was raised to 300 �C directly.

2.4. Products separation and extraction

The separation procedure for production of liquefaction prod-ucts is illustrated in Fig. 1.

The aqueous solutions derived from the SEQHTL steps were col-lected individually. Ethanol was added to each solution at a 1:4 ra-tio (water:ethanol, v/v) in order to precipitate the polysaccharides.The water mixture was subsequently centrifuged and the pelletwas air dried for 36 h and then weighed. Ethanol was removedusing a rotary evaporator under vacuum at 40 �C. The remainingwater was lyophilized and the dried residue was then quantifiedas water extractives. The reaction mixture obtained after the300 �C 1 h reaction, including the supernatant, was treated with300 ml CH2Cl2. The CH2Cl2 phase was filtered to collect the CH2Cl2

precipitant which was then placed into a rotary evaporator to re-move the CH2Cl2. The remaining liquid was defined as crude bio-oil. The CH2Cl2 insoluble solid residues were defined as bio-charand dried at 120 �C for 12 h.

Fig. 1. Schematic diagram of the 3-step SEQHTL extraction strategy.

M. Chakraborty et al. / Fuel 95 (2012) 63–70 65

2.5. Optimization of temperature for high yield of polysaccharides

To optimize the yield of polysaccharides polysaccharide was ex-tracted at different temperature ranging from 120 to 200 �C in thesame the procedure as mentioned above.

2.6. Product yield

The products of SEQHTL were crude bio-oil, crude polysaccha-rides, water extractives, and bio-char, while products from DIRHTLincluded crude bio-oil and bio-char. The yields of each productwere calculated on a dry algae mass basis and were calculated asfollows:

Bio-oil ðwt%Þ ¼ Woil

Wdry alage� 100%

Polysaccharide ðwt%Þ ¼WPolysaccharide

Wdry alage� 100%

Water Extractives ðwt%Þ ¼WWater Extractives

Wdry alage� 100%

Bio-Char ðwt%Þ ¼ WBio-Char

Wdry alage� 100%

where Woil is the mass of bio-oil (g), Wdry algae is the total mass ofdry algae (g), Wpolysaccharide is the mass of polysaccharides (g),WWater Extractives is the mass of water extractives (g), and WBio-Char

is the mass of bio-char (g).

2.7. Product analysis

2.7.1. Elemental analysisSamples for carbon, nitrogen, and sulfur elemental analysis

were combusted separately with an elemental analyzer (ECS4010, Costech Analytical, Valencia, CA). N2 and CO2 gases were sep-arated on a 3.0 m GC column at 40 �C, while sulfur dioxide gas wasseparated on a 0.8 m GC column at 105 �C. Concentrations of thegases were quantified with a continuous flow isotope ratio massspectrometer (Delta PlusXP, Thermofinnigan, Bremen) [24] or athermal conductivity detector (TCD) on the elemental analyzer.Samples for oxygen elemental analysis were thermally convertedin the absence of oxygen by means of a high temperature thermalconversion elemental analyzer (TCEA, Thermofinnigan, Bremen)

66 M. Chakraborty et al. / Fuel 95 (2012) 63–70

and the oxygen was quantified from the resulting CO gases sepa-rated on a GC column (0.6 m � 1=4 in. � 4.0 mm, molecular sieve5A, Varian) at 90 �C and analyzed as mentioned above [24,25].The higher heating value (HHV) was calculated with the Dulongformula [26].

HHVMJmg

� �¼ 0:3383Cþ 1:422 H� O

8

� �

where C, H, and O are the weight percentages of carbon, hydrogen,and oxygen in the oil, respectively.

2.7.2. Fatty Acid Methyl Ester (FAME) analysis by gas chromatography(GC)

Fatty acid methyl ester analysis was performed using the methodof O’Fallon et al. [27]. Briefly, 0.1 g of freeze dried sample was placedinto a 16 � 125 mm screw-cap Pyrex culture tube, to which 1.0 ml ofC13:0 internal standard (0.5 mg C13:0 per ml of MeOH), 0.7 ml 10 NKOH in water, and 5.3 ml MeOH were added. The tube was incubatedin a 55 �C water bath for 1.5 h with intermittent vortex-mixing todissolve and hydrolyze the sample. After cooling to room tempera-ture, 0.58 ml of 24 N H2SO4 in water was added. The tube was mixedby vigorous hand shaking and then incubated again in a 55 �C waterbath for 1.5 h with intermittent vortexing. After fatty acid methyl es-ter synthesis, 2 ml water and 3 ml hexane were added to the tube.The tube was vortexed for 5 min on a multi tube vortex. The tubewas centrifuged for 5 min in a tabletop centrifuge and the hexanelayer, containing the fatty acid methyl esters, was placed into a GCvial. The vial was capped and placed at �20 �C until GC analysis.The fatty acid composition of the sample was determined by capil-lary GC on a SP-2560, 100 m � 0.25 mm � 0.20 micrometer capil-lary columns (Supelco) installed on a Hewlett Packard 5890 gaschromatograph equipped with a Hewlett Packard 3396 Series II inte-grator and 7673 controller, a flame ionization detector, and splitinjection. The initial oven temperature was 140 �C, held for 5 min,subsequently increased to 240 �C at a rate of 4 �C per min, and thenheld for 20 min. Helium was used as the carrier gas at a flow rate of0.5 ml/min, and the column head pressure was 275 kPa. Both theinjector and the detector were set at 260 �C. Fatty acids were identi-fied by comparing their retention times with fatty acid methylstandards.

2.7.3. GC/MS analysisThe biooil (1.5 mg) was weighed into a GC glass vial to which a

solution of anthracene (0.05 mg ml�1) in CH2Cl2 (1.5 ml) wasadded as an internal standard (IS). GC–MS analysis was subse-quently performed on a FOCUS-ISQ (Thermoscientific) instrumentin the electron impact mode and the compounds were separatedby a RTx-5MS capillary column (30 m, 0.25 mm £, Reztek) withHe as carrier gas. The temperature program: from 40 to 320 �C, ele-vated at a rate of 5 �C min�1. The injector and MS-transfer line waskept at 325 �C. Data analysis: Xcalibur v2 software package (Ther-moscientific). Compound identification: by comparison of the MSand retention times with standards together with MS librarymatching (2008 NIST MS library).

2.7.4. FTIR analysisAll Fourier transform infrared (FTIR) spectra were collected

using a ThermoNicolet Avatar 370 spectrometer operating in theattenuated total reflection (ATR) mode (SmartPerformer, ZnSecrystal). Each spectrum was taken as an average of 64 scans ata resolution of 4 cm�1. The spectra were ATR and baseline cor-rected using Omnic v7 software prior to spectral analysis. Crudealgal oil samples were prepared by overnight solvent (CH2Cl2)extraction of the freeze dried algae and the extract concentratedto dryness.

2.7.5. NMR analysisNMR analysis was performed using a 600 MHz Varian spec-

trometer to compare the functional groups present in the DIRHTLand SEQHTL bio-oil. Twenty-four milligrams of bio-oil were dis-solved in 600 ll of CDCl3 with 2 ll p-dioxane (internal standard).The 1H spectra were acquired at 600 MHz, with a 45� pulse angleand a sweep width of 6600 Hz.

2.7.6. ESI-MS analysisMolecular weight determination was performed by negative ion

electrospray-mass spectrometry (ESI-MS) using a LCQ-Deca massspectrometer (ThermoQuest). Biooil samples (2 mg) were dis-solved in methanol (2 ml) containing 0.5% acetic acid and intro-duced to the spectrometer at a flowrate of 10 ll/min. The mass-spectra were converted to an excel spreadsheet and number aver-age molecular weight (Mn) calculated using Mn = Rni �Mi/Rni andweight average molecular weight (Mw) calculated usingMw ¼ Rni �M2

i =Rni, where ni and Mi are the intensity and the massof polymer number [28].

3. Results and discussion

3.1. Yield of polysaccharides at different temperatures

Until now only limited research has been conducted on thehydrothermal degradation pathway of algal polysaccharides. Ingeneral it has been observed that amorphous polysaccharides (likehemicellulose) hydrolysis to water soluble products increases rap-idly as the temperature increase from 150 to 190 �C. Whereas,polysaccharides like celluloses remain stable below 200 �C[29,30]. Algal cell wall polysaccharides are extremely complexand diversified. The chemical structures of the polysaccharidesvary from species to species. Due to the inherent tensile strengthand complexity of the algal cell wall it has been difficult to utilizean ambient temperature for algal polysaccharide extraction. Con-ventionally, algal polysaccharides are treated to 6–8 h of Soxhletextraction with boiling water at a temperature range of 80–90 �C[31,32]. Since our aim was to extract the polysaccharides in theirnear native forms, algae were hydrothermally treated at a temper-ature range of 120–200 �C for 20 min under autogenous pressureto identify an ambient temperature for optimum yield.

Addition of ethanol to the water extracted at 120 �C tempera-ture showed no precipitation (data not shown) indicating a lackof polysaccharide extraction at this temperature. The maximalquantity of polysaccharide precipitation by ethanol was obtainedat 160 �C, while the 200 �C extract showed very low polysaccharidecontent (Fig. 2). From these results it can be inferred that a mini-mum of 140 �C is required to break the algal cell wall, and opti-mum polysaccharide extraction will take place at 160 �C. Furtherdetailed research is required to understand the characteristics ofthe extracted polysaccharides and the rate of hydrolysis.

3.2. Product yield

Total bio-oil yield for the two-step SEQHTL was 23.60%, 20.00%for the three-step, and 27.80% for DIRHTL. Two-step SEQHTL seemsto be ideal for the optimum yield of bio-oil and polysaccharideswhereas 160 �C is the optimum temperature for extraction of poly-saccharides. Three-step SEQHTL produces a high amount of poly-saccharides, however, it requires more energy and resultsshowed that after the 160 �C treatment only 6.4% polysaccharideswere left to be further extracted at 180 �C. Table 1 displays theyield of products under the different hydrothermal treatments.

In comparison to SEQHTL, the DIRHTL method has a higher yieldof bio-oil. Loss of biomass during the use of the batch reactor mightbe the possible reason for such a difference in bio-oil yield. In future,

Fig. 2. Yield of Chlorella sorokiniana polysaccharides extracted at different temperature.

Table 1Product yield from hydrothermal processing by SEQHTL and DIRHTL. Yield wascalculated on the basis of dry weight.

Polysaccharides(%)

Water extractives(%)

Bio-oil(%)

Bio char(%)

160 180

DHT 0 / / 27.8 20.82-step

SEQHTL26.3 / 5.6 23.4 7.6

3-stepSEQHTL

28.1 6.4 12 20.0 5.0

Yield was calculated in terms of percentage of biomass dry. Weight taken initiallyfor the experiment.

Table 2Elemental analysis of carbon, oxygen, nitrogen, hydrogen, and sulfur; and higherheating value (HHV) of bio-oil produced by SEQHTL and DIRHTL.

Products C% O% N% S% H% HHV (MJ/kg)

Crude algae 50.4 37.4 2.91 0.21 7.9 21.6SEQHTL 76 11 0.78 0.16 12 40.8DIRHTL 72 11 1.14 0.14 15 43.7

M. Chakraborty et al. / Fuel 95 (2012) 63–70 67

to minimize the biomass loss during SEQHTL processing a continu-ous reactor will be used. One of the most striking characteristics ofthe SEQHTL process is the large reduction in bio-char formation.There is 20.8% bio-char formation in the DIRHTL process, whereasonly 7.6% bio-char is produced by the SEQHTL method. The forma-tion of bio-char was negatively correlated with polysaccharide pro-duction, making the SEQHTL method beneficial over the DIRHTLmethod, as SEQHTL can successfully recover useful polysaccharidesinstead of converting them to less valuable bio-char.

3.3. Elemental analysis and the high heating value of the bio-oil

The carbon content in the bio-oil for SEQHTL and DIRHTL was76% and 72%, respectively. The oxygen content was 11% and thenitrogen content was less than 2% in both cases. Hydrogen contentvaried from 12% to 15% (Table 2). Elemental analysis showed thatthe algal biomass was very low in nitrogen content (data notshown). Heterotrophically grown algal cells are usually low in pro-tein content. According to the study conducted by Miao and Wu[33], heterotrophically grown Chlorella protothecoides cells pos-sessed five times lower protein content than their autotrophic coun-terpart. Heterotrophically grown algae may thus reduce the cost forbio-oil denitrogenation. The carbon, nitrogen, and hydrogen contentof the extracted bio-oil are comparable with crude petroleum [7].However, the oxygen content of the bio-oil was high. Therefore,an upgrading of the bio-oil, for example, hydrogenation, might berequired to produce high quality bio-oil. The heating value of thebio-oil was calculated using Dulong’s formula and that producedby DIRHTL and SEQHTL was 43.7 MJ/kg and 40.8 MJ/kg, respec-tively. These values are similar to those of crude petroleum [7].

3.4. Fatty acid analysis by GC

Fatty Acid Methyl Ester (FAME) analysis was performed onSEQHTL oil, DIRHTL oil, and dry algae (Table 3). Interestingly, the

Table 3Comparative fatty acid analysis of Chlorella sorokiniana biomass and the fatty acids presen

Fatty acid Structure

Palmitic CH3(CH2)14COOHHexadecenoic CH3(CH2)5CH@CH(CH2)7COOHStearic CH3(CH2)16COOHHexadecatetranoic CH3(CH2)2(CH@CHCH2)3(CH2)3COOHHexadecatrienoic CH3CH2(CH@CHCH2)3(CH2)4COOHOleic CH3(CH2)7CH@CH(CH2)7COOHLinoleic CH3CH2(CH@CHCH2)3(CH2)6COOH

a mg/g biomass dry weight;b mg/g of bio-oil extracted by SEQHTL method;c mg/g of bio-oil extracted by DIRHTL method.

SEQHTL and DIRHTL analysis were very similar. Palmitic acid andhexadecatrienoic acid are the major fatty acid constituents. The to-tal fatty acid content for SEQHTL oil was 39.1% and for DIRHTL oil itwas 43%. Both samples had a saturated fat content of 32%, a mono-unsaturated fatty acid (MUFA) content of 6%, and a polyunsatu-rated fatty acid (PUFA) content of 50%. The major fatty acidsidentified in the crude bio-oil were palmitic acid, hexadecanoicacid, stearic acid, isomers of hexadecatrienoic acid, and linoleicacid. There was a reduction in PUFA from the original algal bio-mass. The high heat used in this process might cause an initialisomerization or epi-isomerization of the unsaturated fatty acidsfrom cis to trans form, but these postulated trans isomers werenot observed. Linoleic (C18:3n3) acids essentially decreased tohalf. However, the hexadecatrienoic acid peak (C16:3n3) decreasedvery little. NMR analysis of the bio-oil also did not reveal any transfatty acid formation and therefore do not fully explain the fattyacid conversions that must have occurred.

3.5. GC/MS analysis of the bio-oil

The bio-oil was further characterized to identify volatile com-pounds present. The main compounds detected were fatty acidswith a minor amount of alkanes (that could not be unequivocallyidentified). Furthermore, other volatile components may have beenlost during evaporation of CH2Cl2 extract or if present must havebeen masked by the solvent peak. This phenomenon is also re-ported by [5,34]. However, bio-oil analyzed prior to the evaporationof solvent do not exhibits any quantifiable volatile peak in the GCchromatogram. Total fatty acids obtained by FAME analysis account

t in the bio-oil produced by SEQHTL and DIRHTL.

Biomassa SEQHTLb DIRHTLc

(mg/g) (mg/g)b (mg/g)

57.48 148.50 167.4914.50 25.53 24.94

5.22 12.40 14.020.00 38.51 43.72

74.76 116.83 130.8911.80 200.97 136.9084.00 49.68 49.77

Table 4Identification of compounds found in the bio-oil produced by SEQHTL and DIRHTL at300 �C by GC/MS analysis.

Rt (min) Compound SEQHTL (% of biooil) DIRHTL (% of biooil)

28.83 Anthracene (IS) – –31.73 Hexadecenoic acid 0.70 0.2532.19 Palmitic acid 11.39 6.9635.34 Octadecadienoic

acid1.80 0.47

35.44 Linoleic acid 8.44 4.3735.50 Oleic acid 3.04 3.8135.81 Stearic acid 0.61 0.90Total 25.06 16.76

Fig. 3. FTIR spectra of bio-oil produced by DIRHTL and SEQHTL and the CH2Cl2

extracted total lipid from algal biomass.

68 M. Chakraborty et al. / Fuel 95 (2012) 63–70

for approximately 60% of the bio-oil and the algal biomass seems toconsist of 25% of fatty acid. GC–MS analysis of the bio-oil (non-methylated) (Table 4) shows 25% of the SEQHTL and 16% of the DIR-HTL bio oil is composed of free fatty acids. Therefore, it can be as-sumed that 15% of the fatty acid in SEQHTL and 24% in DIRHTLbio oil are present in mono, di or triglyceride form and some ofthem might polymerized into even more complex structures. FT-IR spectral data shows that acid to ester ration has been increasedby eight fold during bio oil formation this might support the hydro-lysis of triglycerides, diglycerides, and monoglycerides present inthe biomass to free fatty acids during hydrothermal treatmentwhich perhaps, react among themselves in the hydrothermal mediaand further produces high molecular weight, complex, polymerizedstructures yet maintaining their carboxyl functional group. Freefatty acid yield in the DIRHTL bio oil is lower in comparison toSEQHTL. Direct liquefaction solubilizes all the carbohydrate andprotein of the biomass this might result in high volatile yield inthe DIRHTL bio oil. As a result during solvent extraction, extractionration of free fatty acid to volatile might be lower. This could be apossible reason for the lower yield of free fatty acid in DIRHTL biooil. A most interesting feature of the GC/MS analysis is the absenceof nitrogenous compounds of protein origin. This fact is also corrob-orated by elemental analysis, which shows the presence of very lowamounts of nitrogen in the bio-oil. Conversion of protein in the algalcells to very small volatile compounds may be one explanation forthe nitrogen absence. Alternatively, the nitrogenous component of

Fig. 4. 1H NMR spectra of: (A) DIRHTL and (B) SEQHT

the bio-oil may be found in heavy dimeric or trimeric structures andbe beyond the range of GC/MS analysis.

3.6. FTIR analysis of bio-oil

FTIR Spectra of the bio-oil extracted by DIRHTL and SEQHTL dis-play intense absorbances between 2850 and 3000 cm�1, indicatingthe CAH stretching vibration of CH3 and CH2 (Fig. 3). The intensepeak near the 1700 cm�1 region is ascribed to the C@O vibrationdeveloped in the presence of ketones, aldehydes, esters and car-boxylic acids [35]. Most interesting aspect of the FT-IR analysis isthe comparison between CH2Cl2 extract of freeze dried algal bio-mass and the bio-oil around 1700 cm�1 region. Region 1705–1720 cm�1 is assigned for CAO stretching from acids while1720–1750 cm�1 region denotes CAO stretching from esters. Fromthe spectra it is observed that in case of bio-oil highest intensity ofthe peak in this region is at 1710 cm�1 whereas, in the algal CH2Cl2

extract the peak has been shifted to 1750 cm�1 suggesting, duringbio-oil formation majority of the triglycerides has been hydrolyzed

L bio-oil. The internal standard (IS) is p-dioxane.

0

5E+10

1E+11

0 500 1000 1500 2000m/z

SEQHTL

0

1E+11

2E+11

3E+11

0 500 1000 1500 2000m/z

DIRHTL

Fig. 5. Mass plots of weight average molecular weight distribution by ESI-MS ofDIRHTL and SEQHTL derived bio-oils.

M. Chakraborty et al. / Fuel 95 (2012) 63–70 69

into acid. Intensity ratio shows 83–85% of the triglycerides duringbio-oil formation have been converted to fatty acids. The weakpeaks in the 1250–1500 cm�1 region are due to the presence ofCAO bending vibrations followed by CAH bending vibrations, fur-ther indicating the presence of a minor amount of esters [36] inbio-oil. These results are consistent with the presence of a collec-tion of fatty acids and alkanes in the total ion chromatogram ofthe GC/MS. One of the characteristic features in the FTIR spectraof the bio-oil from C. sorokiniana is the absence of major peaks inthe 3000–3500 cm�1 region [37], indicating the minimal presenceof characteristic polysaccharides and protein derivatives in the bio-oil. These data are further corroborated by GC/MS and NMR analy-sis. In the case of DIRHTL, the data suggests that whatever polysac-charides or proteins were present in the original algae wereconverted into bio-char and other small volatile molecules,whereas in the case of SEQHTL, the polysaccharides were removedprior to the formation of bio-char. The presence of small bands inthe 700–900 cm�1 region can probably be attributed to the pres-ence of substituted aromatic structure.

Fig. 6. FTIR spectra of: (A) polysaccharides isolated by conventional S

3.7. NMR analysis of the bio-oil

The bio-oil produced by both DIRHTL and SEQHTL exhibit simi-lar fatty acids profiles (Fig. 4). The region 2.5–0.5 ppm representsaliphatic protons, and these protons, along with the peaks at5.5 ppm which indicate the presence of protons on unsaturated car-bon bonds, contribute 33% of the total protons present in SEQHTLbio-oil, and 35% of the protons in DIRHTL bio-oil, indicating a highaliphatic content in the bio-oil. No significant peaks were observedin the aromatic region of the spectrum (2.5–3.5 ppm), which sug-gests the absence of aromatic compounds in the bio-oil, and thisfact is consistent with the GC/MS and FTIR analyzes.

3.8. ESI-MS analysis

As noted above, GC–MS analysis only account for the 26% and16% of the SEQHTL and DIRHTL bio-oil. If we consider the removedvolatiles there is still a considerable amount of unknown com-pounds in the crude bio-oil. It is likely that much of the remainingcompounds are of high molecular weight and do not elute from GCcolumn. Therefore, we used negative ion ESI-MS to obtain an aver-age molecular weight of the two bio-oil samples.

Molecular weight profile of the bio oil suggests that the ex-tracted bio-oils are composed of high molecular weight com-pounds. The Mn and Mw of the SEQHTL and DIRHTL bio oil are1002 and 1172 and 829 and 1022 g/mol, respectively. Fig. 5 showsthe Mw distribution of bio-oil samples. Average molecular weightof the DIRHTL is relatively lower than that of SEQHTL this mightbe due to the presence of less cross-linking in the DIRHTL bio-oilproduced from carbohydrate and protein part of the biomass.

3.9. FTIR analysis of the extracted polysaccharides

The FTIR spectrum of the polysaccharides isolated by conven-tional Soxhlet extraction appears to be identical with that of theSEQHTL polysaccharide spectrum (Fig. 6). The polysaccharides ex-tracted by SEQHTL exhibit a strong absorbance between 3300 and2700 cm�1 due to CAH stretching vibrations. The characteristicband at 1700–1600 cm�1 represents the amide-I band due to CAOstretching and the 1600–1500 cm�1 peak represents the amide-IIband due to NAH bending. One of the major monosaccharide in C.sorokiniana is glucosamine, an amino sugar, and therefore the

oxhlet extraction and (B) polysaccharides produced by SEQHTL.

70 M. Chakraborty et al. / Fuel 95 (2012) 63–70

presence of the amide bands might represent this amino sugar.Alternatively, these bands may indicate the co-precipitation of algalprotein with the polysaccharides [38,39]. The 1200–900 cm�1 re-gion displays CAO, CAC, and CAOAC stretching vibrations of poly-saccharides [40–42].

The fate of polysaccharides during hydrothermal liquefaction isnot clear. It is believed that during DIRHTL processing some of thepolysaccharides are converted to oil, some to small aqueous prod-ucts, some to gases, with the majority converted into bio-char [43].Removal of polysaccharides by SEQHTL prior to bio-oil productiondoes not significantly influence the yield of bio-oil or its chemicalcharacterization as determined by NMR and FTIR. Both SEQHTL andDIRHTL bio-oils had similar heating values. Thus, the removal ofpolysaccharides does not seem to be detrimental to bio-oil produc-tion. The removal of polysaccharides by the SEQHTL process priorto bio-oil formation allows for the production of high valued co-products along with bio-oil production. Currently, our lab is focus-ing on optimizing the SEQHTL method for more efficient extractionof polysaccharides and bio-oil and to minimize energy consump-tion. We have observed that the water to biomass ratio, tempera-ture of extraction, and equilibration time are the threeparameters that play crucial roles in the sequential extraction ofpolysaccharides and bio-oil.

Acknowledgements

This work is supported partially by the Department of Energy(DOE) and by Washington State University Agricultural ResearchCenter. The authors would like to thank Dr. James V. O’Fallon andMrs. Kathleen M. Dorgan for their extensive editing of thismanuscript.

References

[1] Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25:294–306.[2] Dote Y, Sawayama S, Inoue S, Minowa T, Yokoyama S-y. Recovery of liquid fuel

from hydrocarbon-rich microalgae by thermochemical liquefaction. Fuel1994;73:1855–7.

[3] Minowa T, Yokoyama S-Y, Kishimoto M, Okakura T. Oil production from algalcells of Dunaliella tertiolecta by direct thermochemical liquefaction. Fuel1995;74:1735–8.

[4] Yang YF, Feng CP, Inamori Y, Maekawa T. Analysis of energy conversioncharacteristics in liquefaction of algae. Resour Conserv Recycl 2004;43:21–33.

[5] Brown TM, Duan P, Savage PE. Hydrothermal liquefaction and gasification ofNannochloropsis sp. Energy Fuels 1987;24:3639–46.

[6] Anastasakis K, Ross AB. Hydrothermal liquefaction of the brown macro-algaLaminaria Saccharina: effect of reaction conditions on product distribution andcomposition. Bioresour Technol 2011;102:4876–83.

[7] Ross AB, Biller P, Kubacki ML, Li H, Lea-Langton A, Jones JM. Hydrothermalprocessing of microalgae using alkali and organic acids. Fuel 1987;89:2234–43.

[8] Zhou D, Zhang L, Zhang S, Fu H, Chen J. Hydrothermal liquefaction ofmacroalgae Enteromorpha prolifera to bio-oil. Energy Fuels 1987;24:4054–61.

[9] Duan P, Savage PE. Upgrading of crude algal bio-oil in supercritical water.Bioresour Technol 2011;102:1899–906.

[10] Duan P, Savage PE. Hydrothermal liquefaction of a microalga withheterogeneous catalysts. Ind Eng Chem Res 2010;50:52–6.

[11] Biller P, Riley R, Ross AB. Catalytic hydrothermal processing of microalgae:decomposition and upgrading of lipids. Bioresour Technol 2011;102:4841–8.

[12] Biller P, Ross AB. Potential yields and properties of oil from the hydrothermalliquefaction of microalgae with different biochemical content. BioresourTechnol 1987;102:215–25.

[13] Kalinichev AG, Churakov SV. Size and topology of molecular clusters insupercritical water: a molecular dynamics simulation. Chem Phys Lett1999;302:411–7.

[14] Marcus Y. On transport properties of hot liquid and supercritical water andtheir relationship to the hydrogen bonding. Fluid Phase Equilib 1999;164:131–42.

[15] Demirbas A. Mechanisms of liquefaction and pyrolysis reactions of biomass.Energy Convers Manage 2000;41:633–46.

[16] Matsumura Yukihiko, Sasaki Mitsuru, Okuda Kazuhide, Takami Seiichi, OharaSatoshi, Umetsu Mitsuo, et al. Supercritical water treatment of biomass forenergy and material recovery. Philadelphia, PA: Taylor & Francis; 2006 [ETATS-UNIS].

[17] Moreno J, Vargas MA, Olivares H, Rivas J, Guerrero MG. Exopolysaccharideproduction by the cyanobacterium Anabaena sp ATCC 33047 in batch andcontinuous culture. J Biotechnol 1998;60:175–82.

[18] Yalcin I, Hicsasmaz Z, Boz B, Bozoglu F. Characterization of the extracellularpolysaccharide from freshwater microalgae Chlorella sp. Lebensm WissenTechnol 1994;27:158–65.

[19] Huheihel M, Ishanu V, Tal J, Arad S. Activity of Porphyridium sp Polysaccharideagainst herpes simplex viruses in vitro and in vivo. J Biochem Biophys Methods2002;50:189–200.

[20] Itoh H, Noda H, Amano H, Cun Z, Mizuno T, Ito H. Antitumor activity andimmunological properties of marine algal polysaccharides especially fucoidanprepared from Sargassum thunbergii of Phaeophyceae. Anticancer Res1993;13:2045–52.

[21] Yang FM, Shi Y, Sheng JC, Hu QH. In vivo immunomodulatory activity ofpolysaccharides derived from Chlorella pyrenoidosa. Eur Food Res Technol2006;224:225–8.

[22] Guzmán S, Gato A, Lamela M, Freire-Garabal M, Calleja JM. Anti-inflammatoryand immunomodulatory activities of polysaccharide from Chlorellastigmatophora and Phaeodactylum tricornutum. Phytother Res2003;17:665–70.

[23] Kuhl A, Lorenzen H. Handling and culturing of Chlorella. Methods Cell Physiol1964;1:159–87.

[24] Brenna JT, Corso TN, Tobias HJ, Caimi RJ. High-precision continuous-flowisotope ratio mass spectrometry. Mass Spectrom Rev 1997;16:382 [vol. 16,1997, p. 227].

[25] Gehre M, Strauch G. High-temperature elemental analysis and pyrolysistechniques for stable isotope analysis. Rapid Commun Mass Spectrom2003;17:1497–503.

[26] Xu CB, Lad N. Production of heavy oils with high caloric values by directliquefaction of woody biomass in sub/near-critical water. Energy Fuels2008;22:635–42.

[27] O’Fallon JV, Busboom JR, Nelson ML, Gaskins CT. A direct method for fatty acidmethyl ester synthesis: application to wet meat tissues oils and feedstuffs. JAnim Sci 2007;85:1511–21.

[28] Parees DM, Hanton SD, Cornelio Clark PA, Willcox DA. Comparison of massspectrometric techniques for generating molecular weight information on aclass of ethoxylated oligomers. J Am Soc Mass Spectrom 1998;9:282–91.

[29] Bobleter O. Hydrothermal degradation of polymers derived from plants. ProgPolym Sci 1994;19:797–841.

[30] Lai YZ. Chemical degradation. In: David N-S, Hon NS, editors. Wood andcellulosic chemistry. New York: Marcel Dekker Inc; 2001. p. 443–512 [Basel].

[31] Yu G, Hu Y, Yang B, Zhao X, Wang P, Ji G, et al. Extraction, isolation andstructural characterization of polysaccharides from a red alga Gloiopeltisfurcata. J Ocean Univ China 2010;9:193–7.

[32] Hsu H-Y, Jeyashoke N, Yeh C-H, Song Y-J, Hua K-F, Chao LK.Immunostimulatory bioactivity of algal polysaccharides from Chlorellapyrenoidosa activates macrophages via toll-like receptor 4. J Agric FoodChem 2009;58:927–36.

[33] Miao X, Wu Q. High yield bio-oil production from fast pyrolysis by metaboliccontrolling of Chlorella protothecoides. J Biotechnol 2004;110:85–93.

[34] Valdez PJ, Dickinson JG, Savage PE. Characterization of product fractions fromhydrothermal liquefaction ofNannochloropsis sp. and the influence of solvents.Energy Fuels 2011;25:3235–43.

[35] Qian YJ, Zuo CJ, Tan H, He JH. Structural analysis of bio-oils from sub-andsupercritical water liquefaction of woody biomass. Energy 2007;32:196–202.

[36] Zou SP, Wu YL, Yang MD, Li C, Tong JM. Thermochemical catalytic liquefactionof the marine microalgae Dunaliella tertiolecta and characterization of bio-oils.Energy Fuels 2009;23:3753–8.

[37] Matsui T-O, Nishihara A, Ueda C, Ohtsuki M, Ikenaga N-O, Suzuki T.Liquefaction of micro-algae with iron catalyst. Fuel 1997;76:1043–8.

[38] Burczyk J, Smietana B, Terminska-Pabis K, Zych M, Kowalowski P. Comparisonof nitrogen content amino acid composition and glucosamine content of cellwalls of various chlorococcalean algae. Phytochemistry 1999;51:491–7.

[39] Jarusutthirak C, Amy G, Croué J-P. Fouling characteristics of wastewatereffluent organic matter (EfOM) isolates on NF and UF membranes.Desalination 2002;145:247–55.

[40] Dumas P, Miller L. The use of synchrotron infrared microspectroscopy inbiological and biomedical investigations. Vib Spectrosc 2003;32:3–21.

[41] Wolkers WF, Oliver AE, Tablin F, Crowe JH. A fourier-transform infraredspectroscopy study of sugar glasses. Carbohydr Res 2004;339:1077–85.

[42] Yee N, Benning LG, Phoenix VR, Ferris FG. Characterization of metal-cyanobacteria sorption reactions: a combined macroscopic and infraredspectroscopic investigation. Environ Sci Technol 2004;38:775–82.

[43] Minowa T, Zhen F, Ogi T. Cellulose decomposition in hot-compressed waterwith alkali or nickel catalyst. J Supercrit Fluids 1998;13:253–9.