in situ processing of nannochloropsis oculata algal...

Research article

In situ processing of Nannochloropsis oculata algal biomassusing a mixed bed ion-exchange resin as a heterogeneouscatalyst

Yousuf Jamal,1* Guofan Luo2 and Bryan O. Boulanger3

1Institute of Environmental Sciences and Engineering, School of Civil and Environmental Engineering, National University of Sciences and Technology,Sector H-12, Islamabad-44000, Pakistan2Environmental and Water Resources Division, Zachry Department of Civil Engineering, Texas A&M University, 3136 TAMU, College Station, TX77843, USA3Department of Civil Engineering, Ohio Northern University, Ada, OH 45810, USA

Received 7 February 2014; Revised 22 April 2014; Accepted 5 May 2014

ABSTRACT: Ion-exchange resins have the ability to convert microalgae-derived lipids into esters and other recoverableorganics. The presented research evaluated the use of mixed bed ion-exchange resins for in situ processing of Nannochloropsisoculata algal biomass to recoverable organics within a co-solvent system. The impact of various processing conditions (resinloading, resin type, biomass drying, solvent type, solvent volume, and biomass sonication) on the reaction yield of recoverableorganics was investigated. Multiple analytical chemical approaches were used to characterize lipids within the biomass and toidentify the converted reaction products. Each reaction system studied produced more than 20 recoverable organics in additionto the normally desired ester products. Phytol was observed as predominant reaction product in each system. The highest esteryield (approaching 60% biomass lipid to ester conversion) was observed with air-dried algae processed at 50 °C at a mixing rateof 550 rpm for 2 h. © 2014 Curtin University of Technology and John Wiley & Sons, Ltd.

KEYWORDS: Nannochloropsis oculata; dried algal biomass; co-solvent extraction; mixed bed ion-exchange resin; in situreaction yield

Additional supporting information may be found in the online version of this article at the publisher’s web site.

INTRODUCTION

Production of biofuels from microalgae reducesgreenhouse gas emissions, dependence on food cropsused for biofuel production, and land-water requirementsrequired for plant-based fuel production.[1,2] Microalgaegrow in all temperature zones during all seasons andabsorb more atmospheric CO2 than plant crops.[3,4]

Microalgae also demonstrate the potential to producemore lipids than vegetable plants and seed crops makingmicroalgae a valuable feed stock for biofuelproduction.[5,6]

Conventionally, biofuel production from microalgaeinvolves a multistep process using solvents to extractlipids from microalgae biomass and converting theextracted lipids into the desired biofuel throughreaction of the extracts with homogenous basic oracidic catalyst in the presence of a primary alcohol.[7]

Depending upon the composition of lipids in theextracts (triglycerides, diglycerides, monoglycerides,phospholipids, sterols, glycolipids, and free fatty acids)additional purification of the extracts may be requiredto increase biodiesel yield and to meet AmericanSociety for Testing and Materials requirements forbiodiesel composition.[8,9]

A sustainable net positive energy gain for theproduction of alkyl esters (biodiesel) from microalgaeis directly related to the extraction efficiency of lipidsfrom biomass.[10] The two most commonly reportedmethods for extracting lipids from microalgae are theFolch and Bligh and Dryer methods, which use achloroform-methanol co-solvent mixture.[11,12] Suitablelipid extraction efficiencies are also observed whenmethyl-tert-butyl ether, dichloromethane, and hexanereplace the use of chloroform in the methanol co-solventsystem.[7,9,13] Drying the algae and disrupting thecells through thawing, bead beating, sonication,mechanical disruption, microwave irradiation, and useof supercritical conditions also increase the lipidextraction yield.[4,7,14–19]

*Correspondence to: Yousuf Jamal, Institute of EnvironmentalSciences and Engineering, School of Civil and EnvironmentalEngineering, National University of Sciences and Technology, SectorH-12, Islamabad, Pakistan. E-mail: [email protected]

© 2014 Curtin University of Technology and John Wiley & Sons, Ltd.Curtin University is a trademark of Curtin University of Technology

ASIA-PACIFIC JOURNAL OF CHEMICAL ENGINEERINGAsia-Pac. J. Chem. Eng. (2014)Published online in Wiley Online Library(wileyonlinelibrary.com) DOI: 10.1002/apj.1827

Even with increased extraction efficiency,conventional processing of lipids based upon amultiple step process is time and energy intensive.Approximately 90% of processing energy consumedduring the conversion of microalgae into biofuels isfrom the extraction step.[1] Moreover, extraction byorganic solvents results in incomplete extraction.[20]

Therefore, in situ processing of microalgae to alkyl estersis an attractive alternative to conventional processingbecause this method simultaneously extracts andconverts the recoverable biomass biomolecules intobiofuel within a single reaction system. In situ processingreduces processing time and cost while demonstratingsubstantial reaction yield.[21]

The same co-solvent mixtures used in conventionalprocessing are also used in in situ processing; however,the homogeneous catalyst and alcohol are addedsimultaneously.[22,23] Because homogeneous catalysts areconsumed during their reaction, the uses of heterogeneouscatalysts within in situ reaction systems are of interest.Heterogeneous catalysts have the ability to be recoveredand reused, further decreasing the processing costs whileobtaining substantial reaction yields.[24]

Li et al. report on the use of in situ heterogeneoustransesterification of algae lipids using an amendedSoxhlet extractor with a methanol-dichloromethane co-solvent system in the presence of a magnesium-zirchonium basic solid catalyst.[9] However, the amendedSoxhlet system was not true in situ processing as thesolvent was recirculated through the algae biomass whilethe transesterification reactions occurred in a separatevessel in the Soxhlet system. Similarly, a two-step processof algae lipids solvent extraction and esterification withheterogeneous acid catalyst and then transesterificationof the deacidified oil with KOH in a separate vessel isalso reported in the literature.[8] Umdu et al. reported a23% ester conversion of Soxhlet hexane extractedNannochloropsis oculata derived lipids at 80 °C, whileusing 80% mixed oxide (Al2O3/CaO or MgO) catalystloading for transesterification reaction in a separate vesselat 50 °C.[25] While these efforts are notable, the approachdecoupled the extraction and conversion.The purpose of this work was to investigate in situ

conversion products and ester yield when N. oculataalgal biomass was reacted withmixed bed heterogeneousion-exchange resin within a single reaction vesseloperated under various reaction conditions. Thepresented research is a step toward establishing a truesingle-step processing method to convert microalgaebiomass to recoverable organics with a broad rangeof applications.

METHODS

A summary schematic highlighting the processingsteps carried out within the experimental approach is

provided in Figure S1 of the supporting information.Detailed descriptions of the materials and methodsfollowed throughout the research are presented withinthis Methods section.

Materials

Air-dried and sun-dried N. oculata algal biomasssamples were supplied by Algeternal Technologies,LLC (TX, USA). The algal biomass samples wereimmediately pulverized using a mortar and pestle andused in experiments without further drying. A portionof each sample was evaluated for initial moisturecontent by drying the sample in an oven operated at105 °C for 24 h. A second portion of each sample wascharacterized for lipid content. High-performanceliquid chromatography (HPLC) grade methanol, HPLCgrade hexane, HPLC grade heptanes, HPLC gradediethyl ether, HPLC grade acetone, and glacial aceticacid were purchased from VWR International(Sugarland, TX, USA). Methyl heptadecanoate(internal standard), 1 amp lipid mixture, oleic acid(≥99% purity), and thin-layer chromatography (TLC)plates were purchased from Sigma Aldrich (St. Louis,MO, USA).The two types of mixed bed resin systems utilized in

this research, a macroporous resin system consisting ofequal mass portions of Amberlyst A26 OH and DowexMonosphere M-31 and a gelular resin systemconsisting of Dowex Monosphere MR-450 UPW, werepurchased from Sigma Aldrich (St. Louis, MO, USA).Nitrogen gas (99% purity) was purchased fromBOTCO (Bryan, TX, USA). The 500mg silica solid-phase extraction (SPE) cartridges were purchased fromSilicaPrepTM (Quebec, Canada).

Algae characterization

The polar lipid fraction, non-polar lipid fraction, andlipid profiles within each fraction of all received algalsamples were characterized through solvent and SPE,gravimetric measurements, TLC, and matrix-assisted layerdesorption/ionization–time-of-flight (MALDI-TOF).The polar lipid fraction of each sample was

determined by extracting 1 g algae in 10mL methanolin a 15mL flat bottom Erlenmeyer flask topped with aglass stopper. The flask was placed on a shaker for5min and then heated in a water bath for 30min at65 °C. The resulting supernatant was decanted andpoured into a pre-weighed clean glass vial. Thesupernatant was then dried under a gentle stream ofnitrogen. The weight of the remaining product wasrecorded as the total polar lipid fraction. The non-polarlipid fraction was determined in a similar manner,except 10mL of a 3 : 1 hexane:chloroform solventmixture was added following decantation of themethanol. The polar and non-polar dried fractions were

Y. JAMAL, G. LUO AND B. BOULANGER Asia-Pacific Journal of Chemical Engineering

© 2014 Curtin University of Technology and John Wiley & Sons, Ltd. Asia-Pac. J. Chem. Eng. (2014)DOI: 10.1002/apj

then evaporated to dryness with nitrogen, re-suspendedin 0.5mL solvent (chloroform for TLC and hexane forMADLI-TOF), and pooled together.Thin-layer chromatography and MALDI-TOF were

then used to explore the lipid profile within the pooledextracts. TLC plates were spotted with an increasingamount of lipid standard mix (1–10μg) to produce astandard curve. 5μL of the re-suspended extracts werespotted onto the TLC plates and developed with ahexane:diethyl ether:glacial acetic acid (80 : 20 : 1)solvent mixture to resolve triglycerides, diglycerides,monoglycerides, and fatty acids. The developed TLCplate was placed in a glass visualization chamber withiodine crystals in order to visualize the separatedcomponents. The area of the algae lipids spotted onthe developed TLC plate was determined using ImageJ software by noting down spot pixel densities. Bealet al. validated the use of Image J software fordetermination of various algae lipid composition.[26]

This extraction and video imaging technique was alsoreported earlier for phospholipid presence andquantification in other biological samples.[27]

Matrix-assisted layer desorption/ionization wasperformed on both SPE separated and non-separatedpooled lipid extracts to characterize the lipid profileswithin the combined samples and within each fraction(polar and non-polar). SPE separation was achievedby loading 300μL of the pooled extract onto a500mg silica SPE cartridge that was pretreated with1mL hexane. The cartridge was then eluted with2mL of 80 : 20 : 1 hexane:diethyl ether:glacial aceticacid to separate out the non-polar fraction. Thecollected fraction was evaporated to dryness under aflow of nitrogen and reconstituted in 50μL of acetone.The polar lipid fraction was eluded from the cartridgeusing 2mL acetone, and the eluent was collected.The two collected fractions (both in acetone) werethen analyzed using MALDI-TOF to examine thelipid profile in each fraction. MALDI was performedon a Voyager STR equipped with a nitrogen laser(337 nm, 3 ns pulse, and 20Hz maximum firing rate).The instrument was operated in the reflectron modewith 2,4,6-trihydroxyacetophenone monohydrate asthe matrix.

Co-solvent extraction capacity

Triplicate samples of pulverized algal biomass were co-solvent extracted to evaluate the lipid extraction yieldoccurring during in situ conversion. For each extraction4mL of either 60 : 40, 40 : 60, and 20 : 80 (volume:volume), methanol:hexane co-solvent was used. Allindividual reactions were carried out in 10mL pre-cleaned glass vial with inserted stir bars. The reactorswere placed into a stirring water bath operated at50 °C at atmospheric pressure with a 550 rpm mixingrate. Samples were extracted for either 2, 4, 6, or 10 h.

Following extraction, the supernatant of eachreactor was collected via a syringe and passed througha 5–10μm filter (Fisher Brand Qualitative P5) into anew pre-cleaned 10mL vial. The amount of filtratecollected was weighed and the entire sample wasevaporated. The resulting dried sample was thenweighed to determine the solvent extractable mass.The extractable yield was then presented as the solventextractable mass normalized to the initial mass of thepulverized algae used in the extraction. Additionalfiltered supernatant for the 40 : 60 methanol:hexaneextraction co-solvent was then stored for recoverableorganic analysis.

In situ conversion experiments

In situ conversion experiments were carried out in4mL of 40 : 60 methanol:hexane co-solvent with aspecified mass loading of mixed bed resin added tothe 10mL pre-cleaned glass reaction vial with aninternal stir bar. Prepared reactors were placed in thestirring water bath and operated at 50 °C at atmosphericpressure with a 550 rpm mixing rate.Experiments were conducted to determine the effect

of (i) sonication, (ii) co-solvent volume, (iii) the dryingtechnique, and (iv) mixed bed resin structure (gelularversus macroporous resin system) on in situ conversionyield. The effect of sonication was carried out bysonicating triplicate one gram samples of pulverizedair-dried algae in 4mL of 40 : 60 methanol:hexane co-solvent. The samples were sonicated for 1 h and thenreacted for 1 h in the presence of 20%, 40%, or 60%Dowex Monosphere MR-450 UPW resin loading.The in situ reaction yield of the sonicated sampleswas then compared with the yield of non-sonicatedair-dried algae samples.The effect of co-solvent volume on in situ yield was

evaluated by increasing the 40 : 60 methanol:hexaneco-solvent from 4 to 10mL for a triplicate set ofsamples. One gram samples of pulverized sun-driedalgae in 4mL and 10mL of 40 : 60 methanol:hexaneco-solvent were reacted in the presence of 20%, 40%,or 60% Dowex Monosphere MR-450 UPW resinloading. The in situ reaction yield of the increased co-solvent volume samples was then compared to the yieldof low volume samples.The effect of the algae drying process on in situ yield

was evaluated by processing triplicate sun-dried algaesamples with 4mL of 40 : 60 methanol:hexane co-solvent with a 20%, 40%, or 60% resin loading. Theresulting in situ yield was then compared to the yieldobserved for the air-dried algae. Finally, the effect ofmixed bed resin structure was evaluated by comparingthe performance of the gelular against the macroporousmixed bed resin. One gram of pulverized algae wasreacted with 20%, 40%, or 60% mixed bed resinloading in 4mL of 40 : 60 methanol:hexane co-solvent.

Asia-Pacific Journal of Chemical Engineering IN SITU PROCESSING OF N.OCULATA BY ION-EXCHANGE RESINS


The resulting in situ yields from both resin types arecompared. The experiments were carried out for bothair-dried and sun-dried algae.Supernatant realized from each reactor in the

experiments mentioned in the preceding paragraphswere syringe filtered and placed into a clean 4mL vialand stored at 4 °C until fatty acid methyl esters(FAME) analysis is conducted. The resulting FAMEanalysis of each sample was then used to calculate thein situ conversion yield. The resulting yields for eachparameter evaluation (sonication, co-solvent volume,algae drying technique, and resin structure) across resinloadings were compared for statistical significance ofthe means using a student’s t-test at a 95% confidenceinterval. All statistical analysis was carried out usingSPSS version 20.Experiments were also carried out to determine

reaction product formation as a function of reactionduration for duplicate one gram samples of sun-driedpulverized algae in co-solvent with a 20% macroporousresin loading. The reaction product experiments wereconducted at intervals of 2, 4, 6, and 10 h of reactionwith and without the presence of mixed bed resin.

Fatty acid methyl esters analysis

The ester content in all evaluated samples wasdetermined according to according to the EuropeanStandard method EN 14103.[28,29] A Thermo Trace GCUltra gas chromatograph (Thermo Electron Corporation)coupled to a Thermo DSQ II mass spectrometer wasused to chromatographically resolve and quantify theester content within injected samples using an internalstandard method.A 1μL splitless injection was introduced on a Restek

RxiTM-5ms column (60m× 0.25mm ID× 0.25μmfilm thickness) at an inlet temperature of 225 °C withhelium as the carrier gas (flow rate of 1.5mL/min).The oven temperature gradient operated from 50 °Cheld for 5min, ramped linearly to 320 °C at a rate of20 °C/min, and held at 320 °C for 5min. The ion sourceand transfer line temperatures weremaintained at 250 °C.For analysis, 600μL aliquots of syringe filtered

supernatant from individual samples were transferredto a pre-cleaned 4mL vial for evaporation under agentle stream of nitrogen. The resulting dried extractmass was weighed, and additional 50μL aliquots ofsupernatant were transferred to the vial and re-evaporated until the final evaporated mass of theextracted sample was 10mg. The 10mg mass ofextracted sample was then prepared for analysis toestablish the recoverable organics content within the10mg of dried extract.The 10mg of dried extracts were reconstituted using

1mL of n-heptane to establish a resulting solutionconcentration of 10mg dried extract per mL n-heptane.Methyl heptadecanoate was then spiked into the

reconstituted extracts at a concentration of 0.04mg/mLfor use as an internal standard. Xcaliber version 2.0.7was used to calculate the peak area, height of theidentified ester peaks, and the resulting concentration ofeach individual FAME within the reconstituted sample.The resulting percent in situ yield was then calculatedfollowing FAME analysis.

Identification and analysis of recoverableorganics

Analysis of other recoverable organics present in thereaction solution was conducted similarly to theanalytical method used to quantitate esters. Thismethod was considered semi-quantitative in nature,because the response factors for all recoverableorganics identified within a reaction solution were notknown against the internal standard.The peak area and peak height of identifiable peaks

were used to approximate a concentration of theidentified analyte within the reaction solution assuminga uniform response against the internal standard.Because of the semi-quantitative nature of the methodand the assumption of uniform response factors, theidentified compounds were grouped into type prior todetermining the percent of each observed group in asample. The groups included alcohols, ketones,aldehydes, and alkanes (broadly defined). Subgroupsof particular interest included esters and phytols.

RESULTS AND DISCUSSION

Algae characterization results

The initial moisture content of the supplied algalbiomass was found 7.0 and 4.0 mass% for the air-driedand sun-dried algae, respectively. The mean polar lipidand non-polar lipid mass fraction of investigated N.oculata samples was determined to be 113 ± 9.1 and11.4 ± 1.3mglipid/galgae, respectively. Converti et al.reported similar total lipid content 134mglipid/galgae inN. oculata at growth conditions of 15 °C and 0.150 g/LNaNO3.

[6]

Because the air-dried and sun-dried algae used in theexperiments were grown under the same conditions,the effect of the drying method was evident within thisdata set. Air-dried algae demonstrated a higher totallipid yield (mean yield = 142.0mglipid/galgae) comparedwith sun-dried algae (mean yield = 120.0mglipid/ galgae).Literature sources also demonstrate that sun dryingreduces the lipid content within algae.[24,30]

Thin-layer chromatography analysis was repeated ontwo occasions to determine the initial triglyceride,diglycerides, monoglycerides, and fatty acid contentwithin the air-dried and sun-dried samples. Anexample TLC plate result is provided in the



supportinginformation (Figure S2). TLC analysisshowed a distribution within the samples betweentriglycerides (15.6mgtriglyceride/galgae for air driedand 14.1mgtriglyceride/galgae for sun dried), digly-cerides (10.2mgdiglycerides/galgae for air dried and14.7mgdiglyceride/galgae for sun dried), and fatty acids(49.4mgFFA/galgae for air dried and 37.5mgFFA/galgaefor sun dried). Monoglyceride content was notincluded within the distribution due to a complicationwith standardization; however, it appears thatmonoglycerides was prevalent in greater amounts inthe air-dried sample based upon TLC spottinganalysis (Figure S2).Matrix-assisted layer desorption/ionization–time-of-

flight analysis was used to identify the compositionof lipids within the separated extract fractions.Representative mass spectra for both the polar andnon-polar lipid extracts are provided in Figure S3 ofthe supporting information. Because a TOF instrumentwas used, spectral libraries were used to determine theexact mass of the triglycerides and fatty acids present inthe extracts. The composition profile based uponspectral library analysis for the polar and non-polarextracts is provided in Table 1.[31,32]

Co-solvent extraction capacity

The co-solvent extraction capacity for air-dried algae wasdetermined to be 5.5× 10� 2± 1.7 × 10� 3glipid/galgaefor the 60 : 40 methanol:hexane co-solvent, 6.0 × 10� 2-± 9.1 × 10� 3 glipid/galgae for the 40 : 60methanol:hexaneco-solvent, and 1.4 × 10� 2±6.8 × 10� 4 glipid/galgaefor the 20 : 80 methanol:hexane co-solvent. The highestlipid extraction occurred for the 40 : 60 methanol:hexaneco-solvent system. A 40 : 60 methanol:hexane solventratio was also observed to yield the highest lipidextraction in Li et al. work with Chlorellapyrenoidosa.[22] Therefore, the 40 : 60 methanol:hexaneco-solvent system was used for the remainder of theexperiments. The 40 : 60 methanol:hexane co-solventextraction capacity for sun-dried algae, sonicated air-dried algae, and for 10mL extraction of sun-dried algaewas determined to be 3.9 × 10� 2±9.1 × 10� 3,9.85 × 10� 2 ± 3.5 × 10� 3, and 19 × 10� 2 ± 4.2 ×10� 3 glipid/galgae, respectively.The resulting co-solvent extraction capacity can also

be used to evaluate the maximum practical yield basedupon the extraction solvent used in the system. Themean practical conversion yield from the algae wascalculated to be 63mgester/galgae for air dried and41mgester/galgae for sun dried. This amount is 44%and 34% of the theoretical yield (respectively) thatwas based upon the total lipid extraction usingmethanol and a mixture of hexane and chloroform.The 40 : 60 methanol:hexane solvent extraction mixtureused for this research provides methanol also for thecatalysis reaction and reduces the amount of extractable

material available from the algal biomass for thereaction thus resulting in lower ester yield mgester/galgaewith mixed bed resin system.

In situ extraction

Table 2 provides a summary of factors effecting the insitu conversion of algae biomass into esters on a massbasis. The method used to dry the algae (forced air vssun drying), the structure of the resin (gelular vsmacroporous), sonication, and solvent volume wereall observed to have an effect on mean ester contentof the reaction product solution. The observeddifference in ester yield as a function of algaedrying was caused because sun drying algae not onlyimpact the composition but also the content of lipidswithin the sun-dried algal biomass.[24,30] Table 3provides the calculated percent in situ reaction yieldunder the same reaction conditions for air-dried andsun-dried algae.

Table 1. Lipid profile within non-polar and polar solid-phase extraction extracts as determined throughmatrix-assisted laser desorption/ionization–time-of-flight.

Extractfraction Identified components (nominal m/z)

Polar Fatty acids• Hendecanoic acid (207.5)• c-9,12-octadecadienoic acid (301.6)• Tricosanoic acid (375.6)Diglycerides• CyM, LaCa, or PCo (433.5)Triglycerides• CoCoCy, BuBuLa,VVCa or CyCyV(427.8)• SSCo, PPCa, LaLaS, or MMM (744.1)

Non-polar Fatty acids• Hendecanoic acid (207.5)• c-9,12-octadecadienoic acid (301.6)• Tricosanoic acid (375.6)• Eicosanoic acid (335.7)Diglycerides• LnS or LO (623.9)Triglycerides• VVLn (543.7)• VVS, CoCoP, EnEnM, CyCyLa, CaCaCy,BuBuA, or LaLaBu (549.8)• LaLaEn (591.9)• EnEnL (601.8)• LLS or OOL (904.1)• SSO (911.1)• SSA or AAP (940.1)

A, eicosanoic acid; Bu, butanoic acid; Ca, decanoic acid; Co,hexanoic acid; Cy, octanoic acid; En, heptanoic acid; L, c-9,12-octadecadienoic acid; La, dodecanoic acid; Ln, c-9,12,15-octadecatrienoic acid; M, tetradecanoic acid; O, c-9-octadecenoic acid;P, hexadecanoic acid; S, octadecanoic acid; and V, pentanoic acid.



Sonication and solvent volume were both observedto have a statistically significant effect on ester yieldduring in situ processing of algae with the mixed bedresin system. Sonicating the algae in the presence ofsolvent and increasing the solvent volume both actedto increase the amount of lipids released into the bulksolution by increasing lipid extraction efficiency.[4]

Increasing the extraction solvent volume, however,did not enhance ester yield. One possible explanationfor this result offered within the literature suggests thatincreased yield was not observed within our reactionsystem because the esters further reacted with theacidic resin. This behavior was previously noted forthe reaction of phytol in a similar test system.[33,34]

Another interesting observation was a statisticallysignificant decrease in ester yield as the catalyst loadingincreased. At first, this decreasing trend was proposedto be caused by esters either adsorbing onto the resin orabsorbing into the resins. While this may be occurring,the data also indicated that the esters could be reactionintermediates in the pathway to other organics as estersappear to undergo additional reaction with the resin.In consideration of both sorption and reactive

intermediates as likely explanations for the decreasingtrend of ester production as a function of catalyst

loading, the reactive intermediate explanation appearsto be more plausible as the ester yield decreases morefor gelular resins than for macroporous resins as afunction of catalyst loading. Because gelular resinshave limited porosity to facilitate pore site reactions,if sorption were of significance in this system, thegelular resin should have higher ester yield at highercatalyst loadings. However, based upon the data, thereverse appears to be true. Presence of additional esterbased metabolites was also observed in the reactionsolution; favoring that the observed trend is based uponthe role of esters as a reactive intermediate.

In situ conversion reaction products

Table 4 (text) and Figure S4 (supporting information)provide information concerning the reaction productsobserved during the in situ reaction of N. oculata withmixed bed resin at a 20% catalyst loading at 50 °Cand 550 rpm. The evolved products show formationof esters, alcohols, and ketones. Interestingly, phytolrepresents one of the largest single peaks in the systemoccurring at a retention time of approximately17.33min. The unexpected presence of phytol, otheralcohols, and ketones within the reaction solutionpointed out the complexity of the underlying multi-component reactions occurring in the system. Futureadditional research work could examine the reactionof each individual product with the acidic or basicfunctional group in the mixed bed resin system.Figure 1 provides the change in relative contribution

of each ester peak during the course of the reaction fora 20% resin loading. C11–C15 esters were present inthe reaction solution at each time step, but at verylow concentrations (below 1% of the total estercontent). The reaction solution was dominated by thepresence of C18 esters and also contains C16 esters.C18 esters decreased over time, hinting at the additionalreactivity of the esters to the mixed bed resin.

Table 2. Ester produced per dry weight of air and sun-dried algae (mgester/galgae) at different catalyst loadings.

Experiment Resin type

Algaldryingmethod

Solventvolume(mL) Sonicated

Catalyst loading (%) Statisticallysimilarmeans20 40 60

Effect of algae drying Gelular Air 4 No 37.2 9.7 1.0 NoGelular Sun 4 No 1.4 0.9 0.2Macroporous Air 4 No 20.3 9.8 3.2 NoMacroporous sun 4 No 8.9 4.0 2.4

Effect of resin type Gelular Air 4 No 37.2 9.7 1.0 NoMacroporous Air 4 No 20.3 9.8 3.2Gelular Sun 4 No 1.4 0.9 0.2 NoMacroporous Sun 4 No 8.9 4.0 2.4

Effect of sonication Gelular Air 4 Yes 39.8 24.5 14.6 NoGelular Air 4 No 37.2 9.7 0.6

Effect of Solvent Gelular Sun 10 No 2.5 1.4 1.4 NoGelular Sun 4 No 1.4 0.9 0.2

(Ester produced, mgester/galgae)

Table 3. Percent in situ reaction yield at differentcatalyst loadings for air and sun-dried algae.

Resin type

Algaldryingmethod

Solventvolume(mL)

Catalyst loading (%)

20 40 60

Gelular Air 4 59.3 15.4 1.6Gelular Sun 4 3.4 2.2 0.5Macroporous Air 4 32.4 15.4 5.1Macroporous Sun 4 21.9 9.8 6.0

Percent in situreaction yield



Several possible explanations for the decrease ofesters observed in the reactors over time are providedin the literature.[34,35] A hydrolytic effect in thepresence of both acidic and basic resin is documentedby Chung et al.[34] When moisture is present, esters canfurther react to other by-products including alcohols,

ketones, and ethers.[36–39] Additionally, evolved glycerolmay reacts with the fatty acids and the acidic resinforming monoglycerides that impact ester yield.[40]

Interestingly, Fig. 2 identifies several by-products ofadditional ester reaction observed within the reactionsystem (including alcohols, ketones, and alkanes). These

Table 4. Representative identified reaction products of in situ reaction ofN.oculata with 20% mixed bed resin in methanol:hexane co-solvent.

Peak retentiontime (min) %Area Identified analyte

14.13 0.01 β-ionone14.26 0.37 Butylated hydroxytoluene15.14 0.08 3-Heptadecene15.24 2.57 Heptadecane15.33 0.04 2,6,10-Trimethyltetradecane15.37 0.59 Tetradecanoic acid methyl ester15.65 0.51 4,8,12-Trimethyl tridecanoic acid methyl ester15.71 0.09 Pentadecanoic acid methyl ester15.76 0.02 12-Methyl tetradecanoic acid methyl ester15.91 0.03 Pentadecanoic acid methyl ester16 0.56 3,7,11,15-Tetra methyl-2-hexadecene-1-ol16.05 7.12 2-Pentadecanone16.13 0.11 3,7,11,15-Tetramethyl-2-hexadecen-1-ol16.22 0.23 3,7,11,15-Tetra methyl-2-hexadecene-1-ol16.27 2.35 Unidentified peak16.3 0.43 7,10-Hexadecadienoic acid methyl ester16.34 1.54 7,10,13-Hexadecatrienoic acid methyl ester16.43 15.23 Hexadecanoic acid methyl ester16.49 0.07 Unidentified peak16.55 0.07 Unidentified peak16.83 0.13 Unidentified peak16.9 1.15 Heptadecanoic acid methyl ester*17.2 0.09 10,13-Octadecadiynoic acid methyl ester17.27 8.9 9,12-Octadecadienoic acid methyl ester17.29 4.12 9-Octadecenoic acid methyl ester17.31 10.64 9,12,15-Octadecatrienoic acid methyl ester17.34 37.89 Phytol18.08 0.12 Unidentified peak18.9 1.74 Unidentified peak19.74 2.96 Unidentified peak20.44 0.18 Unidentified peak

*Internal standard.

Figure 1. Changes in ester formation over time during the insitu processing of N. oculata algal biomass with mixed bedion-exchange resin.

Figure 2. Change in reaction products during the course ofthe reaction.



additional by-products increased in concentration whileester content decreased at later stages of the reaction.

CONCLUSIONS

Simultaneous extraction and conversion of algalbiomass recoverable organics to esters and otherreaction products is achievable in a methanol:hexaneco-solvent system with mixed bed ion-exchange resin.The algae drying method, resin type and characteristics,sonication, and solvent volume all observed to have animpact on the mean ester content and reaction yield.This presented research also demonstrates that the

availability of acidic sites near or at the resin surface alsoimpacts ester formation. Higher rates of ester formationwere shown to occur in the mixed bed gelular-type ion-exchange resins compared with the macroporous resin.The maximum ester formation over ion-exchange resinsalso was documented to occur in the first 2 h of thereaction with a nearly constant yield in the remaininghours. This observation is caused by the variation ofthe molar ratio of alcohol and lipids within the systemat any point of time.[41] Finally, a decreasing trend inester yield was observed as catalyst loading and reactiontime was increased. Decrease in esters, while increase inother reaction products (including alcohols and ketones)indicates that the esters can undergo further reactivityover mixed bed resin.

Acknowledgements

The authors are thankful to the US Department of State,Bureau of Educational and Cultural Affairs FulbrightProgram and the Texas Engineering Experiment(project number 32296-19386) Station for providingthe research grant to support this research.

REFERENCES

[1] L. Lardon, A. Hélias, B. Sialve, J.P. Steyer, O. Bernard.Environ. Sci. Technol., 2009; 43, 6475–6481.

[2] C. Lohrey, V. Kochergin. Bioresour. Technol., 2012; 108,76–82.

[3] L. Gouveia, A. Oliveira. J. Ind. Microbiol. Biotechnol., 2009;36, 269–274.

[4] E. Ryckebosch, K. Muylaert, I. Foubert. J. Am. Oil Chem.Soc., 2012; 89, 189–198.

[5] S.Y. Chiu, C.Y. Kao, M.T. Tsai, S.C. Ong, C.H. Chen, C.S.Lin. Bioresour. Technol., 2009; 100, 833–838.

[6] A. Converti, A.A. Casazza, E.Y. Ortiz, P. Perego, M. DelBorghi. Chem. Eng. Process. Process Intensif., 2009; 48,1146–1151.

[7] B.J. Krohn, C.V. McNeff, B. Yan, D. Nowlan. Bioresour.Technol., 2011; 102, 94–100.

[8] T. Dong, J. Wang, C. Miao, Y. Zheng, S. Chen. Bioresour.Technol., 2013; 136, 8–15.

[9] Y. Li, S. Lian, D. Tong, R. Song, W. Yang, Y. Fan, R. Qing,C. Hu. Appl Energ, 2011; 88, 3313–3317.

[10] S. Haye, C.S. Hardtke. Trends Plant Sci., 2009; 14, 409–412.

[11] J. Folch, M. Lees, G.H. Sloane Stanley. J. Biol. Chem., 1957;226, 497–509.

[12] E.G. Bligh, W.J. Dyer. Can. J. Biochem. Physiol., 1959; 37,911–917.

[13] V. Matyash, G. Liebisch, T.V. Kurzchalia, A. Shevchenko,D. Schwudke. J. Lipid Res., 2008, 49.

[14] S.J. Lee, B.D. Yoon, H.M. Oh. Biotechnol. Tech., 1998; 12,553–556.

[15] F. Adam, M. Abert-Vian, G. Peltier, F. Chemat. Bioresour.Technol., 2012; 114, 457–465.

[16] P. Prabakaran, A.D. Ravindran. Lett. Appl. Microbiol., 2011;53, 150–154.

[17] J. Cheng, T. Yu, T. Li, J. Zhou, K. Cen. Bioresour. Technol.,2013; 131, 531–535.

[18] P. Patil, H. Reddy, T. Muppaneni, S. Ponnusamy, Y. Sun,P. Dailey, P. Cooke, U. Patil, S. Deng. Bioresour. Technol.,2013; 137, 278–285.

[19] R. Halim, B. Gladman, M.K. Danquah, P.A. Webley.Bioresour. Technol., 2011; 102, 178–185.

[20] J. McNichol, K.M. MacDougall, J.E. Melanson, P.J. McGinn.Lipids, 2012; 47, 195–207.

[21] E.A. Ehimen, Z.F. Sun, C.G. Carrington. Fuel, 2010; 89,677–684.

[22] P. Li, X. Miao, R. Li, J. Zhong. J. Biomed. Biotechnol., 2011;2011, 8.

[23] B.D. Wahlen, R.M. Willis, L.C. Seefeldt. Bioresour.Technol., 2011; 102, 2724–2730.

[24] R.K. Balasubramanian. Heterogeneous catalysis of plantderived oils to biodiesel, National University of Singapore,Singapore, 2010; pp. 234.

[25] E.S. Umdu, M. Tuncer, E. Seker. Bioresour. Technol., 2009;100, 2828–2831.

[26] C.M. Beal, R.E. Hebner, D. Romanovicz, C.C. Mayer,R. Connelly. Renew. Energy, 2013; 50, 86–93.

[27] K. Dyńska Kukulska, W. Ciesielski. Rev. Anal. Chem., 2012;31, 43–56.

[28] F. Munari, D. Cavagnino, A. Cadoppi. Determination ofTotal FAME and Linolenic Acid Methyl Ester in PureBiodiesel (B100) by GC in Compliance with EN14103, Thermo Fisher Scientific Milan, Italy, 2007;pp. 4.

[29] W. Ralf, A. Bähren. Application Book Volume 4: BiodieselQuality Control, Vol. 4, Shimadzu Europa Gmbh, Duisburg,Germany, 2008; pp. 26.

[30] A. Widjaja, C.C. Chien, Y.H. Ju. J. Taiwan Inst. Chem. Eng.,2009; 40, 13–20.

[31] W.C. Byrdwell. Taking Lipid Analysis into the 21st Century.http://www.byrdwell.com/Triacylglycerols/ Accessed on July17, 2012.

[32] M.A. Danielewicz, L.A. Anderson, A.K. Franz. J. Lipid Res.,2011; 52, 2101–2018.

[33] N.I. Tracy, D.W. Crunkleton, G.L. Price. Fuel, 2010; 89,3493–3497.

[34] E.A. Chung, C.W. Cho, K.H. Ahn. J. Org. Chem., 1998; 63,7590–7591.

[35] T. Danuthai, S. Jongpatiwut, T. Rirksomboon, S. Osuwan,D.E. Resasco. Appl. Catal., A, 2009; 361, 99–105.

[36] M.A. Harmer, Q. Sun. Appl. Catal., A, 2001; 221, 45–62.[37] A. Corma, M. Renz, C. Schaverien. ChemSusChem, 2008; 1,

739–741.[38] J.C. Serrano-Ruiz, A. Pineda, A.M. Balu, R. Luque, J.M.

Campelo, A.A. Romero, J.M. Ramos-Fernández. Catal.Today., 2012; 195, 162–168.

[39] D.M. Alonso, J.Q. Bond, J.C. Serrano-Ruiz, J.A. Dumesic.Green Chem., 2010; 12, 992–999.

[40] S. Abro, Y. Pouilloux, J. Barrault. Stud. Surf. Sci. Catal.,1997; 108, 539–546.

[41] A. Galia, O. Scialdone, E. Tortorici. J. Supercrit. Fluids.,2011; 56, 186–193.

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