Green Chemistry

PAPER

Cite this: DOI: 10.1039/c5gc00187k

Received 27th January 2015,Accepted 19th February 2015

DOI: 10.1039/c5gc00187k

www.rsc.org/greenchem

Promoting catalysis and high-value productstreams by in situ hydroxyapatite crystallizationduring hydrothermal liquefaction of microalgaecultivated with reclaimed nutrients†

Griffin W. Roberts,a Belinda S. M. Sturm,b Umar Hamdeh,a Gabriel E. Stanton,a

Alejandra Rocha,a Tiffany L. Kinsella,a Marie-Odile P. Fortier,b Shayan Sazdar,a

Michael S. Detamorea and Susan M. Stagg-Williams*a

Although algal biofuels hold great potential for renewable energy, production costs limit widespread tech-

nology adoption. Co-producing high-value products can ensure economic viability. We have discovered

subcritical water will simultaneously convert algae, grown with reclaimed nutrients, into pure-phase sub-

stituted hydroxyapatite nanocrystals and a petroleum-like biocrude. The hydroxyapatite contains substi-

tutions of carbonate, silicate, and magnesium, and can be easily modified to produce varying ratios of

hydroxyapatite and tricalcium phosphate. The crystallization process is shown to undergo a nanoscale

hierarchical order from long hexagonal crystals which aggregate to bundles, sheets, and flower-like

microstructures. The hydroxyapatite promotes in situ catalytic upgrading of the biocrude product, par-

ticularly, the dehydration of fatty acid amides. Overall, in situ oil upgrading provides a superior quality bio-

crude by reducing the oxygen content to <6.7% and resulting in >96% of the oil boiling below 600 °C.

Major compounds found within the biocrude include phenolics and unsaturated hydrocarbons. In

addition to heterogeneous catalysis, the hydroxyapatite product has significant promise for biomedical

engineering applications. Herein, we demonstrate live cell-adhesion of human Wharton’s jelly cells

through extended filopodia on the hydroxyapatite product. This discovery establishes a new paradigm for

water and nutrient reclamation systems and algae-based fuels and chemicals by producing a versatile

high-value product, substituted hydroxyapatite, which can integrate into multiple markets and rapidly

improve economic feasibility for algae biofuels.

Introduction

Multiple pathways have been investigated to understand theviability of algae as a commercial-scale biofuel feedstock.1–3

Thermochemical conversion of the whole cell can producehigh yields while minimizing dewatering and extraction costs.4

Hydrothermal liquefaction (HTL) utilizes subcritical waterchemistry to convert biomass to a carbon-rich biocrude.4,5 Sub-critical water properties include a decreased dielectric con-stant, increased ionic product, and decreased density,6

providing an acid–base reaction media.7 During HTL of algae,lipids, proteins and carbohydrates are converted to smallermolecules that can be recovered as biocrude with similar com-position and properties to petroleum crude.5,8,9 A variety ofalgal species have been converted to biocrude through HTL,4

but there is no fundamental understanding of how algalgrowth conditions impact HTL yields and product quality.

Many studies use algae from commercial-scale farms thatprimarily cultivate algae for the nutraceutical industry at amarket price that is unrealistic for a fuel and chemical feed-stock. Beyond capital cost, large-scale algal cultivationdemands significant amounts of water and nutrients, whichare major concerns for biofuel sustainability as identified bythe National Research Council of the National Academies.3

Reclamation water and/or seawater can be used to lower theenvironmental costs of cultivation,10,11 particularly when algalgrowth is coupled to nutrient removal from wastewater,referred to from here on as reclaimed nutrient algae (RN-algae). However, the growth conditions typical of municipal

†Electronic supplementary information (ESI) available: Information and figuresof the ultimate analysis of the HAp product, complete list of compounds identi-fied by GC-MS including the chromatogram of the derivatized biocrude. SeeDOI: 10.1039/c5gc00187k

aChemical and Petroleum Engineering, University of Kansas, 1530 W 15th Street,

Room 4162 Learned Hall, Lawrence, KS 66045, USA. E-mail: smwilliams@ku.edubCivil, Environmental and Architectural Engineering, University of Kansas, 1530 W

15th Street, Room 4162 Learned Hall, Lawrence, KS 66045, USA

This journal is © The Royal Society of Chemistry 2015 Green Chem.

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

.

View Article OnlineView Journal

wastewater affect the algal composition, resulting in a lowerlipid content and a higher inorganic fraction compared toalgae grown in water supplemented with fertilizers.5,10,12,13

These characteristics are traditionally presumed to be undesir-able properties as a biofuel feedstock because of lower oil andhigher solids (biochar) yield.

Biochar can be burned for energy production,5 used toremove water contaminants,14 provide a source of carbonsequestration and soil amendments,15 and has been evaluatedfor potential catalytic effects.16 The composition of a specificbiochar depends upon its starting material and conditions ofhow it is produced. Recent studies evaluating HTL of RN-algaeindicate that the inorganics are largely retained in the solidproduct. The inorganics were hypothesized to aid in theremoval of oxygen5 and nitrogen13 heteroatoms in the crudeoil. The findings reported herein provide a mechanistic under-standing of these observations.

In this study, we demonstrate that HTL of RN-algae crystal-lizes a pure-phase substituted hydroxyapatite (HAp) which pro-motes in situ catalytic upgrading producing a superior qualitybiocrude. HAp is a calcium orthophosphate (Ca10(PO4)6(OH)2)primarily used as a bio-medical material for bone and teethregeneration, however, HAp has previously been shown to bean effective green catalyst for acid–base chemistries for produ-cing butanol17,18 and acrylic acid,19,20 where crystallographicsubstitutions, morphology, and size contribute to desired cata-lytic activity.21

This study specifically details the crystallization process ofHAp during the HTL of RN-algae. We characterize the struc-ture, ionic substitutions, and morphology of the HAp productand describe the catalytic implications it has on the biocrudefraction. In addition to the actual discovery of in situ HAp crys-tallization during biocrude formation, this work is the first touse a single biogenic source for all elements required in HApor substituted HAp crystallization. This work is the first toreport on biomaterials from water reclamation systems anddemonstrates high-value co-products, including catalysts andbiomaterials, from algal biofuels.

Overall, this discovery will advance the economic viabilityand environmental sustainability of large scale algal biofuelplants. Wastewater treatment plants or water reclamation andresource recovery facilities, as defined by the Water EnvironmentFederation, can recovery nutrients and generate revenue by pro-ducing substituted nanocrystals of HAp and quality algae bio-crude. The HAp product is suitable for various industries withincreasing market values: fertilizers, heterogeneous catalysis,and bio-medical materials. This allows the HAp product to haveoptimal market flexibility with the value proportional to qualitycontrol. The algal fuel & chemical industry can now create high-value products alongside a higher quality biocrude in situ.

ExperimentalAlgae cultivation and biomass characterization

Algae cultivation using reclaimed water and nutrients were per-formed during the summer months of 2013 at the Lawrence,

KS Wastewater Treatment Plant using effluent from the sec-ondary clarifier, before disinfection, as the starting growthmedia. The same open pond system was used as previouslydescribed5 with one alteration, the four bioreactors were set-up as duplicate sets of two ponds in series, in which thereclaimed effluent was sent to the first growth pond, whoseeffluent was sent to a second growth pond. Each pond had ahydraulic retention time (HRT) of 10 days resulting in a totalof 20 day HRT for each set in the 2 × 2 system. Biomass grownin the second pond of one set was used in this work. Theaverage growth conditions for the second pond in series onthe day of harvest were measured with a Ca2+ concentration of76.2 mg L−1, pH of 8.8, alkalinity of 121.3 mg L−1 as CaCO3,total nitrogen of 21.0 mg N L−1, and total phosphorus of3.0 mg P L−1.

Algae slurry was collected,5 freeze dried, and characterizedto acquire proximate and ultimate analysis, and inorganicelemental distribution. Thermogravimetric analysis (TGA),TA Instruments SDT 600, was used for obtaining the proximateanalysis of the biomass. A method was developed to simul-taneously obtain moisture, volatile, fixed-carbon and ashcontents of biomass. The moisture and volatile amountswere collected by undergoing pyrolysis with nitrogen flow.During pyrolysis, the temperature was ramped to 90 °Cwhere it enters a stepwise function until 110 °C. During thestepwise function, the temperature was either isothermal orincreasing at 1 °C min−1 depending upon the derivativeweight change. When the derivative weight change was>0.01 wt% min−1 the sample was held isothermally and when<0.01 wt% min−1 temperature increased. This allowed accuratedetermination of a precise weight loss at a particular tempera-ture. After the first stepwise function the temperature wasramped at a constant rate, 5–10 °C min−1 to 800 °C andentered another stepwise function until 850 °C. Moisture andvolatile matter was then determined at the point where theirweight change was essentially zero at ∼100 °C and 850 °C,respectively. Within the same program, the sample was cooledto 120 °C and flow was switched to air for a post pyrolysiscombustion phase. Under oxygen, the post pyrolysis combus-tion proceeded to 800 °C followed by another stepwise func-tion to 850 °C, determining the fixed-carbon and ash contentof the biomass. The ultimate analysis was performed usinga Perkin Elmer CHN Analyzer. Fig. 1 details the abovementioned characterization and shows TGA plots of thepyrolysis and post pyrolysis combustion stages including theircorresponding derivative weight changes. The inorganicdistribution of the biomass was determined by inductivelycoupled plasma with an optical emission detector (ICP-OES;Agilent) after acid microwave-assisted digestion. Algal ash wasobtained by placing a sample of freeze dried algae, ∼100 mg,in a homemade quartz tube furnace with a programmabletemperature controller. The sample was combusted under aflow of 100 mL min−1 of air during 2 °C min−1 ramp to 600 °Cand held for 8 h. The combustion procedure was performedthree times to ensure all organic material was removed.The RN-algae ash was then analyzed by Fourier transform

Paper Green Chemistry

Green Chem. This journal is © The Royal Society of Chemistry 2015

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

inferred spectroscopy (FTIR) and powder X-ray diffraction(XRD).

FTIR was performed on a Varian 600 series FTIR equippedwith a GladiATR™ (PIKE Technologies) at a constant tempera-ture of 105 °C. Absorbance spectra were averaged over 254scans, collected from 4000 to 500 cm−1 with resolution andsensitivity of 4 cm−1 and 8 cm−1, respectively. The XRD spec-trum(s) were collected using monochromated CuKα radiation(λ = 1.54178 Å) on a Bruker Proteum Diffraction Systemequipped with Helios multilayer optics, an APEX II CCD detec-tor and a Bruker MicroStar microfocus rotating anode X-raysource operating at 45 kV and 60 mA. Sample preparationincluded mixing powder with Paratone N oil and placing it ina <0.5 mm nylon loop mounted on a goniometer head. ABruker Apex2 V2010.3-0 software package was used to collectand merge three, 60 s. 180° φ-scans with the detector at 2θ =30°, 60° and 90° using a sample-to-detector distance of50.0 mm. The raw data was processed with Bruker EVA powderdiffraction software.

Reaction, recovery, yield determination, and analytics

Hydrothermal processing was conducted in a 450 mL 4560series mini bench top reactor (Parr; Moline, IL) equipped witha 4848 model controller (Parr). The reactor was charged with50 mL Milli-Q water and 5.0 g freeze–dried algae. Oxygen waspurged from the system with nitrogen. The reaction was heldat a set-point of 350 °C for 1 h while stirring at 150 rpm. Post-reaction, the reactor was quenched in a water bath and allowedto equilibrate to room temperature. Product isolation followedthe sequence presented in Fig. 2. Once the reactor contentshad equilibrated the gasses formed were vented and the con-tents were vacuum filtered separating the aqueous co-product

(ACP) and solids/oil mixture. The solids/oil mixture was placedback into the reactor vessel with the addition of 50 mL dichloro-methane (DCM; Fisher) and heated to 40 °C with stir rate of600 rpm for 30 min. Upon cooling the vessel again in a waterbath, the contents were gravity filtered and washed withadditional DCM until no additional oil was extracted from thesolids. The solid product was dried and the oil–solvent mixturewas distilled to remove solvent and obtain biocrude product.

Commencing product analysis directly after separations iscritical for obtaining reliable yields, particularly for the bio-crude fraction. After the majority of the DCM solvent was dis-tilled off, a vacuum was applied to lower solvent content to<10 wt%. The weight of the biocrude was recorded in thetarred flask and immediately placed an ice bath. Simulateddistillation (SimDist) and ultimate analysis was immediatelyperformed on the biocrude using the same TGA instrumentand CHN analyzer as used for the proximate and ultimate ana-lysis of the algae biomass, respectively. The SimDist was per-formed under nitrogen flow with a steady 10 °C min−1 ramp to800 °C; with step-wise function employed from 35–45 °C toprecisely quantify the amount of DCM solvent remaining inthe sample, i.e., the sample stays ∼40 °C until the weightchange reaches a threshold of <0.01 wt% min−1. Yield and ulti-mate analysis (using known molecular weight, carbon, andhydrogen content of DCM) are adjusted by the correspondingamount of residual solvent for each experiment individually.

The dried solid product was weighed and analyzed for proxi-mate (TGA; ESI Fig. 1†) and ultimate (CHN) contents, func-tional groups (FTIR), and crystallography (XRD), along withbulk and trace metal compositions (ICP-OES). Yield of thesolids was determined on a dry weight (dw) and an ash freedry weight (afdw) basis. The aqueous co-product underwentanalysis of total organic carbon (TOC), total nitrogen (TN), andtotal phosphorous (TP). The TOC and TN were simultaneouslymeasured by Torch TOC/TN analyzer.5 The measured volume

Fig. 1 Proximate (TGA) and ultimate (CHN Analyzer) analysis of algaecultivated with reclaimed nutrients. TGA plot indicates pyrolysis (blue)and post pyrolysis combustion (green) weight changes with corres-ponding derivative weight changes, (purple) and (red), respectively. Ulti-mate analysis is provided on a ash free dry weight percent (afdw%).*Oxygen calculated by difference after normalized to the ash content:O (afdw%) = C − H − N [afdw%].

Fig. 2 Schematic of product handling from HTL of RN-algae.

Green Chemistry Paper

This journal is © The Royal Society of Chemistry 2015 Green Chem.

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

of ACP and TOC were used for determining the afdw% yield.Any gasses that were formed during HTL were not analyzedand the yield was determined by a difference and/or using theideal gas law assuming all CO2, headspace volume andrecorded post-reaction temperature and pressure.5 Post analy-sis of the elemental balance for carbon, hydrogen, nitrogen,and phosphorus for the feedstock and products were per-formed. The equations used for determining afdw% productyields are as followed:

Biocrude yield ¼ wtbiocrude 1�%DCMð Þalgaeafdw

� 100 ð1Þ

Solids yield ¼ wtsolids 1�%ashsolidsð Þalgaeafdw

� 100 ð2Þ

ACP yield ¼ volACP � TOCalgaeafdw �%carbonafdw

� 100 ð3Þ

Gas yield ¼ 100� eqn ð1Þ � eqn ð2Þ � eqn ð3Þ ð4Þwhere, algaeafdw is the weight of algae used in the reactorminus the amount of moisture and ash determined by TGA,wtbiocrude is the measured weight after distillation, %DCM isthe fraction of solvent remaining found with SimDist, wtsolidsis the measured weight of the solids fraction after gravity fil-tration, %ashsolids is the ash content of the solid productmeasured from TGA, volACP is the measured volume of the ACPafter vacuum filtration, TOC is the measured concentration oforganic carbon in the ACP, and %carbonafdw is the amount ofcarbon in the algae normalized to the organic fraction ofbiomass. Fig. 3 shows the bulk yields and characterizationfrom the HTL process.

Gas chromatography-mass spectrometry (GC-MS) analysisof biocrude was performed using an Agilent 6890 Series GC–FID system with a 5973 network mass selective detector

equipped with a Varian CP8956 column (5% phenyl methylsiloxane mobile phase; 30.0 m length; 320 μm diameter; and0.50 μm thickness). The method used an injection volume of1.0 μL with a split ratio of 1 : 1, an initial temperature of 40 °Cwith a holding time of 15 min, ramp of 10 °C min−1 to 100 °C,holding for 1 min, followed by a 1 °C min−1 ramp to 200 °C,with 15 min holds at 150 °C and 200 °C, followed by a 5 °Cmin−1 ramp to 270 °C, and final hold for 5 min. Samples wereprepared for the GC-MS analysis in two ways: (1) 10 mg of bio-crude in 1 mL of DCM and (2) 10 mg of biocrude derivatizedwith MSTFA (n-methyl-n-(trimethylsilyl)trifluoroacetamide;Agilent). The derivatized sample was prepared using 10 mg ofbiocrude, weighed in a GC vial, with 100 μL of MSTFA and100 μL of pyridine [high-performance liquid chromatography(HPLC) grade, 99.5+%, Alfa Aesar] added, and diluted with800 μL of n-heptane (99+%, Sigma).

Cell culturing and live/dead assay

Tissue-culture treated 24-well plates were coated, via aqueoussuspension, with HAp product calcined at 600 °C. The plateswere dried at 42 °C overnight and subsequently sterilized withethylene oxide. Human umbilical cord Wharton’s jelly cells(hWJCs) were trypsinized with 0.05% trypsin and seeded onplates at 6 × 104 ± 2 × 104 hWJCs per well. hWJCs were culturedwith Dulbecco’s Minimal Essential Medium (DMEM) suspen-sions with 10% mesenchymal stem cell qualified fetal bovineserum (FBS) and 1% penicillin/streptomycin at 37 °C with 5%CO2. hWJCs were isolated from Wharton’s jelly of a humanumbilical cord (#157) obtained from Lawrence Memorial Hos-pital (LMH) (LMH IRB approval #LMH 08-2). The umbilicalcord came from a male born at full term under normal con-ditions. Growth medium was changed every 2 days. After 10days, a viability assay was performed using LIVE/DEAD® Viabi-lity/Cytotoxicity Kit (Molecular Probes® LIVE/DEAD assay;from Invitrogen) and analyzed through fluorescencemicroscopy (Zeiss AX10). Images were processed using ZENlite 2012 (blue edition) software.

Results and discussionIn situ crystallization of substituted hydroxyapatite duringHTL of RN-algae

HAp preparation typically proceeds through a well-controlledsynthesis combining reagents from individual phosphorusand calcium sources. Synthesis procedures follow co-precipi-tation, hydrothermal, or various wet and dry methods resultingin a variety of morphologies and ionic substitutions.18,21–23

Certain marine algae (Corallina officinalis and Amphiroa ephe-drathe) have be used as a calcium source for HAp crystalliza-tion, but only after prior removal of organics and using anexternal phosphorus source.24,25 We have used RN-algae tocrystallize HAp in situ, and act as both the calcium and phos-phorus source without pre-treatment.

The RN-algae cultivated in open ponds at the LawrenceWastewater Treatment Plant, Lawrence, KS resulting in mixed

Fig. 3 Product Yields of HTL of RN-algae on ash free dry weightpercent (afdw%). The organic recovery represents the fate of algalcarbon and nitrogen toward each perspective product fraction. *Oxygencalculated by difference; O = 100 − C − H − N − ash [wt%].

Paper Green Chemistry

Green Chem. This journal is © The Royal Society of Chemistry 2015

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

algal cultures as previously identified.5 Calcium hydroxide, orlime, is added by the treatment facility in significant quan-tities daily (2000 lbs per day) as a buffering agent to raise alka-linity and pH as activated sludge, or bacteria, performsnitrification in the water column. Elemental composition ofthe growth pond, RN-algae, and HTL solids all measured byinductively coupled plasma, is shown in Table 1.

In the growth ponds calcium, phosphorous, magnesium,sulfur, potassium, and sodium had the highest observableconcentrations; silicon, although not measured by ICP due toanalytical limitations, is expected from the presence ofdiatoms in the mixed algal cultures. ICP analysis of RN-algaeand the resulting HTL solid product indicates certain elementsare preferentially retained within the solid product such as Ca,P, Mg, Fe, and Al, whereas others (S, K, & Na) are not (Table 1).Literature suggests algae that have higher sodium and potass-

ium and lower calcium concentrations produce less solidsduring HTL and retain more inorganics in the aqueousproduct.8,9,26

The general HAp structure, Ca10(PO4)6(OH)2, allows bothcationic and anionic substitutions, including magnesium,silicon, and carbonate substitutions, which have been shownto improve HAp performance as biomaterials.27–30 We deter-mined the structure of the HTL-produced solids using XRDand FTIR, shown in Fig. 4a,b. XRD reveals a monophasic HApaccording to JCPDS #73-0293, and is confirmed by FTIR. TheFTIR spectrum also indicates the following: residual organicsremain from HTL processing, B-type (PO4) carbonate substi-tutions occur, and significant hydroxyl deficiencies and phos-phorus bound hydroxyls are present. To confirm that the HApwas not formed via precipitation during the cultivationprocess, but rather from the HTL reaction, we compared theXRD and FTIR patterns of the solid product and the RN-algaeash. The ash was solely tricalcium phosphate (TCP) andcalcium oxide. A previous study31 demonstrated combustion ofcalcium and phosphorus sources results in an amorphousmaterial which, upon calcination at 600 °C, crystallizes to formTCP. However, the combustion process at 600 °C is notsufficient for significant transformation of HAp to TCP.32,33

Thus, HAp formation does not occur during algae cultivationand is formed during the HTL of RN-algae.

RN-algae, as the biogenic source for HAp production,provide elements that favor ionic substitutions during calciumorthophosphates crystallization. The main substitutions in theHAp product are shown by scanning electron microscopy-

Table 1 Elements measured by ICP-OES (ND refers to not detectable)

Growth pond(ppm)

RN-algae(wt%)

% Retentionin HTL solids

Ca 78.49 5.70% 89%P 3.07 2.30% 95%Mg 12.83 0.42% 100%S 29.20 0.82% 19%Fe 0.01 0.13% 84%K 16.18 0.36% 24%Al NDa 0.06% 79%Na 23.15 0.06% 44%

aDetected in the feed water to growth ponds.

Fig. 4 Solid product from the HTL of RN-algae is pure phase HAp. (a), Integration of powder XRD of HTL solid product (top) and combustion-pro-duced algal ash (bottom) at a 2θ range of 20° to 70°; peaks for HAp (green square), calcium oxide (red triangle) and TCP (blue circle), JCPDS #70-2005, are identified. (b) FTIR spectrum of HTL solid product and RN-algae ash; all peaks present in the range from 500–3700 cm−1 are shown. RN-algae ash exhibit vibration bands for PO4

3− (947 and 983 cm−1; as TCP), P2O74− (1218 cm−1), and surface hydroxyl (3639 cm−1; from calcium oxide).

The solid HTL product exhibit distinct absorption bands for HAp; including PO43− vibrations, v1 (964 cm−1), v3 (1087 and 1017 cm−1), v4 (603 and

561 cm−1), hydroxide stretching (3564 cm−1 and 3677 cm−1; lattice -OH and P–OH, respectively), CO32− (1417 and 1452 cm−1; carbonate B-Type sub-

stitution), and either HPO42− or CO3

2− (871 cm−1). Residual organics from HTL processing show bands for C–O (1563 cm−1) and C–H (2852 and2922 cm−1) groups. (c) SEM-EDS imaging of a single particle of solid HTL product, scale bar equals 10 µm, with elemental mapping of calcium, phos-phorus, oxygen, magnesium, and silicon; corresponding EDS spectrum is found in ESI Fig. 2.†

Green Chemistry Paper

This journal is © The Royal Society of Chemistry 2015 Green Chem.

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

energy dispersive X-ray spectroscopy (SEM-EDS), Fig. 4c, whichinclude Mg and Si for Ca and P, respectively. Additionalelements preferentially retained (Table 1) in the HAp productwould be present as minor substitutions and show relativelysmall peaks on the EDS spectrum (ESI Fig. 2†). Mg substi-tution is indicated by the hydroxyl shift in FTIR spectrum,from the typical 3572 cm−1 to the observed 3564 cm−1. Pre-viously, this was seen when x = 2 in Mg substituted HAp of theform Ca10−xMgx(PO4)6(OH)2.

34 Mg results in lower IR absor-bance of lattice –OH,27,28 consistent with Fig. 4b. The presenceof silicate can be confirmed by the FTIR bands at 656 and782 cm−1 (Si in HAp) in the HAp product and at 800 cm−1

(amorphous silica) in the RN-algae ash.28

Ionic substitutions in HAp can promote thermal transform-ation to TCP at temperatures above 600 °C.32,33,35 Multiphasiccalcium orthophosphates (mixtures of HAp and TCP) as bio-materials are known to have high osteogenicity and osteo-inductive properties,35 particularly in achieving an optimumbalance between the more stable HAp and more soluble TCP.36

Thermal treatment of the HAp product steadily transforms thematerial to TCP with intermediate temperatures giving variablebiphasic mixtures. FTIR and XRD analysis of thermally treatedHAp product in 100 °C increments from 500 °C to 900 °C aredepicted in Fig. 5. Residual organics from HTL processing areremoved at 500 °C, as observed by the disappearance of the C–O peak at 1563 cm−1 and the C–H bands at 2852 & 2922 cm−1

(not shown) in the FTIR spectra. However, the XRD spectrumindicates that thermal treatment at 500 °C does not alter theHAp structure. The FTIR at 500 °C retains significant carbon-ate peaks at 1417 & 1452 cm−1, indicative of B-type, or PO4,substitution. These ionic substitutions can distort the latticeand compensate charge by creating hydroxyl vacancies result-ing in a reduction of FTIR stretching intensity,27 as seen withHAp product here.

A clear decrease in the FTIR carbonate bands (1417 &1452 cm−1) is observed with increasing thermal treatment. TheXRD spectra defines the onset of transition from HAp to TCPat 600 °C and corresponds to the loss of carbonate bands, indi-cating that carbonate decomposition plays a role within phasetransformation. The TGA for the ultimate analysis of the HApproduct (ESI Fig. 1†) indicates there are four main areas ofdecomposition between 600–850 °C. Three regions in the XRDpattern can be used to monitor phase transformation; indi-cated in Fig. 5 by dashed red lines. First, the peak at 2θ = 30.5°is unique to TCP, initially observed at 600 °C, and grows withincreasing temperature. Second, the primary HAp peak (2θ =32.0°) shifts to 31.5°, characteristic of TCP. Third, the peaks at2θ = 47° & 50° for HAp shift closer together to 47.5° & 48.5°,indicative of the TCP structure. The transition from HAp toTCP is nearly complete after thermal treatment at 900 °C,when comparing the XRD patterns with databases.

Significant broadening of the phosphate shoulder peaks at800–1000 cm−1 in the FTIR patterns occur with increasingthermal treatment. Broad shoulders with multiple peaks areunlike typical FTIR patterns for TCP, which have a sharppeak near 983 cm−1 and small shoulder at 947 cm−1, similar to

RN-algae ash spectra shown in Fig. 4b. These shoulders areattributed to SiO4

4− substitution, which have been seen inpure Si-HAp.28 However, the XRD profiles suggest that theSiO4

4− is actually within the TCP phase, suggesting the Sipresent in HAp is retained during the transformation to TCP.If the Si was not integrated in the TCP structure, FTIR peaksfor silicate or poly-silicate would be observed, similar to theRN-algae ash, with increasing thermal treatment. However,no characteristic peak for amorphous silica is observed at800 cm−1.

Fig. 5 HAp product transforms to tricalcium phosphate in stages up to900 °C. Powder XRD integration of 2θ from 20° to 70°; TCP begins toform at 600 °C until it is the primary phase at 900 °C. Dashed red linesindicate characteristic peaks for TCP. FTIR spectrum(s) showing hydroxyl(3300–3800 cm−1), carbonate (1300–1600 cm−1) and phosphate(500–1300 cm−1) regions. TCP retains silica substitution dissimilar to theTCP from combustion of RN-algae.

Paper Green Chemistry

Green Chem. This journal is © The Royal Society of Chemistry 2015

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

HAp crystallization promotes in situ upgrading of biocrude

HAp is increasingly studied for its acid–base catalytic capabili-ties. Typical catalytic applications of HAp include dehydration(acid catalyzed) and dehydrogenation (base catalyzed),19,37

Guerbet coupling reactions,38 and use as active metal supportmaterials.39 While biocrude oil can be generated from a varietyof feedstocks, it typically needs to be upgraded in order to beeasily integrated into current infrastructure for refining andtransportation. Therefore, creating a biocrude with anincreased amount of lower boiling point distillate fractionsand fewer heteroatoms is of significant interest. The HAp co-produced by HTL of RN-algae provides in situ upgrading inboth accounts. Complete analysis of molecular profile and dis-tillate fractions are found in Fig. 6 and 7, respectively. The bio-crude product resulted in 80 wt% direct fuels and vacuum gasoil, and overall 96 wt% could be distilled below 600 °C. GC-MSanalysis on the biocrude indicate the direct fuels are highlyaromatic with primary components including phenol, cresolisomers, dimethyl and ethyl phenols, and indoles, along withsignificant amounts of straight chain and branched aliphatics.The vacuum gas oil primarily consists of free fatty acids and>C20 straight chain aliphatics (complete list of all identifiedcompounds are in ESI Tables 1 and 2†). Significant amountfree fatty acids and methyl amine are shown in the MSTFA-derivatized GC-MS chromatogram (ESI Fig. 3†). Only one fattyacid amide was present, in small amounts, and no nitriles.Studies have shown both thermal40 and catalytic41 upgradingtreatments to reduce fatty acid amides and nitriles while pro-ducing lower boiling point aromatics and aliphatics. Roussiset al.40 explains a thermal upgrading pathway from amides tonitriles to aliphatics and aromatics at thermal treatment above400 °C. The small amounts of fatty acid amides, with nonitriles, and the presence of methyl amine suggests that HApproduct is catalyzing the dehydration pathway from amides toaliphatics and aromatics. Further evidence of HAp catalysiscan be found by comparing results to Bai et al. catalytically

treated biocrude.41 Their findings also included reduction offatty acid amides and nitriles, and upon upgrading they beganto form benzenamine and carbozole. Both of these com-pounds are found within the RN-algae biocrude (Fig. 6, peak23: benzenamine and ESI Fig. 3,† peak E′: carbozole).Additionally, Bai et al. determined catalytic treatment wasrequired to achieve total distillation of >90 wt% at or below

Fig. 6 GC-MS of RN-algae biocrude produced in the presence of HAp crystallization.

Fig. 7 Simulated distillation via TGA of the biocrude and correspondingdistillate fractions found in the biocrude produced in the presence ofHAp crystallization.

Green Chemistry Paper

This journal is © The Royal Society of Chemistry 2015 Green Chem.

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

600 °C, similar to the biocrude produced during HApcrystallization.

The distillate fractions from the RN-algae biocrude co-pro-duced with HAp, presented in Fig. 7, more closely resembleRoussis et al. thermally-upgraded biocrude at 400 °C,especially in the lower boing point distillates. The amount ofvac. gas oil and residual are more similar to Roussis et al.350 °C upgraded oil, however, this is misleading becausemuch higher boiling point molecules are cracking whichproduce higher amounts of >C20 aliphatics.

HAp is primarily a non-porous crystal which exhibits acid–base activity on external crystal surfaces. Amount and strengthof the active sites are a function of Ca/P ratio, substitutions,and crystal morphology; each altering which atoms areexposed on the crystal surface. Silvester et al.21 contribute P–OH as Brønsted acid sites and the Ca2+ and OH vacancies asLewis acids. Yan et al.19 describe when organics adsorb on theHAp surface, a P–OH complex is formed and catalytically dehy-drates lactic acid to acrylic acid. Once the molecule desorbsfrom the surface the P–O− is regenerated. The closer view ofthe FTIR hydroxyl stretching area of the HAp product in Fig. 5show two things; (1) the lattice hydroxyl peak is extremelysmall from considerable hydroxyl vacancies (or Lewis acids),and (2) the there is a pronounced P–OH complex peak.Removal of organics at 500 °C results in no changes to latticehydroxyls, however, the P–OH complex is lost suggesting theoriginal P–O− is regenerated. These results demonstrate thatHTL of RN-algae results in the in situ synthesis of a HAp cata-lyst which, contain Brønsted and Lewis acid sites and catalyti-cally upgrade the biocrude produced. Further, regenerationcan be achieved through minimal heat treatment and create apure phase HAp product stream.

HAp morphology and hierarchical structure

Catalytically active sites on HAp surfaces are also dictated bytheir crystallite morphology. The exposed crystal face deter-mines which type of active site is more dominant, acid orbase; typically longer crystals are more acidic and globularmorphologies are more basic.18,21 Similarly to HAp catalysts,morphology of HAp biomaterials is also important. Naturalbone consists of poorly crystalline ion-substituted HAp wherethe micro-42,43 and hierarchical structure44 play pivotal roles inboth natural- and engineering-assisted growth and regener-ation. Therefore, commercial production of HAp gives signifi-cant effort to generating calcium orthophosphates with bothcontrolled nanoparticle rod and hierarchical morphologies.23

Previous studies have demonstrated that HAp nanorods canaggregate into bundles and, through Oswald ripening, formsheets and hierarchical flower structures.45,46 To facilitatenanoparticle rod and hierarchical morphologies, surfactantshave been incorporated during commercial HAp crystalliza-tion, including an array of molecules such as fatty acids,47

amino acids,48,49 and a variety of organic polymers.23,46 Thehydrothermal synthesis of HAp has also been demonstratedutilizing a biogenic phosphate source (adenosine 5′-triphos-phate or ATP) as both morphological control and a phos-

phorus source using external calcium source.50 RN-algaecontain phosphorus in many biomolecules within the cell,including DNA, RNA, ATP, and phospholipids, thus providingbiomolecules that can contribute to hierarchical structureswithout the addition of external calcium sources.

Morphology and hierarchical structures are shown throughtransmission electron (TEM) and scanning electron micro-graphs (SEM) of the HAp product directly after HTL formation(Fig. 8a–c) and after thermal treatment at 900 °C (Fig. 8d).Hexagonal nanocrystals (Fig. 8a,b) are present up to 1 micronin length along the c-axis and 15 nm about the a and b-axis;these crystals then form into bundles along the c-axis, Fig. 8b.Each bundle can then form hierarchical sheets and flower-likestructures, Fig. 4c. Unlike literature studies, which havesuggested homogenous formation of the nanoparticles andhierarchical morphologies in the presence of a single surfac-tant, the in situ crystallization of HAp during HTL of RN-algaeresults in heterogeneous morphologies. The heterogeneity ofthe material could be due to the short reaction times com-pared to conventional HAp synthesis techniques, or the pres-ence of variable growth mechanisms due to localizedbiomolecules (i.e., fatty acids, amino acids, phospholipids)being present at different concentrations during the HTL pro-cessing. Transformation to the TCP phase induces globularmorphological changes, Fig. 8d. The combination of bothphase and morphological changes that occur through simpleheat treatments of the HAp product allows tunability withineach market, catalysis and/or biomedical.

Preliminary cell culturing and attachment

The substituted HAp structure formed with HTL of RN-algaeshows great promise for biomedical and bioengineering appli-cations. Although natural bone is mainly Ca2+ and PO4

3−, italso incorporates other ions of CO3

2−, SiO44−, and Mg2+. These

substitutions, and optimizing HAp/TCP ratio through calcina-

Fig. 8 HAp product consists of hexagonal nanorods, which assembleinto hierarchal structures from bundles to sheets to flowers. TEM imagesof (a) well-defined hexagonal nanocrystals which (b) aggregate tobundles along the c-axis. SEM images of (c) ∼40 µm particle with sheetand flower-like hierarchical morphologies and (d) TCP globular mor-phologies after calcination at 900 °C.

Paper Green Chemistry

Green Chem. This journal is © The Royal Society of Chemistry 2015

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

tion, can help HAp behave more like natural apatite in humanbone and increase bioactivity to allow steady bone regener-ation over an extended period of time.33,51 Our first stage ofevaluating efficacy for bone scaffolds or regenerative materialsis to determine whether human Wharton’s jelly cells (hWJCs)will attach to the material. We observed within 1 day ofseeding hWJCs on the HAp product calcined at 600 °C, cellsbegan to attach and showed beginning stages of morphologi-cal changes. After 10 day incubation, we performed a live/deadassay; the images in Fig. 9 show living cells which are clearlyattached to the HAp product through extended filopodia.These promising results strongly warrant further investigationto understand if the HAp product will promote a geneticresponse toward an osteogenic lineage. If so, this processcould provide a cheap and effective alternative to producingbone-like apatite.

Conclusions

In summary, algae cultivated during nutrient reclamationcan be converted to a high-value product, hydroxyapatite,while co-producing a crude oil displacement for biofuel pro-duction. This discovery will enable commercial viability toachieve a cost-competitive biofuel and a cheap alternativeroute for crystallizing hydroxyapatite for both heterogeneouscatalysis and tissue engineering applications. Our datadescribes a novel synthesis with a distinct synergy betweenin situ crystallization and oil upgrading, benefiting both pro-ducts. The oil produced has desired properties, including anincreased amount of lower boiling point distillate fractionsand lower heteroatom content. The hydroxyapatite containsbone-like substitutions, with the ability to tailor the degree ofbiphasic orthophosphate(s) for particular applications ineither catalysis or bone regeneration. Crystallization of hydroxy-apatite directly from algae using hydrothermal liquefactionrepresents a new value stream for traditional biofuel pro-

duction. We found a reasonably small municipal water recla-mation plant (serving population of ∼80 000) could produceup to 18 barrels of biocrude and 2000 kg of refined pure phasesubstituted hydroxyapatite per day, while simultaneouslyreclaiming nitrogen and phosphorous. Our data suggests thata commercial scale hydrothermal liquefaction plant can nowhave the ability to synthesize a catalyst within the process,utilize its catalytic ability in situ and upon regeneration,produce a high-value product stream. The HAp product pro-duced has the ability to be easily tailored in both its phase andmorphologies to accompany any desired application for acid–base chemistries in heterogeneous catalysis or bio-activity forregenerative medicines.

This novel approach for creating high-value productstreams from water reclamation sources can have significantimpacts to tax payer dollars, creating profit for a traditionallyregulatory industry while contributing to biofuels, green cataly-sis, and medicine. These findings provide the basis for newdevelopments in algae-based fuels, chemicals, and biopro-ducts at both the fundamental and macro commercializationlevels, where the full impacts into heterogeneous catalysis andtissue engineering applications are widely unknown.

Acknowledgements

We thank the staff with the City of Lawrence WWTP, particu-larly David Corliss, Dave Wagner, Mark Hegeman, and CarolynWoodhead, who hosted our pilot work at the wastewater treat-ment plant. Jay Barnard, Jerry deNoyelles and Scott Campbellprovided critical field assistance setting up the pilot reactors.This work was supported by the University of Kansas Transpor-tation Research Institute Grant no. DT0S59-06-G-00047,funded by the U.S. Department of Transportation Researchand Innovative Technology Administration; by the NationalScience Foundation EPSCOR Grant no. EPS-0903806, andmatching support from the State of Kansas through KansasTechnology Enterprise Corporation; by a Department of EnergyGrant no. DE-EE0000408; by the NASA EPSCOR Grant no.NNX07AO27A; by the National Science Foundation REU Grantno. NSF0065504 (funding for Shayan Sadzar) and SSTEM grantno. EPS-0903806 (funding for Umar Hamdeh); and the Univer-sity of Kansas Madison and Lila Self Graduate Fellowship(funding for Griffin W. Roberts).

Notes and references

1 E. Stephens, I. L. Ross, Z. King, J. H. Mussgnug, O. Kruse,C. Posten, M. A. Borowitzka and B. Hankamer, Nat. Biotech-nol., 2010, 28, 126–128.

2 L. Brennan and P. Owende, Renewable Sustainable EnergyRev., 2010, 14, 557–577.

3 National Research Council of the National Academies,National Academy Press, Report from the Committee onthe Sustainable Development of Algal Biofuels, 2012.

Fig. 9 Human Wharton’s jelly cells attached to HAp product. Live/deadassay micrograph after 10 day incubation on HAp product calcined at600 °C. Live cells fluoresce in green, dead cells fluoresce in red. TheHAp product interacted with the florescent components to induce autofluorescence, primarily in the red. Scale bars are 100 µm.

Green Chemistry Paper

This journal is © The Royal Society of Chemistry 2015 Green Chem.

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online

4 D. López Barreiro, W. Prins, F. Ronsse and W. Brilman,Biomass Bioenergy, 2013, 53, 113–127.

5 G. W. Roberts, M.-O. P. Fortier, B. S. M. Sturm andS. M. Stagg-Williams, Energy & Fuels, 2013, 27, 857–867.

6 M. Möller, P. Nilges, F. Harnisch and U. Schröder, Chem-SusChem, 2011, 4, 566–579.

7 G. Felício-Fernandes and M. Laranjeira, Quim. Nova, 2000,23, 441–446.

8 D. C. Elliott, T. R. Hart, A. J. Schmidt,G. G. Neuenschwander, L. J. Rotness, M. V. Olarte,A. H. Zacher, K. O. Albrecht, R. T. Hallen andJ. E. Holladay, Algal Res., 2013, 2(4), 445–454.

9 P. J. Valdez, M. C. Nelson, H. Y. Wang, X. N. Lin andP. E. Savage, Biomass Bioenergy, 2012, 46, 317–331.

10 B. S. M. Sturm and S. L. Lamer, Appl. Energy, 2011, 88,3499–3506.

11 I. Woertz, A. Feffer, T. Lundquist and Y. Nelson, J. Environ.Eng., 2009, 135, 1115–1122.

12 B. S. M. Sturm, E. Peltier, V. Smith and F. deNoyelles,Environ. Prog. Sustainable Energy, 2012, 31, 10–16.

13 W.-T. Chen, Y. Zhang, J. Zhang, G. Yu, L. C. Schideman,P. Zhang and M. Minarick, Bioresource Technol., 2013, 152,130–139.

14 D. Mohan, R. Sharma, V. K. Singh, P. Steele andC. U. Pittman, Ind. Eng. Chem. Res., 2012, 51, 900–914.

15 N. Borchard, A. Wolf, V. Laabs, R. Aeckersberg,H. W. Scherer, A. Moeller and W. Amelung, Soil UseManage., 2012, 28, 177–184.

16 N. Muradov, B. Fidalgo, A. C. Gujar, N. Garceau andA. T-Raissi, Biomass Bioenergy, 2012, 42, 123–131.

17 T. Tsuchida, T. Yoshioka, S. Sakuma, T. Takeguchi andW. Ueda, Ind. Eng. Chem. Res., 2008, 47, 1443–1452.

18 T. Tsuchida, J. Kubo, T. Yoshioka, S. Sakuma, T. Takeguchiand W. Ueda, J. Catal., 2008, 259, 183–189.

19 B. Yan, L.-Z. Tao, Y. Liang and B.-Q. Xu, ACS Catal., 2014,1931–1943.

20 V. C. Ghantani, S. T. Lomate, M. K. Dongare andS. B. Umbarkar, Green Chem., 2013, 15, 1211.

21 L. Silvester, J.-F. Lamonier, R.-N. Vannier, C. Lamonier,M. Capron, A.-S. Mamede, F. Pourpoint, A. Gervasini andF. Y. Dumeignil, J. Mater Chem. A, 2014, 2(29), 11073–11090.

22 Y. Matsuura, A. Onda and K. Yanagisawa, Catal. Commun.,2014, 48, 5–10.

23 M. Sadat-Shojai, M. T. Khorasani, E. Dinpanah-Khoshdargiand A. Jamshidi, Acta Biomater., 2013, 9, 7591–7621.

24 P. J. Walsh, G. M. Walker, C. A. Maggs and F. J. Buchanan,J. Mater. Sci. Mater. Med., 2010, 21, 2281–2286.

25 E. Spassova, S. Gintenreiter, E. Halwax, D. Moser,C. Schopper and R. Ewers, Materialwiss. Werkstofftech.,2007, 38, 1027–1034.

26 P. J. Valdez, J. G. Dickinson and P. E. Savage, Energy Fuels,2011, 25, 3235–3243.

27 T. Huang, Y. Xiao, S. Wang, Y. Huang, X. Liu, F. Wu andZ. Gu, J. Thermal Spray Technol., 2011, 20, 829–836.

28 D. Marchat, M. Zymelka, C. Coelho, L. Gremillard,L. Joly-Pottuz, F. Babonneau, C. Esnouf, J. Chevalierand D. Bernache-Assollant, Acta Biomater., 2013, 9,6992–7004.

29 N. Y. Mostafa, H. M. Hassan and O. H. Abd Elkader, J. Am.Ceram. Soc., 2011, 94, 1584–1590.

30 A. M. Pietak, J. W. Reid, M. J. Stott and M. Sayer, Biomater-ials, 2007, 28, 4023–4032.

31 S. Loher, W. J. Stark, M. Maciejewski, A. Baiker,S. E. Pratsinis, D. Reichardt, F. Maspero, F. Krumeich andD. Günther, Chem. Mater., 2004, 17, 36–42.

32 K. Zorn, U. Gbureck, D. Mitró, F. A. Müller andE. Vorndran, Bioinspired, Biomimetic Nanobiomater., 2013,2, 11–19.

33 M. H. Fathi, A. Hanifi and V. Mortazavi, J. Mater. Proc.Technol., 2008, 202, 536–542.

34 S. Diallo-Garcia, D. Laurencin, J.-M. Krafft, S. Casale,M. E. Smith, H. Lauron-Pernot and G. Costentin, J. Phys.Chem. C, 2011, 115, 24317–24327.

35 S. V. Dorozhkin, Acta Biomater., 2012, 8, 963–977.36 G. Daculsi, Biomaterials, 1998, 19, 1473–1478.37 A. Rahmanian and H. S. Ghaziaskar, J. Supercrit. Fluids,

2013, 78, 34–41.38 J. T. Kozlowski and R. J. Davis, ACS Catal., 2013, 3, 1588–

1600.39 A. Indra, C. S. Gopinath, S. Bhaduri and G. Kumar Lahiri,

Catal. Sci. Technol., 2013, 3, 1625.40 S. G. Roussis, R. Cranford and N. Sytkovetskiy, Energy Fuels,

2012, 26, 5294–5299.41 X. Bai, P. Duan, Y. Xu, A. Zhang and P. E. Savage, Fuel,

2014, 120, 141–149.42 H. Zhou and J. Lee, Acta Biomater., 2011, 7, 2769–

2781.43 S. J. Hollister, R. D. Maddox and J. M. Taboas, Biomaterials,

2002, 23, 4095–4103.44 P. Fratzl, H. S. Gupta, E. P. Paschalis and P. Roschger,

J. Mater. Chem., 2004, 14, 2115.45 M. G. Ma, Int. J. Nanomed., 2012, 7, 1781–1791.46 A. J. Nathanael, S. S. Han and T. H. Oh, J. Nanomater.,

2013, 2013, 1–8.47 X. Wang, J. Zhuang, Q. Peng and Y. D. Li, Adv. Mater., 2006,

18, 2031–2034.48 E. Boanini, M. Fini, M. Gazzano and A. Bigi, Eur. J. Inorg.

Chem., 2006, 2006, 4821–4826.49 H. Cui, H. Liu, J. Qin, Y. Li, H. Tang and X. Yang, Micro

Nano Lett., 2012, 7, 1292–1295.50 C. Qi, Q.-L. Tang, Y.-J. Zhu, X.-Y. Zhao and F. Chen, Mater.

Lett., 2012, 85, 71–73.51 A. F. Khan, M. Saleem, A. Afzal, A. Ali, A. Khan and

A. R. Khan, Mater. Sci. Eng. C, 2014, 35, 245–252.

Paper Green Chemistry

Green Chem. This journal is © The Royal Society of Chemistry 2015

Publ

ishe

d on

20

Febr

uary

201

5. D

ownl

oade

d by

Uni

vers

ity o

f K

ansa

s on

02/

03/2

015

18:0

6:24

. View Article Online