Effects of inoculum size, light intensity, and dose of

Algal Research 19 (2016) 278–290

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Effects of inoculum size, light intensity, and dose of anaerobic digestion centrate on growth and productivity of Chlorella and Scenedesmus microalgae and their poly-culture in primary and secondary wastewater Pavlo Bohutskyi a,⁎, Debora Cynamon Kligerman a,b, Natalie Byers a, Laila Khaled Nasr a, Celine Cua a, Steven Chow a, Chunyang Su a, Yuting Tang c,d, Michael J. Betenbaugh c, Edward J. Bouwer a a

Department of Geography & Environmental Engineering, Johns Hopkins University, 3400 North Charles Street, Baltimore, MD 21218-2686, USA Departamento de Saneamento e Saúde Ambiental, Escola Nacional de Saúde Pública, Fundação Oswaldo Cruz, Rua Leopoldo Bulhões, 1480, 21041-210 Rio de Janeiro, RJ, Brazil c Department of Chemical & Biomolecular Engineering, Johns Hopkins University, 3400 North Charles Street, Baltimore, MD 21218-2686, USA d Department of Chemical Engineering, Nanjing Forestry University, No. 159 Longpan Street, Nanjing, JS 210037, PR China b

a r t i c l e

i n f o

Article history: Received 13 February 2016 Received in revised form 23 June 2016 Accepted 7 September 2016 Keywords: Microalgal-bacteria poly-culture FAME composition Sustainable biofuel-phytoremediation process Nitrogen to phosphorus ratio Macronutrients and trace elements Chemical fertilizer replacement

a b s t r a c t Scale-up of microalgal biofuel technology is challenged by availability of nitrogen and phosphorus fertilizers and the potential negative impact vast increases in chemical fertilizer demand would have on conventional agriculture. The current study investigated replacement of chemical fertilizers with nutrients sourced from primary and secondary wastewater effluents and anaerobic digestion centrate (ADC). Although primary wastewater effluent possessed a high optical density (OD) and bacterial contamination, it was a superior growth medium for microalgal cultivation than nutrient-scarce secondary effluent. Chlorella sorokiniana and Scenedesmus acutus f. alternans showed higher growth rates, productivities, and robustness than other species or poly-cultures of five species. While supplementing with 5–10% nutrient-rich ADC increased wastewater OD, it also enhanced microalgal growth rates from 0.2–0.3 d−1 to 0.7–0.9 d−1 and biomass productivity from 10 to 20 mg L−1 d to 40–60 mg L−1 d with greater improvements for secondary effluents. Supplementation with ADC also increased nutrient concentrations (N, P, Mn, B, Zn, Co by N 100% and S, Mg, Ca, Mo by 20–60%) and improved the nitrogen to phosphorus (N:P) ratio. Higher ADC dose of 20% inhibited microalgae growth potentially due to ammonia toxicity. Elevation of inoculum doses and light intensity increased final biomass density and productivity, with intensities b140 μmol photon m−2 s−1 limiting algal growth rates. Inoculum doses of ≥2.5 × 105 cell mL−1 were most favorable for cultivation of all tested microalgae and for FAME content and composition for a newly characterized strain of Chlorella sorokiniana. Overall, ADC represents an economical fertilizer substitute providing various nutrients needed for microalgal growth and enhancing biofuel sustainability. © 2016 Elsevier B.V. All rights reserved.

1. Introduction Renewable and carbon-neutral alternatives to fossil energy sources are critical in order to mitigate climate change and to maintain fuel and food price stability. Algal-based technologies offer great promise as sources of biofuels, animal feeds and added value bioproducts [1– 3]. A number of technological challenges, including sufficient fresh water and nutrient availability must be overcome to make microalgal cultivation and microalgal-based biofuels environmentally sustainable and economically competitive [4–6]. Photoautotrophic cultivation of algae requires a light source, carbon, nutrients, and water. Several studies have estimated that up to 2600–

⁎ Corresponding author at: Biological Sciences Division, Earth & Biological Sciences Directorate, Pacific Northwest National Laboratory, Richland, WA, USA. E-mail addresses: [email protected], [email protected] (P. Bohutskyi).

http://dx.doi.org/10.1016/j.algal.2016.09.010 2211-9264/© 2016 Elsevier B.V. All rights reserved.

3400 gal of water are required per gallon of generated biofuel if no recycling is applied [7,8]. While the actual consumptive water loss can be reduced to 240–600 gal per gal of fuel through water reuse [8,9], complete recycling is impossible due to evaporative losses and accumulation of salts, cell remains, and other inhibiting compounds [10–13]. Given the critical role that water plays in microalgal cultivation and the impact its use has on process sustainability, potential alternatives to fresh water are urgently needed. The application of wastewater is particularly appealing from an economic and sustainability prospective [14,15]. Moreover, wastewater contains numerous components including nitrogen (N), phosphorus (P), and other essential macro- and micronutrients that can potentially replace industrially produced fertilizers. Indeed, the production of one gallon of microalgal lipids, precursors of biodiesel, requires 0.33–1.5 and 0.071–0.21 kg of N and P, respectively, depending on the algal lipid content and other parameters [8,9]. At the same time, uncontrolled discharge of these nutrients in improperly treated wastewater represents a major threat to coastal

P. Bohutskyi et al. / Algal Research 19 (2016) 278–290

regions and water resources; ironically often in the form of uncontrollable algal blooms [16]. Certain sewage treatment processes, including anaerobic digestion (AD) of sewage sludge and livestock manure, generate wastewater with particularly high levels of N, P, and other nutrients. The amount of sewage sludge isolated during municipal wastewater treatment in the U.S. is estimated at nearly 7.2 million dry tons per year (mdt yr−1) [17]. With average composition of 4.2% of N and 3.0% of phosphorus [18] this sewage sludge contains roughly 300,000 and 220,000 tons per year of N and P, respectively. Moreover, animal feeding operations in the U.S. generate an even higher amount of nutrient-rich manure approximated as 500 mdt yr−1 [19]. The liquid effluent from sewage sludge or manure anaerobic digestion could be used as one source of nutrients and a low-cost substitute for chemical fertilizers for production of valuable algal biomass [20,21]. Some reports have demonstrated the potential of cultivating microalgae, especially from Chlorella and Scenedesmus, genera in the anaerobic digestion centrate (ADC) generated during processing of domestic sewage sludge [20,22, 23], swine manure [24–27], cattle manure [28–31], and co-digested animal manure [32,33]. However, limited data are available on optimizing microalgal growth in wastewater mixed with ADC or on the effect of critical process parameters including wastewater pretreatment, ADC concentration, light intensity and microalgal inoculum dose on culture growth rate, biomass density and productivity. Since wastewater, especially primary effluent, contains high numbers of bacteria, the ecological interactions between microalgae and wastewater bacteria are important factors to consider, as they could be competitive or mutually beneficial [34–38]. When microalgae utilize solely inorganic carbon (CO2) they are not competing with heterotrophic bacteria for a carbon source [39]. Moreover, the presence of bacteria in an algae-bacteria system may improve carbon conversion efficiency [40]. However, bacteria may compete with algae for nutrients such as N and P, especially when they become limited [39]. In fact, there are few studies focused on growth of microalgae with the mixture of wastewater-originated microorganisms and confirming the interference for microalgal growth [41]. The aims of this study were to determine the influences of multiple process parameters including ADC supplement dose, light intensity, and initial concentration of inoculum on algal growth rate, productivity, and lipid composition in either primary or secondary wastewater. To assess this goal, the growth parameters of five different microalgae species including strains of Chlorella sorokiniana (CCTCC M209220, UTEX B 3010 and and UTEX 1230), Chlorella vulgaris (UTEX 2714), Scenedesmus acutus f. alternans (UTEX B 72), and Scenedesmus dimorphus (UTEX 1237) were evaluated at ADC dilution rates from 0 to 20%, light intensities from 30 to 200 μmol photon m−2 s− 1, and microalgal inoculum concentrations ranging from 104 to 106 cells mL−1. Since sterilization of wastewater and centrate are problematic to implement at large scale, this study was performed using unsterilized wastewater spiked with various amounts of the ADC. We found that certain species are more robust in bacteria-contaminated wastewater than others and observed that all these parameters are critically important for microalgal growth and optimizing biomass productivity. 2. Materials and methods 2.1. Algal strains and growth medium The microalgal cultures Chlorella vulgaris (UTEX 2714), Chlorella sorokiniana (UTEX 1230), Scenedesmus acutus f. alternans (UTEX B 72) and Scenedesmus dimorphus (UTEX 1237) were received from the Culture Collection of Algae at the University of Texas at Austin (UTEX). The Chlorella sorokiniana strain UTEX B 3010 was isolated from wastewater, identified, and deposited into the UTEX Algal Collection by the authors [20]. Finally, Chlorella sorokiniana (CCTCC M209220) was kindly provided by Minxi Wan, a visiting student at Johns Hopkins University. All species were maintained on sterile Bold's Basal Medium (BBM) agar

279

plates. The components of the BBM medium were as follows (per liter): 250 mg NaNO3, 176 g KH2PO4, 75 mg K2HPO4, 75 mg MgSO4·7H2O, 50 mg Tetrasodium EDTA, 31 mg KOH, 25 mg CaCl2·2H2O, 25 mg NaCl, 11.4 mg H3BO3, 10 mg FeSO4·7H2O, 8.83 mg ZnSO4·7H2O, 1.84 mg H2SO4, 1.57 mg CuSO4·5H2O, 1.44 MnSO4·H2O, 0.71 mg MoO3, 0.49 mg Co(NO3)2·6H2O. Before the start of the experiments, microalgal inocula were grown in 125 mL flasks containing 50 mL of autoclaved BBM medium using a fluorescent light source (T5/HO grow light bulbs, 6500 K, 14/10 h light/dark) with an intensity 140 μmol photon m− 2 s−1, unless otherwise stated, and mixed by a 20 mm magnetic stir bar at 350 rpm. The conversion between measured luminance (Model EA30 EasyView™ Wide Range Light Meter by Extech Instruments, Nashua, NH, USA) and photon flux density was 0.0185 μmol photon m−2 s−1 per 1 lx [42].

2.2. Wastewater media and cultivation experiments The wastewater and anaerobic digestion centrate (ADC) were obtained from the Back River Wastewater Treatment Plant (Baltimore, MD) and used the same day for cultivation experiments. Primary wastewater (wastewater I) was collected after the primary sedimentation tank but before the activated sludge process. Secondary wastewater (wastewater II) was collected after the secondary sedimentation tank following activated sludge treatment. The liquid fraction of the ADC was obtained by centrifugation of the anaerobic digestate consisting of a mix of the primary sewage sludge and surplus activated sludge. The physicochemical and microbiological characteristics of wastewater and ADC are shown in Tables 1 and 2. Three series of cultivation experiments were performed in order to evaluate the effects of ADC supplement, light intensity, and algal inoculum on biomass growth and productivity. The wastewater media for cultivation experiments were prepared by spiking either wastewater I or II with a specific amount of ADC. Microalgae were then cultivated in custom-built 440 mL cylindrical bioreactors open to the atmosphere and vertically exposed to light using a fluorescent light source described above (Supplementary information Fig. S2). First, 400 mL mix of freshly collected wastewater and ADC were carefully measured and transferred into bioreactors. Then, the inoculum cells grown in BBM were harvested by centrifugation in 50 mL centrifuge tubes (4200 ×g for 10 min), resuspended in corresponding wastewater medium and added into bioreactors to achieve target initial algal cell dose. Uninoculated wastewater media were used as controls in all experiments. All cultivations were performed in duplicate. In the first series of cultivation experiments, the level of ADC supplement to wastewater ranged from 0 to 20% (v/v). The microalgal cultures

Table 1 Physicochemical characteristics of primary and secondary wastewater (wastewater I & II respectively), anaerobic digestion centrate (ADC) and Bold's Basal Medium (BBM) medium. Parameter

Primary (I)

Secondary (II)

ADC

BBM1

pH OD680nm COD, mg L−1 TS, mg L−1 VS, mg L−1 TSS, mg L−1 VSS, mg L−1 PO4 – P, mg L−1 TP, mg L−1 NH3 – N, mg L−1 NO3 – N, mg L−1 TN, mg L−1 Inorganic N:P ratio

7.5–7.8 0.15 ± 0.05 270 ± 60 700 ± 90 250 ± 45 80 ± 30 66 ± 15 5.5 ± 3 8.6 ± 3.5 16 ± 5 0.9 ± 0.4 23 ± 5 3.1

7. 5–8.0 (8 ± 3) 10−3 38 ± 10 480 ± 50 130 ± 30 8±3 6±2 1.3 ± 0.3 1.6 ± 0.3 0.3 ± 0.2 5±1 7±3 4.1

6.9–7.2 0.6 ± 0.1 1600 ± 400 2950 ± 320 1500 ± 210 1400 ± 350 1050 ± 320 40 ± 7 51 ± 10 800 ± 100 5±1 880 ± 65 22

7.05 0.0 0.0 321 0.0 0.0 0.0 53.3 53.3 0.0 41.2 41.2 0.77

1

According to BBM recipe.

280


Table 2 Microbiological characteristics of primary & secondary wastewater.

(B)

Parameter

Primary

Secondary

(A)

Total coliforms, MPN1 L−1 Escherichia coli, MPN L−1 Enterococcus, MPN L−1 Pseudomonas aeruginosa, MPN L−1

108 107 106 106

106 105 104 104

(C)

(D)

(E)

(F)

1

Most probable number.

were grown with a light intensity 140 μmol photon m−2 s−1 using initial algal cell density 104 cell mL−1. In the second series of cultivation experiments, the light intensity was adjusted to five different levels: 30, 70, 100, 140 and 200 μmol photon m− 2 s− 1 while using constant ADC and inoculum doses of 5% (v/v) and 104 cell mL−1, respectively. In the third series of experiments, the algal cell inocula were added into the media in order to achieve the following initial cell doses (cell mL−1): 104; 105; 2.5 × 105; 5 × 105; and 106. The cultures were grown in either primary or secondary wastewater spiked with 5% ADC using light intensity of 140 μmol photon m−2 s−1. An additional cultivation experiment was performed to assess algal biomass fatty acid methyl esters (FAME) content using 4 L spinner flasks containing 3 L of either primary or secondary wastewater spiked with 5% ADC. The initial inoculation densities of C. sorokiniana UTEX B 3010 were 104; 2.5 × 105; 5 × 105; and 106 cell mL−1. The spinner flasks were incubated under a fluorescent light source (~180 μmol photon m−2 s−1, 14/10 h light/dark), supplied with ambient air at 2 ccm and mixed by a 40 mm magnetic stir bar at ~150 rpm. At the end of experiment, algal biomass was harvested by centrifugation in 50 mL centrifuge tubes (4200 ×g for 10 min) and used for fatty acid methyl esters analysis. 2.3. Determination of the microalgal growth Algal biomass density was estimated using sample optical density at 680 nm (OD680 algae). It was calculated by subtracting the optical density of the wastewater control (OD680 control, with no algal inoculum added) from the optical density of the algal culture in wastewater for the purpose of accounting for wastewater and ADC background color (Eq. (1)). 680 680 OD680 algae ¼ ODculture −ODcontrol

ð1Þ

Fig. 1. Relationship between microalgal biomass density as AFDW and optical density at 680 nm, linear regression slope k ± 95% CI and Pearson correlation coefficient R. Each data point corresponds to OD at 680 nm and ash free dry weight (AFDW) for microalgal cultures cultivated either in primary or secondary wastewater spiked with a specific amount of anaerobic digestion centrate ranging from 0 to 20%.

The average volumetric algal productivity was calculated for the cultivation period from 3.5 to 8.5 days (unless specified) by the following Eq. (4): PV ¼

∑ðAFDW i−1 −AFDW i Þ=ðt i−1 −t i Þ n

ð4Þ

where AFDWi−1 and AFDWi are the algal culture densities in mg L−1 at time ti−1 and ti, and t is the time of sampling in days. The error bars represent the 95% confidence interval calculated using Excel. The average areal algal productivity (g m− 2 d− 1) was estimated from the volumetric productivity (mg L−1 d−1) by applying the following Eq. (5): PA ¼ 0:145 P V

ð5Þ

where 0.145 is the depth of the bioreactor (m). A linear relationship between OD680 algae and ash free dry weight (AFDW) in mg per liter (mg L−1) was determined for every strain used in this study and applied to estimate the algal biomass density as AFDW at every time point (Eq. (2)). AFDW ¼ k OD680 algae

ð2Þ

where k is the slope of the linear regression (Fig. 1). The regression statistics were calculated using Excel. Each data point on the growth curves indicates the average of two independent biological replicates with calculated standard deviation. The specific growth rate (k1, d−1) was calculated by using the least squares method to fit the culture density as AFDW for the exponential parts of the growth curve of two independent cultures with the following Eq. (3): ln AFDW talgal ¼ k1 t þ ln AFDW 0algal

ð3Þ

where AFDWtalgal and AFDW0algal are the algal culture densities at day t and 0, and t is the time interval (in days) between AFDWtalgal and AFDW0algal. The regression statistics were calculated using Excel.

2.4. Analytical methods Wastewater samples were analyzed to determine the total suspended solids (TSS, mg L− 1) and volatile suspended solids (VSS, mg L− 1) according to the Standard Methods for the Examination of Water and Wastewater [43]. The chemical oxygen demand (COD), total −3 nitrogen (Ntot), ammonia (NH3), nitrate (NO− 3 ), phosphate (PO4 ), and total phosphorus (Ptot) were measured using Hach Kits (Hach, USA). The wastewater and ADC concentration of microelements was assayed using ICP-MS. Briefly, the elements were extracted from the biomass through microwave-assisted digestion (CEM Mars Xpress, Matthews, NC, USA) using concentrated nitric acid (TraceMetal™ Grade, Fisher Chemical) [44,45]. First, the samples were transferred (1 mL) into acid-washed Teflon digestion vessels. Then, concentrated nitric acid (10 mL) was added to vessels (the accurate weight of the sample and added nitric acid was measured gravimetrically using density 1.41 kg L−1). Next, the vessels were placed in the microwave and heated to 200 °C for 5 min in order to dissolve the biomass in the concentrated nitric acid completely. Finally, the samples were diluted with Milli-Q® water to obtain 2% HNO3 and the concentration of the selected elements were analyzed using an Elan® DRC™ II ICP-MS (PerkinElmer/Sciex,


281

primary treatment Influent

Grit removal

Screening

wastewater I

Primary settling primary sludge

Anaerobic digestion

biogas Sludge dewatering

CHP Plant

Screening

Grit removal

secondary treatment (biological)

primary sludge

RAS WAS

Sludge thickener

(B)

Activated sludge

Primary settling

Algal Secondary wastewater II cultivation settling

activated sludge

Anaerobic digestion

AD centrate biogas Sludge dewatering biosolids

CO2 rich flue gases

Influent

Algal biomass processing

power & heat

biosolids primary treatment

algal slurry

CO2 rich flue gases

AD centrate

Sludge thickener

(A)

Effluent Algal harvesting

Algal cultivation

CHP Plant

Algal Effluent harvesting algal slurry Algal biomass processing

power & heat

Fig. 2. Schematic diagram of the integrated phycoremediation and mirocalgal biomass production process. (A) Cultivation of microalgae in the primary effluent (wastewater I) spiked with ADC. (B) Cultivation of microalgae in the secondary effluent (wastewater II) spiked with ADC. (WAS – waste activated sludge; RAS – recycled activated sludge; CHP – combined heat and power or cogeneration plant).

Concord, ON, Canada). The calibration curve was prepared using TraceCERT® multielement standard solution (Sigma-Aldrich, St. Louis, USA) and the procedure was verified with Standard Reference Materials® (SRM2976; mussel tissue) purchased from the U.S. National Institute of Standards and Technology (Gaithersburg, MD, USA). The optical density of the algal cultures and controls were measured using a spectrophotometer (UV-1800, Shimadzu, Japan). The algal biomass density as AFDW (mg L−1) was estimated according to the Standard Methods for the Examination of Water and Wastewater [43]. The culture pH was determined daily using accumet AB15 Basic pH Meter (Fisher Scientific, USA). Fatty acid methyl esters (FAME): The in situ transesterification of the lipids was achieved by placing 34 mg of freeze-dried algal biomass sample into a round-bottom flask with Dimroth condenser and adding 3 mL of methanol containing 5% HCl (v/v). Additionally, 1 mL of 0.5 mg mL−1 heptadecanoic acid was added to each test tube as an internal standard. Samples were stirred in a water bath at 70 °C for 90 min. After that, samples were cooled to room temperature, 3 mL of hexane and 1 mL of water were added to extract the FAME. Then, samples were centrifuged

(A)

(B)

Fig. 3. Influence of ADC content on the (A) optical density (filled markers) and transmittance (opened markers) through 1 cm of primary (circles) and secondary (triangles) wastewater. (B) N:P ratio for primary and secondary wastewater with shaded area representing the optimal N:P values.

and the upper layer of hexane phase containing FAME was removed for analysis with a glass transfer pipette. FAME analysis was performed using a Gas Chromatograph (Shimadzu 2010, Shimadzu Corporation,

(A)

(B)

(C)

(D)

(E)

(F)

Fig. 4. Concentration of macro- and micronutrients in the primary (circles) and secondary (triangles) wastewater media as a function of ADC content. Arrows point to the corresponding Y-axis. The error bars represent the standard deviation among 4–5 wastewater samples collected on different days during three months.

282


(B.II)

(B.I)

(A.I)

(A.II)

(C.I)

(C.II)

(D.II)

(D.I) (E.I)

(F.II)

(E.II)

(F.I)

Fig. 5. Microalgal growth curves for different levels of AD centrate supplement. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with a certain amount of AD centrate (I + AD or II + AD, respectively).

(A)

(B) (A)

(B)

(D)

(C)

(D) (C) (E)

(E)

(F)

(F)

Fig. 6. Pseudo-first-order initial growth rate constants (k1) at different levels of AD centrate supplement. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with a certain amount of AD centrate (I + AD or II + AD, respectively).

Fig. 7. The average volumetric and areal productivities based on the amount of ADC added during days 3 to 9 of cultivation (I + AD or II + AD, respectively).


Japan) equipped with discharge ionization detector. The oven was equipped with a Stabilwax-DA 30 m × 0.25 mm × 0.5 μm column (Restek Corporation, USA). The injector temperature of the GC was set at 250 °C and the detector was set at 260 °C. The temperature program was started at 50 °C, increased to 190 °C at a rate of 20 °C min−1, and held constant for 1 min. The rate was next decreased to 4 °C min− 1 from 190 to 220 °C and then held constant for 14 min. Helium was used as the constant carrier gas. The identity of each FAME peak was determined by comparing the retention times of the compounds to the retention time of the individual FAME standards obtained from SigmaAldrich (St. Louis, MO, USA). The heptadecanoic methyl ester (SigmaAldrich, St. Louis, MO, USA) was used as an internal standard to quantify each FAME peak by relative area. FAME degree of unsaturation (DU) parameter and predicted biodiesel cetane number (CN) were calculated using empirical equations (Eqs. (6) & (7)) [46]: DU ¼ ðmonounsaturated; wt%Þ þ 2 ðpolyunsaturated; wt%Þ

ð6Þ

CN ¼ Σ Xi CNi

ð7Þ

where: Xi is the weight percentage of each methyl ester (wt%); CNi is the cetane number of a particular methyl ester. 3. Results and discussion 3.1. Influence of wastewater treatment and AD centrate content on media characteristics Conventional wastewater reclamation processes have two main process steps: primary (wastewater I) and secondary (wastewater II)

283

treatment. Microalgal cultivation might be incorporated into the treatment process through two alternative scenarios. In the first configuration (Fig. 2A), wastewater I obtained after the primary treatment step, involving only the physical separation of suspended solids, is used as the feed medium to cultivate microalgae. In the second configuration (Fig. 2B), the wastewater is processed sequentially in primary and secondary treatment steps, involving physical separation and biological treatment, respectively, and wastewater II is used as the algal medium afterwards. While both alternative designs have certain advantages, there are distinct disadvantages for each of them as well. For example, wastewater I has a relatively high optical density (OD) due to suspended particles that scatter and/or absorb light and thereby reduce its availability for the microalgae (Table 1). The light intensity for wastewater I diminishes 25 to 35% over 1 cm compared to only 2.5% for wastewater II (0% AD centrate in Fig. 3A). In addition, wastewater I has significantly higher bacterial contamination that can be deleterious for microalgae and compete for organic carbon and other nutrients (Table 2). In contrast, wastewater II has a lower concentration of bacteria but it has nearly 3 times less nitrogen and 4 times less phosphorus (Table 1). This reduction in nutrients may significantly limit algal productivity and biomass density grown in wastewater II. Additionally, the nitrogen to phosphorus ratio (N:P) for wastewaters I and II are 3.1 and 4.1, respectively. In contrast, the optimal N:P value for microalgal cultivation was estimated to be 10 for freshwater green microalgae [47] or in the range from 6 to 8.5 (based on the phytoplankton cell composition) [48]. Thus, both wastewaters I and II are limiting in nitrogen content for optimal microalgal growth. The nitrogen balance in wastewater and the N:P ratio could be potentially improved by supplementing with nutrient-rich side streams, such as ADC generated from the dewatering of anaerobically digested

Fig. 8. Microalgal growth curves for different light intensities. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).

284


The impact of ADC concentration on growth rate and biomass productivity was measured for five microalgal strains and their poly-culture. Primary and secondary wastewaters were spiked with various

amounts of ADC to have media with an ADC concentration ranging from 0 to 20%. The effect of ADC content on the density of the algal cultures measured as TSS over time is shown in Fig. 5. As anticipated, addition of the nutrient-rich ADC was advantageous for algal growth and resulted in higher density for almost all tested cultures. This positive effect was stronger for cultures cultivated in wastewater II spiked with ADC as compared to cultures grown in wastewater I with ADC added. The positive influence was greater for wastewater II likely due to the removal of nutrients during secondary treatment and resulting nutrient scarcity as discussed previously. The highest biomass concentration reached only 200 mg L−1 for Chlorella sorokiniana and as low as 100 mg L−1 for other microalgae when cultivated in the wastewater II alone. In contrast, the maximum concentration of microalgal biomass in wastewater II + ADC reached up to 350–380 mg L− 1. Therefore, ADC might represent an inexpensive substitute to chemical fertilizers for algae grown in wastewater or other media with an insufficient nutrient content. Additionally, centrate utilization for microalgal cultivation reduces the load on the wastewater treatment plant and therefore the cost of reclamation. However, the effects of ADC supplements were uneven for the different microalgae. In general, the final biomass density increased by 1.5–3.4 times due to addition of centrate. In contrast, ADC supplement had no positive effect on the Scenedesmus acutus f. alternans growth. Various microalgae likely have different nutrient requirements and/or unequal capabilities for scavenging nutrients from the solution. Therefore, it will be important to determine the optimal ADC concentration for every type of cultivated microalgae prior to process scale-up. The effects of ADC on initial growth rate and biomass productivity are shown in Figs. 6 and 7. The initial growth rate and productivity improved with the addition of the centrate from 0% to 5% followed by a plateau from 5 to 10% and decreased with further addition of the AD centrate for most of the species. Indeed, the growth of all strains except Chlorella sorokiniana was inhibited at high (20%) concentrations of

Fig. 9. Pseudo-first-order initial growth rate constants (k1) at different light intensities. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).

Fig. 10. Volumetric and areal productivities at different light intensities (calculated as average of daily productivities from day 3 to day 9 of cultivation). The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).

sludge (see Fig. 2). Addition of 5 to 20% (v/v) ADC to wastewater I or 1 to 5% (v/v) to wastewater II shifts the N:P ratio into the optimal range (Fig. 3B). In addition, ADC can be used to supplement wastewater with additional macro- and micronutrients essential for algal growth and potentially improve algal productivity. The influence of ADC dose in wastewaters I and II on the content of selected elements is shown in Fig. 4. First, it is important to point out that the content of N, P, Mn, B, Zn, and Co in wastewater I is N50% higher than wastewater II, which comes after the biological wastewater treatment. Second, addition of ADC to both primary and secondary wastewaters significantly enhances (by more than on 100%) the content of N, P, Fe, Mn, and Co when compared to wastewater alone. The concentrations of S, Mg, Ca, and Mo increased moderately, by 20–60%. Also, there was no increase in the levels of the B and Cu when adding ADC, which suggests that ADC is not a good source for these elements. Finally, even with addition of ADC at levels as high as 20%, the concentrations of some trace elements (B, Cu, Zn, Mo, and Co) in the wastewater media are substantially lower than in defined growth media commonly used for microalgal cultivation. For comparison, the composition of Bold's Basal Medium (BBM) typically used to grow green microalgae is N 41 mg L−1, P 53 mg L−1, S 12.1 mg L− 1, Mg 7.4 mg L−1, Ca 6.8 mg L−1, B 2 mg L−1, Zn 2 mg L−1, Fe 1 mg L−1, Mo 470 μg L−1, Mn 400 μg L−1, Cu 400 μg L−1 and Co 100 μg L−1. Furthermore, supplementing with greater amounts of ADC can significantly increase the wastewater OD and reduce light transmittance (Fig. 3A). As a result, the amount of light available for microalgae can be attenuated up to 50% and 25% in 1 cm of wastewaters I and II, respectively, when spiked with the 20% (v/v) of the ADC. 3.2. Influence of ADC level on microalgal growth


centrate. This negative impact could be caused by high ammonia content in the ADC (nearly 800 mg L−1) and/or the potential presence of propionate (typically below 100 mg L− 1 but may spike above 1000 mg L−1 for overload events [49–51]). It has been shown previously that ammonia at concentrations higher than 50–100 mg L− 1 as N could inhibit growth of some microalgae including Chlorella and Scenedesmus [52,53]. Also, it has been demonstrated that ammonium toxicity strongly depends on the solution pH and distribution between the toxic ammonia and nontoxic ionized ammonium [54]. Therefore, the control of pH by addition of CO2 can be a potential solution to mitigate the ammonia toxicity for microalgae [55]. Other results suggest that propionate present in the ADC can inhibit growth of Chlorella and Scenedesmus microalgae [30,56]. Importantly, the C. sorokiniana strain used in the current study did not exhibit any inhibition of growth even at high concentrations of the ADC. Indeed, some previous studies showed that certain microalgae have better tolerance [57,58] or ability to acclimatize [59,60] to high ammonia concentrations. Identification and application of the strains with natural low sensitivity and/or high adaptability to the presence of ammonia represents one approach to avoid limitations in processing due to toxicity effects. The maximum biomass productivity observed for C. sorokiniana reached 60 mg L−1 d−1 or 8.7 g m−2 d−1, which is favorable compared to the maximum productivity of other species of approximately 40 mg L−1 d−1 or 5.8 g m−2 d−1. These measured productivity values are in the same range as peak productivities reported previously for microalgae grown in secondary municipal wastewater (33 mg L−1 d−1) [61], untreated carpet industry wastewaters (41 mg L−1 d−1) [62], anaerobically digested dairy manure (5.5 g m−2 d−1) [31,63], pig manure (29 mg L−1 d−1) [64], and poultry litter (37–76 mg L−1 d−1) [65]. However, other studies have demonstrated that a biomass productivity of

285

200–600 mg L−1 d−1 can be achieved when using ADC as the sole source of nutrients (reviewed by Xia and Murphy [66]). Further enhancement in microalgal productivity would likely require optimization of carbon supply. Some studies indicated that supplementing gas enriched with 2–5% CO2 could increase the algal biomass density and productivity 1.8 to 2.5 fold [25,62]. The strategy of bubbling algal cultures with CO2-rich flue gases may be advantageous from both economic and sustainability perspectives. However, further increases in the CO2 concentration to 10– 15% of the supplied gas can inhibit the microalgal growth [25] and requires identification and application of the strains tolerant to extreme CO2 levels [67–69]. Another strategy to achieve higher productivity would be to take advantage of the organic carbon present in the wastewater and ADC. Indeed, productivity of microalgae capable of mixotrophic and heterotrophic growth can be up to 400–500 mg L−1 d−1 in concentrated municipal wastewater rich in organic carbon [70]. Importantly, it was demonstrated that species of C. sorokiniana and Scenedesmus sp. possess a unique carbon metabolism capable of autotrophy, mixotrophy, and heterotrophy, taking advantage of fixed carbon in wastewater and ADC [71–73]. However, methods to suppress the bacterial growth other than sterilization by autoclaving would be necessary in order to make mixotrophic/heterotrophic cultivation economically feasible. Finally, photoautotrophic and mixotrophic microalgal growth may be limited by light availability. Given its importance to algae, the influence of the irradiance intensity on algal growth in wastewater media was assessed next. 3.3. Influence of light intensity on microalgal growth The biomass density in the algal cultures over time at various light intensities is illustrated in Fig. 8. The algal growth rate and biomass density were very poor at low light intensity (30 μmol photon m−2 s−1).

Fig. 11. Growth curves for different initial doses of microalgal inoculum. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).

286


Thus, algal cultivation may be inefficient in areas with frequent overcast days and sunlight intensity below 40 μmol photon m−2 s−1. Increasing the light intensity resulted in higher biomass concentrations for most of the tested microalgal species. Interestingly, photoinhibition was not observed even at high values of intensity between 140 and 200 μmol photon m−2 s−1. However, the initial growth rate was limited in light only in the range from 30 to 140 μmol photon m−2 s−1 (Fig. 8). The plateau in the growth rate with further increases in the light intensity above approximately 140 μmol photon m−2 s−1 reflects the irradiance saturation for the studied species. Previous studies indicate saturation irradiance can vary over a wide range from 30 to 280 μmol photon m−2 s−1 for aquatic photosynthetic microorganisms with an average of 100 ± 50 μmol photon m−2 s−1 (identified for 24 species, primarily dinoflagellates and diatoms) [74]. The lowest reported saturation irradiance values for Chlorella species reported were nearly 20 μmol photon m−2 s−1 for Chlorella pyrenoidosa [75], 110 μmol photon m−2 s−1 for Chlorella sp. [76] and 250 μmol photon m−2 s−1 for Chlorella kessleri [77]. Saturation light intensity measured in the current study falls within this same range. While different microalgae are phenotypically acclimated to habitats with various amounts of light, the photoinhibition and saturation irradiance also strongly depends on other environmental parameters such as temperature and availability of CO2, water, and nutrients [78]. Limitations in CO2 as a sink of excitation energy through carbon reduction and/or nitrogen as one of the main building blocks for RuBisCO enzyme may cause saturation and photoinhibition at lower irradiances compared to cultures with elevated CO2 and nitrogen contents [79]. In addition, microalgae have the ability to photo-acclimatize to extreme irradiance levels [80]. Finally, it is important to note that the outdoor sunlight irradiation is usually higher than the range of intensities investigated in the current study and can reach up to 370 μmol photon m−2 s−1 for diffused sunlight and to N1800 μmol photon m−2 s−1 for direct sunlight on a clear day. Therefore, the photoinhibition and saturation irradiance levels have to be closely monitored prior to scaling up a process for each species under varying environments and cultivation conditions. For most microalgal species, both growth rate and productivity increased over most of the range of intensities but slowed or stalled above 150 μmol photon m− 2 s−1 (Figs. 9 and 10). Unlike the initial growth rate, however, the productivity of other select species (i.e., C. sorokiniana and an microalga species isolated from local wastewater) continued to increase for all ranges of tested intensities (Fig. 10). These species were saturated in light at low cell density of the culture but were light-limited at high biomass density, perhaps due to the self-shading effect. Furthermore, the productivity in wastewater II spiked when ADC was higher than in wastewater I spiked with the ADC for most species and their poly-culture. One potential reason for this result is the high concentration of bacteria in the primary wastewater that may be deleterious for microalgae and consume essential nutrient resources. In contrast, productivities for C. sorokiniana and the local microalgal species were quite similar in both wastewaters I and II. It is possible that these species are resistant to adverse effects from bacteria and are able to compete effectively for nutrients. Maximum productivity measured in the current study was up to 45 mg L− 1 d− 1 or 6 g m−2 d−1.

of algal cells has been found to be beneficial for their survival in the presence of harsh and toxic compounds, which may be present in wastewater or ADC [81–84]. As previously discussed, nutrient-rich ADC contains high concentrations of ammonia, which can be toxic for microalgae [71–74]. Among all tested strains, the highest growth rates of nearly 1.2 d−1 were observed for C. sorokiniana strains UTEX B 3010 and CCTCCM 209220, followed by UTEX 1230, (1.1 d−1) another C. sorokiniana strain (Fig. 12). While the highest biomass densities of the cultures were observed when using primary effluent, the highest growth rates were obtained in secondary effluent at the lowest initial inoculum dose of 104 cells mL−1. Although the primary effluent had a high nutrient content, the higher optical density and lower light penetration efficiency led to lower initial growth rates than the secondary effluent. Both Scenedesmus species exhibited lower growth rates than the Chlorella species with the peak rates of 0.5 d− 1 at inoculum concentrations of 105–2.5 × 105 cells mL−1 (Fig. 12E and F). However, Scenedesmus cultures yielded more biomass than Chlorella spp., which has been previously observed [20,85]. Specifically, S. dimorphus UTEX 1237 achieved the highest biomass density of 350 mg L−1 followed by S. acutus UTEX B72 (300 mg L−1) and then C. sorokiniana UTEX B 3010 (275 mg L−1), which achieved these levels at the highest initial inoculum dose. All other Chlorella strains tested in this study (CCTCCM 209220, UTEX B 3010, UTEX 1230 and UTEX 2714) also reached their highest biomass density with the largest initial inoculum of 106 cells mL−1 and in the primary effluent spiked with 5% AD centrate to confirm the importance of both inoculum size and the role of nutrients [85,86]. Interestingly, Scenedesmus sp. achieved nearly 40% higher biomass density in dry weight per volume than Chlorella even at similar levels of culture optical density [85]. Scenedesmus cells (crescent-shaped, 3 μm × 10 μm) are larger in size than C. sorokiniana cells (spherical, 3–4 μm) and exhibit a natural predisposition to form coenobia (colonies of 4 cells attached side by side) (see Supplemental Materials Fig. S1). Increasing the

3.4. Influence of inoculum dose on microalgal growth of all tested species and lipid content and profile of newly characterized C. sorokiniana UTEX B 3010 The influence of inoculum dose on microalgal growth in wastewater was tested in the next cultivation experiment using initial algal concentrations of 104, 105, 2.5 × 105, 5 × 105 and 106 cells mL−1. As shown in Fig. 11, the biomass density increased along with the inoculum dose throughout the cultivation time for all algal cultures. Increasing the algal inoculum may have a significant effect on relations between microalgae and wastewater-borne bacteria, especially in competition for organic carbon and nutrients. In addition, a high initial concentration

Fig. 12. Microalgal growth rate constant for different initial doses of microalgal inoculum. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).


Fig. 13. Microalgal productivity for different initial doses of microalgal inoculum. The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).

inoculum size also had a positive effect on volumetric and areal productivities (Fig. 13). Although the trend was quite similar for most tested species and all cultivation conditions it appeared to be stronger for Chlorella strains cultivated in secondary effluent than in primary effluent. However, the collapse of some select Scenedesmus cultures after 3–4 days of cultivation in supplemented secondary effluent, perhaps due to an adverse impact from bacterial contamination, led to a nearly zero-order productivity for these cultures over the nine days of cultivation.

287

In order to determine the lipid content, productivity and fatty acid methyl esters (FAME) composition important to algal biofuel feasibility, the C. sorokiniana UTEX B 3010 strain was cultivated with inoculum densities of 104, 105, 5 × 105 and 106 cells mL−1 in photobioreactors with a larger volume to generate sufficient algal biomass for lipid analysis. The C. sorokiniana UTEX B 3010 growth curves and culture pH achieved in the scaled-up photobioreactors are shown in Fig. 14. As observed previously (Fig. 11) in the smaller volume photobioreactors, the biomass density was directly related to algal inoculum density. The highest algal biomass density of 380 mg L−1 was achieved with the highest inoculum dose and cultivation in primary effluent spiked with 5% AD. The biomass lipid content, composition and predicted fuel characteristics such as kinematic viscosity (KV), cetane number (CN) and oxidative stability (OS) are shown in Table 3. The major methyl esters representing nearly 75% of all esters were composed of palmitic, linoleic and linolenic acids, which offer advantageous viscosity characteristics for the biodiesel generated. However, the presence of unsaturated methyl esters containing three double bonds (linolenic and hexadecatrienoic represented nearly 30% of FAME), while favoring the viscosity properties, may affect the fuel oxidative stability. In addition, these unsaturated esters have a CN below the standard values recommended for biodiesel and petrodiesel by the American Society for Testing and Materials (ASTM) and European Committee for Standardization (CEN) [87,88]. Importantly, the estimated average CN of the FAME mixture was within the recommended standard value. While the lowest lipid content and the lowest average CN were observed at the smallest inoculum dose, increasing inocula concentrations above 105 cell mL−1 did not strongly benefit the lipid content or improve lipid composition. High concentration of inoculum allowed microalgae to compete more successfully with wastewater bacteria for nutrients and organic carbon present in wastewater but high inoculum did not shift microalgal metabolism towards lipid accumulation. Interestingly, few previous studies noted that presence of bacteria in algal culture might either decrease or increase the biomass lipid content [89–91]. Lu et al. suggested that dense microalgal cultures originating from high inocula doses might be light limited and accumulate less energy storage products such as lipids [92]. Indeed, a minor decrease in lipid content was observed at the highest inoculum dose in the current study as well. Therefore, there may be an optimal inoculum loading in terms of lipid composition. Indeed, previous reports from our and

Fig. 14. Microalgal growth curves and culture pH for different initial doses of inoculum (cultivation in 4 L spinner flasks). The cultures were cultivated either in primary (I) or secondary (II) wastewater spiked with 5% of AD centrate (I + 5% AD or II + 5% AD, respectively).

288


Table 3 C. sorokiniana UTEX B 3010 biomass FAME composition, content and properties at various inoculum density. FAME content (% dwb,) at various inoculum doses in cell mL−1 Primary effluent+5% AD Fatty acid methyl estera Palmitic Palmitoleic Hexadecadienoic Hexadecatrienoic cis-10-heptadecenoic Stearic Oleic Linoleic Linolenic Total FAME, % dw Saturated, %f Monounsaturated, % Polyunsaturated, % Degree of unsaturation Average predicted kinematic viscosity at 40 °C, mm2 s−1 Average predicted CN

C16:0 C16:1 C16:2 C16:3 C17:1 C18:0 C18:1ω9 C18:2ω6 C18:3ω3

Secondary effluent+5% AD

104

105

2.5 × 105

106

104

105

2.5 × 105

106

KV c, mm2 s−1

CN d

OS e, h

1.49 0.0 0.0 0.88 0.67 0.0 0.0 1.89 1.30 6.2 23.9 10.8 65.3 1.41 3.63 49.5

1.87 0.0 0.0 0.77 0.76 0.0 0.38 2.18 1.55 7.5 24.9 15.2 59.9 1.35 3.72 51.3

2.16 0.0 0.0 0.93 0.77 0.0 0.38 2.34 1.48 8.1 26.9 14.3 58.9 1.32 3.73 51.9

2.02 0.0 0.0 0.93 0.52 0.0 0.41 2.16 1.04 7.1 28.5 13.1 58.3 1.30 3.74 52.6

1.25 0.0 0.0 0.38 0.75 0.0 0.0 1.04 1.48 4.9 25.6 15.2 59.2 1.34 3.68 50.7

1.69 0.0 0.0 0.63 0.86 0.0 0.0 1.79 1.70 6.7 25.4 12.9 61.8 1.36 3.67 50.5

1.88 0.0 0.0 0.74 0.87 0.0 0.0 2.08 1.71 7.3 25.8 11.9 62.3 1.36 3.67 50.6

1.84 0.0 0.0 0.83 0.72 0.0 0.0 2.24 1.41 7.0 26.1 10.2 63.7 1.38 3.67 50.6

4.38 3.67 3.10 2.70 4.09 5.85 4.51 3.65 3.14

80.9 56.1 38.9 29.1 59.3 90.1 62.2 42.8 31.8

N24 2.11 n.a.g n.a. n.a. N24 2.79 0.94 0.00

a

The amount of other FAMEs is less than 5% of total FAME content. Dry weight percent. c Measured and predicted kinematic viscosity at 40 ° C. The standard values are 1.9–6.0 mm2 s−1 (ASTM D6751, USA) and 3.5–5.0 mm2 s−1 (EN 14214, EU) [87,88]. d Cetane number estimated by using quadraticcorrelation with the number of carbon atoms and the number of double bonds [101]. The standard values are 47 (ASTM D6751, USA) and 51 (EN 14214, EU) [87,88]. e Oxidative stability at 110 °C, h (Rancimat test). The standard values are 3 min (ASTM D6751, USA) and 6 min (EN 14214, EU) [87,88]. f As percent from total FAME content. g Not available. b

other laboratories have found that nutrient limited conditions can enhance the lipid content for some of the C. sorokiniana and Scenedesmus species tested in the current study [30,71,91,93–100]. Therefore, follow-up investigations will be useful to achieve both maximal biomass and biodiesel precursor content. 4. Conclusions Unsterilized primary and secondary wastewaters were shown to be suitable substrates for cultivation of Chlorella and Scenedesmus strains. Microalgal cultivation in primary wastewater was observed to be superior despite stronger contamination by potentially detrimental bacteria due to higher levels of nutrients and less pretreatment. However, both primary and secondary wastewaters were found to lack a balanced nutrient content needed to support augmented algal growth. The addition of 5–10% anaerobic digestion centrate (ADC) was shown to be an excellent supplement for improving microalgal growth and productivity by enhancing the levels of essential macro- and micronutrients and improving the nitrogen to phosphorus ratio. This enhancement in growth was found to be strain-dependent with species such as Chlorella sorokiniana and Scenedesmus acutus f. alternans especially appropriate for growth in supplemented wastewater feeds. Incident light intensity proved to be another critical factor controlling algal growth even when an excess of nutrients is provided. The growth of all tested microalgae was increased as the light intensity was raised from 30 to 140 μmol photon m−2 s−1 with further increases not providing further improvements in algal growth. In addition, supplying a sufficient dose of algal inoculum (above 2.5 × 105 cell mL−1) was critical for achieving maximum final densities of algal cultures and therefore biomass productivity. Presumably, increasing the initial microalgal inoculum concentration enables microalgae to compete more efficiently with wastewater borne bacteria for nutrients and organic carbon present in wastewater. The inoculum dose was also observed to have a minor effect on the FAME content and composition of harvested biomass of C. sorokiniana UTEX B 3010 with the highest FAME content at an inoculum dose of 2.5 × 105 cell mL−1, supporting a potential limit on lipid production at high cell densities. The most abundant methyl esters were represented by palmitic, linoleic and linolenic acids offering advantageous

viscosity and cetane number characteristics for the generated biodiesel. In addition, follow-up studies would be worthwhile to better understand the effects of parameters such as inoculum dose and bacterial content on microalgal culture overall FAME content and composition across a range of prospective algal candidates. Acknowledgements The authors gratefully acknowledge the financial support from the U.S. Environmental Protection Agency P3 Award Program (Grant No. SU835318), from the U.S. NSF CBET Program (Grant No. 1236691), and from the Bureau of Education and Cultural Affairs of U.S. Department of State through an International Fulbright Science and Technology Award to Pavlo Bohutskyi. Also, the authors would like to thank Nick Frankos and Marshall Phillips for their assistance in collection of wastewater and anaerobic digestion effluent samples from the Back River Wastewater Treatment Plant in Baltimore that were used in this study. Finally, we would like to thank Minxi Wan for sharing the Chlorella sorokiniana (CCTCC M209220) strain. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.algal.2016.09.010. References [1] Y. Tang, J.N. Rosenberg, P. Bohutskyi, G. Yu, M.J. Betenbaugh, F. Wang, Microalgae as a feedstock for biofuel precursors and value-added products: green fuels and golden opportunities, 2015. [2] E. Uggetti, B. Sialve, E. Trably, J.-P. Steyer, Integrating microalgae production with anaerobic digestion: a biorefinery approach, Biofuels Bioprod. Biorefin. 8 (2014) 516–529. [3] R.W. Nicol, A. Lamers, W.D. Lubitz, P.J. McGinn, Evaluation of algal biomass and biodiesel co-products for bioenergy applications, J. Biobased Mater. Bioenergy 8 (2014) 429–436. [4] J.N. Rogers, J.N. Rosenberg, B.J. Guzman, V.H. Oh, L.E. Mimbela, A. Ghassemi, M.J. Betenbaugh, G.A. Oyler, M.D. Donohue, A critical analysis of paddlewheel-driven raceway ponds for algal biofuel production at commercial scales, Algal Res. 4 (2014) 76–88.

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