Bioprocess Biosyst Eng DOI 10.1007/s00449-014-1303-5

ORIGINAL PAPER

High-productivity lipid production using mixed trophic state cultivation of Auxenochlorella (Chlorella) protothecoides Hamid Rismani-Yazdi • Kristin H. Hampel • Christopher D. Lane Ben A. Kessler • Nicholas M. White • Kenneth M. Moats • F. C. Thomas Allnutt

•

Received: 28 May 2014 / Accepted: 7 October 2014 Ó Springer-Verlag Berlin Heidelberg 2014

Abstract A mixed trophic state production process for algal lipids for use as feedstock for renewable biofuel production was developed and deployed at subpilot scale using a green microalga, Auxenochlorella (Chlorella) protothecoides. The process is composed of two separate stages: (1) the photoautotrophic stage, focused on biomass production in open ponds, and (2) the heterotrophic stage focused on lipid production and accumulation in aerobic bioreactors using fixed carbon substrates (e.g., sugar). The process achieved biomass and lipid productivities of 0.5 H. Rismani-Yazdi
K. H. Hampel
C. D. Lane
B. A. Kessler
N. M. White
K. M. Moats
F. C. Thomas Allnutt Phycal Inc., 51 Alpha Park, Highland Heights, OH 44143, USA

and 0.27 g/L/h that were, respectively, over 250 and 670 times higher than those obtained from the photoautotrophic cultivation stage. The biomass oil content (over 60 % w/DCW) following the two-stage process was predominantly monounsaturated fatty acids (*82 %) and largely free of contaminating pigments that is more suitable for biodiesel production than photosynthetically generated lipid. Similar process performances were obtained using cassava hydrolysate as an alternative feedstock to glucose. Keywords Auxenochlorella (Chlorella) protothecoides
Algae biofuel
Heterotrophic
Photoautotrophic
Fermentation

K. H. Hampel e-mail: [email protected]

Introduction

C. D. Lane e-mail: [email protected]

There has been a surge of activity in recent years focused on new methods and their validation for the production of economically and environmentally viable biofuels using microalgae. A contributing factor to the increased activity around algal biofuels was the increasing price of petroleum, which peaked at US$145 per barrel in July of 2008. R&D activity continues and improvements to the unit processes surrounding algal biofuel production have occurred, such as improvement in light penetration for algae production in ponds [1], novel harvesting methods [2], extraction of lipids [3, 4], hydrothermal liquefaction [5], innovative bioreactors [4], and catalytic gasification [6]. Nonetheless, recent analyses indicate that technical and economic hurdles remain (as well as political obstacles) to profitable commercialization [7]. Continued innovation and research are required to overcome the remaining technological and commercial barriers. It is clear that there is a need to replace fossil fuels with renewable and sustainable fuels and that

B. A. Kessler e-mail: [email protected] N. M. White e-mail: [email protected] K. M. Moats e-mail: [email protected] F. C. Thomas Allnutt e-mail: [email protected] Present Address: H. Rismani-Yazdi (&) Novozymes North America Inc., P.O. Box 576, Franklinton, NC 27525, USA e-mail: [email protected] Present Address: F. C. Thomas Allnutt BrioBiotech LLC, P.O. Box 26, Glenelg, MD 21737, USA

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algal biofuels still have tremendous potential to contribute to this positive outcome if these hurdles can be overcome. Mixed trophic state background Traditionally, algal mass culturing is photoautotrophic, with a complete reliance on solar energy and inorganic carbon (carbon dioxide or bicarbonate) for conversion into sugars and complex carbon compounds. Photoautotrophic growth is, by its nature, self-limiting due to shading that occurs as cell density increases (limiting light penetration) and has other associated problems, such as pond water loss through evaporation and insufficient gas exchange [8]. While the majority of algae are photoautotrophic, a surprising number are capable of heterotrophic or mixotrophic growth [9–13]. This led to efforts to look at the heterotrophic or mixotrophic production of biofuels from algae [10]. Production of biomass through heterotrophy, mixotrophy, or using mixed trophic states provides an opportunity for improved productivity but requires an inexpensive source of fixed carbon usable by algae (e.g., hydrolysates or partially purified sugars) to be commercially relevant. With the increasing focus on the production of ethanol from cellulosic feedstock and starch, it is interesting to note that algae represent a way to produce triglycerides from these same sugar sources that, once transesterified, can be burned directly as transportation fuels or act as an ideal feedstock for the production of dropin replacements for existing transportation fuels (i.e., having equivalent energy content and properties). Employing microalgae cultivation systems that use renewable inputs to generate biofuels is one potential approach to improving the ability to reach economic targets for production of algal biofuels. The production of algal biomass based on mixotrophic growth, simultaneously using light and fixed carbon for growth, is well known [14– 17]. Mixotrophic production systems for algae, based on deep ponds and supplied with fixed carbon (e.g., glucose), have been applied commercially for the production of Chlorella for the health food markets [18]. Alternatively, a mixed trophic cultivation process has been recently proposed for the production of biofuels [19–21]. The mixed trophic state systems use multiple trophic processes that are separated either by time or location (for example, photoautotrophic growth then heterotrophic growth). A mixed trophic state system is the focus of this work and will be referred to as the Heteroboost process (HTB). This HTB process has a photoautotrophic stage of algae cultivation in open ponds that is followed by a concentration step and then a heterotrophic growth stage in an aerobic bioreactor. The process takes advantage of the ability of the sun to provide cheap energy to photosynthetically fix inorganic carbon from either the atmosphere or industrial emissions,

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yet is able to rapidly accumulate oil to high density during the heterotrophic growth using renewable inputs from agricultural production of a fixed carbon feedstock (e.g., sugar, acetate or glycerol). The advantages anticipated in the HTB process are the spatial and temporal separation of inexpensive mass culture of algae on a continuous basis (the photoautotrophic stage) and rapid lipid accumulation in a more concentrated and constrained system (the heterotrophic bioreactor). The HTB process contrasts with traditional photoautotrophic mass culturing systems where the cells are grown to the highest density possible under photoautotrophic conditions then subjected to stress (e.g., nutrient limitation such as nitrogen) to induce lipid synthesis. The traditional photoautotrophic approach is limiting in that it cannot be run as a continuous process and requires additional time to induce lipid synthesis through induction of the nutrient stress. The HTB process uses the photoautotrophic step for biomass generation in open ponds, which can be run in a continuous manner at the optimized maximal production rate; this is an important difference from the straight photoautotrophy and nutrient stress approach traditionally used. The ability of the microalga Auxenochlorella (Chlorella) protothecoides to grow both photoautotrophically and heterotrophically, when combined with its high photosynthetic efficiency of 8 % [22] and ability to accumulate large quantities of neutral lipids (up to 70 %) [23] at relatively high cell densities using a range of fixed carbon substrates [24], make it a suitable candidate for the HTB process. A recent review of the genus Chlorella discovered that species traditionally ascribed to Chlorella were spread over two classes of chlorophytes, the Trebouxiophyceae and the Chlorophyceae [25]. The main species used in the current study was placed in the Trebouxiophyceae and renamed as Auxenochlorella protothecoides. An extensive academic and patent background has been developed around the heterotrophic growth and lipid production in A. protothecoides [10, 21, 26]. The objective of this study was to develop and evaluate a mixed trophic state cultivation strategy for high productivity mass culturing and lipid production by A. protothecoides at subpilot scale.

Materials and methods Organism and medium composition Auxenochlorella (Chlorella) protothecoides KRT1009, a single-celled green alga belonging to the phylum Chlorophyta, was selected from UTEX 25 obtained from the University of Texas Culture Collection. UTEX 25 as received contained mixed cell types, a clonal line of A. protothecoides was isolated from this mother culture and

Bioprocess Biosyst Eng Fig. 1 A schematic diagram of the HTB mixed trophic state process and the seed train for production of inoculum for ponds in the subpilot plant

verified using the cleaved amplified polymorphic sequence techniques [27]. For photoautotrophic growth, the medium was composed of 0.5 g NH4Cl, 1.44 g K2HPO4, 0.72 g KH2PO4, 50 mg tetrasodium EDTA, 20 mg MgSO4
7H2O, 10 mg CaCl2
2H2O, 10 mg FeCl3
6H2O, 5 mg H3BO3, 1.25 mg ZnSO4
7H2O, 0.38 MnSO4
H2O, 0.25 mg CoCl2
6H2O, 0.25 mg Na2MoO4
2H2O, 0.08 mg CuSO4
5H2O, and 100 lg thiamine hydrochloride per liter of water. Subculture and seed cultivation Seed cultures were grown in an inoculum train as shown in Fig. 1. Seed cultures were maintained on agar plates and transferred regularly to new plates such that the age of the cells on the plate at the time of starting the first flask was consistent. The stages of the inoculum train are presented in Table 1: (1) 25 mL volume in a 250-mL flask, (2) 250 mL in a 500-mL flask, (3) 2 L volume in a 2-L flatbottom flask, and (4) 20 L volume in a carboy. Based on historical data, cultures were transferred to maintain an exponentially growing population up to the carboy stage. Stages 1 and 2 were maintained at 28 ± 2 °C, a constant 140 ± 10 lmol/m2/s light intensity from an Eye Hortilux metal halide bulb (Iwasaki MT1000B-D/HOR/HTL-BL1) in a SunTubeTM six-inch reflector (Sunlight Supply, Vancouver, WA), 5,000 ppm ambient CO2 concentration, and

180 rpm shaking speed. Stage 3 was maintained at the same light and temperature levels, but was bubbled with a sterile-filtered air/CO2 mix (approximately 1 % (w/w) CO2) at 1 L/min. The gas flow provided both the mixing and carbon source. Aseptic technique was used to maintain an aseptic culture until the carboy stage. The Stage 4 carboy was maintained at the same temperature, light, and CO2 conditions as Stage 3, but with a higher total gas flow rate that was set manually to provide optimal mixing. Flasks and carboys were grown in triplicate vessels. Photoautotrophic growth in raceway ponds The carboy volumes were combined and mixed before being redistributed into two 2,400 L raceway ponds that were approximately 4.5 m long, 1.2 m wide and 0.45 m deep, and were operated at 15 cm volume depth with 800 L working volume. Raceway ponds were grown in duplicate. Mixing was provided through a paddlewheel with a liquid linear velocity of no less than 0.2 cm/s throughout the pond. Eye Hortilux metal halide bulbs (Iwasaki) in SunTubeTM six-inch reflectors (Sunlight Supply) were used to provide the culture with an average of 700 lmol/m2/s incident light intensity. Carbon dioxide was provided through a sintered metal sparger into each pond based on a feedback loop controlling a solenoid valve. The valve was

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Bioprocess Biosyst Eng Table 1 Conditions for photoautotrophic production of A. protothecoides biomass

Inoculum stages

Vessel

Vessel capacity (L)

Culture volume (L)

Light intensity (lmol/m2/s)

Starting density (A750)

Final density (A750)

Duration (days)

Stage 1

Erlenmeyer flask

0.25

0.025

140

0.08 ± 0.0

1.37 ± 0.08

1.8

Stage 2

Erlenmeyer flask

0.50

0.25

140

0.19 ± 0.0

0.75 ± 0.03

1.0

Stage 3

Flat-bottom flask

2

2

140

0.08 ± 0.01

3.06 ± 0.75

4.2

Stage 4

Carboy

20

20

140

0.31 ± 0.02

2.67 ± 0.67

7.1

Pond

Raceway pond

2,400

800

700

0.34 ± 0.08

1.26 ± 0.11

5.4

opened when the pH of the culture was above the pH setpoint value of 6.7. Temperature was controlled at 28 ± 2 °C using an immersion chiller. The two ponds were operated for 5.4 days and achieved a volumetric productivity of 0.04 ± 0.01 g/L/day and an aerial productivity of 4.9 ± 2.0 g/m2/day, before being harvested. Biomass dewatering Photoautotrophically grown cells in the ponds (1,600 L) were harvested and transferred to a centrifuge feed tank. A stacked disc centrifuge (Alfa Laval MAPX207) was used to dewater and concentrate the cells. The centrifuge had 7.6 L of solids capacity and operated at 8009g at 24 ± 2 °C. Centrifuge discharge frequency was adjusted to obtain the desired biomass concentration for the heterotrophic growth stage. The concentrated biomass was collected and transferred rapidly to the bioreactors. Heterotrophic growth (Fed-batch operation) Heterotrophic growth of A. protothecoides was conducted in 10 L Biostat B fermentors (Sartorius, Bohemia, NY). The bioreactor was filled with 5 L of the concentrated cell slurry, and the pH was adjusted and set to 6.5, using 3 M NaOH. The cultivation temperature was controlled at 28 °C, and dissolved oxygen (DO) was set at C20 %. The aeration flow rate was kept at 4 L/min. Agitation was controlled automatically in response to the DO level. A proprietary organic antifoam DF 204 (BASF, Florham Park, NJ) was used for foam control as needed. A concentrated glucose or cassava starch hydrolysate solution (500–600 g glucose-equivalent/L) was fed manually into the bioreactor using a peristaltic pump to maintain the residual glucose concentration in the bioreactor between 10 and 30 g/L throughout the fermentation. Stock solutions were continuously stirred with a magnetic stirrer. Samples from the bioreactor were collected periodically for cell

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growth determination, extraction.

glucose

analysis,

and

lipid

Preparation of cassava starch hydrolysate Pure cassava starch was used for preparing the cassava starch hydrolysate as an alternative substrate to lab-grade glucose for A. protothecoides fermentation. The liquefaction and saccharification processes were done in a 50 L Letsch stainless steel reactor (Letsch Corp., Springfield, MO). For the liquefaction, the substrate slurry (20 % w/v on dry starch basis, DSB), was titrated to pH 5.7 with 6 N NaOH, and alpha amylase at 0.02 % w/w DSB (Spezyme Alpha, Genencor, Palo Alto, CA) was added to the slurry. The mixture was heated to 85 °C and held for 90 min under continuous agitation. The liquefied starch was then saccharified (pH 4.3 at 55 °C for 24 h) using glucoamylase at 0.05 % w/w DSB (Optidex L-400, Genencor, Palo Alto, CA). Samples of the mixture were analyzed by HPLC, as described below, to determine the glucoseequivalent sugar content of the mixture. The syrup was diluted to the desired glucose-equivalent concentration with water and sterilized by autoclaving before being used in the fermentation. Biomass measurement Cell growth was estimated by measuring absorbance at 750 nm (A750) or the dry cell weight (DCW, g/L). The DCW was determined by removing a sample that would provide at least 10 mg of DCW (estimated empirically). This sample was filtered through a pre-weighed 0.4 lm glass fiber filter (Whatman), rinsed three times with 5 mL ammonium bicarbonate solution (125 mM), and weighed wet. The sample was then put into a Shimadzu MOC 120H electronic moisture balance (Shimadzu Corp, Kyoto, Japan) and run until the dry weight was constant. The DCW was determined by correction for the weight of the filter and the volume of sample filtered.

Bioprocess Biosyst Eng

Glucose analysis For glucose analysis, the culture broth supernatant or cassava hydrolysate was diluted to the appropriate concentration using HPLC grade water and filtered through 0.45 lm cellulose-acetate filters (Whatman). Samples were analyzed on a Thermo Scientific HPLC system equipped with a Surveyor LC pump plus quaternary gradient pump, Surveyor Autosampler, Surveyor IR Plus detector, and ChromQuest 5.0 software (Thermo/Finnigan, Waltham, Massachusetts, USA). Separation was achieved on a 300 mm 9 7.8 mm, 9 lm particle size Aminex HPX-87H column (Bio Rad, Hercules, CA, USA). The column was held at 55 °C using a heating blanket. The mobile phase was 5 mM H2SO4 with a flow rate of 0.6 mL/min. Injection was achieved using a 20-lL injection loop. Glucose quantification was achieved through comparison to a glucose standard curve created from ultra-pure glucose (Sigma-Aldrich, St. Louis, MO) at concentrations ranging from 0.5 to 10 g/L glucose. The glucose calibration curve was generated by plotting the glucose chromatogram peak areas against the glucose concentration, and the linearity of the line was evaluated using least squares regression. Lipid extraction Lipid extraction was conducted using a modified Bligh and Dyer method [28]. Briefly, lyophilized cell pellet (50–300 mg) was mixed with a 1:2 chloroform/methanol solution in a 50-mL glass centrifuge tube and five 6.35 mm glass beads were added. The samples were vortexed for 1 h. After mixing, an additional aliquot of water was added to each sample to bring the ratio to 1:2:0.8 chloroform/ methanol/water and samples centrifuged at 560 9g for 10 min at room temperature using a Beckman G6B centrifuge. The chloroform layer was collected, and the solvent was removed by evaporation under N2. Extracted material was weighed using a high precision analytical balance (Sartorius ED124S) with 0.1 mg sensitivity to provide a gravimetric measurement of total lipid. Determination of fatty acid methyl esters (FAMES) Transesterification of the lipid extract was achieved by placing a lipid sample into a screw-capped glass test tube, adding 5.3 mL of methanol and 0.58 mL of 12 M sulfuric acid. Additionally, 1 mL of a solution containing 2.2 mg/ mL (final concentration of 0.74 mg/mL) nonadecanoic acid was added to each test tube as an internal standard. Samples were capped and placed in a dry heat bath at 75 °C for 90 min. Every 20 min the samples were removed and mixed vigorously for 20 s. Samples were cooled to room temperature and neutralized using KOH. Hexane was

added to extract the FAMEs from the methanol phase. The hexane phase was filtered through anhydrous sodium sulfate. FAME analysis was performed using a Trace GC Ultra Gas Chromatography system equipped with flame ionization detector, AS 3000 auto sampler, and ChromQuest 5.0 software (Thermo Fisher). The oven was equipped with a 30 m 9 0.32 mm 9 0.5 lm BPX-90 column (SGE Analytical Science, USA). The identity of each FAME peak was determined by comparing the retention times of the compounds to the retention time of FAME standards (Sigma-Aldrich, St. Louis, MO). The internal standard (nonadecanoic methyl ester) method was used to quantify each FAME peak by relative area. All analytical measurements for biomass, glucose, lipid and FAME were performed in duplicate and results are presented as Mean ± SD. Nile red staining and microscopy Cells were stained with Nile Red dye and imaged using a fluorescent microscope. Nile Red stain is a neutral oil stain that permeates cell membrane and targets triglycerides, or neutral oils [29]. Microscopy was performed with a Nikon Eclipse E200 light microscope equipped with an Amscope MD900 digital camera. The dye was excited with a Mercury lamp and measured with an absorbance and emission filter sets of 450–490 and 500–515 nm, respectively.

Results and discussion A mixed trophic state cultivation strategy for mass culturing and lipid production by A. protothecoides was designed and tested at subpilot scale. The process, referred to here as the HTB process, consists of a photoautotrophic open pond cultivation of algae that is followed by concentration and a heterotrophic stage (Fig. 1). The advantages anticipated in such an approach are the spatial and temporal separation of inexpensive mass culture of algae on a continuous basis (the photoautotrophic stage) and high productivity lipid accumulation in a more concentrated and constrained system (the heterotrophic bioreactor). The HTB process uses the photoautotrophic step for mass culture in open ponds, which can be run in a continuous manner, and then provides a concentrated algal suspension that is fed with fixed carbon (e.g., glucose) to induce rapid lipid accumulation. The inoculum is produced in aseptic culture then aseptically and sequentially transferred to progressively larger photoautotrophic culture systems until it reaches the carboy stage where asepsis is no longer maintained (Fig. 1). The carboy is transferred to open ponds where the bulk of the photoautotrophically produced biomass is generated.

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Photoautotrophic growth A representative photoautotrophic growth profile of A. protothecoides in open raceway ponds is shown in Fig. 2a, and key performance indicators of the process are listed in Table 2. Photoautotrophic cultivation of A. protothecoides in open ponds resulted in a final biomass concentration of 0.3 ± 0.1 g/L and an aerial biomass productivity of 4.9 ± 2.0 g/m2/day (Table 2). During the photoautotrophic growth in open ponds, A. protothecoides biomass

Fig. 2 Photoautotrophic and heterotrophic growth of A. protothecoides KRT1009. a Representative growth of A. protothecoides KRT1009 in an open 800 L raceway pond with 15 cm depth and 2.6 m2 culture area. Pond bacterial contamination was 120 CFU/mL, and areal productivity was 7.3 g/m2/day. b Representative growth and lipid production by A. protothecoides KRT1009 using glucose as the source of substrate in a 10 L bioreactor. The bioreactor was seeded with photoautotrophically grown cells (40 g/L) concentrated by centrifugation from open ponds. Bioreactor initial volume was 5 L, and temperature, pH and dissolved oxygen were controlled at 28 °C, 6.5 and 22 %, respectively. The bioreactors were run as a fed-batch for 75 h

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concentration reached 0.3 ± 0.1 g/L, with the lipid content of 17.7 ± 2.3 %, on the basis of dry cell weight, and the average volumetric lipid productivity of 0.4 ± 0.1 mg/L/h. At the end of the photoautotrophic stage, algal biomass from the ponds was harvested by centrifugation to obtain a highly concentrated seed for the heterotrophic phase of the process. Heterotrophic growth During the heterotrophic stage of the HTB process, concentrated biomass of photoautotrophically grown A. protothecoides was added as inoculum to 10 L bioreactors and grown under fed-batch conditions using glucose as the sole fixed carbon substrate. A representative growth profile for photoautotrophically cultivated A. protothecoides grown on glucose in the bioreactor is shown in Fig. 2b, and key performance indicators of the process are listed in Table 3. Under heterotrophic growth conditions, cells changed color from green to yellow through degradation of chlorophyll and produced on average 116.7 ± 15.5 DCW/L biomass containing over 60 % lipid (Table 2). The accumulated lipids were deposited as intracellular lipid bodies, which could easily be detected by a fluorescent probe, Nile red (Fig. 3). The volumetric rate of lipid production during the heterotrophic stage was 0.7 ± 0.2 g/L/h, which was more than three orders of magnitude higher than that achieved during photoautotrophic cultivation stage. Similarly, the biomass productivity in the heterotrophic stage of the process was 600-fold higher than that achieved under photoautotrophic conditions (Table 2). These significantly higher biomass and lipid productivities during the heterotrophic stage of the process lend the HTB process significant performance advantages over growing oleaginous algae solely photoautotrophically. The cumulative biomass and lipid productivities of the HTB process are presented in Table 2 and were over 250 and 600 times higher, respectively, than those obtained from the photoautotrophic cultivation stage. The concept of using a mixed trophic state system for production of biofuels was described in a recent patent [19]. The methods described in the Oyler patent were poorly defined and no data were provided with any algal strain. Subsequent patent applications by Sayre [30] and Wu and Xiong [21] provided more detail on a mixed trophic state production process. The process described by Sayre provided data using a mixed trophic growth system at shake flask scale where the cells were grown aseptically photosynthetically, concentrated, then grown heterotrophically on glycerol [30]; no scaled up data were provided. The process of Wu and Xiong [21] described shake flask and small fermentor scale mixed trophic growth of A. protothecoides. The phototrophic growth was done in the

Bioprocess Biosyst Eng Table 2 Key performance indicators of the individual photoautotrophic and heterotrophic stages, and the entire HTB process Process stage Photoautotrophic

Biomass concentration (g/L) 0.3 ± 0.1

Biomass productivity (g/L/h)

Areal productivity (g/m2/d)

Lipid content (% DCW)

Lipid productivity (g/L/h)

Lipid yield (g/g glucose)

2.0 ± 0.8 9 10-3

4.9 ± 2.0

17.7 ± 2.3

0.4 ± 0.1 9 10-3

na

Heterotrophic

116.7 ± 15.5

1.2 ± 0.3

na

60.1 ± 6.5

0.7 ± 0.2

0.21 ± 0.03

HTB (cumulative)

116 ± 15.5

0.5 ± 0.1

na

60.1 ± 6.5

0.27 ± 0.07

0.21 ± 0.03

Results represent Mean ± SD of at least 16 subpilot-scale HTB process runs

Table 3 Effect of different initial biomass concentrations on key performance indicators of the heterotrophic stage of the HTB process

Initial biomass concentration (g/L)

Final biomass concentration (g/L)

Biomass productivity (DCW/L/h)

Biomass yield (DCW/ g glucose)

Lipid content (% DCW)

Lipid productivity (g/L/h)

Lipid yield (g/g glucose)

12.4

94.7

0.81

0.32

63.1

0.60

0.18

25.7

121.9

1.09

0.46

56.0

0.65

0.24

47.3

136.9

1.1

0.45

61.7

0.83

0.26

Fig. 3 Microscopic images of A. protothecoides harvested at the end of the HTB process. The micrograph shows intercellular lipid bodies stained with Nile Red, a fluorescent dye, indicative of significant lipid accumulation by A. protothecoides during the HTB process

presence of glycine (optimal at 5 g/L); cells were settled for 12 h and then centrifuged prior to growth in a 5 L bioreactor. This second procedure most closely resembles that described in the current study. However, the current study provides an open photosynthetic stage cultivation at greater than 1,000 L scale; no amino acids were provided during the raceway photosynthetic growth, cells were collected by centrifugation, and fermentation was done in 15 L bioreactors at high cell density. Another recent study describes a mixed trophic system using A. protothecoides where the photosynthetic stage of the process used the carbon dioxide generated by the fermentation [31]. This is a very promising approach but needs to be further scaled before its findings will have relevance to the production of biofuel. Issues surrounding the flow rate of the off gases needed to maintain the bioreactor at sufficient oxygen content need to be addressed. The HTB grown A. protothecoides cells had a different fatty acid composition than that possessed by the photoautotrophically grown cells (Fig. 4). The lipid extracted

from cells harvested at the end of HTB consisted of mainly oleic (C18:1, 67 %), linoleic (C18:1, 15 %), palmitic (C16:0, 10 %), and stearic acids (C18:0, 4 %). However, the lipid from cells grown photoautotrophically in open ponds (before inoculation into the bioreactors) was characterized by lower oleic acid (C18:1, 14.2 %) and higher contents of arachidonic acid (C20:4, 41.3 %), linoleic acids (C18:2, 35.9 %), and palmitic acids (C16:0, 28 %). The methyl esters of monounsaturated fatty acids (e.g., oleic acid) are considered to be better feedstock for biodiesel production than polyunsaturated ones (e.g, arachidonic acid) due to having higher octane number and iodine value without adverse effects on biodiesel cold temperature properties [32]. These results demonstrated that heterotrophic cultivation of A. protothecoides changes the fatty acid profile of cells toward a composition (*82 % monounsaturated fatty acids) that is more suitable for biodiesel production. Under optimal growth conditions, microalgae synthesize fatty acids principally for esterification into glycerol-based

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Bioprocess Biosyst Eng Fig. 4 Total content and fatty acid composition of lipids extracted from A. protothecoides cells harvested at the end of photoautotrophic stage compared to cells that have gone through the heterotrophic stage of the HTB process

membrane lipids, which constitute about 5–20 % of their dry cell weight (DCW). When exposed to stress conditions or under unfavorable growth environments, oleaginous microalgae change their lipid biosynthetic pathways toward the formation and accumulation of large quantities of neutral lipids (20–50 % DCW), mainly in the form of triacylglycerides (TAG) [33]. Synthesis and accumulation of TAGs under stress conditions is accompanied by considerable changes in composition of lipids and fatty acids. The most commonly used stress condition to induce lipid accumulation in oleaginous microalgae is nitrogen starvation which is also referred to as a high C/N ratio. During the heterotrophic stage of the HTB process, the only nutrient fed to the bioreactors is glucose, and the amount of nitrogen carried over from the ponds via the cell slurry was minimal (data not shown), resulting in high C/N ratios. Subsequently, nitrogen starvation or some other metabolic factor brought on by a trophic shift resulted in a significant increase in the synthesis of lipids and fatty acids. A fed-batch bioreactor strategy has been previously used successfully to achieve high cell density cultures of A. protothecoides [34]. During the conventional algal fed batch fermentation, maximum logarithmic growth rate ((l) = ln(X2/X1)
(t2-t1)-1?D) typically occurs through the first 75–80 h post inoculation. During this time, the oil volumetric productivity (Qp = g oil
(L
h)-1) is very low. Due to nutritional limitations, the culture growth slows and shifts from logarithmic to a more linear growth (linear growth rate = (X2-X1)
(t2-t1)-1?D), where there is a measured increase in Qp. During this stage, a metabolic overflow of intracellular carbon intermediates increases. This growth condition results in a rapid increase in the fatty acid production rate. In contrast, the standard logarithmic growth rate of normal fermentations does not occur in the HTB

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process because of the high cell densities at inoculation. Therefore, the metabolic overflow condition is occurring much earlier thereby more rapidly moving the HTB process toward oil production. This earlier start for oil production results in higher cumulative oil volumetric productivities. The percentage of oil in the algal biomass typically increases from *17 % (DCW) in the pond (photosynthetically generated) to about 60 % (DCW) following the heterotrophic step. The accumulated lipids during the heterotrophic stage were predominantly non-polar (data not shown) and were largely free of chlorophyll and carotenoids. This is due to a degreening (i.e., bleaching) that occurs during the heterotrophic step of the process when the chloroplasts with their associated pigments are degraded [35]. The reduced pigment composition in the extracted lipid is a significant advantage for the downstream oil recovery process as it can be simplified and less expensive. Preliminary results (data not shown) suggest a gentle cell disruption combined with an aqueous extraction process reduces the need for further refining of the extracted oil. This compares favorably to oil extracted from algae grown exclusively under photosynthetic conditions that has less lipid and more pigment, which mandates extensive extraction methods based on solvent extraction of the lipid and further distillation/purification (huge cost drivers). As a result, the HTB process will require an oil recovery strategy that is a low energy and cost effective recovery process, yielding a very stable product with minimum need for secondary purification with its associated cost. In the HTB process described here, dewatering of algal biomass generated in the pond was performed by centrifugation, assuring viable and consistent starting algal biomass for the fermentation in the bioreactor. The supernatant resulting from the centrifugation, once supplemented with

Bioprocess Biosyst Eng

nutrients and cleared from potential growth inhibitors, can be recycled back to the ponds for many cycles without additional water treatment, resulting in significant reduction in water treatment cost. While this has not been done yet at large scale, small-scale recycling of spent medium using 6.5 L aquaria was used to demonstrate this ability. Starting with fresh medium, cells were grown and then collected by centrifugation and the spent medium reused in the same system for growth of A. protothecoides. These experiments were performed in triplicate and compared to aquaria provided only with fresh medium. Using three cycles of growth and reuse, there was no significant difference in the recycled medium growth rate (8.0 g/m2/d) and that using the fresh medium (7.9 g/m2/d). Microbial contamination of algal biomass is inevitable when the culture is grown in non-aseptic conditions, such as raceways and open ponds. The expectation is that bacterial contamination will be transferred from the open photosynthetic culture to the bioreactor by way of the concentrated inoculum. It was a pleasant surprise that in the bioreactors the A. protothecoides dominated and productivity was not negatively impacted by bacterial load. The bacteria are still present and had an average daily doubling time of 0.62 (STD 0.01; % CV 1.3 %), while the A. protothecoides had a doubling time of 1.35 (STD 0.11; % CV 8.15 %). In the scaled up process outlined in this study, on harvest from the bioreactor, the contaminating organisms present in the broth have been very low, to the extent that no significant differences in key performance indicators were observed when results were compared with those obtained from bioreactors seeded with axenic cultures (data not shown). These results are consistent with reports of a recent study demonstrating that A. protothecoides is able to grow well in non-sterilized wastewater with no effects of bacterial contamination on its growth rate [36]. An interesting observation is that when this process was run with Chlorella vulgaris the bioreactor was overrun with bacteria that crashed the culture. Effect of initial biomass concentration (i.e, inoculum size) on fermentation Lipid production by A. protothecoides during the heterotrophic stage of the HTB process was carried out with different inoculum sizes to determine the effect of initial biomass concentration on key performance indicators of the fermentation process. The initial inocula sizes evaluated were 12, 25, and 47 g/L at the starting volume of 5 L in 10 L bioreactors. Figure 5 follows the changes in biomass concentration and lipid content during the course of 120 h fermentation at various inoculum sizes, and the key performance indicators of the fermentation are summarized in Table 3. The final biomass concentration increased with

Fig. 5 Effect of different initial biomass concentrations on biomass and lipid production by A. protothecoides KRT 1009 during the heterotrophic stage of the HTB process in a 10-L bioreactor. The bioreactors were seeded with photoautotrophically grown cells (12.4, 25.7, and 47.3 g/L) concentrated by centrifugation from open ponds. Bioreactor was run as fed-batch with the glucose concentration maintained between 10 and 30 g/L. Bioreactor initial volume was 5 L, and temperature, pH and dissolved oxygen were controlled at 28 °C, 6.5 and 22 %, respectively. The bioreactors were run as a fedbatch for 120 h

increase in the initial inoculum size. The increase in biomass concentration corresponded to higher biomass productivities and higher yield of biomass on glucose at larger inoculum sizes. The 25 and 47 g/L inoculum sizes resulted in similar biomass productivities (1.09 vs. 1.1 g/L/h) and biomass yields (0.46 vs. 0.45 g/g), which were higher than the values achieved with the 12 g/L inoculum size, 0.81 g/ L/h and 0.32 g/g, respectively (Table 3). The late (*80 h) decline in biomass concentration from the lowest inoculum size tested (i.e. 12 g/L) could be due to competition for nutrients with contaminating bacteria. This, however, was not verified experimentally. The inoculum size affected the lipid production, especially in terms of lipid productivity and lipid yield. Lipid productivity and lipid yield increased considerably with an increase in inoculum size, where the highest values achieved for these parameters were 0.83 (g/ L/h) and 0.26 (g/g), respectively, by the 47 g/L inoculum size (Table 3). The highest lipid content (63 % DCW) was obtained by 12 g/L inoculum size, whereas the 25 g/L inoculum resulted in the least lipid accumulation (56 % DCW) by the cells. Lipid production by oleaginous organisms (e.g., A. protothecoides) via HTB growth must have the primary objective of maximizing the lipid yield on the substrate (i.e., glucose), because the cost of the carbon source is the

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Fig. 6 Biomass and lipid production by A. protothecoides KRT 1009 during the heterotrophic stage of the HTB process in a 10-L bioreactor, comparing cassava sugar as an alternative substrate to glucose. The bioreactors were seeded with photoautotrophically grown cells (23 g/L) concentrated from open ponds. Bioreactor initial volume was 5 L, and temperature, pH and dissolve oxygen were controlled at 28 °C, 6.5 and 22 %, respectively. Bioreactors were run for 95 h as fed-batch with the glucose concentration maintained between 10 and 30 g/L

dominant production cost. In addition, the process productivity is an important parameter in that it corresponds to effective equipment utilization. Here the results show that increasing the inoculum size from 12 to 47 g/L significantly improved the productivity and yield of lipid production when photoautotrophically grown A. protothecoides cells are used as the seed culture in the bioreactors.

starch hydrolysate with glucose-equivalent concentration of 650 g/L was fed-batched into the bioreactor. A control bioreactor was operated in parallel using glucose as the substrate. Bioreactor operating conditions were the same as described above. The results on fermentation of cassava starch hydrolysate in comparison with that of glucose during the heterotrophic stage of the HTB process are shown in Fig. 6, and key performance indicators are presented in Table 4. Biomass growth with cassava sugar followed the same profile as that with glucose, resulting in similar biomass productivity of 1.16 vs. 1.17 g/L/h, respectively, and the identical biomass yield of 0.34 g DCW/g glucose, after 95 h of heterotrophic growth (Table 4). Lipid production was also very similar for the two substrates, though the final lipid content and yield on substrate were slightly higher with glucose (Table 4). In addition, relatively higher lipid productivities were achieved with glucose (0.61 g/L/h) than with cassava hydrolysate (0.55 g/L/h). These results demonstrated that A. protothecoides could produce almost the same amount of oil when cultivated with the cassava starch hydrolysate as when grown in the presence of glucose. These findings are consistent with previous reports on production of lipids by oleaginous microalgae using cassava starch hydrolysate as the organic substrate [37]. Due to high fermentable carbohydrate yield, low cost of production and drought resistance, cassava represents an alternative source of substrate for production of biofuels and value-added chemicals [38]. Our results suggest that the use of cassava as feedstock for heterotrophic growth of oleaginous microalgae could be a costeffective way for producing biofuel precursors in regions where cassava is abundant.

Conclusions Cassava sugar as an alternative carbon source Lipid production from cassava starch hydrolysate by the HTB process was evaluated at a subpilot plant scale. Pure cassava starch was hydrolyzed using a low-temperature dual enzyme process at ambient pressure. Photoautotrophically grown A. protothecoides, concentrated as described above with the initial concentration of 23 g/L, was used as the seed in the bioreactor. A concentrated cassava Table 4 Key performance indicators of the heterotrophic stage of the HTB process with cassava hydrolysate in comparison with glucose as the sources of substrates

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This study evaluates a two-stage photoautotrophic-to-heterotrophic process for high-productivity algal lipid as feedstock for biofuel production at subpilot-scale using Auxenochlorella protothecoides. The process uses photoautotrophically grown algal biomass as a high-density inoculum for an aerobic bioreactor focused on lipid accumulation using fixed carbon substrates. The process achieves high volumetric lipid productivity (0.8 g/L/h),

Substrate

Final biomass concentration (DCW/L)

Biomass productivity (DCW/L/h)

Biomass yield (DCW/g glucose)

Lipid content (% DCW)

Lipid productivity (g/L/h)

Lipid yield (g/g glucose)

Cassava sugar

110.6

1.16

0.34

48.3

0.55

0.18

Glucose

111.6

1.17

0.34

53.1

0.61

0.19

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lipid contents (63 % w/DCW), and yield (yp/s 0.26), which compares favorably to conventional heterotrophic algae growth (yp/s 0.10–0.15). The process reduces overall cost and improves oil quality compared to the strictly photoautotrophic process. Future efforts must focus on optimizing lipid yields, improving productivity, and reducing feedstock cost.

14.

15.

16. Acknowledgments This research project was supported under the Department of Energy grant DE-FE-0000888 awarded by National Energy Technology Laboratory. Additional support was supplied by a large group of excellent support staff at Phycal in the R&D group who worked tirelessly to help enable this process.

17.

Conflict of interest All authors were former employees of Phycal Inc. and paid in part from a grant supplied by the US Department of Energy. 18.

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