Mixed Trophic State Production Process for Microalgal Biomass with High Lipid Content for Generating...

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Bioprocess and BiosystemsEngineering ISSN 1615-7591 Bioprocess Biosyst EngDOI 10.1007/s00449-014-1303-5

High-productivity lipid productionusing mixed trophic state cultivationof Auxenochlorella (Chlorella)protothecoides

Hamid Rismani-Yazdi, KristinH. Hampel, Christopher D. Lane, BenA. Kessler, Nicholas M. White, KennethM. Moats & F. C. Thomas Allnutt

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ORIGINAL PAPER

High-productivity lipid production using mixed trophic statecultivation 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

� 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

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 predomi-

nantly 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

Introduction

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 com-

mercialization [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

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

K. H. Hampel

e-mail: [email protected]

C. D. Lane


B. A. Kessler


N. M. White


K. M. Moats




Present Address:

H. Rismani-Yazdi (&)

Novozymes North America Inc., P.O. Box 576, Franklinton,

NC 27525, USA


Present Address:


BrioBiotech LLC, P.O. Box 26, Glenelg, MD 21737, USA

123

Bioprocess Biosyst Eng

DOI 10.1007/s00449-014-1303-5

Author's personal copy

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 sur-

prising number are capable of heterotrophic or mixotrophic

growth [9–13]. This led to efforts to look at the hetero-

trophic or mixotrophic production of biofuels from algae

[10]. Production of biomass through heterotrophy, mixo-

trophy, or using mixed trophic states provides an oppor-

tunity for improved productivity but requires an

inexpensive source of fixed carbon usable by algae (e.g.,

hydrolysates or partially purified sugars) to be commer-

cially 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 drop-

in 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 pro-

posed 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, photo-

autotrophic 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,

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 het-

erotrophic 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 syn-

thesis. The traditional photoautotrophic approach is limit-

ing 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 (Chlo-

rella) protothecoides to grow both photoautotrophically

and heterotrophically, when combined with its high pho-

tosynthetic efficiency of 8 % [22] and ability to accumulate

large quantities of neutral lipids (up to 70 %) [23] at rel-

atively 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 Chloro-

phyta, 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


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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 flat-

bottom 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, Van-

couver, 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 car-

boy 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 Sun-

TubeTM 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

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


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opened when the pH of the culture was above the pH set-

point 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 produc-

tivity 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 trans-

ferred 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 con-

centrated 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

growth determination, glucose analysis, and lipid

extraction.

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 lique-

faction 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 glucoamy-

lase 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 glucose-

equivalent 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.

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


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Glucose analysis

For glucose analysis, the culture broth supernatant or cas-

sava hydrolysate was diluted to the appropriate concen-

tration 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. Injec-

tion was achieved using a 20-lL injection loop. Glucose

quantification was achieved through comparison to a glu-

cose 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 cen-

trifuge. The chloroform layer was collected, and the sol-

vent 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. Sam-

ples 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 sul-

fate. FAME analysis was performed using a Trace GC

Ultra Gas Chromatography system equipped with flame

ionization detector, AS 3000 auto sampler, and Chrom-

Quest 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 pre-

sented 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 Mer-

cury 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 cul-

turing 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 con-

centration and a heterotrophic stage (Fig. 1). The advan-

tages 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 cul-

ture 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

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, con-

centrated biomass of photoautotrophically grown A. prot-

othecoides 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 hetero-

trophic stage of the process lend the HTB process signifi-

cant 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, respec-

tively, than those obtained from the photoautotrophic cul-

tivation 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 tro-

phic 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 heterotro-

phically 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

Fig. 2 Photoautotrophic and heterotrophic growth of A. prototheco-

ides 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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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 photo-

autotrophically 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 char-

acterized 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 heterotro-

phic cultivation of A. protothecoides changes the fatty acid

profile of cells toward a composition (*82 % monoun-

saturated fatty acids) that is more suitable for biodiesel

production.

Under optimal growth conditions, microalgae synthesize

fatty acids principally for esterification into glycerol-based

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

Table 2 Key performance indicators of the individual photoautotrophic and heterotrophic stages, and the entire HTB process

Process stage Biomass

concentration (g/L)

Biomass

productivity (g/L/h)

Areal productivity

(g/m2/d)

Lipid content (%

DCW)

Lipid productivity

(g/L/h)

Lipid yield (g/g

glucose)

Photoautotrophic 0.3 ± 0.1 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


123


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 con-

siderable changes in composition of lipids and fatty acids.

The most commonly used stress condition to induce lipid

accumulation in oleaginous microalgae is nitrogen starva-

tion 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

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 (photosyn-

thetically generated) to about 60 % (DCW) following the

heterotrophic step. The accumulated lipids during the het-

erotrophic stage were predominantly non-polar (data not

shown) and were largely free of chlorophyll and carote-

noids. 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 degra-

ded [35]. The reduced pigment composition in the extrac-

ted 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 strat-

egy that is a low energy and cost effective recovery pro-

cess, 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 centrifu-

gation, assuring viable and consistent starting algal biomass

for the fermentation in the bioreactor. The supernatant

resulting from the centrifugation, once supplemented with

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


123


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 pro-

vided 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 bac-

terial 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 pro-

ductivity 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 cul-

tures (data not shown). These results are consistent with

reports of a recent study demonstrating that A. prototh-

ecoides 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 hetero-

trophic 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 evalu-

ated were 12, 25, and 47 g/L at the starting volume of 5 L

in 10 L bioreactors. Figure 5 follows the changes in bio-

mass 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

increase in the initial inoculum size. The increase in bio-

mass concentration corresponded to higher biomass pro-

ductivities 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

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 fed-

batch for 120 h


123


dominant production cost. In addition, the process pro-

ductivity 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 signifi-

cantly improved the productivity and yield of lipid pro-

duction when photoautotrophically grown A.

protothecoides cells are used as the seed culture in the

bioreactors.

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. Photoautotro-

phically grown A. protothecoides, concentrated as descri-

bed above with the initial concentration of 23 g/L, was

used as the seed in the bioreactor. A concentrated cassava

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 hydroly-

sate in comparison with that of glucose during the het-

erotrophic 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 repre-

sents 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 hetero-

trophic growth of oleaginous microalgae could be a cost-

effective way for producing biofuel precursors in regions

where cassava is abundant.

Conclusions

This study evaluates a two-stage photoautotrophic-to-het-

erotrophic process for high-productivity algal lipid as

feedstock for biofuel production at subpilot-scale using

Auxenochlorella protothecoides. The process uses photo-

autotrophically grown algal biomass as a high-density

inoculum for an aerobic bioreactor focused on lipid accu-

mulation using fixed carbon substrates. The process

achieves high volumetric lipid productivity (0.8 g/L/h),

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

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

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


123


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 photo-

autotrophic process. Future efforts must focus on opti-

mizing lipid yields, improving productivity, and reducing

feedstock cost.

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.

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.

References

1. Heinrich JM, Niizawa I, Botta FA, Trombert AR, Irazoqui HA

(2012) Analysis and design of photobioreactors for microalgae

production II: experimental validation of a radiation field simu-

lator based on a Monte Carlo algorithm. Photochem Photobiol

88(4):952–960. doi:10.1111/j.1751-1097.2012.01149.x

2. Xu L, Guo C, Wang F, Zheng S, Liu C-Z (2011) A simple and

rapid harvesting method for microalgae by in situ magnetic

separation. Bioresour Technol 102(21):10047–10051. doi:10.

1016/j.biortech.2011.08.021

3. Halim R, Danquah MK, Webley PA (2012) Extraction of oil from

microalgae for biodiesel production: a review. Biotechnol

Advances 30(3):709–732. doi:10.1016/j.biotechadv.2012.01.001

4. Mercer P, Armenta R (2011) Developments in oil extraction from

microalgae. Eur J Lipid Sci Technol. doi:10.1002/ejlt.201000

455

5. Faeth J, Valdez P, Savage P (2013) Fast hydrothermal liquefac-

tion of Nannochloropis sp. to produce biocrude. Energy Fuels

27:1391–1398

6. Onwudili JA, Lea-Langton AR, Ross AB, Williams PT (2013)

Catalytic hydrothermal gasification of algae for hydrogen pro-

duction: composition of reaction products and potential for

nutrient recycling. Bioresour Technol 127:72–80. doi:10.1016/j.

biortech.2012.10.020

7. Rampton R, Zabarenko D (2012) Algae biofuel not sustainable

now, review says. msnbccom

8. Leite GB, Abdelaziz AEM, Hallenbeck PC (2013) Algal biofuels:

challenges and opportunities. Bioresour Technol 145:134–141.

doi:10.1016/j.biortech.2013.02.007

9. Samejima H, Myers J (1958) On the heterotrophic growth of

Chlorella pyrenoidosa. J Gen Microbiol 18(1):107–117

10. Miao X, Wu Q (2006) Biodiesel production from heterotrophic

microalgal oil. Bioresour Technol 97(6):841–846. doi:10.1016/j.

biortech.2005.04.008

11. Azam F, Hemmingsen BB, Volcani BE (1974) Role of silicon in

diatom metabolism. V. Silicic acid transport and metabolism in

the heterotrophic diatom Nitzschia alba. Arch Microbiol

97(2):103–114

12. Lewin J, Hellebust JA (1975) Heterotrophic nutrition of the

marine pennate diatom Navicula pavillardi Hustedt. Can J

Microbiol 21(9):1335–1342

13. Chojnacka K, Noworyta A (2004) Evaluation of Spirulina sp.

growth in photoautotrophic, heterotrophic and mixotrophic

cultures. Enzyme Microb Technol 34(5):461–465. doi:10.1016/j.

enzmictec.2003.12.002

14. Fabregas J, Garcia D, Lamela T, Morales ED, Otero A (1999)

Mixotrophic production of phycoerythrin and exopolysaccharide

by the microalga Porphyridium cruentum. Cryptogamie Algolo-

gie 20(2):89–94

15. Kamjunke N, Tittel J (2009) Mixotrophic algae constrain the loss

of organic carbon by exudation. J Phycol 45(4):807–811. doi:10.

1111/j.1529-8817.2009.00707.x

16. Sforza E, Cipriani R, Morosinotto T, Bertucco A, Giacometti GM

(2012) Excess CO2 supply inhibits mixotrophic growth of

Chlorella protothecoides and Nannochloropsis salina. Bioresour

Technol 104:523–529. doi:10.1016/j.biortech.2011.10.025

17. Hu B, Min M, Zhou W, Du Z, Mohr M, Chen P, Zhu J, Cheng Y,

Liu Y, Ruan R (2012) Enhanced mixotrophic growth of mi-

croalga Chlorella sp. on pretreated swine manure for simulta-

neous biofuel feedstock production and nutrient removal.

Bioresour Technol 126C:71–79. doi:10.1016/j.biortech.2012.09.

031

18. Ogbonna JC, Masui H, Tanaka H (1997) Sequential hetero-

trophic/autotrophic cultivation—an efficient method of producing

Chlorella biomass for health food and animal feed. J Appl Phycol

9:359–366

19. Oyler JR (2008) Two-stage process for producing oil from mic-

roalgae. USA Patent, 8,475,543

20. Sayre R, Pereira S (2010) Molecuar approaches for the optimi-

zation of biofuel productions. USA Patent, 20100317073

21. Wu Q, Xiong W (2009) Method for producing biodiesel from an

alga. USA Patent, 20090298159

22. Krohn BJ, McNeff CV, Yan B, Nowlan D (2011) Production of

algae-based biodiesel using the continuous catalytic Mcgyan

process. Bioresour Technol 102(1):94–100. doi:10.1016/j.bior

tech.2010.05.035

23. Heredia-Arroyo T, Wei W, Hu B (2010) Oil accumulation via

heterotrophic/mixotrophic Chlorella protothecoides. Appl Bio-

chem Biotechnol 162(7):1978–1995. doi:10.1007/s12010-010-

8974-4

24. Xu H, Miao X, Wu Q (2006) High quality biodiesel production

from a microalga Chlorella protothecoides by heterotrophic

growth in fermenters. J Biotechnol 126(4):499–507. doi:10.1016/

j.jbiotec.2006.05.002

25. Huss V, Frank C, Hartmann E, Hirmer M, Kloboucek A, Seidel

B, Wenzeler P, Kessler E (1999) Biochemical taxonomy and

molecular phylogeny of the genus Chlorella sensu lato (Chloro-

phyta). J Phycol 35:587–598

26. Miao X, Wu Q (2004) High yield bio-oil production from fast

pyrolysis by metabolic controlling of Chlorella protothecoides.

J Biotechnol 110(1):85–93. doi:10.1016/j.jbiotec.2004.01.013

27. Konieczny A, Ausubel FM (1993) A procedure for mapping

Arabidopsis mutations using co-dominant ecotype-specific PCR-

based markers. Plant J 4(2):403–410

28. Bligh EG, Dyer WJ (1959) A rapid method of total lipid

extraction and purification. Can J Biochem Physiol

37(8):911–917

29. Greenspan P, Mayer EP, Fowler SD (1985) Nile red: a selective

fluorescent stain for intracellular lipid droplets. J Cell Biol

100(3):965–973

30. Sayre RT (2009) Optimization of biofuel production. USA Patent

US20090181438

31. Santos CA, Nobre B, Lopes da Silva T, Pinheiro HM, Reis A

(2014) Dual-mode cultivation of Chlorella protothecoides

applying inter-reactors gas transfer improves microalgae biodie-

sel production. J Biotechnol 184:74–83. doi:10.1016/j.jbiotec.

2014.05.012

32. Ramos MJ, Fernandez CM, Casas A, Rodriguez L, Perez A

(2009) Influence of fatty acid composition of raw materials on


123


http://dx.doi.org/10.1111/j.1751-1097.2012.01149.x

http://dx.doi.org/10.1016/j.biortech.2011.08.021


http://dx.doi.org/10.1016/j.biotechadv.2012.01.001

http://dx.doi.org/10.1002/ejlt.201000455

http://dx.doi.org/10.1002/ejlt.201000455






http://dx.doi.org/10.1016/j.enzmictec.2003.12.002

http://dx.doi.org/10.1016/j.enzmictec.2003.12.002

http://dx.doi.org/10.1111/j.1529-8817.2009.00707.x

http://dx.doi.org/10.1111/j.1529-8817.2009.00707.x






http://dx.doi.org/10.1007/s12010-010-8974-4

http://dx.doi.org/10.1007/s12010-010-8974-4

http://dx.doi.org/10.1016/j.jbiotec.2006.05.002





biodiesel properties. Bioresour Technol 100(1):261–268. doi:10.

1016/j.biortech.2008.06.039

33. Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert

M, Darzins A (2008) Microalgal triacylglycerols as feedstocks for

biofuel production: perspectives and advances. Plant J

54(4):621–639. doi:10.1111/j.1365-313X.2008.03492.x

34. Xiong W, Li X, Xiang J, Wu Q (2008) High-density fermentation

of microalga Chlorella protothecoides in bioreactor for microbio-

diesel production. Appl Microbiol Biotechnol 78(1):29–36.

doi:10.1007/s00253-007-1285-1

35. Hortensteiner S, Chinner J, Matile P, Thomas H, Donnison IS

(2000) Chlorophyll breakdown in Chlorella protothecoides:

characterization of degreening and cloning of degreening-related

genes. Plant Mol Biol 42(3):439–450

36. Ramos Tercero EA, Sforza E, Morandini M, Bertucco A (2014)

Cultivation of Chlorella protothecoides with urban wastewater in

continuous photobioreactor: biomass productivity and nutrient

removal. Appl Biochem Biotechnol 172(3):1470–1485. doi:10.

1007/s12010-013-0629-9

37. Lu Y, Zhai Y, Liu M, Wu Q (2010) Biodiesel production from

algal oil using cassava (Manihot esculenta Crantz) as feedstock.

J Appl Phycol 22(5):573–578. doi:10.1007/s10811-009-9496-8

38. Shetty J, Chotani G, Gand D, Bates D (2007) Cassava as an

alternative feedstock in the production of renewable transporta-

tion fuels. Intl Sugar J 109(1307):663–677


123




http://dx.doi.org/10.1111/j.1365-313X.2008.03492.x

http://dx.doi.org/10.1007/s00253-007-1285-1

http://dx.doi.org/10.1007/s12010-013-0629-9

http://dx.doi.org/10.1007/s12010-013-0629-9

http://dx.doi.org/10.1007/s10811-009-9496-8

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