Biotechnology Reports 25 (2020) e00392

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Biotechnology Reports
journal homepage: www.elsevier.com/locate/btre

Coproduction of lipids and extracellular polysaccharides fromthe novel green alga Parachlorella sp. BX1.5 depending on
cultivation conditions
Mitsuki Sasakia , Akari Takagia , Shuhei Otab,d , Shigeyuki Kawanob,e , Daisaku Sasakic,
Munehiko Asayamaa,f,*aCollege of Agriculture, Ibaraki University, 3-21-1 Ami, Ibaraki, 300-0393, Japan
Department of Integrated Sciences, Graduate School of Frontier Sciences, University of Tokyo, 5-1-5 Kashiwanoha, Kashiwa, Chiba, 277-0882, Japanc
BioX Chemical Industries Co. Ltd., 2-20-11 Inokuchidai, Nishi-ku, Hiroshima 733-0844, JapandCenter for Environmental Biology and Ecosystem Studies, National Institute for Environmental Studies, Tsukuba, Ibaraki, 305-8506, Japane
Future Center Initiative, The University of Tokyo, 178-4-4 Wakashiba, Kashiwa, Chiba, 277-0871, Japanf
United Graduate School of Agricultural Science, Tokyo University of Agriculture and Technology, 3-5-8 Fuchu, Tokyo, 183-8509, Japanb

A R T I C L E I N F O

A B S T R A C T

Article history:
Received 15 June 2019
Received in revised form 3 October 2019
Accepted 29 October 2019

A novel strain of microalga Parachlorella sp. BX1.5 was isolated and its unique properties of producing
lipids and extracellular polysaccharides (EPS) characterized. The cells could extracellularly produce alarge amount of acidic EPS, when cultured in nitrogen-deﬁcient BG110 medium (BG11–N) with 2 % CO2air supply. The main component of intracellularly accumulated lipids was triacylglycerol (TAG),
depending on the different cultivation conditions of BG11, BG11–N, BG11–P (phosphate depleted), and
BG11–N–P (nitrogen and phosphate depleted). Fatty-methyl-esters (FAMEs), methyl-esteriﬁcation of
total lipids, consisted of abundant saturated C16 and unsaturated C18 fatty acids under the culture
conditions. Cell spot assays on BG11 plates revealed the resistance of cells to pH 2–11, high temperaturesof 50–70  C, ultraviolet irradiation, and drought, under different culture conditions, thereby suggesting
the biological signiﬁcance of lipid and EPS accumulation. The prospects of BX1.5 as a dual producer has
also been discussed for bioreﬁneries.
© 2019 Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://
creativecommons.org/licenses/by-nc-nd/4.0/).

Keywords:
BiofuelBioreﬁnery
Dual production
Parachlorella
Polysaccharide

1. Introduction
Microalgae are model microorganisms that have strong
photosynthetic capabilities and are used as a resource inbioreﬁneries to produce fuels, fertilizers, feed, ﬁber, and food
[1,2]. Microalgae include prokaryotes (cyanobacteria) and eukaryotes (green algae, diatoms, stonewort, dinoﬂagellates, andeuglena). In biofuel production, cyanobacteria are employed as
producers of alkane/alkene, which is used as jet fuel, or kerosene,
which is a mixture of hydrocarbons [3–5].
Some green algae (e.g., Chlorella, Nannochloropsis, Nephroselmis,
and Chlamydomonas) have been applied in the production of
triacylglycerol (TAG), a neutral lipid that may be used as biodiesel
or food oil [6–11]. In such cases, properties of the oil depend on the
structure determined by the number of carbons and the position/* Corresponding author at: College of Agriculture, Ibaraki University, 3-21-1 Ami,
Ibaraki, 300-0393, Japan.
E-mail address: munehiko.asayama.777@vc.ibaraki.ac.jp (M. Asayama).

number of double bonds. In lipid biosynthesis, synthesis of fatty
acids is terminated once at the stage of palmitic acid (C16:0) as asaturated fatty acid, followed by the addition of sequentiallysynthesized acetyl-CoA to the substrates by fatty-acid synthase.
Thereafter, palmitic acid is extended to stearic acid (C18:0) by
fatty-acid elongation enzyme, resulting in further synthesis of C18derivatives as oleic acid (C18:1), linoleic acid (C18:2), or α-linolenicacid (C18:3). Therefore, the productivities of C16/C18 fatty-acid
derivatives in cells could be important when oil production is
examined for bioreﬁnery.
Regarding polysaccharide production, cyanobacteria and green
algae have been used as good producers in bioreﬁneries. For
example, sacran or spirulan from cyanobacteria Aphanothecesacrum (called suijenji-nori as Japanese) or Arthrospira (Spirulina)
sp. is an acidic extracellular polysaccharide (EPS) that can function
as useful material such as water-retention reagent or additive inthe ﬁelds of food/medical industries [12,13]. Rhamnan sulfate, asacidic EPS extracted from the green alga Monostroma nitidum, hasalso been characterized as a possible material for anti-bloodclotting, antiviral infection, or anti-cancer metastasis [14–16].

https://doi.org/10.1016/j.btre.2019.e00392
2215-017X/© 2019 Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

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M. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

Although useful EPS from microalgae have been reported, EPS fromthe green alga Parachlorella sp. has not been reported till date.
When we consider practical application of such usefulmaterials, the cost of extraction, processing, and manufacture is
critical. Therefore, effective productivities of such useful materials
need to be extensively investigated. In this study, a novel green alga
Parachlorella sp. strain BX1.5 was isolated and characterized as adual producer of useful oils (lipids) and EPS that accumulate
simultaneously inside and outside the cell, respectively. The
usefulness of this strain and possible outdoor cultivation have also
been discussed for practical bioreﬁneries.
2. Materials and methods
2.1. Isolation of green alga BX1.5
The Parachlorella sp. BX1.5 strain was isolated from an outdoor
cell culture, consisting of natural water collected from amountainous area in Hiroshima Japan, using a mixed and diluted
overlay method on BG11 agar plates [17].
2.2. Identiﬁcation of the BX1.5 strain
The 18S rDNA region of BX1.5 was ampliﬁed by PCR with theprimers 1F (50 - AACCTGGTTGATCCTGCCAG -30 ) and ITS2_r (50 TCCTCCGCTTATTGATATGC -30 ) (Fig. 1A). The ampliﬁed DNA
fragment was cloned into the cloning vector pUC119B to give
pBX18S. Nucleotide sequence of the 2.5-kb 18S rDNA region
was veriﬁed by a dideoxy method with the universal primers
1F, 93F (50 - CTGCGAATGGCTCATTAAAWCAG-30 ), 1528R (50 CCTCTAGGTGGGAGGGTTTAATG-30 ), 2346R (50 - GMAACCTTGTTACGACTT-30 ), ITSF (50 - AAGTCGTAACAAGGTTTCCG -30 ), and ITS2_r.
The 2.5-kb 18S rDNA sequence (DDBJ accession number LC473527)
was aligned by CLUSTAL W, integrated into the package MEGA7[18], for phylogenetic analysis and cell identiﬁcation [19].

2.3. Cultivation conditions
Cells were pre-cultivated in 50 mL of BG11 in a 100-mL
Erlenmeyer ﬂask for 5 days. The ﬂasks were placed in a CF-415
cultivation chamber (TOMY Co., Ltd., Tokyo, Japan) that supplied 2% CO2-containing air, and were subjected to continuous reciprocating shaking (40 rpm) under continuous white light conditions at
100 mmol photons/m2 s. Five milliliters of the cell culture wascollected and cells suspended in 30 mL of new BG11 [20], BG11–N(= BG110 as a nitrogen-deﬁcient condition without NaNO3), BG11–P(as a phosphorous-deﬁcient condition without K2HPO4), and
BG11–N–P (as nitrogen- and phosphorous-deﬁcient condition
without NaNO3 and K2HPO4) media in 300-mL Erlenmeyer ﬂasks.
These ﬂasks were placed in the same cultivation chamber to induce
production of ESP and lipid. During cultivation, a part of the culture
was sequentially harvested and cell turbidity measured at OD730
using MULTISKAN GO (Thermo scientiﬁc Co. Ltd., Tokyo, Japan)[17]. Cells were collected on the 6th day, during the main
cultivation, for cell staining or lipid extraction. Cells were also
cultured under static (standing) conditions with 0.04 % CO2-air in
50 mL BG11 in a 100-mL Erlenmeyer ﬂask, in which pre-cultured
0.5-mL cells were used as an inoculation, for standard cultivation
as a control of the above induction culture.
2.4. Cell staining and microscopic observations
To detect extracellular polysaccharides, a 100-mL aliquot of thecell culture, harvested from the main cultivation, was suspended in
20 mL India ink (Daiso Co., Ltd., Japan) and observed under an
optical microscope (BX53: Olympus, Tokyo, Japan) at DIC(differential interference contrast) conditions with a shutter speedof 0.48 s. Alcian blue (#015-13805: Wako Co., Ltd., Osaka, Japan)
staining was conducted for detecting acidic polysaccharides [21–23]. PAS (#164-19705: Wako Co., Ltd.) staining was performed for
detecting neutral polysaccharides [24]. Cells stained with Alcian

Fig. 1. Identiﬁcation of isolated Parachlorella sp. BX1.5. (A) The 18S rDNA-ITS1-5.8S-ITS2 region on the genome and nucleotide-sequence alignment of the region. The region(2,505 bp, DDBJ accession number LC473527) was PCR-ampliﬁed with a set of primers, 1F and ITS2_r, cloned, sequenced, and subjected to database analysis. Asterisks show
distinct nucleotide sequences from Parachlorella kessleri, which is the most homologous strain to BX1.5. (B) Phylogenetic analysis (MEGA 7 software) of the 18S rDNA
sequences resulted in classiﬁcation of the BX1.5 cells. Respective cases are shown with accession numbers. A bootstrap test was performed with 1,000 replicates. The 18S
rDNA sequence of Chlamydomonas moewusii was utilized as an out-group. The scale bar indicates 1 % sequence divergence.

M. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

blue or PAS were observed using BX53 (Olympus) at DIC conditions
with a shutter speed of 0.48 s. Regarding intracellular lipid staining[25], a 1-mL aliquot of the cell culture was collected from the main
cultivation and mixed with 20 mM of Nile red (#144-08811: Wako
Co., Ltd.). Samples were observed for morphology using an opticalphotomicroscope at a high resolution with a Nomarski prism orusing the ﬂuorescence microscope, BX53/DP72, ﬁtted with a
special ﬁlter, U-FBW (Olympus) for excitation (460–495 nm) andemission (510 nm). The exposure time for photography was 0.2 s.
2.5. Electron microscope observations
For transmission electron microscopy (TEM), cells werecollected from 20 mL cell culture, and ﬁxed with the reagents 2% formaldehyde (PFA) + 2 % glutaraldehyde (GA) in 0.05 M
cacodylate butter (pH 7.2) for pre-ﬁxation, and 2 % osmium
tetraoxide in 0.1 M cacodylate butter for post-ﬁxation. The
resulting cells that stained with uranyl acetate and lead solution
were observed under TEM (JEM-1200EX: Jeol, Tokyo, Japan) using
an ultrathin sectioning method [26].
2.6. Thin-layer chromatography (TLC)
To analyze the lipids accumulating in cells, samples wereprepared as follows. Cells were collected from the main cultivationand freeze-dried. Twenty milligrams of dried cells were transferred
into a 15-mL glass tube, to which 2 mL of a solvent (chloroform:
methanol = 2:1) was added, and lipids extracted using the UD-100
sonicator (TOMY, Tokyo, Japan: set at an output of 70 for 10 min). The
resultant samples were left overnight in a cool dark place to improveextraction. Subsequently, they were centrifuged at 3,000  g for
10 min and the supernatants were collected. To wash the samples,
equal amount of 0.9 % aqueous KCl solution was added to thesupernatant, mixed, and subjected to centrifugation at 2,000  g for
5 min. Approximately 2 mL of the bottom fraction was collected as
the organic layer, transferred to a 15-mL glass tube, and placed in arotary vacuum evaporator (V-850: BUCHI, Flawil, Switzerland). The
resultants were dissolved in 200 mL chloroform, and eventuallyused for TLC. A 4-mL aliquot of each sample was spotted onto theorigin (ori) of a silica gel plate (#60F254: Merck Millipore,
Darmstadt, Germany). Lipids on the plate were developed with
an eluent (hexane:diethylether:acetic acid = 80:20:1) and stainedwith 5 % (v/w in EtOH) phosphomolybdic acid solution (#32186-00:
Kanto Kagaku Co., Ltd., Tokyo, Japan).
2.7. Lipid measurements by gas chromatography/ﬂame-ionization
detection (GC/FID)
To measure the intracellular accumulation of lipids, samples
were prepared as follows [27]. Ten milliliters of the cell culture
harvested from the main cultivation was transferred into a 50-mL
plastic tube. Cultures from the respective media were directly
freeze-dried to obtain total fractions of cells with EPS. Dry-cellweight (DCW) of the fractions was measured to calculate theamount (mg) of fatty acyl methyl esters (FAMEs) in DCW (g). Dried
fractions were added to 4 mL of solvent (chloroform: methanol = 2:1), and n-Eicosane (#44-2673, Sigma Aldrich, St. Louis, USA:stock solution as 10,000 ppm in ethyl acetate) was used as aninternal standard up to a ﬁnal concentration of 100 ppm. Lipids
were extracted from the resultants, using the same procedure,with the UD-100 sonicator (see the section above). Aftercentrifugation at 3,000  g for 10 min, approximately 4 mL of
the supernatant was collected and an equal amount of 5 % NaClaqueous solution was added to wash the sample. After mixing,
samples were centrifuged at 3,000  g for 10 min, and the bottom
fraction, as the organic layer, was transferred into a 4-mL glass

3

tube. Samples were dried using the same method, with the rotary
vacuum evaporator. Dried samples were dissolved in 1 mL of
toluene, transferred to a 20-mL glass tube with a lid, and 2 mL of
3 M HCl-methanol (#33050-U: Sigma Aldrich) was added to theextract. After mixing, samples were allowed to react in a water bathat approximately 90  C for 3 h to convert lipids to FAMEs. The
resultants were transferred to a 50-mL plastic tube. After cooling toroom temperature, 1 mL of sterilized water and 2 mL of hexane
were added. After mixing, samples were centrifuged at 3,000  g
for 10 min, and the organic layer was transferred thereafter to a
3-mL glass tube. Samples were concentrated and dried using
a rotary vacuum evaporator. The resultants were transferred into a
1.5-mL vial and dissolved in 1 mL hexane. This sample was thensubjected to TLC analysis, as described above. A 1-mL aliquot of
each sample was subjected to GC/FID analysis using a GC-2014
system (Shimadzu Co., Ltd., Kyoto, Japan), wherein, a capillary tube
was purged with helium gas (80 kPa), hydrogen (50 kPa), and air(50 kPa), and the injector temperature was maintained at 250  C.
Temperature of the column (Stabiliwax-DA: Shimadzu GLC, Tokyo,
Japan) was initially maintained at 140  C for 5 min, increased to220  C at an interval of 2.5  C per min, and maintained at 220  C for
7 min. n-Eicosane (C20:0, #44-2673: Sigma Aldrich) as an internal
standard or a C16:0/C18:0/C18:1/C18:2/C18:3 mixture (GLC-10,#1891-1AMP: Sigma Aldrich, St. Louis, USA), cis-9-hexadecenoic
acid (C16:1, #373-49-9: Tokyo Chemical Industry Co., Ltd, Tokyo,
Japan), cis-7, 10-hexadecadienoic acid (C16:2, #10-1622-4: Funakoshi, Tokyo, Japan), and all-cis-7, 10, 13-hexadecatrienoic acid
(C16:3, #10-1403: Funakoshi) were obtained as standards from therespective companies.
2.8. Spot assay for stress resistance
BX1.5 cells were pre-cultivated in 50 mL of BG11 medium in the
2 % CO2 gas-supplying incubator for 5 days (see the section on
Cultivation Conditions in the Materials and methods). A 5-mL
aliquot from a pre-inoculated cell culture was harvested,transferred into 30 mL of new BG11, BG11–N, or BG11P medium,and cultivated thereafter under the same conditions as the main
cultivation for 6 days. The cells were then collected and exposed
subsequently to the respective stress conditions described. For
heat stress, 0.25 mL of the cell culture was harvested, transferred
into a 1.5-mL microtube, and then exposed to 50, 60, or 70  C for
5 min. For acidic or alkaline stress, 0.25 mL of the cell culture washarvested and transferred into a 1.5-mL microtube. A 25-mL aliquotof 1 N hydrochloric acid (cell culture as pH 2) or 0.5 N sodiumhydroxide (cell culture as pH 11) was added to the culture. For
heat-dry (HD) stress, 1 mL of the cell culture was harvested, cellpellet collected by centrifugation at 15,000  g, and subsequently
exposed to 42  C for a day. One milliliter of new BG11 medium wasthen added to the dried cell pellet and mixed well. For freeze-dry(FD) stress, 1 mL of the cell culture was harvested, cell pellet
collected by centrifugation at 15,000  g, and cells kept at atemperature -80  C until next use. This sample was vacuum freezedried using V-850 (BUCHI) for 90 min under a 0.002 mbar setting.
One milliliter of new BG11 medium was added to the dried cellpellet and the sample mixed well. For ultraviolet (UV) stress,
100 mL of cell culture was harvested, transferred to a 96-well plate,and UV-irradiated (100 mmol photons/m2 s, with a 260-nm light)
for 5 min at a distance of 8 cm from the UV light source. Fivemicroliter aliquots of cell culture samples exposed to therespective stresses were spotted onto BG11 agar plates just after
the stress treatment or after standing for a day following the stresstreatment. HD samples were also spotted onto BG11 agar plates
after the stress treatment. These cells were incubated in 0.04 %CO2-air condition with 30 mmol photons/m2 s of white-ﬂuorescentlight at 30  C for 6 days.

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3. Results and discussion
3.1. Isolation and identiﬁcation of BX1.5
An aliquot of cell culture from an open pond, at a 1.5-ton scalebioreactor in Hiroshima, Japan, was harvested and the algal cells
isolated. These cells were analyzed by comparing the phylogenetic
relationships across 11 microalgal strains based on the 2.5-kbp
nucleotide sequences of the 18S rDNA-ITS1-5.8S-ITS2 region(Fig. 1). Results revealed the BX1.5 sequence (accession number
LC473527) to be homologous at 99.9 % (2,502 bp/2,505 bp  100 %)to that (accession number FR865655) of Parachlorella kessleri.
However, the cells possessed three distinct nucleotide sequencesat the regions of ITS1 and ITS2 (Fig. 1A). Therefore, this strain was
designated as green alga Parachlorella sp. BX1.5, a novel species (orsubspecies) belonging to the order Chlorellales in the class
Trebouxiophyceae, and phylum Chlorophyta (Fig. 1B).
3.2. Cultivation condition and culture proﬁleParachlorella sp. BX1.5 was cultured in the laboratory under avariety of controlled conditions in order to examine the potential ofthis new alga for use in bioreﬁnery applications. This investigation
involved different concentrations of CO2 and different types ofnutrient-deﬁcient media. Especially, speciﬁc growth conditions
supplying high concentration of 2 % CO2-air were efﬁcient inprompting the induction of overproduction of microalgae for
useful materials [19,28,29], compared to normal cultivation
conditions supplying 0.04 % CO2-air (Materials and methods).
The resulting culture color and cellular turbidity proﬁles are shownin Fig. 2. When cells were cultivated in BG11 media, under static
conditions, with 0.04 % CO2-air, cell turbidities with green color
were gradually increased, exhibiting a standard curve (Fig. 2A, B).
In contrast, when the cells were cultivated in BG11 media undershaking conditions with 2 % CO2-air, a darker green color wasobserved and turbidity reached a maximum value by the 6th day(Fig. 2A, B and Fig. S1); it remained almost constant for 21 days
although the color changed from green to yellow. Under the same
conditions, a yellow color was observed for the BG11–N media.
However, a greenish-yellow color was observed for the BG11P
and BG11–NP cultivated cells. Cell turbidities of nutrientdeﬁcient culture were relatively lower than that of BG11 culture(Fig. 2B). These results implied that cell properties are distinct intheir respective cultures; both cell size and chlorophyll a (Chl a)content of cells differed signiﬁcantly depending on the culture
conditions (supporting information in Figs. S1 and S2). From theseresults, the optimal efﬁcient period in this study was set as 6 days
under induction cultivation with 2 % CO2-air (Fig. S1) for usefulmaterial production.
3.3. Acidic EPS production
First, we conﬁrmed the validity of India ink staining method byusing several microalgae, which have been known to produce
extracellular polysaccharides (EPS) (data not shown). Thereafter,
the BX1.5 cells grown under different cultivation conditions, as
shown in Fig. S1, were stained with India ink for EPS and observed
under the microscope (Fig. S3). Large amount of EPS accumulated
in the culture with 2 % CO2-air, especially in the samples from
BG11–N and/or BG11–NP medium. Therefore, the cells cultivatedin the BG11–N medium were further characterized by staining with
India ink for EPS, with Alcian blue for acidic polysaccharides, andwith PAS for neutral polysaccharides. The results are shown in
Fig. 3. It was interesting that a thick layer observed outside the cell,possibly EPS, remained unstained by India ink (Fig. 3A). Althoughmost EPS peeled off during the course of Alcian blue staining, an

Fig. 2. Culture and cell-turbidity proﬁles of BX1.5. Following cultivation in 2 % CO2air for 21 days (see details in Materials and methods), an aliquot of 0.2 mL wascollected from the culture, transferred into a 96-well plate (A), and cell turbidities
measured as OD730 (B). Control cells were also cultivated under standard conditions
with 0.04 % CO2-air, collected, and the turbidities measured. The OD730 values areshown with standard deviations (n = 3 or 2).

acidic EPS layer that retained the blue color existed outside the cell(Fig. 3B). In contrast, the outer layer of the cell failed to stain withPAS (Fig. 3C). These results suggested that acidic EPS is selectively
over-produced under BG11–N growth conditions. Carbon metabolic pathways linked to lipid synthesis are suggested to be
activated under nutrient-depletion culture conditions [10,19].
Thus, there is a possibility that the same situation occurred for the
overproduction of EPS under N/P  starved conditions with 2 %CO2-air supply.
Acidic EPS is referred to as rhamnan sulfate when produced
from the green alga Monostroma nitidum. Rhamnan sulfate has
been reported to confer a blood-clotting effect as well as antiviraleffect [14–16]. Acidic EPS is also known as sacran (acidic sugar)
when extracted from the cyanobacterium Aphanothece sacrum[30]. This cyanobacterium is edible and marketed as Suijenji-nori
in Japan. Sacran is also utilized for product development (e.g.,
metal absorber, absorbent polymer, etc.) due to high water
retentivity. We had recently reported a novel ﬁlamentous
cyanobacterium Limnothrix sp. SK1-2-1 that can co-produce
pentadecane and acidic EPS, thereby contributing to cell ﬂocculation [19]. Therefore, microalgal EPS might contribute to manyﬁelds such as medical, food, and industry. Although it is very
interested in a mechanism for production and a molecular
structure of BX1.5 EPS, they are research agenda for elucidation.
3.4. Oil production in cells
BX1.5 cells were next utilized to simulate oil production withinthem. Nile red, which is a common reagent used for lipid-staining

M. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

5

BG11–N, or BG11P, respectively (Fig. S2B). These values seemed tosupport the photosynthesis-induced cell color observed in thisstudy, such as green, yellow, or yellow-green, as shown in Figs. 2,
S1, and 4 (top panels).
3.5. TEM observation of the Parachlorella sp. BX1.5 cells

Fig. 3. Acidic polysaccharide accumulation on the cell exterior and oil production in
the interior. The BX1.5 cells were cultivated under BG11N conditions shown in
Fig. S1. An aliquot of 100 mL culture was collected and subjected to staining with
India ink (A), Alcian blue (B), or PAS (C) (see details in Materials and methods). The
cells were then subjected to microscopic observation. Bar, 10 mm.

in microalgae, was utilized to stain the cells [8,31,32]. Cells
cultivated in BG11 medium with 2 % CO2-air were stained for 5 min
on glass slides and then observed under the microscope. Perhaps
due to oil drops existing in the cells, almost all the cells with adiameter of 5.23  0.04 mm (Fig. S2A) were insufﬁciently stainedwith Nile red. However, when these cells were stained for 30 min
on glass slides, larger cell size was observed, as the color changed
from red (by autoﬂuorescence with photosynthesis pigments) toyellow (by oil production) in some parts of the cells (Fig. 4, left).
Further, when cells were cultivated in the BG11N medium with 2% CO2 and stained for 5min on glass slides, microscopic observation
revealed oil drops in the cells, with signiﬁcantly smaller cells withan average diameter of 4.420.03 mm (Fig. S2A), which also failed
to stain sufﬁciently, indicative of possible inhibition of EPS
accumulation. Following 30-min staining, the sample again
resulted in larger cells, changing color from thin red (byautoﬂuorescence with fewer photosynthesis pigments) to strong
yellow (by signiﬁcant oil production) either in the cells or outside
the lysed cells (Fig. 4, center). This phenomenon might haveoccurred due to cellular desiccation, causing damage to the cells
and oozing of oil, gradually over time, on the glass. A similartendency in staining was observed when cells were cultivated in
the BG11P medium with 2 % CO2 (Fig. 4, right). The cell size
increased, producing larger cells, with a diameter of 6.450.06mm(Fig. S2A), which were signiﬁcantly larger than that of the BG11
medium. In this situation, cells from the BG11P culture contained
signiﬁcant oil and a certain amount of photosynthetic pigment
corresponding to the yellow-green color of cells. Thus, we further
investigated this reaction by measuring and evaluating the amountof chlorophyll a present in cells harvested from BG11, BG11–N, and
BG11P. Chlorophyll a levels (mg/mL of cell culture) were observed
as mean values: 2.7  0.07, 0.09  0.009, or 0.23  0.02 for BG11,

Starch and oil production in BX1.5 cells was examined via TEM
observations (Fig. 5), in order to visualize the cell contents of thisnovel alga. Cells were observed following cultivation undercontrolled conditions in the BG11 medium with 0.04 % CO2 for
13 days (Figs. 2B and 5 A). A characteristic feature of this alga is thatthylakoid membranes are spread throughout the cytoplasm when
the cells are cultivated under standard culture conditions (Fig. 5A).
Another of its characteristics was the absence of clear chloroplasts,
and the cell proﬁle seemed to be similar to those of cyanobacteria
or green algae from Chlamydomonas. Starch had visibly accumulated along multiple layers of thylakoid membranes, a pyrenoid
body for storage, and starch formation was also observed.
Moreover, the cells also produced starch (white particles) andoil (gray particles) when they were cultivated under variableconditions in BG11 media at 2 % CO2-air (Fig. 5B). This might
indicate that supplementing CO2 into the environment effectively
conferred oil production ability to the cells. Cultivation in nitrogendeﬁcient BG11–N medium with 2 % CO2-air resulted in mass
production of oil in the cell, which occurred due to thedegeneration of thylakoid membranes (Fig. 5C). Oil storage
accounted for over 70 % of the cross-sectional area of a cell. Suchoverproduction of oils was also observed in the hypertrophic cells
cultivated under phosphorus-deﬁcient conditions in BG11P
medium with 2 % CO2-air (Fig. 5D). Interestingly, although oil
storage accounted for over 55–65 % of the cross-sectional area in acell, the area of thylakoid membranes involved with starch
production was more prominently visible in the representative
cells (Fig. 5D). These observations were consistent with the test
results in Figs. 2, S1, and 4. The results from Fig. 5 also indicatedthat starch or oil production and cell morphology depend
signiﬁcantly on cell culture condition, including CO2-gas concentration and type of nutrient depletion in the medium.
Overproduction of oil was observed to be inversely correlatedwith the reduction in the accumulated amount of starch in thethylakoid membranes of cells cultivated under 2 % CO2 conditions(Fig. 5B–D). In contrast, starch accumulated, as did clear thylakoid
membranes, in the cells cultivated at 0.04 % CO2 conditions in BG11
medium (Fig. 5A). In the cell, pyrenoid was conﬁrmed to occur,
which can in turn confer an ability of CO2 ﬁxation for generating
and maintaining a rich environment of CO2 concentration in
combination with the photosynthetic enzyme ribulose-1,5bisphosphate carboxylase/oxygenase (RuBisCO) found in the
carboxysome of cyanobacteria. Therefore, carbon ﬂow from starchmight supply a part of a precursor for synthesizing lipids or EPS. In
green alga, Chlamydomonas, starch and lipids also accumulated
along the spaces of thylakoid membranes in the cells, and thecontent proﬁles appeared to be similar to those seen in the BX1.5
cells [7,33].
3.6. TLC analysis for the lipids
Thin Layer Chromatography (TLC) revealed the lipid composition of cells, and the results are depicted in Fig. 6. Samples wereprepared from cells shown in Fig. 2. Remarkable accumulation of
TAG, whose Rf value was 0.35 on a silica plate [7,10], was observed
from the unit per dry-cell weight (DCW), cultivated in BG11, BG11–
N, and BG11–P media at 2 % CO2-air. The order corresponding to theaccumulation rate of TAG in cells in each medium was as follows:
BG11–P>BG11–N>BG11. Apparent accumulation of wax ester

6

M. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

Fig. 4. Oil production in the cell interior. BX1.5 cells were cultivated under respective conditions shown in Fig. S1. Following this, an aliquot of 1 mL was collected from theculture, transferred into a new microtube, and Nile red was added to stain the lipids accumulated in the cells. Cell morphology was observed using the opticalphotomicroscope, at a high resolution, with a Nomarski prism (Top: optics, differential interference contrast) or with a ﬂuorescence microscope (Bottom). The respective
times of 5 or 30 min are the sequential times when the sample was put on a glass slide. Bar, 10 mm.

(WE), whose Rf value was 0.77 on the silica plate [7,10], was also
observed in the cells. In contrast, only a minute amount ofaccumulated TAG was observed in cells cultivated in the BG11
medium at 0.04 % CO2-air. We also conﬁrmed that lipids (Fig. 6,left) were totally converted to fatty-acid methyl esters (FAMEs:
Fig. 6, right), Rf value being 0.57 on the silica plate for GC/FID
analysis.
3.7. FAME content in the cells
The results for C16 and C18 FAMEs are shown in Fig. 7. The total
amount of BX1.5 dry-cell weight (DCW) harvested from 1 L of cell
culture grown on BG11, BG11–N, BG11P, and BG11–NP media
was approximately 5.70, 1.77, 1.40, and 1.60 g, respectively. This
indicates that the order of biomass production per unit of cell
culture is BG11 > BG11–N > BG11NP > BG11P. Total amountsof FAME (lipids) were 16 %, 27 %, 32 %, or 31 % in DCW of BG11,
BG11N, BG11–P, or BG11NP, respectively, in media (Fig. 7A).
Lipid (oils) composition and FAMEs were also evaluated and listed
in Supplementary Table S1. Interestingly, the range of proportionally integrated unsaturated fatty acids for C18:1 + C18:2 + C18:3
was 68 %, 67 %, 66 %, or 71 % in the FAMEs from cells cultivated in
BG11, BG11N, BG11P, or BG11NP media, respectively,
indicating that BX1.5 could signiﬁcantly accumulate useful lipids
utilized as food oils. The ability of lipid production was converted
and shown per unit of cell-culture (Fig. 7B). The total productivities
of FAMEs were 900, 473, 441, or 491 mg from one-liter of cell
culture grown on BG11, BG11N, BG11P, or BG11NP media,
respectively. These results indicated that abundant unsaturated

C18 FAMEs, which are useful food oils, could be obtained from
BX1.5 cultivation in BG11 medium.
TAG production by the green alga Parachlorella kessleri is usuallycharacterized by its growth on sulfur-resource-deﬁcient liquidmedium [9,34,35]. The production range of total lipid values wasreported to be approximately 0.25–1.9 g/L or 30–55% in dry-cellweight, whose biomass was given as 0.04–2 g/L of medium per day.
In this report, the values are approximately 0.44 to 0.90-g/Lmedium or 16 (27) to 32 % in dry-cell weight whose biomass wasgiven as 0.23 to 0.95 g/L per day from BX1.5 at cultivation (Fig. 7,
Table S1). The production values of 27 or 32 % in DCW did notmatch the extensive oil accumulation found in BG11N or BG11Pthat accounted for 70 % or 65 %, respectively, of the cross-sectional
area of cells during cultivation (Fig. 5). DCW involving EPS
eventually resulted in increased biomass, correlating with lower
values of lipid production rate per DCW. Therefore, the lipid
production value per liter of culture may be appropriate for
evaluating the BX1.5 case. In fact, 0.9-g lipid production from oneliter of BX1.5 cell-culture in the BG11 medium could be high among
all Parachlorella cases (Fig. 7, Table S1).
C16 and C18 FAMEs derived from TAG are generally known to be
detected in green algae [8,9,31,35]. The ingested α-linolenic acid
(C18:3) is further converted to n-3 (v3)-essential fatty acids as EPA(C20:5) and DHA (C22:6) for useful oils. At this time, no clear
accumulation of unsaturated polyvalent fatty acid C20:5 or C22:6
FAMEs, derived from the BX1.5 cells, was observed (data notshown). Further analysis of long-chain unsaturated fatty acidaccumulation is still awaited. Till date, oil production potential ofgreen algae has mainly been discussed as a good example ofM. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

7

Fig. 5. Electron microscopic observation of the BX1.5 cells. Cells were cultivated under static conditions in BG11 medium with 0.04 % CO2-air for 13 days (A) or under shaking
conditions with either BG11 (B), BG11N (C), or BG11P (D) media and 2 % CO2-air for 6 days (Fig. S1), as shown in Fig. 2. Resulting cells were harvested, treated with reagents,
and then observed with a transmission electron microscope (TEM). C/P, carboxysome in pyrenoid; N, nucleus; O, oil body; S, starch; TM, thylakoid membrane. Bar, 1 mm.

biodiesel fuel production due to the high productivity andabundant renewable oil content of algae [6,36]. However, theBX1.5 strain could be a candidate for food production as well, since
it produces useful food oil and EPS acidic sugar, thus suggesting anovel characteristic, never seen before, in Parachlorella spp. strains.

We have utilized BG11 medium for both cyanobacteria andgreen algae. Therefore, it was remarkable to discover that thismedium, when utilized for the cultivation of the new alga, wascapable of producing several useful materials by simply altering
the growth conditions. During this study, the conditions of 2 % CO2air, nutrient (N/P)-depletion, and high-light irradiation at
100 mmol photons/m2 s resulted in overproduction of oil and
EPS. A dilution of liquid culture, resulting in cell density, or airdrying of cells might be important strategies for future research[8,10,32]. In addition, trials using other media or reinforcing BG11
medium by adding sodium acetate might be advantageous [37,38].
Moreover, addition of carbon-source supplements or sugars
created by heterotrophic cultivation [39,40], may also help further
in effective production. Use of genetically modiﬁed BX1.5 strains
may also create an opportunity for increased production potential
for the future since genes related to TAG production have beenreported [7,31,33,41,42].
3.8. Spot assay for cell resistance to stressFig. 6. TLC analysis of lipids. Cells were harvested following cultivation described inFig. S1, and then freeze-dried. Lipid fractions were extracted and prepared from theunit per dry-cell weight (DCW) (see details in Materials and methods). An aliquot of
the 4-mL extract was spotted onto origin (ori) in 4 lanes at the left of a silica gel plate.
FAMEs were also prepared from the lipid fractions (Materials and methods). An
aliquot of the 4-mL extract was also spotted onto origin in 3 lanes at the right of thesilica gel plate. This plate was subjected to thin-layer chromatography (TLC) with adeveloper eluent, and then stained with phosphomolybdic acid solution. Positions
referring to respective lipids are shown as wax ester (WE), triacylglycerol (TAG),
possible diacylglycerol (DG), and fatty acid methyl esters (FAMEs), on the right. The
Rf values are shown on the left.

Stress tolerance of the cell is an important factor to consider
while developing large-scale cultivation, particularly in an outdoor
bioreﬁnery. Therefore, we trialed resistance to environmental
stress factors using BX1.5 cells, and the results are shown in Fig. 8.
Cell samples were spotted onto the plate once immediately
following stress tests (Fig. 8A) and again 24 h later (Fig. 8B).
Following the stress treatments, photographs of cell proﬁles weretaken after 6-days of cultivation. Signal intensities of respective
cell spots on the plate were quantiﬁed and presented here as a
Supplementary ﬁgure (Fig S4). The data relating BX1.5-cellresistance to stress are characterized by pH resistance; the
BG11-cultivated cells exhibited tolerance over a wide range ofpH, from pH 2 to pH 11. Notably, the instability of BG11-cultivated8

M. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

Fig. 7. FAME contents in the BX1.5 cells. Samples were prepared, as described inMaterials and methods, from 10 mL culture of the cells, as shown in Fig. S1, and thensubjected to GC/FID analysis in which eicosanoic acid (C20: n-0) methyl-ester wasused as an internal standard substance. (A) The amount (mg) of FAMEs in dry-cellweight (g) are shown. (B) Content (mg) of FAMEs from 1 L of cell-culture is also
shown. Data with standard deviations (n = 3) were obtained from independent
triplicate experiments.

cell on the acidic side was reinforced by utilizing the cells
cultivated in BG11N/P medium. Cells overproducing oil andacidic EPS may become resistant to acidic conditions. Interestingly,
these cells also exhibited a relatively strong tolerance to alkalinestress conditions at pH 11. This knowledge is advantageous to thecultivator since they can facilitate the penetration of CO2 gas into
an alkaline liquid growth medium and utilize this method tosuppress contamination by other microorganisms in ﬁeld culture.
Similarly, cyanobacterium Spirulina tolerated highly alkaline
growth SOT medium with a pH of 10–11 and withstood higher
levels of sodium bicarbonate (NaHCO3) during outdoor cultivation[43]. pH of the medium at the start of BG11 culture was about 6.5;
the supply of 0.04 % CO2-air increased the pH of liquid culture to
8–10 for 6 days until the conclusion of the trial (Fig. S2C). Thistendency resulted in pH 7–8 under 2 % CO2-air supply in theculture, indicating that uptake of a high concentration of CO2
gas will lead to a shift in pH from alkaline to the acidic side
(Fig. S2C).
The BG11-cultivated cells exhibited tolerance at high temperatures of 50–70  C. It would favor the sterilization of othercontaminants in cultivation or processing of BX1.5 cells to utilize
them as health supplements or as functional foods. UV-irradiation
resistance testing of cells found the tolerance effect to be BG11 >BG11P> BG11N, correlating well with the accumulation rates ofchlorophyll a (Fig. S2B). We considered, based on the results in
Figs. 2, 8, S1, and S2, that the extent of UV tolerance might notcontradict the chlorophyll a (and EPS) amounts accumulated in(and outside) the cells. Most types of stress involving photolesions(e.g., high-light or UV irradiation) have been postulated to inhibit
the ﬁxation of CO2 within the resultant ROS (reactive oxygen
species), thereby inactivating the photochemical reaction center of
PSII, consisting of photosynthesis pigments [44]. For dry resistance, the BG11, BG11N, or BG11P cultivated cells exhibitedtolerance to heat-dry (HD) and freeze-dried (FD) stress factors.
Regarding drought resistance, although the cells survived the HD
and FD stresses, instability of the BG11P cell was observed in caseof FD stress (Figs. 8 and S4). Phosphate is known to be an essential
element for cells. Its depletion in the BG11 medium could cause
reduced strength of biological membranes since phospholipids area necessary component of the cell membrane, resulting in enlarged
and lysed cells during microscopic observation (Figs. 4 and 5). The
nature of these altered cells, despite being more vulnerable,
allowed for easy extraction of sugars and oils, hence resulting inlower extraction cost. Moreover, the altered BX1.5 cells may beeffectively absorbed into the digestive system in vivo when they areeaten. These results together suggested that the Parachlorella sp.
BX1.5 cells possess high resistance to environmental stress,
exhibiting characteristics that facilitate its use in an outdoor
cultivation for bioreﬁneries.
4. Conclusions

Fig. 8. Spot assay for stress resistance. Cultured cells were harvested, exposed tostress factors, and 5 mL aliquots were spotted onto a BG11 plate (see details inMaterials and methods). The cells were incubated for 6 days to observe stressresistance. (A) Samples were spotted onto the plate immediately after the stresstreatments. (B) Samples were spotted onto the plate 24 h after the stresstreatments. UV, ultraviolet irradiation; HD, heat dry; FD, freeze dry.

A novel green alga Parachlorella sp. BX1.5 can produce highyields of both lipids and extracellular polysaccharides, without
conﬂicting with the internal space required for other processes,
even under cultivation conditions utilizing the BG11 liquid media.
The BX1.5 cells also showed signiﬁcant stress tolerance, and theircharacteristics drastically changed with cultivation conditions,which regulated the amount and content of potential usefulﬁnished products or materials, thus promising outdoor cultivation
for bioreﬁneries.
Declaration of Competing Interest
The authors declare no conﬂict of interest.

M. Sasaki et al. / Biotechnology Reports 25 (2020) e00392

Acknowledgments
This work was partially supported by a propulsive project from
Ibaraki University (to MA), Ministry of Agriculture, Forestry and
Fisheries, Japan (to DS and MA), Japan Society for Bioscience,
Biotechnology, and Agrochemistry (to MA and DS: AcademicIndustrial R&D Support for Small-to-Medium-sized Enterprises,
SME-3), and Japan Society for the Promotion of Science (to MA and
DS: A-STEP, JPMJTM19BT; OPERA).
Appendix A. Supplementary data
Supplementary material related to this article can be found, inthe online version, at doi:https://doi.org/10.1016/j.btre.2019.
e00392.
References[1] L. Brennan, P. Owende, Biofuels from microalgae–a review of technologies for
production, processing, and extractions of biofuels and co-products, Renew.
Sustain. Energy Rev. 14 (2010) 557–577.
[2] M.P. Caporgno, A. Mathys, Trends in microalgae incorporation into innovative
food products with potential health beneﬁts, Front. Nutr. 5 (2018) 58.
[3] R.C. Coates, S. Podell, A. Korobeynikov, A. Lapidus, P. Pevzner, D.H. Sherman, E.
E. Allen, L. Gerwick, W.H. Gerwick, Characterization of cyanobacterialhydrocarbon composition and distribution of biosynthetic pathways, PLoS
One 9 (2014) e85140.
[4] N. Quintana, F.V. Kooy, M.D. Rhee, G.P. Voshol, R. Verpoorte, Renewable energy
from Cyanobacteria: energy production optimization by metabolic pathway
engineering, Appl. Microbiol. Biotechnol. 91 (2011) 471–490.
[5] A. Schirmer, A.R. Mathew, X. Li, E. Popova, S.B. Cardayre, Microbial biosynthesisof alkanes, Science 329 (2010) 559–562.
[6] Y. Chisti, Biodiesel from microalga, Biotech Adv. 25 (2007) 294–306.
[7] J. Fan, C. Andre, C. Wu, A chloroplast pathway for the de novo biosynthesisof triacylglycerol in Chlamydomonas reinhardtii, FEBS Lett. 585 (2011) 1985–
1991.

[8] X. Li, P. Pribyl, K. Bišová, S. Kawano, V. Cepák, V. Zachleder, M. Cížková,
I.
Brányiková, M. Vítová, The microalga Parachlorella kessleri––a novel highly
efﬁcient lipid producer, Biotechnol. Bioeng. 110 (2013) 97–107.
[9] Y. Mizuno, A. Sato, K. Watanabe, A. Hirata, T. Takeshita, S. Ota, N. Sato, V.
Zachleder, M. Tsuzuki, S. Kawano, Sequential accumulation of starch and lipid
induced by sulfur deﬁciency in Chlorella and Parachlorella species, Bioresour.
Technol. 129 (2013) 150–155.
[10] T. Shiratake, A. Sato, A. Minoda, M. Tsuzuki, N. Sato, Air-drying of cells, thenovel conditions for stimulated synthesis of triacylglycerol in a green alga,
Chlorella kessleri, PLoS One 8 (2013) e79630.
[11] Z. Xue, F. Wan, W. Yu, J. Liu, Z. Zhang, X. Kou, Edible oil production frommicroalgae, Eur. J. Lipid Sci. Technol. 120 (2018) 6.
[12] J.H. Choi, S. Kim, S.J. Kim, Spirulan from blue-green algae inhibits ﬁbrin andblood clots: its potent antithrombotic effects, J. Biochem. Mol. Toxicol. 29(2015) 240–248.
[13] N.R. Ngatu, M.K. Okajima, M. Yokogawa, R. Hirota, M. Eitoku, B.A. Muzembo, N.
Dumavibhat, M. Takaishi, S. Sano, T. Kaneko, T. Tanaka, H. Nakamura, N.
Suganuma, Anti-inﬂammatory effects of sacran, a novel polysaccharide from
Aphanothece sacrum, on 2,4,6-trinitrochlorobenzene-induced allergic
dermatitis in vivo, Ann. Allergy Asthma Immunol. 108 (2012) 117–122.
[14] S. Karnjanapratum, S. You, Molecular characteristics of sulfatedpolysaccharides from Monostroma nitidum and their in vitro anticancer andimmunomodulatory activities, Int. J. Biol. Macromol. 48 (2011) 311–308.
[15] J.B. Lee, S. Koizumi, K. Hayashi, T. Hayashi, Structure of rhamnan sulfate fromthe green alga Monostroma nitidum and its anti-herpetic effect, Carbohydr.
Polym. 81 (2010) 572–577.
[16] W.J. Mao, F. Fang, H.Y. Li, X.H. Qi, H.H. Sun, Y. Chen, S.D. Guo, Heparinoid-active
two sulfated polysaccharides isolated from marine green algae Monostroma
nitidum, Carbohydr. Polym. 74 (2008) 834–839.
[17] T. Nishizawa, T. Hanami, E. Hirano, T. Miura, Y. Watanabe, A. Takanezawa, M.
Komatsuzaki, H. Ohta, M. Shirai, M. Asayama, Isolation and molecularcharacterization of a multicellular cyanobacterium, Limnothrix/Pseudanabaena
sp. strain ABRG5-3, Biosci. Biotechnol. Biochem. 74 (2010) 1827–1835.
[18] K. Tamura, G. Stecher, D. Peterson, A. Filipski, S. Kumar, MEGA6: molecular
evolutionary genetics analysis version 6.0, Mol. Biol. Evol. 30 (2013) 2725–
2729.

9

[19] T. Sugawara, M. Chinzei, S. Numano, C. Kitazaki, M. Asayama, Flocculation andpentadecane production of a novel ﬁlamentous cyanobacterium Limnothrix sp.
strain SK1-2-1, Biotechnol. Lett. 40 (2018) 829–836.
[20] R. Rippka, Isolation and puriﬁcation of cyanobacteria, Methods Enzymol. 167(1989) 3–27.
[21] A.A. Tuffery, Light and Electron microscopy of the sheath of a blue-green alga, J.
Gen. Microbiol. 57 (1969) 41–50.
[22] V.A. Shepherd, M.J. Beilby, The effect of an extracellular mucilage on the
response to osmotic shock in the charophyte alga Lamprothamnium
papulosum, J. Membr. Biol. 170 (1999) 229–242.
[23] P. Whiteman, Quantitative measurements of Alcian Blue-glycosaminoglycan
complexes, Biochem. J. 131 (1973) 343–350.
[24] J.F. McMANUS, Histological demonstration of mucin after periodic acid, Nature
158 (1946) 202.
[25] P. Greenspan, E.P. Mayer, S.D. Fowler, Nile red: a selective ﬂuorescent strain for
intracellular lipid droplets, J. Cell Biol. 100 (1985) 965–973.
[26] C. Kitazaki, S. Numano, A. Takanezawa, T. Nishizawa, M. Shirai, M. Asayama,
Characterization of lysis of the multicellular cyanobacterium Limnothrix/
Pseudanabaena sp. strain ABRG5-3, Biosci. Biotechnol. Biochem. 77 (2013)
2339–2347.
[27] G. Breuer, P.P. Lamers, D.E. Martens, R.B. Draaisma, R.H. Wijffels, The impact ofnitrogen starvation on the dynamics of triacylglycerol accumulation in nine
microalgae strains, Bioresour. Technol. 124 (2012) 217–226.
[28] S. Hondo, M. Takahashi, T. Osanai, M. Matsuda, T. Hasunuma, A. Tazuke, Y.
Nakahira, S. Chohnan, M. Hasegawa, M. Asayama, Genetic engineering andmetabolite proﬁling for overproduction of polyhydroxybutyrate incyanobacteria, J. Biosci. Bioeng. 120 (2015) 510–517.
[29] S. Yoshida, M. Takahashi, A. Ikeda, H. Fukuda, C. Kitazaki, M. Asayama,
Overproduction and easy recovery of biofuels from engineered cyanobacteria,
autolyzing multicellular cells, J. Biochem. 157 (2015) 519–527.
[30] M.K. Okajima, T. Bamba, Y. Kaneso, K. Hirata, E. Fukusaki, S. Kajiyama, T.
Kaneko, Supergiant ampholytic sugar chains with imbalanced charge ratio
form saline ultra-absorbent hydrogels, Macromolecules 41 (2008) 4061–4064.
[31] A. Sato, R. Matsumura, N. Hoshino, M. Tsuzuki, N. Sato, Responsibility of
regulatory gene expression and repressed protein synthesis for triacylglycerol
accumulation on sulfur-starvation in Chlamydomonas reinhardtii, Front. Plant
Sci. 5 (444) (2014) 1–12.
[32] K. Hirai, T. Hayashi, Y. Hasegawa, A. Sato, M. Tsuzuki, N. Sato, Hyperosmosis
and its combination with nutrient-limitation are novel environmental
stressors for induction of triacylglycerol accumulation in cells of Chlorella
kessleri, Sci. Rep. 6 (25825) (2016) 1–12.
[33] C.H. Hung, K. Kanehara, Y. Nakamura, Isolation and characterization of amutant defective in triacylglycerol accumulation in nitrogen-starved
Chlamydomonas reinhardtii, Biochim. Biophys. Acta 186 (2016) 1282–1293.
[34] S. Ota, K. Oshima, T. Yamazaki, S. Kim, Z. Yu, M. Yoshihara, K. Takeda, T.
Takeshita, A. Hirata, K. Bišová, V. Zachleder, M. Hattori, S. Kawano, Highly
efﬁcient lipid production in the green alga Parachlorella kessleri: draft genome
and transcriptome endorsed by whole-cell 3D ultrastructure, Biotechnol.
Boifuels 9 (2016) 13.
[35] T. Takeshita, S. Ota, T. Yamazaki, A. Hirata, V. Zachleder, S. Kawano, Starch andlipid accumulation in eight strains of six Chlorella species under comparatively
high light intensity and aeration culture conditions, Bioresour. Technol. 158(2014) 127–134.
[36] G.R. Stansell, V.M. Gray, S.D. Sym, Microalgal fatty acid composition:
implications for biodiesel quality, J. Appl. Phycol. 24 (2012) 791–801.
[37] S.J. Lee, B.D. Yoon, H.M. Oh, Rapid method for the determination of lipid fromthe green alga Botryococcus braunii, Biotechnol. Techniq. 12 (1998) 553–556.
[38] T. Tanoi, M. Kawachi, M.M. Watanabe, Effect of carbon source on growth andmorphology of Botoryococcus brauni, J. Appl. Phycol. 23 (2011) 25–33.
[39] T. Marudhupandi, R. Sathishkumar, T.T.A. Kumar, Heterotrophic cultivation of
Nannochloropsis salina for enhancing biomass and lipid production,
Biotechnol. Rep. 10 (2016) 8–16.
 ska, J. Tys, Enrichment of Parachlorella kessleri biomass[40] A. Piasecka, I. Krzeminwith bioproducts: oil and protein by utilization of beet molasses, J. Appl.
Phycol. 29 (2017) 1735–1743.
[41] F. Daboussi, S. Leduc, A. Maréchal, G. Dubois, V. Guyot, C. Perez-Michaut, A.
Amato, A. Falciatore, A. Juillerat, M. Beurdeley, D.F. Voytas, L. Cavarec, P.
Duchateau, Genome engineering empowers the diatom Phaeodactylum
tricornutum for biotechnology, Nat. Commun. 29 (2014) 3831–3837.
[42] C.H. Hung, M.Y. Ho, K. Kanehara, Y. Nakamura, Functional study of
diacylglycerol acyltransferase type 2 family in Chlamydomonas reinhardtii,
FEBS Lett. 587 (2013) 2364–2370.
[43] A. Vonshak, S. Boussiba, A. Abeliovich, A. Richmond, Production of Spirulina
biomass: maintenance of monoalgal culture outdoors, Biotechnol. Bioeng. 25(1983) 341–349.
[44] N. Murata, S. Takahashi, Y. Nishiyama, S.I. Allakhverdiev, Photoinhibition ofphotosystem II under environmental stress, Biochim. Biophys. Acta 1767
(2007) 414–421.

Table S1. Sasaki et al.

Table S1. Lipid composition in the BX1.5 cells.
BG11

−N

−P

−N−P

6 days6 days6 days6 days

C16:0

21.9±0.02

22.7±0.11

23.8±0.05

20.3±0.38

C16:1

1.0±0.05

0.8±0.15

0.7±0.02

0.6±0.01

C16:2

2.5±0.15

1.0±0.01

1.1±0.05

1.1±0.02

C16:3

0.8±0.04

0.9±0.02

1.1±0.02

0.9±0.05

C18:0

5.5±0.23

7.8±0.08

7.1±0.06

6.5±0.19

C18:1

13.0±0.52

17.9±0.18

19.5±0.08

18.3±1.89

C18:2

45.2±0.33

32.1±0.05

39.0±0.09

43.6±1.26

C18:3

10.0±0.23

16.9±0.14

7.7±0.148.8±0.10

FAMEs

Lipid composition as FAMEs in the BX1.5 cells cultivated in
the BG11, BG11−N, BG11−P, and BG11−N−P medium shownin Figure 7 was evaluated and displayed as percentage values
with standard deviations.

Fig. S1. Sasaki et al.

2% CO2

0.04% CO2

A

BG11

BG11

–N

–P

–N–P

6
days

B

3.5
2% BG112% -N
2% -P
2% -N-P
0.04% BG11

Absorbance (OD)

3.0
2.5
2.0
1.5
1.0

0.5
0.0
300

400

500
600
700
Wavelength (nm)

800

900

Fig. S1.
Culture profile and absorption spectrum. (A) Cells were
harvested on the 6th day under induction cultivation conditions, as
shown in Fig. 2A. (B) This sample was directly subjected to analysis byabsorption spectrophotometry. The main peak at OD674 for chl a and
OD730 for cell turbidity measurement (Fig. 2B) are shown with arrows.

Fig. S2. Sasaki et al.

Cell size (μm)

A

Chl a (μg/mL)

B

3
2.5
2
1.5
1
0.5
0

pH

C

0 6 110 6 110 6 110 6 110 6 11

0.04%

2%

0.04%

2%

0.04%

BG11

−N

0 6 11 days
2% CO2−P

Fig. S2. Cell size, chlorophyll a, and pH of BX1.5. (A) Diameters of thecells cultivated under the supply of 2% CO2-air shown in Fig. S1 andS3, and that with standard deviations from the photography profiles areshown in Fig. 4. Sample numbers: BG11, n = 28; BG11–N, n = 13, and
BG11–P, n = 19, respectively. (B) The amount of chlorophyll a from thecell culture in panel A was measured by the methanol-extraction method[17]. (C) pH of the cell culture shown in Fig. 2 was sequentiallymeasured for 0, 6, and 11 days under the induction cultivation condition
supplying with 2% CO2-air or under the standard condition of 0.04%
CO2-air.

Fig. S3. Sasaki et al.

6 days

0.04%
CO2

BG11

BG11

−N
2%
CO2
−P

−N−P

Fig. S3. Cells were harvested on the 6th day under induction
cultivation condition as shown in Fig. 2A or Fig. S1. The samples were
subjected to microscopic observations without (left) or with (right)
India-ink stain. More details are described in the text.

Relative values (CSS)

Fig. S4. Sasaki et al.

BG11
−N
−P

Fig. S4. Cell resistance to respective stresses. Relative cell-spot
signal intensities (CSS) for the cell resistance on the BG11 plate,shown in Fig. 8B, were evaluated by a densitometry image analyzer(Bio1D Optical Density: Vilber Lourmat, Wielandstrasse, Germany) andpresented as a profile with relative values. More details are describedin the text.