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Algal Research
journal homepage: www.elsevier.com/locate/algal
Demonstration of the potential of Picochlorum soloecismus as a microalgal
platform for the production of renewable fuels
C. Raul Gonzalez-Esquer
a
, Kimberly T. Wright
a
, Nilusha Sudasinghe
a
, Carol K. Carr
a
,
Claire K. Sanders
a
, Aiko Turmo
b
, Cheryl A. Kerfeld
b,c,d,e
, Scott Twary
a
, Taraka Dale
a,⁎
a
Bioscience Division, Bioenergy and Biome Sciences Group, MS M888, Los Alamos National Laboratory, Los Alamos, NM 87545, United States of America
b
Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824, United States of America
c
MSU-DOE Plant Research Laboratory, Michigan State University, East Lansing, MI 48824, United States of America
d
Environmental Genomics and Systems Biology and Molecular Biophysics and Integrated Bioimaging Divisions, Lawrence Berkeley National Laboratory, Berkeley, CA
947206, United States of America
e
Berkeley Synthetic Biology Institute, Berkeley, CA 94720, United States of America
ARTICLE INFO
Keywords:
Picochlorum
Biofuel feedstock
Halotolerant
Genetic engineering
Genome analysis
ABSTRACT
Photosynthetic microbes are considered promising biofactories for transforming inorganic carbon from the at-
mosphere into a renewable source of chemicals and precursors of industrial interest; however, there continues to
be a need for strains that demonstrate high productivity, environmental robustness, and the potential to be
genetically manipulated. Genome sequencing and biochemical characterization of promising culture collection
microalgae strains, as well as the isolation of previously unidentified strains from the environment or mixed
cultures, bring us closer to the goal of decreasing the cost-per-gallon of algal biofuels by identifying new and
promising potential production strains. The halotolerant alga Picochlorum soloecismus was isolated from the
culture collection strain, Nannochloropsis salina CCMP 1776. Here, we show that P. soloecismus accumulates
moderate levels of fatty acids and high levels of total carbohydrates and that it can effectively grow in a range of
salinities. In addition, we make use of its sequenced genome to compare it to other biofuel production platforms
and to validate the capacity for engineering this strain's genome. Our work shows that Picochlorum soloecismus is
a candidate production strain for the generation of renewable bioproducts.
1. Introduction
Through the utilization of CO
2
as a carbon source, photosynthetic
microbes such microalgae and cyanobacteria have been recognized as
suitable candidate feedstocks for renewable chemical production. Algae
may overcome some environmental challenges faced by traditional
plant-based feedstocks, in that algae have greater rates of biomass
production than plant crops and do not have to compete with food
crops for arable land [1–4]. Furthermore, algae can utilize various
marginal water sources (brackish, waste, or seawater) [5] and can be
leveraged to utilize fertilizer runoffbefore it reaches important water
reservoirs. While the most traditional algae components for fuels are
intracellular lipids and fatty acids for biodiesel [4,6,7], certain micro-
algal storage carbohydrates (including starch) can also be used for the
production of biofuels or bioproducts [8,9]. Moreover, there are on-
going efforts toward valorizing the protein fraction of the biomass, ei-
ther for food, feed, or chemicals [10–12].
In spite of the above potential, challenges for sustainable commer-
cial-scale production of algae commodities remain [13], including a
continued need for further increases in strain productivity and en-
vironmental robustness. The characterization of microalgae strain
phenomes, genomes, and commensurate gene expression patterns per-
mits a deeper understanding of growth, carbon storage, and environ-
mental tolerance [14,15], while the development of molecular tools
will permit the manipulation of these phenotypes in order to further
reduce the costs of renewable products from algae [16,17].
The Picochlorum genus has received much attention recently
[18–23], as new strains have recently been identified and character-
ized, adding to the relatively few examples of Picochlorum from pre-
vious years [24–29]. Picochlorum strains have faster exponential growth
rates than other commonly used microalgae (i.e. Dunaliella, Nanno-
chloropsis)[23,30], are broadly halotolerant [19,31], can withstand
temperatures ranging from 0 to 40 °C [21,32] and can accumulate
20–58% lipids on a dry weight basis [28,30,33]. The analysis of newly
https://doi.org/10.1016/j.algal.2019.101658
Received 23 January 2019; Received in revised form 30 August 2019; Accepted 4 September 2019
⁎
Corresponding author.
E-mail address: tdale@lanl.gov (T. Dale).
Algal Research 43 (2019) 101658
2211-9264/ © 2019 The Authors. 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/).
T
generated Picochlorum genomes has shed light on the potential mole-
cular mechanisms behinds the genus' environmental stress resilience
[18,22], which may translate into approaches for improved outdoor
large-scale growth. We have described the genome of Picochlorum so-
loecismus (previously Picochlorum sp. [26]orPicochlorum soloecismus
DOE101 [34]) and suggested it could be a strain of interest for pro-
duction, due to its lipid accumulation and ability to be transformed
[26]. Here we expand the biochemical characterization of this strain to
include the carbohydrate and protein fractions, examine its salinity
tolerance, conduct an in silico analysis of the strain's metabolic po-
tential, and demonstrate an improved method for genetic engineering.
Taken together, the inherent features of P. soloecismus described here
demonstrate its potential as a biotechnological production platform.
2. Materials and methods
2.1. Strain and culture conditions
The strain Picochlorum soloecismus (previously known as Picochlorum
sp. strain DOE101) was isolated at Los Alamos National Laboratory, Los
Alamos, NM from a culture of Nannochloropsis salina CCMP 1776. P.
soloecismus was grown on a modified version of f/2 media [35,36],
which contained: 0.75 g NaNO
3
, 0.035 g NaH
2
PO
4
·H
2
O, 3.15 mg
FeCl
3
·6H
2
O, 4.36 mg Na
2
EDTA·2H
2
O, 10 μg CuSO
3
·5H
2
O, 6 μg
Na
2
MoO
4
·2H
2
O, 22 μg ZnSO
4
·7H
2
O, 10 μg CoCl
2
·6H
2
O, 180 μg
MnCl
2
·4H
2
O, 1 μg vitamin B
12
,1μg biotin, 200 μg thiamine HCl and
38 g Instant Ocean™(I.O.) seawater salt mix (Blacksburg, VA) per liter
(hereafter f/2). Stock cultures were maintained at 20 °C and 10 μmoles
photons m
−2
s
−1
. For solid media preparation, f/2 was adjusted to
19 g/l of I.O. seawater mix and supplemented with 1.5% phytoagar
(Gold Bio, St. Louis, MO) as the solidifying agent.
2.2. Light and Transmission Electron Microscopy
30-ml cultures were grown in Erlenmeyer flasks at 1% CO
2
, 16:8
light:dark cycle, 300 μmoles photons m
−2
s
−1
of light intensity, 25 °C
and shaken at 130 rpm. For light microscopy, a 1-ml aliquot of cells at
OD
750 nm
≈0.7 were spun down at 775 rcf for 5 min, concentrated and
visualized using an Olympus BX51 fluorescence microscope (Tokyo,
Japan). Time courses of cell diameters (n = 150 per time point) were
measured in micrographs using ImageJ software [37] and cell counts
(n = 3 per time point) were measured by hemocytometer. For Trans-
mission Electron Microscopy, cells were harvested at OD
750 nm
≈0.4
(one day of growth) and OD
750
≈3.2 (four days of growth). Each 2-ml
aliquot was spun down at 775 rcf for 5 min. The pellet was resuspended
in 2.5% glutaraldehyde in 0.1 M Millonig's buffer, pH 7.2, overnight at
4 °C. The cell pellet was washed two times with 100 mM phosphate
buffer (pH 7.2), embedded in 1% agarose, and secondary fixed with 1%
osmium tetroxide in 50 mM Na-PO
4
buffer for 2 h. The cell pellet was
then washed three times with H
2
O. Additionally, the pellets were wa-
shed with 2% uranyl acetate for one hour and washed with H
2
O three
times. Afterwards, the cell pellet was dehydrated in increasing acetone
series (acetone % in water: 30, 50, 70, 80, 90, 95, 3 × 100). Finally, the
cells were infiltrated in epoxy resin (Spurr's resin, firm mixture), and
70 μm-thin sections were post-stained with uranyl acetate/lead citrate
as described previously [38] and visualized on a JEOL (Tokyo, Japan)
JEM 100CX II transmission electron microscope equipped with a Gatan
(Lakewood Ranch, FL) Orium SC200–830 charge-coupled device
camera.
2.3. Baseline growth
For wild type growth determination, cells were grown in triplicate
in f/2 media, magnetically stirred and supplemented with CO
2
on-de-
mand to maintain a pH = 8.25, in 2.8 liter Erlenmeyer flasks under a
16:8 light:dark cycle, at 25 °C and outfitted with surrounding white LED
lights set at 800 μmoles photons m
−2
s
−1
(hereafter “spin”flask).
Samples were collected daily, 2 h after the beginning of the light period,
through day 13 of the culture. Nitrogen in cell-free media (culture su-
pernatant after centrifugation) was quantified by ion chromatography
using a Dionex ICS-100 (Sunnyvale, CA) as described previously [25].
From day 13 to day 24 of the culture, three additional samples were
removed. Optical density was measured at the wavelength of 750 nm
using a Varian Cary 300 Bio UV–Visible Spectrophotometer (Palo Alto,
CA). For the measurement of dense aliquots, cells were diluted with
media in order to fall within the linear range of the spectrophotometer.
2.4. Biochemical characterization
Total fatty acid content was determined as fatty acid methyl esters
(FAMEs) by gas chromatography coupled with flame ionization detec-
tion (GC/FID) according to Van Wychen et al. [39]. Acid-catalyzed
transesterification to quantify the total fatty acid content in the biomass
was performed by treating 5–10 mg of freeze-dried biomass containing
25 μL of 10 mg/mL methyl tridecanoate (C13:0ME) as the internal
standard with 200 μL of chloroform:methanol (2:1, v/v) and 300 μLof
0.6 M HCl:methanol. The samples were heated at 85 °C for 1 h and
FAMEs were back extracted with 1 ml of hexane. The hexane extracts
were analyzed with an Agilent 7890A Series GC/FID. 2-μl injections at a
10:1 split ratio were loaded onto a DB-WAX column (30 m length x
0.25 mm inner diameter x 0.25 μmfilm thickness, (Agilent Technolo-
gies, Santa Clara, CA)). Helium was used as the carrier gas at a flow rate
of 1 ml/min. Initial column temperature was 100 °C. Then, the tem-
perature was ramped to 200 °C at 25 °C/min and held for 1 min and
again ramped to 242 °C at 1.5 °C/min and held for 1 min (35 min total).
The inlet and the detector temperatures were 250 and 280 °C, respec-
tively. Chromatographic signals were compared with those of the GLC
461C 30-component FAME standard mix (Nu-Chek Prep, Inc., Elysian,
MN), and FAME quantification was performed using C13:0ME as the
internal standard.
Total carbohydrates (including starch and non-storage carbohy-
drates) were quantified by 3-methyl-2-benzothiazolinone hydrazine
(MBTH) method according to Van Wychen et al. [40]. The method
involved a two-step sulfuric acid hydrolysis to break down carbohy-
drates into their monomeric subunits, followed by quantification of the
monosaccharides by spectrophotometry in a Varian Cary 300 Bio
UV–Visible Spectrophotometer. Briefly, ~25 mg of freeze-dried biomass
was first hydrolyzed with 250 μL of 72% (w/w) sulfuric acid. The hy-
drolysate was diluted to a concentration of 4% (w/w) sulfuric acid with
deionized in a water and autoclaved at 121 °C for 1 h. The solids were
separated from the acidic hydrolysate by filtration and the mono-
saccharides were quantified using D (+) glucose as a calibration
standard.
Protein content was determined by measuring the elemental ni-
trogen content in freeze-dried biomass using a Thermo Fisher
(Waltham, MA) FlashEA 1112 elemental analyzer and multiplying the
nitrogen content by a nitrogen-to-protein conversion factor of 4.78
[41].
Biochemical composition was normalized to dry weight (lyophilized
biomass), except on the salinity tolerance experiment. In this case, the
biomolecule composition was normalized to ash-free dry weight
(AFDW), performed according to Van Wychen [42].
2.5. Salinity tolerance
To assess the natural salinity tolerance of P. soloecismus, the cultures
were grown in triplicate in f/2 media with varying salinities, prepared
by adjusting the amount of dissolved I.O. seawater salt mix to the fol-
lowing levels: 1%, 10%, 50%, 100%, 150%, 200%, and 400%. “100%”
is the salinity of Instant Ocean™when made per the manufacturer's
instructions, which equals 35 parts per thousand (ppt). The actual
salinities of each media variation were acquired using a YSI Pro30
C.R. Gonzalez-Esquer, et al. Algal Research 43 (2019) 101658
2
Conductivity-Salinity Instrument (Yellow Springs, OH) and determined
to be: 1% ≈0.35 ppt; 10% ≈3.5 ppt; 50% ≈17.5 ppt; 100% ≈35 ppt;
150% ≈52.5 ppt; 200% ≈70 ppt; 400% ≈140 ppt. 30 ml of salinity-
adjusted media in 125 ml Erlenmeyer flasks were inoculated with P.
soloecismus conditioned for growth in 35 ppt salinity f/2 media. Flasks
were grown in 1% CO
2
under a 16:8 light:dark cycle, 300 μmoles
photons m
−2
s
−1
of light intensity, 25 °C and shaken at 130 rpm using a
Benchmark Orbi-Shaker™(Sayreville, NJ). Samples for optical density
(750 nm) measurements were collected daily 2 h after the start of the
light period and were measured using a Biotek Synergy H4 Hybrid
Microplate Reader (Winooski, VT) and diluted accordingly to fall
within the linear range of the plate reader. Nitrate levels in the media
were monitored by EMD Millipore MQuant Nitrate Test Strips
(Burlington, MA). Cells were harvested 10 days after nitrogen depletion
and freeze-dried prior to biochemical composition analysis.
Growth rates (per day) were determined by calculating the ln
(OD
750
) for each time point and graphing those data as a function of
time. The linear part of the curve was fit using linear regression and the
slope noted as the growth rate per day. The growth rate of each bio-
logical replicate was calculated separately, with the exception of
140 ppt which grew too poorly to calculate. For 0.35 ppt salinity, the
linear portion of the curve was found to be after a lag of 10–13 days,
depending on the replicate. The remaining salinities showed lag times
of 1–2 days. The statistical significance of pairwise comparisons of each
salt condition was determined using Tukey's multiple comparisons test
(post-hoc analysis of a one-way ANOVA).
2.6. Bioinformatic analysis
Genome sequence data and protein predictions from Picochlorum
soloecismus [34], Picochlorum SENEW3 [43], Chlamydomonas reinhardtii
[44] and Chlorella sorokiniana UTEX 1230 [45] were obtained from the
Greenhouse website (https://greenhouse.lanl.gov). Gene/protein
homologs were identified through BLAST search [46] by comparison
with known homologs from C. reinhardtii using the pre-set parameters
from the Greenhouse Blast tool and a cutoffvalue of < 1e-55 for po-
sitive homology. Protein domains were identified through Pfam search
[47], through batch extraction using the pre-set parameters at the Pfam
database website (http://pfam.xfam.org).
2.7. Vector design
Two plasmids were used for P. soloecismus transformation. The first
plasmid consisted of a dual gene expression vector (Pico-Dual) in which
two fluorescent protein genes were each expressed as a fusion (via the
2A peptide [48] as shown previously for C. reinhardtiii [49]) to a re-
sistance marker gene (sh-Ble for Zeocin antibiotic resistance and neo for
G418 antibiotic resistance) (Supplementary Figs. 6A and 7). The
mCherry-2A-Neo fusion gene was controlled by the RbcS promoter/
terminator pair and the GFP-2A-shBle fusion by a native nitrate re-
ductase (NR) promoter/terminator pair. The second plasmid (Pico-Zeo)
was a modified version from the pTY100 plasmid in Unkefer et al. [26]
consisting of a synthetic fusion formed by the P. soloecismus Diglyceride
Acyltransferase (DGAT1) gene, 2A peptide and sh-ble gene under the
control of the native NADP-dependent glyceraldehyde-3-phosphate
dehydrogenase (GAPDH) promoter/NR terminator pair (Supplementary
Figs. 6B and 7). Synthetic genes were not codon optimized, as the codon
usage table for P. soloecismus does not show any specific codon bias
(Supplementary Table 1).
2.8. Electroporation
P. soloecismus was grown in Erlenmeyer flasks in 1% CO
2
under a
16:8 light:dark cycle, 300 μmoles photons m
−2
s
−1
of light intensity,
25 °C and shaken at 130 rpm using a Benchmark Orbi-Shaker™. A 50 ml
aliquot at early stationary phase was diluted to OD
750
= 0.3–0.5 with
fresh media and grown overnight. Afterwards, the culture was spun
down at 1500 rcf for 5 min on a tabletop centrifuge. The cell pellet was
washed two times with sterile 375 mM ice-cold sorbitol and re-
suspended in 5 ml of 375 mM cold sorbitol. The cell suspension was
kept on ice until electroporation. For transformation, 250 μl of the cell
suspension were mixed with ~1 μg linearized plasmid DNA and placed
it in a pre-chilled 4 mM gap electroporation cuvette. The DNA/cell mix
was pulsed three times (~10 s between each pulse) in a Biorad
(Hercules, CA) Gene Pulser Xcell using the program setting “time con-
stant”with the following settings: 20 ms pulse, 1600 V. Electroporated
cells were incubated on ice for 5 min and then resuspended in 5 ml f/2
at room temperature in a culture tube. After an overnight incubation
(20 μmoles photons m
−2
s
−1
, 20 °C in shaker), cells were spun down at
1500 rcf for 5 min and resuspended in 500 μl of f/2 media. The resulting
cell suspension was spread on three f/2 solid agar plates supplemented
with antibiotic at the appropriate concentration and incubated at 60
μmoles photons m
−2
s
−1
. Colonies appeared on the agar plates after
2–3 weeks. To determine the ideal antibiotic marker, kill curves were
performed by spotting 100 μl of a cell suspension of known cell number
into f/2 solid media supplemented with various antibiotic concentra-
tions (0, 10, 20, 40, 60, 80 and 100 μg/ml of paromomycin, G418 and
zeocin, respectively). After the aliquot dried, the plates were incubated
in 60 μmoles photons m
−2
s
−1
for one week to determine growth ca-
pacity.
2.9. Mutant screening
DNA from a loop-full of cells grown on solid f/2 media plus anti-
biotic was extracted using the Zymo Research Quick-DNA Fungal/
Bacterial Kit (Irvine, CA) following the manufacturer's protocol. PCR
screening was performed as described previously [50] using the fol-
lowing primer pair combination not found in P. soloecismus wildtype
genome: for Pico-Dual: GFP fwd primer (5′-GTGCAGCTGGCCGACCAC
TACCAG-3′) and NR term rev (5′-CGATAGCACGCTTCTGAAGCTTGCA
TGC-3′); for Pico-Zeo: GAPDH fwd primer (5′-GTTTGTGGTATGATGT
AAGGCAGTCGTC-3′) and DGAT rev (5′-CAGTCGCGCAATATCTCCGA
CCGAG-3′). The identity of the PCR amplicons was confirmed by Sanger
sequencing through Genewiz (South Plainfield, NJ). For western blot,
50-ml aliquots from cells at OD
750
≈1 were spun down and pellets
were resuspended in 60 μl of NuPAGE LDS sample buffer. These were
heated to 96 °C for 5 min. Afterwards, samples were centrifuged at 8610
rcf for 20 min. Supernatants were run on a NuPAGE 4–12% Bis/Tris gel
using the MES-SDS buffer system from Thermo Fisher. The resulting gel
was blotted on nitrocellulose membrane and the western blot was
performed following Biorad's general protocol for western blotting
(Bulletin 6376). We identified the GFP protein in Pico-Dual using
mouse anti-GFP Antibody (B-2) conjugated to HRP from Santa Cruz
Biotechnology (Dallas, TX) and developed using the 1-Step TMB-Blot-
ting Substrate Solution from Thermo Fisher.
3. Results and discussion
3.1. Morphology of Picochlorum soloecismus
P. soloecismus is a small unicellular coccoid microalgae (Fig. 1).
Imaging analysis of bright field micrographs using ImageJ [37] re-
vealed that the cells have a diameter of 2.2 ± 0.3 μm under N replete
conditions and 4.4 ± 0.5 μm 11 days after N =0 (Supplementary
Fig. 1A), and morphologically, this strain correlates well with other
Picochlorum species [14,31]; for example, cells generally present a
single chloroplast with no pyrenoids. Cells from two different time
points in growth were examined by Transmission Electron Microscopy.
There were clear ultrastructural feature changes observed between the
samples, with cellular inclusions occupying a greater portion of the
cellular volume in the later phase of the culture. The single chloroplast
accumulates starch granules over time, while lipid bodies accumulate in
C.R. Gonzalez-Esquer, et al. Algal Research 43 (2019) 101658
3
the cytoplasm (Fig. 2 and Supplementary Fig. 2), consistent with the
physiological changes observed in other green algae upon nitrogen
depletion [51-53].
3.2. Growth and biochemical composition of Picochlorum soloecismus
When cultivated under a 16:8 light:dark cycle (800 μmoles photons
m
−2
s
−1
) with pH controlled by CO
2
delivery on-demand, Picochlorum
soloecismus doubled exponentially during the first four days, at a growth
rate of 0.77 day
−1
(Fig. 3, Supplementary Table 3).
Although the nitrogen in the medium depleted on day 7 of culti-
vation, the optical density of the cultures continued to increase until
day 12 and OD
750
≈14. Cell counts compared to optical density in-
dicate that the cells stop dividing upon nitrogen depletion
(Supplementary Fig. 1). After N depletion the cells increase in diameter
up through Day 17, which may in part explain the continued increase in
OD
750
from Day 7–12. (Supplementary Fig. 1). Also, carbohydrate and
fatty acid levels began to increase before the media was completely
depleted of N, reaching 30% and 10% of dry biomass weight (DW) for
carbohydrates and fatty acid methyl esters (FAMEs), respectively, by
day 7. These data indicate that P. soloecismus begins to store carbon in
the late replete phase, which may be leveraged in a production scenario
to shift biochemical composition while maintaining biomass pro-
ductivity. Also, because the carbon storage response begins when ni-
trate levels are between 40 and 60 ppm (Fig. 3), it may be that P. so-
loecismus can detect when nitrate concentrations in the medium have
declined to a certain level or possibly detect the rate of nitrogen de-
pletion, which may be a different response than other strains whose
carbon storage response begins when (or shortly after) the nitrogen
concentration in the media reaches zero [51-54].
Overall, carbohydrates were observed to be the main carbon storage
compound, which peaked at 38% of DW by day 11 and decreased once
the culture reached stationary phase. Fatty acids (measured as FAMEs)
increased over time and leveled at 22% DW, whereas protein content
decreased to 12% DW by day 20 (Fig. 2). Taken together, these results
show that Picochlorum soloecismus can grow rapidly to high cell den-
sities and can also accumulate 51% ( ± 2.2; days 8–24) of biomass dry
weight in carbon storage molecules (FAMEs + carbohydrates) after
nitrogen depletion (Supplementary Fig. 3).
The fatty acid profile of Picochlorum soloecismus also varied as the
culture progressed. Fatty acids in nitrogen replete cultures were mainly
unsaturated (the major fatty acid at N replete was γ-linolenic acid with
40.7% ± 10 of total fatty acids) but shifted to saturated fatty acids after
nitrogen depletion (the major fatty acid at N deplete was palmitic acid
with 37.6% ± 12.4) (Supplementary Table 2). A similar change was
observed previously in the marine strain Nannochloropsis oceanica [55].
Since fatty acid chain lengths and the degree of unsaturation have an
effect on the resulting biodiesel properties (i.e. the Cetane Number
[56,57]), knowing the progression the fatty acid profile in P. soloecismus
could be useful for the optimization of biomass harvesting, especially
for balancing maximum biomass collection with desired feedstock
Fig. 1. Representative bright field image of Picochlorum soloecismus. Scale bar:
5μm.
Fig. 2. Cell morphology of Picochlorum soloecismus. Transmission electron micrographs depicting the change in ultrastructural features from an A) early stage culture
to a B) later stage culture. White arrowheads: Thylakoids; Black arrowhead: Starch granules; Black arrow: Lipid body. Scale bar: 200 nm.
C.R. Gonzalez-Esquer, et al. Algal Research 43 (2019) 101658
4
characteristics.
3.3. Effect of salinity on P. soloecismus
Picochlorum strains are robust and capable under a variety of growth
conditions [19,21,28,30–33]. However, other factors –such as water
usage and nutrient recycling–must be considered for commercial fea-
sibility [13]. P. soloecismus was isolated from a marine culture and
grows on a seawater-based medium; thus, it does not compete with
freshwater for human consumption or traditional agriculture. None-
theless, at large-scale in open systems, broad changes in salt con-
centration may occur due to water evaporation, precipitation, or vo-
lume replenishing after biomass harvesting [58,59]. Therefore, we
assessed the native tolerance of P. soloecismus to changes in salinity,
testing a range from 0.35 to 140 ppt. Apart from any adaptation that
may have occurred during the 13 day experiment, these results describe
the tolerance to “shocking”the culture with a variety of salinities.
Fig. 4A shows the growth of P. soloecismus in seven different sali-
nities, and Fig. 4B shows growth rates calculated during exponential
growth (Supplementary Fig. 4) after the respective lag for each condi-
tion, except 140 ppt (rate values and statistics in Supplementary
Table 3). Although OD
750
appeared to increase exponentially for days
3–5 for the 0.35 ppt and 140 ppt cultures, the overall optical densities
for these cultures were so low that we did not calculate growth rates for
these days. For the 0.35 ppt we noted a small improvement in growth at
the end of the time course (after a lag of 10–13 days), which was used to
calculate the growth rate. For the 140 ppt culture, we deemed the
growth to be insufficient to calculate a rate. At 17.5 ppt, P. soloecismus
grew similarly to the 35 ppt culture. Notably, even though this strain
has been considered a marine strain, P. soloecismus also adjusted quickly
to the 3.5 ppt medium and grew rapidly after a short lag. Both of these
results indicate that P. soloecismus can thrive on brackish waters, in-
cluding sources with quite low salinity. Growth in conditions of sali-
nities above average seawater salinity (52.5 and 70 ppt) also demon-
strated a short lag relative to the 35 ppt culture, as well as a
compromise in end-point biomass accumulation. For example, for the
52.5 and 70 ppt cultures, the maximum OD
750
was 84% and 77%, re-
spectively, of the average maximum OD
750
of the 0.35–3.5 ppt cultures
at day 10. However, once the extra day of lag was overcome, the growth
rates at 52.5 and 70 ppt were similar to 35 ppt. Thus, apart from some
small differences in culture lag and more notable compromises in total
biomass accumulation for higher salinities, P. soloecismus growth rates
were found to be similar from 3.5 ppt to 70 ppt (not statistically dif-
ferent, Supplementary Table 3).
Fig. 3. Time course of growth and biochemical composition of P. soloecismus. Cells were grown in “spin flasks”in f/2 media at 35 ppt salinity (see Methods), 25°C,
16:8 light:dark cycle and 800 μmoles photons m
−2
s
−1
. Data shown corresponds to the mean of triplicate cultures; error bars are the standard deviation of the mean.
Fig. 4. Comparison of P. soloecismus growth in dif-
ferent salinity concentrations, upon shifting from a
starting concentration of 35 ppt (100% Instant Ocean
seawater mix in f/2, ppt = parts per thousand) to
each salinity shown in the legend. A) Growth ob-
served as an increase in optical density (750 nm), B)
Growth rates at the exponential phase for each sali-
nity except 140 ppt, which grew too poorly to cal-
culate a growth rate. Cells were grown in Erlenmeyer
flasks in a growth chamber at 1% CO
2
and 25 °C
under a 16:8 light:dark cycle, 300 μmoles photons
m
−2
s
−1
of light intensity. Data shown corresponds
to the mean of triplicate cultures; error bars are the
standard deviation of the mean.
C.R. Gonzalez-Esquer, et al. Algal Research 43 (2019) 101658
5
Interestingly, while a relatively tight range of performance was
observed between 3.5 and 70 ppt, a large drop in growth was observed
both from 3.5 ppt to 0.35 ppt and from 70 ppt to 140 ppt (Fig. 4A).
Notably, after 10 days of incubation, the 0.35 ppt culture began to
grow, although the growth rate was significantly lower than the 35 ppt
growth rate (p < 0.001, Supplementary Table 3). Meanwhile, the
140 ppt culture did not increase in OD during the time frame of the
experiment. Overall, these results show that P. soloecismus is broadly
halotolerant, including brackish waters as well as hypersaline water (up
to twice the salinity of average seawater), consistent with the haloto-
lerant nature observed in other Picochlorum strains [27,43] and sug-
gesting that changes in the salinity of outdoor ponds should not greatly
affect culture growth.
While varying salinity did not have a major effect on growth, it
could have provoked changes in the biochemical composition (quality)
of the biomass. As expected, the increase in salinity correlates with an
increase in ash content (Fig. 5A), which can affect the quality of bio-
crude oil from algae [60], and also affects the proximate analysis.
Therefore, to quantify lipids and carbohydrates in the biomass for each
salinity (collected 10 days after nitrogen depletion for each flask), we
normalized each value to Ash-Free Dry Weight (AFWD). Carbohydrates
reached a maximum at 35 ppt (again, 100% Instant Ocean salts in f/2
media), however maximum lipids accumulated at a salinity of 3.5 ppt
(10% Instant Ocean salts in f/2 media) (Fig. 5B). Overall, the accu-
mulation of lipid and carbohydrates in these experiments accounted
for > 60% of ash-corrected biomass, showing that while variation of
salinity appears to shift the proportion of lipids and carbohydrates,
there was still a large fraction of the biomass accounted for as carbon
storage molecules.
3.4. Protein domain comparison and analysis of Picochlorum soloecismus
We made use of P. soloecismus' sequenced genome to assess its dis-
tinctiveness, relevance, and metabolic potential as a microalgal bio-
factory [61-63]. To do so, we compared protein domains as PFams [47],
because these represent the essential building blocks of proteins and
therefore, metabolism. We tallied and compared the Pfam domains of P.
soloecismus [34] to the model strain Chlamydomonas reinhardtii [44],the
potential production strain Chlorella sorokiniana UTEX 1230 [45] and
the close relative Picochlorum SENEW3 [43]. Over 2000 protein do-
mains (2134) were common to the four strains, while 28 were unique to
P. soloecismus (Fig. 6 and Supplementary Data; expression data can be
found at https://greenhouse.lanl.gov and at [26]). Many of these unique
domains relate to stress response; i.e. Pfam14678 and Pfam14680
(within the NSC_04251 gene) are found in Fanconi anemia group I
proteins, which participate in the repair of DNA double-strand breaks
[64]. Other domains may be involved in regulatory mechanisms, such
as Pfam15279 in NSC_04882 (SOBP; a protein carrying a zinc-finger
domain); zinc-finger regulatory proteins have been the target of en-
gineering to increase carbon partitioning toward lipids [65].
Through the same Pfam search, we also identified several genes
related to the CO
2
-concentrating mechanism (CCM) machinery [66].
We identified one Ribulose-1,5-bisphosphate carboxylase/oxygenase
(RuBisCO) large subunit (NSC_06630; Pfam Pfam02788 and
Pfam00016) and two small subunits (NSC_01826 and NSC_05239;
Pfam00101): one α‑carbonic anhydrase (CA) (NSC_04175; Pfam00194)
and two β-CA (NSC_06162 and NSC_01359; Pfam00484). In addition,
by homology search to the C. reinhardtii proteins, we predict the ex-
istence of one γ-CA (NSC_01478, containing Pfam00132), three trans-
porters homologous to the HLA3 periplasmic bicarbonate pump
(NSC_05588, NSC_00126 and NSC_04805, which contain Pfam00664
and Pfam00005) and a homolog to the chloroplast bicarbonate pump
LciA (NSC_02673 containing Pfam01226). Interestingly, searches for an
EPYC1 homolog (pyrenoid-assembly protein) following the analysis
method outlined in Mackinder et al. [67] were unsuccessful. However,
this is in accordance with the light and electron micrographs (Figs. 1
and 2, and Supplementary Fig. 2), where pyrenoids are seemingly ab-
sent from cells.
Multiple C. reinhardtii homologs involved in starch metabolism [68]
were identified. Among them, the starch synthases NSC_06319 (SSS1),
NSC_06600 (SSS4), NSC_05568, and NSC_05724; the Glucose-1-phos-
phate adenylyltransferases NSC_04450 (STA1), NSC_04684 (STA2) and
NSC_06341 (STA6), and the 4-alpha-glucanotransferase NSC_02924
(STA11). Similarly, we identified various Diglyceride Acyltransferases
Fig. 5. Composition of P. soloecismus biomass in different salinity concentrations
harvested 10 days after N depletion. A) Ash content per dry weight; B) Percent
carbohydrates and lipids in biomass at varied salinities, normalized to ash-free
dry weight (AFDW). Data shown corresponds to the mean of triplicate cultures;
error bars are the standard deviation of the mean.
Fig. 6. Comparison of the abundance of protein domains (Pfams) between P.
soloecismus and the model strains C. sorokiniana UTEX1230, C. reinhardtii and a
close relative P. SENEW3. Details on the domain tallies can be found in the
supplementary materials.
C.R. Gonzalez-Esquer, et al. Algal Research 43 (2019) 101658
6
(DGAT), which are essential for triglyceride production [69]: the type-1
DGAT NSC_05425 and the type-2 DGAT NSC_01244 and NSC_01696.
Notably absent from the P. soloecismus genome were known domain
components of the canonical RNAi machinery [70], such as the DICER
dimerization domain (Pfam03368), Pfam16486 (N-term Argonaut),
Pfam02170 (PAZ) and Pfam02171 (PIWI). It has been previously ob-
served that some algae with genomes sizes between 12.5 and 21.9 Mb
lack the RNAi machinery [70], consistent with the genome size of P.
soloecismus, which is 15.2 Mbp [34].
Overall, P. soloecismus is less complex than traditional microalgae
platforms, such as C. reinhardtii. On one hand, this simplicity may be
unfavorable; for example, an absent RNAi mechanism may restrict
options for artificially fine-tuning gene expression. However, in some
cases, relatively simple genomes are superior. Reduced gene re-
dundancy may be easier to overcome when creating gene deletion
mutants; similarly, competing metabolism is reduced when expressing
heterologous pathways. Therefore, P. soloecismus holds great potential
for genetic improvement.
3.5. Gene overexpression as a proof-of-concept for genetic engineering
capability
Prior transformation efforts of P. soloecismus required enzymatic
pre-treatment of cell walls prior to DNA delivery and achieved an ef-
ficiency of ~10
−7
[26]. Consequently, we set to develop a simple yet
robust P. soloecismus transformation method that does not require cell
wall digestion. First, we tested the sensitivity of P. soloecismus to three
commonly used antibiotics. Strong growth inhibition was obtained with
Zeocin (< 40 μg/ml), but much higher concentrations were required for
cell death when using G418 or paromomycin (> 100 μg/ml) (Supple-
mentary Fig. 5). Linearized Pico-Dual (Supplementary Figs. 6 and 7)
was delivered into P. soloecismus by electroporation. Putative transfor-
mant colonies appeared on f/2 selective media at the concentrations of
40 μg/ml of Zeocin or 150 μg/ml of G418. These colonies were screened
by PCR by amplifying a 1.1-Kbp region between the Green Fluorescent
Protein (GFP) gene and Nitrate Reductase (NR) reductase terminator
(Fig. 7A), which does not occur in wild type P. soloecismus. Similarly,
for method confirmation, the Pico-Zeo vector (Supplementary Figs. 6
and 7) was transformed as above and mutants were screened by am-
plifying a 1-kb region between the GAPDH promoter and the DGAT1
(Fig. 7B). In both cases, PCR positive and false positive colonies were
identified. Amplicons of the expected size were sequenced and con-
firmed as positive. These results show the introduction of the over-
expression construct into P. soloecismus. Transformation efficiency was
similar to previously described transformation methods, without the
need for cell wall pre-treatment. Afterwards, to corroborate transgene
expression of the GFP-2A-ShBle gene, we identified the GFP protein by
Western Blot on one out of four strains tested. We observed cross-re-
activity of the anti-GFP antibody with a protein of approximately
28 kDa, while no interaction was observed on wild type (Fig. 7C).
Therefore, we demonstrate here that enzymatic treatment can be
avoided and that traditional electroporation is a convenient method
that can be utilized for P. soloecismus transformation.
4. Conclusions
In this work we show that Picochlorum soloecismus has the capacity
to grow to high optical densities and accumulate moderate levels of
fatty acids and high levels of carbohydrates. This strain also maintains
relatively fast growth rates when challenged with broad salinity con-
centration changes in its growth media, and these salinity changes can
affect the biochemical composition of algal biomass, but does not
compromise the total level of carbon storage molecules. We have de-
monstrated that P. soloecismus can be genetically engineered and that its
genetic background is favorable for many metabolic adjustments. While
protein overexpression is shown here to be feasible, further research is
required to add elements to its molecular toolbox (i.e. gene repression
or deletion). Thus, in this work, we demonstrate P. soloecismus to be a
resilient strain that holds great potential as a platform for production of
biofuels and bioproducts.
Acknowledgements
C.R.G-E, K.W., N.S., C.S. and T.D. acknowledge funding provided by
Fig. 7. Confirmation of P. soloecismus transformation by electroporation. A) PCR screening of 28 antibiotic resistance colonies transformed with Pico-Dual vector; B)
PCR amplification of 10 colonies transformed with Pico-Zeo vector; C) Western Blot for the identification of GFP-positive strains after Pico-Dual DNA delivery. Red
arrow: Expected size of the PCR amplicon. Green arrow: Expected size of GFP. (For interpretation of the references to color in this figure legend, the reader is referred
to the web version of this article.)
C.R. Gonzalez-Esquer, et al. Algal Research 43 (2019) 101658
7
the U.S. Department of Energy Bioenergy Technologies Office for con-
tract NL0025841, and S.T. acknowledge funding from U.S. Department
of Energy Bioenergy Technologies Office (DE-EE0003046) given to the
National Alliance for Advanced Biofuels and Bioproducts. A.T and
C.A.K. acknowledge support from the Office of Science of the U.S.
Department of Energy DE-FG02-91ER20021 with infrastructure support
from MSU AgBio Research. K.W. would like to thank funding from the
Department of Energy Science Undergraduate Laboratory Internship
(SULI) program. This work has been authored by an employee of Triad
National Security, LLC, operator of the Los Alamos National Laboratory
under Contract No.89233218CNA000001 with the U.S. Department of
Energy. The United States Government retains and the publisher, by
accepting this work for publication, acknowledges that the United
States Government retains a nonexclusive, paid-up, irrevocable, world-
wide license to publish or reproduce this work, or allow others to do so
for United States Government purposes.
Author contributions
C.R.G-E and K.W. contributed to physiological characterization and
transformation experiments. N.S., C.K.C and C.S. contributed to the
biochemical composition analysis. A.T. and C.K contributed with
Transmission Electron Microscopy experiments. S.T. contributed to
strain isolation and identification. T.D. contributed with the experi-
mental design, statistical expertise, and obtaining of funding. All au-
thors contributed to data analysis, the writing of the article, and critical
revisions. The authors declare no conflict of interest. No conflicts, in-
formed consent, or human or animal rights are applicable to this study
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://
doi.org/10.1016/j.algal.2019.101658.
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