Download Introducing extra NADPH consumption ability significantly increases

Metabolic Engineering 38 (2016) 217–227
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Metabolic Engineering
journal homepage: www.elsevier.com/locate/ymben
Original Research Article
Introducing extra NADPH consumption ability significantly increases
the photosynthetic efficiency and biomass production of cyanobacteria
Jie Zhou a,1, Fuliang Zhang a,b,1, Hengkai Meng a,c, Yanping Zhang a, Yin Li a,n
a
CAS Key Laboratory of Microbial Physiological and Metabolic Engineering, Institute of Microbiology, Chinese Academy of Sciences, Beijing, China
University of Chinese Academy of Sciences, Beijing, China
c
School of Life Sciences, University of Science and Technology of China, Hefei, China
b
art ic l e i nf o
a b s t r a c t
Article history:
Received 5 April 2016
Received in revised form
28 June 2016
Accepted 4 August 2016
Available online 4 August 2016
Increasing photosynthetic efficiency is crucial to increasing biomass production to meet the growing
demands for food and energy. Previous theoretical arithmetic analysis suggests that the light reactions
and dark reactions are imperfectly coupled due to shortage of ATP supply, or accumulation of NADPH.
Here we hypothesized that solely increasing NADPH consumption might improve the coupling of light
reactions and dark reactions, thereby increasing the photosynthetic efficiency and biomass production.
To test this hypothesis, an NADPH consumption pathway was constructed in cyanobacterium Synechocystis sp. PCC 6803. The resulting extra NADPH-consuming mutant grew much faster and achieved a
higher biomass concentration. Analyses of photosynthesis characteristics showed the activities of photosystem II and photosystem I and the light saturation point of the NADPH-consuming mutant all significantly increased. Thus, we demonstrated that introducing extra NADPH consumption ability is a
promising strategy to increase photosynthetic efficiency and to enable utilization of high-intensity lights.
& 2016 International Metabolic Engineering Society. Published by Elsevier Inc. All rights reserved.
Keywords:
Photosynthetic efficiency
Light/dark reactions cycle
Energy balance
Extra NADPH consumption
Metabolic engineering
1. Introduction
The continuing decline of arable land and the constantly
growing population pose significant challenges to the food supply.
It is estimated that global food production must achieve a 50%
increase by 2030 to meet the global food demand (Covshoff and
Hibberd, 2012; Singh et al., 2014). Photosynthesis is one of the
fundamental components determining crop productivity and is
perhaps the most important biochemical process on Earth. It
converts solar energy into biochemical energy, while converting
CO2 into organic compounds. Improving photosynthesis efficiency
would significantly contribute to increasing the productivity of
crops and other photosynthetic organisms, i.e. microalgae and
cyanobacteria, which have shown great potential in production of
fuels and chemicals from CO2 (Evans, 2013; Oliver and Atsumi,
2014; Stephenson et al., 2011; Xiong et al., 2015a).
Improving photosynthetic efficiency is always of great interest
in spite of its challenging nature (Evans, 2013; von Caemmerer and
Evans, 2010). On the energy conversion side, researchers have
been working on the optimization of light reactions by expanding
the photosynthetically active radiation spectrum (Blankenship and
n
Correspondence to: Institute of Microbiology, Chinese Academy of Sciences, No.
1 Beichen West Road, Chaoyang District, Beijing 100101, China.
E-mail address: [email protected] (Y. Li).
1
Contributed equally.
Chen, 2013; Gan et al., 2014) and optimizing light-harvesting antenna complexes (LHC) (Blankenship and Chen, 2013; Melis, 2009;
Ort et al., 2011; Work et al., 2012). On the carbon fixation side,
increasing the catalytic efficiency of ribulose-1,5-bisphosphate
carboxylase/oxygenase (Rubisco) has been the most challenging
task (Khan, 2007). Increasing the catalytic efficiency of Rubisco
through molecular engineering (Cai et al., 2014; Liu et al., 2010)
and replacing the native Rubisco in tobacco with a Rubisco from
cyanobacteria (Lin et al., 2014) are recent exciting examples. Alternative strategies to increase carbon fixation include introducing
CO2 concentrating mechanisms, increasing the regeneration of ribulose-1, 5-bisphosphate to accelerate the Calvin cycle and reducing or engineering photorespiration (Covshoff and Hibberd, 2012;
Kebeish et al., 2007; Shih et al., 2014).
Photosynthetic reactions are usually divided into light reactions
and dark reactions; however, light reactions and dark reactions are
not absolutely separated. Light energy is converted into chemical
energy, in the form of ATP and reducing equivalent NADPH, via a
series of photochemical reactions during light reactions. During
dark reactions, CO2 is reduced into organic compounds via the
Calvin cycle at the expense of the ATP and NADPH that are generated from the light reactions. The generation and consumption
of ATP and NADPH are therefore the key for coupling the light and
dark reactions. Theoretical analysis showed that 2.57 ATP/2
NADPH are generated in the light reactions via linear electron
transport (Fig. 1), whereas 3 ATP/2 NADPH are required for CO2
http://dx.doi.org/10.1016/j.ymben.2016.08.002
1096-7176/& 2016 International Metabolic Engineering Society. Published by Elsevier Inc. All rights reserved.
218
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
Fig. 1. Diagram of ATP and NADPH-coupled production in the light reaction and the NADPH consumption pathway-isopropanol synthetic pathway constructed in S. 6803.
The isopropanol synthetic pathway is an acetyl-CoA-dependent pathway. From acetyl-CoA to isopropanol, three exogenous enzymes were required: coenzyme A transferase
(CTFAB, UniProt accession number P33752 and P23673) and acetoacetate decarboxylase (ADC, UniProt accession number P23670), which were obtained from Clostridium
acetobutylicum DSM 1731 to convert acetoacetyl-CoA to acetone; and secondary alcohol dehydrogenase (sADH, UniProt accession number P25984) from Clostridium beijerinckii to convert acetone to isopropanol.
fixation in the dark reactions (Kramer and Evans, 2011; Marcus
et al., 2011). This indicates that the ATP generated in light reactions
(2.57 ATP) is insufficient to meet the energy requirement (3 ATP)
for CO2 fixation in dark reactions. Such an imbalanced energy
supply/requirement between the light and dark reactions might be
one of the rate-limiting factors affecting photosynthetic efficiency
(Kramer and Evans, 2011; Marcus et al., 2011).
Considering from another perspective, if the ATP generated
from the light reactions met precisely the demand of ATP in the
dark reactions, the expected ATP/NADPH ratio in light reactions
would be 3 ATP/2.33 NADPH, as ATP generation is tightly coupled
with NADPH production via linear electron transport in the light
reactions (Kramer and Evans, 2011). This would result in an
NADPH imbalance, as only 2 NADPH will be consumed in dark
reactions. We therefore hypothesized that increasing consumption
of NADPH generated from the light reactions might improve the
coupling between the light and dark reactions, thus increasing the
photosynthetic efficiency. This hypothesis was tested by introducing an NADPH-consuming pathway into an oxygenic, photosynthetic, prokaryote cyanobacterium, Synechocystis sp. PCC 6803
(hereafter termed as S. 6803), with the aim to better balance the
ATP/NADPH between light and dark reactions. A series of biochemical analyses demonstrated that this approach significantly
increased the photosynthetic efficiency and biomass production of
S. 6803.
2. Materials and methods
2.1. Strains and growth conditions
The strains used are listed in Table S1. Escherichia coli strain
DH5a was used as the host for vector construction. Wild-type S.
6803 and its mutants were grown in BG11 medium at 30 °C at an
illumination intensity of approximately 100 μmol photons/m2/s, as
described previously (Nielsen et al., 2013; Zhou et al., 2012).
Chloromycetin (10 μg/ml) and/or kanamycin (10 μg/ml) were added to the medium when necessary.
2.2. Construction of vectors
The plasmids used and constructed in this work are listed in
Table S2. Vectors were constructed to delete the phaCE and pta
genes or to express the ctfAB, adc and sadh genes in S. 6803. All
primers used are listed in Table S3.
Plasmid pSM5 was constructed by inserting the Pcpc560-sadh
expression cassette, which comprised the promoter Pcpc560 and the
structural gene sadh, into the Bam HI site of plasmid pSM1 (Zhou
et al., 2012). The Pcpc560-sadh expression cassette was synthesized
by GENEWIZ. Inc. (China) and the nucleotide sequence of sadh was
optimized for the preferred codon usage of S. 6803.
Plasmid pSM6 was constructed by inserting the Pcpc560-ctfAB
and PrbcL-adc expression cassettes into the Xho I site of pSM2
(Zhou et al., 2012). The Pcpc560-ctfAB and PrbcL-adc expression
cassette were synthesized by GENEWIZ. Inc. (China) and the nucleotide sequences of the Pcpc560-ctfAB and PrbcL-adc were optimized for the preferred codon usage of S. 6803.
2.3. Construction of mutant strains
Mutants of S. 6803 were constructed by transforming S. 6803
with plasmids listed in Table S2. Transformations were performed
as previously described (Lindberg et al., 2010). All constructed
strains are listed in Table S1. Briefly, strain SM6 was constructed by
integration of plasmid pSM6 at the pta locus of SM2 (Zhou et al.,
2012) via double crossover homologous recombination. Strain SM7
was constructed by integration of plasmids pSM5 and pSM6 at the
phaCE and pta loci of S. 6803, respectively. Strain SM7* was constructed by integration of plasmids pSM5 at the phaCE loci of S.
6803.
2.4. Enzyme assays
Crude cell extracts were prepared as previously described
(Zhou et al., 2014c), with some modifications. S. 6803 wild-type
and mutant cells grown to mid-exponential growth phase were
collected by centrifugation (14,000g, 4 °C, 2 min) and washed with
prechilled buffer. The cells were then disrupted using the silicon
carbide disruption method (Zhou et al., 2014a) in prechilled buffer
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
for the enzyme assay described below. The Bradford method (Zhou
et al., 2014c) was used to determine the total protein in the crude
cell extracts.
The specific activity of CoA transferase (CTFAB) was determined
by monitoring the decrease in absorbance at 310 nm, which corresponded to the disappearance of acetoacetyl-CoA, as previously
described (Clark et al., 1989), using a spectrophotometer (Specter
MAX 190, Molecular Devices, CA, USA). The reaction mixtures
contained 110 mM Tris chloride (pH 7.5), 5.5% (vol/vol) glycerol, 20
mMMgCl2, 0.1 mM acetoacetyl-CoA, 0.32 M potassium acetate, and
1-5 μg of protein in the crude cell extracts.
The specific activity assay for acetoacetate decarboxylase (ADC)
was performed as described previously (Kusakabe et al., 2013). The
reaction mixtures contained 30 mM sodium acetate buffer (pH
5.8), 30 mM lithium acetoacetate and 1-5 μg protein in the crude
cell extracts. The consumption of lithium acetoacetate was monitored at 270 nm using a spectrophotometer.
The specific activity assay for secondary alcohol dehydrogenase
(SADH) was carried out as previously described (Hanai et al.,
2007). The total reaction mixture contained 50 mM Tris-HCI (pH
7.5), 1 mM dithiothreitol, 0.2 mM NADPH, crude cell extracts (1–
5 μg protein) and 6.7 mM acetone. The consumption of NADPH
was determined by measuring the decrease in absorbance at
340 nm.
219
molecule, propidium iodide, which can permeate the cellular
membranes of dead cells and bind to nucleic acids, as previously
described (Riccardi and Nicoletti, 2006). Samples were analyzed
using a flow cytometer (FACSCalibur, Becton Dickinson, San Jose,
CA, USA) equipped with a laser emitting at 488 nm and an optical
filter FL1 (530/30 nm). The collected data were analyzed using
FlowJo software (Tree Star, San Carlos, CA, USA).
2.8. Carbon fixation rate determination
To calculate the net CO2 uptake rate by each strain, the initial
and remnant CO2 concentrations after cultivation were measured
as previously described (Xiong et al., 2015b) with modification.
Briefly, cells in the exponential growth phase were harvested,
washed once with BG11, resuspended in 130 ml BG11 at OD730 of
0.2 (approximately 3 mg dry cell weight), and cultivated for an
additional 2 h. Then 130 ml cell suspension was transferred to a
sealed tube with a volume of 140 ml. The total inorganic carbon of
BG11 medium and the culture after two hours cultivation was
released by addition of concentrated HCl (Xiong et al., 2015b). The
CO2 in 1 ml headspace gas was quantified through the RTX-QBOND capillary column (30 m; 0.32 mm inside diameter; 10 mm
film thickness; RESTEK Corporation, Bellefonte, USA) using the GC2014 gas chromatograph (Shimadzu, Kyoto, Japan) equipped with
a TCD detector (Gong et al., 2015).
2.5. Isopropanol production assay
2.9. Chlorophyll fluorescence of PS II analysis
For isopropanol production assay, the wild-type S. 6803 and its
mutants were grown in 250 ml flasks containing 50 ml BG11
medium with an initial OD730 of 0.4. The incubation was carried
out in a shaking incubator (30 °C, 200 rpm) under constant illumination intensity of 100 μmol photons/m2/s for about 3 weeks.
Growth was monitored by measuring the OD at 730 nm every
2 days.
To quantify the isopropanol produced, 0.5 μL of culture supernatant was analyzed using a GC-2014 gas chromatograph (Shimadzu, Kyoto, Japan) equipped with a flame ionization detector
and an Rtx-WAX capillary column (30 m, 0.25 mm inside diameter,
0.25 mm film thickness; Agilent Technologies, Santa Clara, CA, USA)
as previously described (Kusakabe et al., 2013). Isopropanol was
used as the internal standard.
To determine the quantum efficiency of PS II, chlorophyll
fluorescence analysis of S. 6803 and its mutants grown to exponential growth phase was performed on a Dual-PAM-100 instrument (Walz, Germany), according to the manufacturer's instructions. To exclude the effect of phycobiliprotein, a red light
detector (RD) was used. After 4 min of dark adaptation, slow induced curve and light response curve of cell suspension at an
OD730 of 3.5 were measured to get parameters of chlorophyll
fluorescence kinetics, including the maximum quantum yield (Fv/
Fm), the effective quantum yield (Y(II) ¼Fv’/Fm’) and the relative
electron transport rate (rETR(II)) of PS II. The intensity of measure
light and saturation pulse were 0.1 μmol/m2/s and 5000 μmol/m2/
s, respectively.
2.6. NADPH and ATP Assays
2.10. Measurement of P700 signal
Wild-type Synechocystis and mutants cells grown to exponential growth phase were collected by centrifugation (14,000g,
4 °C, 2 min) and washed with prechilled PBS buffer. The cells were
then lyzed using the lysis buffer associated with the Enhanced ATP
Assay Kit, S0027 (Beyotime Biotechnology, Shanghai, China).
The NADPH content was determined by using an NADP/NADPH
Quantification Colorimetric Kit (Biovision, Catalog-K347-100), according to the manufacturer's instructions. The concentration of
NADPH was calculated according to an NADPH standard curve and
expressed as pmol/OD730.
The ATP content was determined by using an Enhanced ATP
Assay Kit, S0027 (Beyotime Biotechnology, Shanghai, China) according to the manufacturer's instructions. The concentration of
ATP was calculated according to an ATP standard curve and expressed as nmol/OD730.
To determine the quantum efficiency of PS I, the redox state of
the reaction center chlorophyll of PS I (P700) was determined from
its absorbance change in the near-infrared (wavelength 870–
830 nm) simultaneously with the measurement of chlorophyll
fluorescence of PS II, using a Dual-PAM-100 instrument (Walz,
Germany), as previously described (Klughammer and Schreiber,
2008; Wientjes and Croce, 2012). After 4 min of dark adaptation,
slow induced curve and light response curve of cell suspension at
an OD730 of 3.5 were measured to obtain PS I maximum quantum
yield (Pv/Pm), PS I effective quantum yield (Y(I) ¼ Pv’/Fm’), the relative electron transport rate (rETR(I)) via PS I, and PS I complementary quantum yields of non-photochemical energy dissipation, Y(ND) and Y(NA). The intensity of measure light and saturation pulse were 0.1 μmol/m2/s and 5000 μmol/m2/s, respectively. Far-red light of 820 nm was used to measure the absorbance
and far-red light of 875 nm was used as control as previously
described (Klughammer and Schreiber, 2008; Pfündel et al., 2008).
2.7. Flow cytometry
Flow cytometry was used to analyze the ratio of dead cells in
the cultures of S. 6803 and its mutants. Cells grown to stationary
phase were collected, washed once and resuspended in 1 ml of
PBS at 1–2 106 cells/ml. Cells were stained with a fluorescent
2.11. Oxygen evolution
The oxygen evolution rate of all strains was measured using a
LED1/W-1503illuminator (Hansatech) and an Oxygraph Plus Clark-
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J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
type electrode (Hansatech) as previously described (Gonzalez-Esquer et al., 2015) with modification. To determine the light response curves of oxygen evolution rate, cultures grown to exponential growth phase were harvested and resuspended in BG11
at an OD730 of 6.0. Two microliter cell suspension was supplemented with 100 mM bicarbonate and the steady state rate of
oxygen evolution was determined at different light intensity at
30 °C for 2 min
3. Results
3.1. Design and introduction of extra NADPH consumption ability
into S. 6803
Two criteria need to be considered when designing the experiments. One is that such perturbation should directly target the
consumption of NADPH to drive the coupling of the light and dark
reactions, while minimally affecting the cellular physiology. The
other consideration is that such perturbation should be sufficiently
strong to disturb the existing ATP/NADPH ratio in S. 6803, which
has been established through long-term evolution and supposes to
be quite stable.
To meet the first criterion, we proposed to introduce a synthetic pathway that only consumes NADPH but does not affect
energy metabolism. The pathway selected was the isopropanol
biosynthetic pathway, which does not exist in S. 6803, but is found
in some heterotrophic microbes, such as Clostridium beijerinckii
(Dai et al., 2012; Ismaiel et al., 1993). To connect the heterogeneous
isopropanol biosynthetic pathway with the central metabolism of
S. 6803, three exogenous enzymes, namely ctfAB, encoding coenzyme A transferase and adc, encoding acetoacetate decarboxylase
from Clostridium acetobutylicum DSM 1731 (Bao et al., 2011), and
sadh, encoding secondary alcohol dehydrogenase from C. beijerinckii NRRLB593 (Dai et al., 2012), need to be introduced (Fig. 1).
The assembled isopropanol pathway would connect with central
metabolism at the acetoacetyl-CoA node, which is already present
in S. 6803. This pathway only consumes NADPH and does not
consume ATP as the secondary alcohol dehydrogenase is solely
dependent on NADPH (Dai et al., 2012).
We calculated the ΔG of the proposed isopropanol biosynthetic
pathway (from acetyl-CoA to isopropanol). The ΔG is 67.6 KJ/
mol, suggesting this pathway is thermodynamically highly favorable as a decarboxylation reaction is included in the pathway. In
addition, we planned to use a strong promoter that was developed
in a previous study (Zhou et al., 2014c) to express the genes involved in this pathway, so as the second criterion can be met.
To introduce a functional isopropanol biosynthetic pathway, we
constructed two mutants, SM6 with two genes (ctfAB þadc), which
was expected to produce acetone only and SM7 with three genes
(ctfABþ adc þsadh), which was expected to produce isopropanol at
the expense of NADPH. As indicated in Fig. 2A, mutant SM6 was
constructed by placing the codon-optimized adc and ctfAB genes
under a promoter PrbcL (Takeshima et al., 1994) and a strong promoter Pcpc560 (Zhou et al., 2014c), respectively. Because knockout
pta gene would not completely block acetate production, this DNA
fragment was inserted into the pta site of the mutant SM2 (Zhou
et al., 2012). Mutant SM7 was constructed by placing the codon-
Fig. 2. The genetic modifications in SM strains for extra NADPH consumption. (A) Genetic modifications. (B) Whole cell PCR using primers for the recombinant cassette at the
phaCE and pta sites; one primer which is 100 bp upstream the recombinant cassette and the other primer which is located in the recombinant cassette were used to confirm
the complete segregation of all mutants. (C) Whole cell PCR with specific primers demonstrating the integration of each gene into the chromosome of individual mutants of
S. 6803.
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
221
Fig. 3. Growth profile analysis of all strains and the time course of isopropanol production by NADPH consumption strain SM7. (A) Growth profile. (B) Cell death rate analysis
by flow cytometry. (C) CO2 fixation rate. (D) Time course of isopropanol production by strain SM7. Error bars indicate standard deviation (SD) of the data from three
independent experiments. For each experiment, three technical replicates were performed.
optimized sadh under Pcpc560, and the Pcpc560-sadh was inserted
into the phaCE site of mutant SM6. Therefore, the sole genomic
difference between mutant SM7 and its direct control mutant SM6
was that the phaCE gene (Zhou et al., 2012) in mutant SM6 was
replaced with the Cmr gene, whereas in mutant SM7, the phaCE
gene was replaced with the Cmr gene and sadh.
Complete segregation and gene insertions were verified by PCR
and sequencing (Fig. 2B and C). Metabolite spectrum analysis indicated that mutant SM6 produced 108.5 mg/L acetone, while
mutant SM7 produced 226.9 7 5.4 mg/L isopropanol, when incubated in BG11 medium at 30 °C under constant white light for
20 days (Figs. 2A and 3D). Enzyme assays showed that the specific
activities of CTFAB, ADC and SADH in mutant SM7 were 8.6 72.4,
140727 and 528 779 μmol/min/mg crude extract, respectively.
These results demonstrated that the introduced isopropanol
pathway was functional. Subsequently, a series of physiological
and biochemical analyses were performed to investigate whether
the introduced isopropanol biosynthetic pathway increased the
photosynthetic efficiency.
3.2. Introduction of extra NADPH consumption ability increased the
ATP/NADPH ratio
First, the intracellular concentrations of the two photochemical
products of the light reactions, ATP and NADPH, were determined
in the wild-type (WT), SM6, and SM7, respectively (Table 1).
Concentration of NADPH in strain SM7 was the lowest among all
the strains tested, which was approximately 40% lower than that
in control strain SM6 and WT. Moreover, the concentration of ATP
in strain SM7 was 77% and 59% higher than that in strain SM6 and
WT, respectively. Taken together, the ATP/NADPH ratio of strain
SM7 (0.55) was significant higher than that of SM6 (0.18) and WT
Table 1
Intracellular concentration of NADPH and ATP in WT and mutant cells of S. 6803.
Strain
NADPH (pmol/OD730)
ATP (nmol/OD730)
WT
SM6
SM7
193.5 7 4.27
190.83 7 7.06
112.83 7 3.94
39.28 7 1.39
35.23 7 1.63
62.53 7 0.89
WT, wild-type; Standard deviation (7 SD) of the data from three independent
experiments. For each experiment, three technical replicates were performed.
(0.20). These data suggested that the introduced synthetic isopropanol pathway not only decreased the intracellular NADPH
level, but also increased intracellular ATP level, therefore resulted
in a significantly increased ATP/NADPH ratio in strain SM7.
3.3. Introduction of extra NADPH consumption ability increased the
biomass production of S. 6803
Growth curves of different mutants were assessed under the
same growth conditions as those described previously (Fig. 3A).
Strain SM7 grew significantly faster than the other strains. At the
first day of cultivation, the growth rate of mutant SM7 was 81%,
42%, and 33% higher than that of the WT strain, mutant SM2 (in
which the pta gene was replaced with the Kmr gene and phaCE
gene was replaced with the Cmr gene) (Zhou et al., 2012), and
mutant SM6, respectively (Fig. S1). At 20 days of cultivation, the
dry cell concentration of strain SM7 was 84%, 62%, and 48% higher
than that of strains WT, SM2, and SM6, respectively (Table S4).
To exclude the possibility that the high OD730 and dry cell
weight of mutant SM7 was resulted from an increased amount of
dead cells, the ratio of dead cells over total cells in each culture
was determined by flow cytometry. The results showed that strain
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J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
SM7 had the least dead cells among the four strains tested during
the whole culture process (Fig. 3B). Notably, the ratio of dead cells
in the cultures of the acetone-producing mutant SM6, the control
strains SM2, and the WT, all increased sharply after 20 days of
cultivation. Rather, the ratio of dead cells in the culture of the
NADPH-consuming strain SM7 only increased slightly after 20
days of cultivation, while the isopropanol concentration also
ceased to increase (Fig. 3D). These suggest that increasing NADPH
consumption can increase biomass production and cellular activity. Furthermore, the carbon-fixation rates of all strains were determined according to the amount of net CO2 uptake during a
period of cultivation (Xiong et al., 2015b). Fig. 3C showed that the
carbon-fixation rate of the NADPH-consuming strain SM7 was
38%, 40%, and 36% higher than that of the control strains WT, SM2
and SM6, respectively.
3.4. Introduction of extra NADPH consumption ability increased
photochemical efficiency of PS II in S. 6803
Light reactions take place in photosystem II (PS II, P680 as the
reaction-center) and photosystem I (PS I, P700 as the reactioncenter). The photochemical efficiency is dependent on the activity
of both PS II and PS I. Chlorophyll fluorescence kinetics indicates
the activity of PS II (Shen and Luo, 2011; Singh et al., 2005; Thamatrakoln et al., 2013). To investigate the effect of introducing
extra NADPH consumption ability on the activity of PS II, parameters of chlorophyll fluorescence kinetics, including the maximum quantum yield (Fv/Fm), the effective quantum yield (Y(II) ¼
Fv’/Fm’), and the relative electron transport rate (rETR(II)) of PS II,
were determined (Table 2). Fv/Fm of all strains were approximately 0.42, showing there was no significant difference among
the maximum quantum yield of PS II of strain SM7 and its control
strains WT, SM2 and SM6. Due to state transitions, the cyanobacterial Fv/Fm is not as accurate as the Fv/Fm of higher plants to
reflect the maximum quantum yield of PS II (Campbell et al., 1998).
Therefore, the Y(II) and rETR(II) of each strain were measured
under normal light condition of 100 μmol/m2/s (Table 2). Y(II) and
rETR(II) of strain SM7 were 16.2% and 15.8% higher than that of
control strains WT, SM2 and SM6, respectively, indicating that
extra NADPH consumption increased the activity of PS II.
To further analyze the effect of higher light intensity on NADPH
consumption strain SM7 and its control strains WT, SM2 and SM6,
the light response curve of rETR(II) was investigated (Fig. 4B). The
rETR(II) of strain SM7 was significant higher than that of its control
strains WT, SM2 and SM6 under the tested light intensity (ranging
from 142 to 2292 μmol photons/m2/s). Interestingly, the rETR(II) of
all control strains reached the highest value (approximately 21)
under 588 μmol photons/m2/s; whereas the rETR(II) of SM7 kept
increasing and reached the highest value (38.68 71.45) under
1182 μmol photons/m2/s. This means that the saturation light
points of all control strains were around 600 μmol photons/m2/s,
whereas the saturation light point of SM7 increased to 1200 μmol
photons/m2/s. Introducing extra NADPH consumption ability thus
resulted in a significant increase of the saturation light point and a
1.84 fold increase of the highest rETR(II) value (rETR(II) of strain
SM7 under 1182 μmol photons/m2/s versus rETR(II) of control
strains under 588 μmol photons/m2/s). Notably, the rETR(II) of all
control strains decreased to approximately 10 under 2292 μmol
photons/m2/s, whereas the rETR(II) of strain SM7 was still significantly higher than the highest rETR(II) of all control strains.
Subsequently, the light response curves of Y(II) of all strains
were plotted under different light intensity (Fig. S2). The Y(II) of
SM7 was 1.8-fold and 2.4-fold that of the control strains WT, SM2
and SM6 under 588 μmol photons/m2/s and 1182 μmol photons/
m2/s, respectively (Fig. S2). Moreover, the Y(II) of all control strains
decreased to nearly 0 under 2292 μmol photons/m2/s, whereas
strain SM7 still exhibited an effective quantum yield, which was
higher than that of all control strains under 1182 μmol photons/
m2/s. These suggested that introducing extra NADPH consumption
ability not only increased the photosynthetic efficiency of PS II, but
also enabled strain SM7 to utilize light with higher intensity.
Photosynthetic oxygen is another sensitive indicator of photosynthetic activity of PS II, and an increased oxygen evolution may
indicate the increased activity of the light reactions (Xiong et al.,
2015b). Light response curve of rETR(II) showed that the saturation
light point of SM7 was approximately 1200 μmol photons/m2/s.
We therefore investigated the effect of introducing extra NADPH
consumption ability on photosynthetic oxygen evolution under
600 μmol photons/m2/s, the semi-light saturation point of SM7.
Fig. 4A shows that there are no significant difference among the
oxygen evolution rate of strain WT, SM2, and SM6, under
600 μmol photons/m2/s. Interestingly, the oxygen evolution rates
of strain SM7 were approximately 86% higher than the rest of the
strains. This shows that strain SM7 exhibits a significantly higher
photosynthetic oxygen evolution activity, indicating the light reactions of strain SM7 was improved.
3.5. Introduction of extra NADPH consumption ability contributed to
energy conversion of PS I in S. 6803
To further investigate the effects of introducing extra NADPH
consumption ability on energy conversion efficiency of PS I, the
maximum quantum yield (Pv/Pm), the effective quantum yield Y
(I), and relative electron transport rate rETR(I) were analyzed
(Klughammer and Schreiber, 2008). Pv/Pm of all strains were approximately 0.94, showing there were no significant difference
among the quantum yield of strain SM7 and control strains WT,
SM2 and SM6. However, Y(I) and rETR(I) of strain SM7 were approximately 14.9% and 15.6% higher than the Y(I) and rETR(I) of
control strains WT, SM2 and SM6, respectively, under normal light
condition of 100 μmol/m2/s (Table S5).
Similarly, we investigated the light intensity response curve of
Table 2
Measurement of chlorophyll fluorescence kinetics of PS II, Fv/Fm and Y(II), rETR(II) of NADPH consumption strain (SM7) and control strains (SM6, SM2 and WT) under
different light intensity.
Light intensity (μmol/m2/s)
Parameter
WT
0
100
100
588
588
Fv/Fm
Y(II)
rETR(II)
Y(II)
rETR(II)
0.420
0.264
10.97
0.097
24.04
7
7
7
7
7
0.014
0.014
0.261
0.003
1.615
SM2
SM6
SM7
0.435 7 0.021
0.263 7 0.025
10.86 7 1.237
0.094 7 0.002
23.18 7 1.386
0.430 7 0.000
0.281 7 0.007
11.77 7 0.049
0.090 7 0.003
22.17 7 0.484
0.440 7 0.028
0.326 7 0.020
13.63 7 0.778
0.153 7 0.003
37.82 7 0.572
Fv/Fm, maximum quantum efficiency of PS II.
Y(II), effective quantum efficiency of PS II.
rETR(II), relative electron transport rate of PS II.
Standard deviation ( 7 SD) of the data from three independent experiments. For each experiment, three technical replicates were performed.
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
223
Fig. 4. Analyses the activity of PS II and PS I of NADPH consumption strain (SM7) and control strains (SM6, SM2 and WT). (A) O2 evolution rate under 600 μmol/m2/s.
(B) Light response curve of rETR(II), relative electron transport rate of PS II. (C) Light response curve of rETR(I), relative electron transport rate of PS I. Error bars indicate
standard deviation (SD) of the data from three independent experiments. For each experiment, three technical replicates were performed.
rETR(I) of all strains (Fig. 4C). The rETR(I) of strain SM7 was also
higher than that of its control strains WT, SM2 and SM6 under the
tested light intensity. The rETR(I) of all control strains reached the
highest value (approximately 38) under 588 μmol photons/m2/s,
whereas the rETR(I) strain SM7 reached the highest value
(86.08 73.05) under 1182 μmol photons/m2/s, which was 2.3 fold
that of all control strains. The light intensity response curve of Y(I)
(Fig. S3) showed that the Y(I) of strain SM7 was approximately 1.3fold and 2.8-fold that of control strains WT, SM2 and SM6 under
588 μmol photons/m2/s and 1182 μmol photons/m2/s, respectively
(Fig. S3). These suggested that introducing extra NADPH consumption ability increased the energy conversion efficiency of PS I.
The quantum yield of PS I can be divided into Y(I), the effective
quantum yield and non photochemical dissipation which includes
Y(ND), the quantum yield of non-photochemical energy dissipation due to donor side limitation; and Y(AD), the quantum yield of
non-photochemical energy dissipation due to acceptor side limitation. Thus, Y(I) þ Y(NA) þ Y(ND) ¼1. To understand the fate of
quantum yield of PS I, the non photochemical dissipation of PS I
quantum yield was analyzed (Table 3). Under normal light intensity of 100 μmol photons/m2/s, both quantum yields of nonphotochemical energy dissipation due to donor side limitation Y
(ND) and acceptor side limitation Y(NA) of strain SM7 were much
lower than that of the control strains, indicating that the introducing extra NADPH consumption ability decreased the quantum yields of non-photochemical energy dissipation of PS I. As a
result, the photochemical energy was significantly increased. Under higher light intensity of 588 μmol photons/m2/s, although the
total quantum yields of non-photochemical energy dissipation of
PS I (Y(ND) þY(NA)) of strain SM7 was much lower than that of the
control strains, Y(ND) of strain SM7 decreased 34% while Y(NA) of
SM7 increased 82%, compared to Y(ND) and Y(NA) of all control
strains, respectively (Table 3).
4. Discussion
In this study, we hypothesized and tested a new strategy for
Table 3
Measurement of redox state of P700 to detect Y(I), rETR(I), Y(ND) and Y(NA) under light intensity 588 μmol/m2/s of NADPH consumption strain (SM7) and control strains
(SM6, SM2 and WT).
Light intensity (μmol/m2/s)
Parameter
WT
588
588
588
588
Y(I)
rETR(I)
Y(ND)
Y(NA)
0.153
37.70
0.853
0.013
7
7
7
7
0.005
1.516
0.043
0.023
SM2
SM6
SM7
0.150 7 0.005
37.16 7 5.666
0.845 7 0.024
0.013 7 0.017
0.148 7 0.010
36.52 7 2.485
0.854 7 0.025
0.005 7 0.012
0.341
84.16
0.637
0.024
Y(I), effective quantum efficiency of PS I.
rETR(I), relative electron transport rate of PS I.
Y(ND), quantum yield of non-photochemical energy dissipation due to donor side limitation.
Y(NA), quantum yield of non-photochemical energy dissipation due to acceptor side limitation.
Standard deviation ( 7 SD) of the data from three independent measurements. For each experiment, three technical replicates were performed.
7
7
7
7
0.006
1.516
0.016
0.015
224
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
improving the photosynthetic efficiency of a photoautotrophic
organism, cyanobacterium S. 6803. The core idea of this strategy
was to better couple the light reactions and dark reactions by introducing extra NADPH consumption capability. Using this strategy, the activity of the light reactions was remarkably increased,
resulting in a significantly increased photosynthetic growth rate.
Strikingly, the photosynthetic oxygen evolution and photochemical efficiency all significantly increased, at a much higher
light intensity, indicating strain SM7 has not only increased its
photosynthesis efficiency, but also acquired ability for utilizing
stronger light.
To address the insufficient supply of ATP in dark reactions, the
first attempt was to increase the absorption of light energy by
increasing light intensity or optimizing the light capture system to
produce more ATP (Blankenship and Chen, 2013; Melis, 2009). As
ATP synthesis is tightly coupled with NADPH production via linear
electron transport chain in a fixed ratio (Fig. 1) (Kramer and Evans,
2011), an increase in ATP synthesis is accompanied by an increase
in NADPH production. However, the demand for NADPH is much
less than that for ATP in the dark reactions. Increasing ATP production using the above approach would result in the accumulation of NADPH, which is toxic to cells. Therefore, directly increasing ATP synthesis via increasing absorption of light energy
would have a limited positive impact on increasing the photosynthetic efficiency (Kozaki and Takeba, 1996; Nogales et al., 2012).
Alternatively, some photosynthetic organisms contain an accessorial cyclic electron transport chain, from which ATP can be
generated without forming NADPH (Allen, 2003). In such a case,
three ATP and two NADPH could be generated in the light reactions via both linear and cyclic electron transport chains (Allen,
2003; Kramer and Evans, 2011), which would possibly satisfy the
energy needs in the dark reactions. However, considering that
photorespiration, nitrogen assimilation and other anabolic processes all consume ATP, the extra ATP generated via the cyclic
electron transport chain might not meet the ATP requirement for
cell growth (Allen, 2003; Kramer and Evans, 2011; Noctor and
Foyer, 1998).
In this work, extra ATP generation in light reactions was
achieved by introducing the capability to consume extra NADPH.
The introduced extra NADPH consumption capability not only
resulted in a significant increase of oxygen revolution, relative
electron transfer rate and effective quantum yield of both PS II and
PS I, but also resulted in a significant increase of cell growth rate,
CO2 fixation rate and dry cell weight. Compared to the directed
control strain SM6, the introduced extra NADPH consumption
capability resulted in an approximately 0.6 g/L extra biomass formation (Table S4). Generally, for photoautotrophic microorganisms, formation of 1 g biomass requires approximately 100 mmol
ATP (Kliphuis et al., 2012). In this work, formation of 0.6 g/L extra
biomass would require 60 mmol/L ATP, indicating introducing
extra NADPH consumption capability stimulated ATP production.
Notably, although the specific growth rate at early stage of
cultivation and the biomass concentration after 20 days cultivation
of strains SM2 and SM6 were significantly higher than that of WT
(Fig. S1; Table S4), the CO2 fixation rates of strains WT, SM2 and
SM6 were the same (Fig. 3C). This indicates that there was no
difference among the photosynthetic efficiency of WT, SM2 and
SM6. The better growth of strain SM2 and SM6 might be attributed
to the disruption of acetate and PHB synthetic pathways in these
mutants, which may contribute to cell growth due to redistribution of carbon flux (Zhou et al., 2014b).
Moreover, through analysis of quantum yields of non-photochemical energy dissipation of PS I, we learned that the introduced
extra NADPH consumption ability mainly decreased the donor side
non-photochemical energy dissipation under high-intensity light
conditions. The increase of acceptor side non-photochemical
energy dissipation in strain SM7 under high-intensity light conditions indicates the energy generated from PS I cannot be completely utilized by carbon fixation, thus extra energy was dissipated as heat in PS I. Namely, it was the acceptor side (carbon
fixation) that limited the energy conversion of PS I. Thus, if carbon
fixation rate of strain SM7 can be improved, the efficiency of light
reactions of strain SM7 under high-intensity light conditions can
be further increased.
Photosynthetic microorganisms usually utilize light with low or
medium intensity (below 600 μmol photons/m2/s). Light intensity
higher than 600 μmol photons/m2/s usually leads to photoinhibition or photodamage (Tikkanen et al., 2014; Vinyard et al., 2014).
However, the light intensity in nature often fluctuates. For instance, the maximum light intensity in daytime often fluctuates
from 990 μmol photons/m2/s to 1486 μmol photons/m2/s (He
et al., 2015). Therefore, capabilities for utilizing fluctuating and
high intensity lights are desired features for photosynthetic organisms. To utilize high-intensity light, the D1 protein of PS II of
Synechococcus elongatus PCC 7942 was engineered by point mutagenesis (Vinyard et al., 2014). Although the oxygen evolution
rate of each mutant strain were all lower than that of wild type,
the authors provided fundamental design principles for engineering photosynthesis with optimal photochemical efficiencies
for growth under low versus high light intensities (Vinyard et al.,
2014). In this work, the mutant strain SM7 functions well in a
broad range of light intensity, and its saturation light point,
1200 μmol photons/m2/s, falls perfectly in the range of the maximum light intensity in nature. This indicates that introducing
extra NADPH consumption ability into photosynthetic organisms
could be a useful strategy for utilizing fluctuating and high-intensity lights.
The isopropanol synthetic pathway was constructed in Synechococcus sp. PCC 7942; however, the effects of NADPH consumption on growth and photosynthesis was not investigated
(Hirokawa et al., 2015; Kusakabe et al., 2013). Because the production of isopropanol under light and aerobic growth conditions
was too low to be detected, isopropanol was produced under dark
and anaerobic conditions and 26.5 mg/L isopropanol was produced (Kusakabe et al., 2013). Subsequently, the isopropanol production was increased to 146 mg/L by optimizing isopropanol
production conditions, shifting cells from dark and anaerobic
conditions to light and aerobic conditions (Hirokawa et al., 2015).
In this study, we optimized the expression of the introduced three
enzymes in S. 6803 and the production of 226.9 mg/L isopropanol
was solely achieved under light and aerobic conditions, while dark
and anaerobic conditions were required for isopropanol production in previous isopropanol works (Hirokawa et al., 2015; Kusakabe et al., 2013).
In terms of isopropanol production, this study is an extension
of our previous acetone work (Zhou et al., 2012). In our previous
acetone work, the production of acetone was very low and only
36 mg/L acetone was detected under dark and anaerobic conditions. It is known that codon optimization and application of
strong promoters for expression of key enzymes can significantly
increase the production of target chemicals (Angermayr and Hellingwerf, 2013; Guerrero et al., 2012; Lan and Liao, 2012; Zhou
et al., 2014b). Therefore, in this work, the three genes required for
construction of the isopropanol pathway were codon-optimized
and placed under a strong promoter Pcpc560 before introducing
into in S. 6803 (Zhou et al., 2014c). Consequently, acetone production was increased to 108.5 mg/L and 226.9 mg/L isopropanol
production was achieved, under light and aerobic growth conditions. The actual titer of isopropanol could be higher, as the evaporated isopropanol during 20 days aerobic cultivation was not
taken into account.
To further investigate whether the increased biomass
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
225
Fig. 5. The time course of isopropanol conversion from acetone by NADPH consumption strain SM7* (A) and growth profile analysis of all strains (B). All strains were
supplied with 2 g/L acetone. In A, square represented acetone assay; circle represented isopropanol assay. Error bars indicate standard deviation (SD) of the data from three
independent experiments. For each experiment, three technical replicates were performed
production was due to the introduced extra NADPH consumption
ability, we constructed a mutant strain SM7*, in which only the
Pcpc560 controlled codon-optimized sadh was inserted into the
phaCE site of S. 6803. Acetone, the direct substrate of the sadh
encoded NADPH dependent secondary alcohol dehydrogenase
(SADH), was added to the medium of mutant strain SM7*, WT and
its direct control strain SM1, in which the phaCE gene was replaced
with the Cmr gene (Zhou et al., 2012) to a final concentration of
1800 mg/L. Isopropanol was detected in the medium of strain
SM7* but not in that of WT and SM1 (Fig. 5A), suggesting acetone
can be converted to isopropanol by strain SM7*. The gradually
decreased concentration of acetone in the medium of strain WT
and SM1 (Fig. 5A) was simply due to evaporation, as the blank
medium containing 1800 mg/L acetone but free of S. 6803 cells
also showed similar acetone profile (data not shown). This also
suggested the depletion of acetone in the culture of strain SM7*
includes evaporated acetone. Growth profile under light and
aerobic conditions showed strain SM7* grew significantly faster
than WT and its direct control strain SM1 (Fig. 5B). At 14 days of
cultivation, the dry cell concentration of strain SM7* was 39% and
49% higher than that of strains WT and SM1, respectively (Table
S6). These data further confirmed that consumption of extra
NADPH stimulated the growth of S. 6803 cells.
As NADPH is the major form of reducing equivalent in cyanobacteria (Takahashi et al., 2008), NADPH was used as a cofactor for
chemicals production in engineered cyanobacteria by introducing
NADPH-dependent enzymes or transhydrogenases to convert
NADPH into NADH to aid the NADH-dependent enzymes (Angermayr et al., 2014; Atsumi et al., 2009; Lan and Liao, 2012; Liu et al.,
2011; Luan et al., 2015; Oliver et al., 2013; Savakis et al., 2013). In
previous efforts to produce chemicals at the expense of NADPH,
for instance, isobutyraldehyde (Atsumi et al., 2009), butanol (Lan
and Liao, 2012), fatty acids (Liu et al., 2011), lactate (Angermayr
et al., 2014), 2,3-butanediol (Oliver et al., 2013; Savakis et al., 2013)
and ethanol (Luan et al., 2015), the growth of mutant cells all
decreased or did not change as compared with the wild type cells.
Among these efforts, Oliver and Machado reported an increased
oxygen evolution of an engineered cyanobacterium, associated
with production of 2.38 g/L 2,3-butanediol (Oliver et al., 2013). The
author simply ascribed the increased oxygen evolution to the
overproduction of the target chemical (Oliver et al., 2013). However, increased cell growth was not observed in the 2,3-butanediol
production work (Oliver et al., 2013). In another work of sucrose
production in cyanobacteria, enhanced biomass accumulation and
photosynthetic activity was observed in sucrose-exporting cells
under high osmolality conditions (Ducat et al., 2012). Although the
author speculated that consumption of excess reducing equivalents during sucrose production might be one of factors (Ducat
et al., 2012), the growth of sucrose-exporting cells was still much
lower than the growth of wild type cells under light and aerobic
growth conditions. Our work represents the first report of having a
significant increase in biomass accumulation upon expressing a
pathway that utilizes NADPH.
Improving photosynthetic efficiency would have a great impact
on agriculture. Our study demonstrated that improving energy
imbalance between light reactions and dark reactions is a simple
but useful strategy to improve the photosynthetic efficiency and to
enable utilization of high-intensity light for cyanobacteria. Compared with other strategies, such as optimization of Rubisco or
introducing carbon concentrating mechanisms into C3 plants, we
propose that up-regulating the activity NADPH-dependent enzymes in C3 plants might contribute to establishing a new ATP/
NADPH balance that would improve its photosynthetic efficiency,
biomass production and utilization of high-intensity light. The
idea of establishing a new ATP balance could be further tested in
other oxygenic, photoautotrophic microorganisms and higher
plants to check whether similar improvements in photosynthesis
could be achieved.
Authors’ contributions
Y.L. and J.Z. designed the research. F.Z., J.Z., H.M. and Y.P.Z.
performed the research. Y.L. and J.Z. wrote the manuscript. All
authors analyzed the data and discussed the results.
Acknowledgments
This work was supported by the Key Research Program of the
Chinese Academy of Sciences (ZDRW-ZS-2016-3), the Strategic
Priority Research Program of the Chinese Academy of Sciences
(XDA07040405), and the National Natural Science Foundation of
China (31470231). We thank Feng Guo, the technical expert of
Dual-PAM 100, for assistance with the chlorophyll fluorescence
and P700 signal analyses.
226
J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227
Appendix A. Supplementary material
Supplementary data associated with this article can be found in
the online version at http://dx.doi.org/10.1016/j.ymben.2016.08.
002.
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