Chemicals used in this study were purchased from Acros Organics (Na₂MoO₄⋅2H₂O), BD (agar), MP Biomedicals (CuSO₄⋅5H₂O), Fisher Scientific (NaCl, MgSO₄⋅7H₂O, KCl, NaNO₃, tris base, H₃BO₃, and kanamycin monosulfate), and Sigma-Aldrich (Na₂EDTA, CaCl₂⋅2H₂O, KH₂PO₄, vitamin B12, FeCl₃⋅6H₂O, MnCl₂⋅4H₂O, ZnCl₂, CoCl₂⋅6H₂O, and spectinomycin dihydrochloride pentahydrate). Primers were synthesized by Integrated DNA Technologies (IDT). DNA purifications were performed using the Plasmid Miniprep, DNA Clean and Concentrator, and Gel DNA Recovery kits from Zymo Research. Restriction enzymes, DNA polymerases, and ligases were purchased from New England Biolabs. Suppliers of all other materials used in this study are described below.

Strains used and constructed in this study are listed in Table 1. The first step in engineering Synechococcus sp. PCC 7002 for FFA production was gene knockout of the long-chain-fatty-acid CoA ligase (fadD, SYNPCC7002_A0675), responsible for FFA recycling (Kaczmarzyk and Fulda, 2010). Homologous regions (~500 bp) upstream and downstream of fadD were cloned using the primers in Table S2 in Supplementary Material and integrated into pSE15 (Ruffing and Jones, 2012) at the AflII/SpeI (5′ fragment) and BglII/XhoI (3′ fragment) restriction enzyme sites, generating the fadD knockout plasmid, pS12. Transformation of pS12 into Synechococcus sp. PCC 7002, followed by spectinomycin selection and PCR screening of fadD, resulted in the construction of strain S01. Transformation of Synechococcus sp. PCC 7002 was performed using a modified protocol based on that proposed by Stevens and Porter (1980). Synechococcus sp. PCC 7002 was grown in medium A⁺ until OD₇₃₀ = 1.0 was reached. Approximately, 0.1 μg of linearized plasmid DNA was added to 1 mL of culture (OD₇₃₀ = 1.0) in a 16 mm glass test tube with ventilated cap. The transformation culture was incubated at 30°C with 150 rpm and 60 μmol photons m⁻² s⁻¹. After 24 h of incubation, the transformation culture was concentrated to 100 μL using centrifugation (5000 × g, 5 min) and spread on medium A⁺/agar plates with 1 mM of sodium thiosulfate and 40 μg/mL of spectinomycin dihydrochloride pentahydrate and 50 μg/mL of kanamycin monosulfate, as required. Colonies were re-streaked a minimum of three times to obtain complete segregants. After antibiotic selection and PCR screening, two positive transformants were tested for FFA production in 400 mL cultures (see Culture Conditions).

For S02 construction, the truncated E. coli thioesterase, ‘tesA, from pSE16 (Ruffing and Jones, 2012) was cloned using primers in Table S2 in Supplementary Material and integrated into pS12 at the EcoRI and BamHI sites following the inducible trc promoter to form pS13. Transformation of pS13 into Synechococcus sp. PCC 7002 resulted in gene knockout of fadD and integration of ‘tesA. Gene knockout of the desaturase gene, desB, was achieved by transforming pSB into S02. pSB was constructed by replacing the NSII homologous regions of pSA (Ruffing, 2013a) with 951 and 936 bp fragments homologous to the sequences upstream and downstream of desB. The fat1 thioesterase from Chlamydomonas reinhardtii was cloned from pSE20 and integrated at the EcoRI and BamHI sites of pS12 to form pS14, using primers listed in Table S2 in Supplementary Material. Strain S03 was constructed by transformation and integration of pS14 in Synechococcus sp. PCC 7002. A truncated fat1 thioesterase (tfat1) was constructed by redesigning the forward primer to eliminate the predicted chloroplast-targeting signal, determined using ChloroP 1.1 (Emanuelsson et al., 1999). Integration of tfat1 into the EcoRI and BamHI sites of pS12 yielded pS17, and subsequent transformation and integration of pS17 into Synechococcus sp. PCC 7002 produced strain S05.

To insert another restriction enzyme site for rbcLS integration, ‘tesA was re-cloned using primer ‘tesAR2, yielding pS18. The small and large subunits of RuBisCO (rbcLS) from S. elongatus PCC 7942 were cloned from pSE18 (Ruffing, 2013a) (primers rbcLSF1/rbcLSR1 in Table S2 in Supplementary Material) and inserted downstream of ‘tesA at the AatII site in pS18 to produce pS19. Transformation of pS19 into Synechococcus sp. PCC 7002 yielded strain S06. To insert the psbA1 promoter from S. elongatus PCC 7942 upstream of rbcLS, the psbA1 promoter was cloned from pSE20 and inserted into pS18 to yield pS20. Subsequent cloning of rbcLS from SE20 using the second rbcLS primer set (rbcLSF2/rbcLSR2) and insertion into pS20 at the BsrGI site produced pS21. Transformation of pS21 into Synechococcus sp. PCC 7002 yielded strain S07.

Escherichia coli DH5α was used for plasmid construction; competent cells were prepared as described in Sambrook and Russell (2001). All E. coli DH5α strains used for plasmid maintenance were grown in LB medium at 37°C and 350 rpm in a VWR incubating mini-shaker. After overnight growth in a test tube containing 4 mL of LB media, the plasmid-containing E. coli strains were harvested for plasmid isolation using the plasmid mini-prep kit from Zymo Research. Plasmids derived from pS12 were linearized using NdeI digestion, and plasmids derived from pSA were linearized using SpeI digestion.

Wild-type and engineered strains of Synechococcus sp. PCC 7002 were grown in medium A⁺ with antibiotic supplementation as required (40 μg/mL spectinomycin dihydrochloride pentahydrate for S01, S02, S02ΔdesB, S03, S05, S06, and S07 and 50 μg/mL of kanamycin monosulfate for S02ΔdesB). Agar plates of medium A⁺ contained 1 mM of sodium thiosulfate. Single transformed colonies were re-streaked at least three times or until full segregants were obtained, as confirmed by PCR. Synechococcus sp. PCC 7002 strains grown on solid media were inoculated into 4 mL of medium A⁺ with antibiotics as necessary and grown at 30 or 38°C and 150 rpm in a New Brunswick Innova 42R shaking incubator with photosynthetic light bank. Alternating cool white and plant fluorescent lights provided ~60 μmol photons m⁻² s⁻¹ of continuous illumination. After 5–7 days of growth, 1 mL of the test tube inoculum was transferred to a 500 mL baffled Erlenmeyer flask containing 100 mL of medium A⁺ with appropriate antibiotics. After 4 days of growth under the aforementioned conditions, the 100 mL culture was used to inoculate 400 mL of medium A⁺ in a 1 L glass media bottle with a three-port lid for sampling, bubbling of 1% CO₂ in air, and ventilation through a 0.22 μm filter. The initial cell concentration of the 400 mL culture vessels was approximately an OD₇₃₀ of 0.1. No antibiotics were added to the large culture vessels to eliminate any potential effects associated with antibiotic supplementation. The large cultures were sampled every 2 days for ~3 weeks. IPTG was added at 100 h to induce expression from the trc and LlacO-1 promoters.

Cell growth was measured using optical density (OD) readings at 730 nm from a PerkinElmer Lambda Bio spectrophotometer. Photosynthetic yield measurements were obtained using a Waltz mini-PAM photosynthetic yield analyzer; for each sample, triplicate technical measurements were averaged due to variability associated with these measurements. Photosynthetic yield or efficiency measurements FV′∕Fm′ are a fluorescence-based measurement of the photochemical efficiency of photosystem II (PSII) in higher plants. Variable fluorescence (F_v) is the difference between the maximum fluorescence (F_m) under saturated light and the initial fluorescence (F₀) under actinic light; the apostrophe indicates that the samples were not dark adapted prior to measurement. Photosynthetic yield FV′∕Fm′ measurements in cyanobacteria are complicated by fluorescence contributions from phycobiliproteins and the low fraction of total chlorophyll associated with PSII; however, FV′∕Fm′ measurements in cyanobacteria have been shown to correlate well with changes in the rate of oxygen evolution from PSII (Campbell et al., 1998). In this study, photosynthetic yield measurements are used as a general indicator of photosynthetic efficiency and overall cell health. FFA measurements were performed as described previously (Ruffing and Jones, 2012) using the FFA Quantification Kit from Biovision. Cell growth, photosynthetic yield, and FFA measurements were taken every 2 days. At day 16, the absorbance spectrum of each culture was measured using a Beckman Coulter DU-800 UV-Vis Spectrophotometer to determine changes in photosynthetic pigment concentrations. Each data point is the average of three biological replicates with error bars indicating the standard deviation.

Synechococcus sp. PCC 7002 was genetically engineered for FFA production and excretion by targeting the long-chain-fatty-acid CoA ligase (fadD, SYNPCC7002_A0675) for gene knockout and overexpression of a truncated thioesterase from E. coli (‘tesA). The resulting engineered strains, S01 (ΔfadD) and S02 (ΔfadD, ‘tesA⁺), are analogous to the previously constructed, FFA-producing strains of S. elongatus PCC 7942, SE01 (Δaas) and SE02 (Δaas, ‘tesA⁺). A schematic of the engineered pathway for FFA production can be found in the Supplemental Material (Figure S1). The FFA concentrations, cell growth profiles, and photosynthetic yields of Synechococcus sp. PCC 7002, S01, and S02 are shown in Figure 1, along with the previous results from S. elongatus PCC 7942, SE01, and SE02 (Ruffing and Jones, 2012). As expected, no FFAs are excreted by the wild-type (7002), yet both S01 and S02 synthesized and excreted FFAs. The FFA concentrations produced by fadD knockout in S01 (Figure 1A) are very low (<6 mg/L), particularly in comparison to the comparably engineered S. elongatus PCC 7942 strain, SE01, which produced up to 43 mg/L of excreted FFAs. Expression of the truncated E. coli thioesterase in S02, however, increased the level of excreted FFAs, producing concentrations similar to SE01 and higher levels than the analogous S. elongatus PCC 7942 strain, SE02 (Figure 1A). While the FFA-producing strains of Synechococcus sp. PCC 7002 (S01 and S02) did not show improved FFA production in comparison to the S. elongatus PCC 7942 strains (SE01 and SE02), there were notable differences in the physiological responses of the hosts. The late exponential growth rates of S01 and S02 showed a slight decrease in comparison to the wild-type 7002 after induction at 100 h (4.17 days) (Figure 1B); this small reduction is expected as FFA production reduces the available pool of acyl-ACP for cell membrane biosynthesis. On the other hand, SE01 and SE02 showed a severe reduction in final cell concentrations, 21 and 59% lower than wild-type (Figure 1B), much greater than expected due to FFA production. Additionally, the photosynthetic yields of S01 and S02 are very similar to the wild-type 7002, while the photosynthetic yields of SE01 and SE02 were significantly reduced compared to the wild-type 7942 strain (Figure 1C). These results suggest that FFA production in Synechococcus sp. PCC 7002 does not compromise the cellular physiology of the host strain, unlike the engineered strains of S. elongatus PCC 7942, which exhibited a stress response to FFA production and excretion.

The FFA production experiments described in the previous section and the results reported in Figure 1 were conducted at a temperature of 30°C for comparison with the engineered S. elongatus PCC 7942 strains, which have an optimal growth temperature of 30°C (Golden and Sherman, 1984). However, the reported optimal growth temperature for Synechococcus sp. PCC 7002 is 38°C (Sakamoto and Bryant, 1997). Therefore, FFA production experiments were also conducted at this higher temperature for 7002, S01, and S02 (Figure 2). As expected, the higher cultivation temperature yielded a slight improvement in FFA concentrations excreted by S01 and S02 (Figure 2A), likely due to the increased thermal kinetics associated with the higher temperature. Unexpectedly, cell growth was reduced at the higher temperature for all three strains, including the wild-type (Figure 2B). To confirm the culture temperature, the aqueous temperature of the culture vessels was measured using a standard glass thermometer. The aqueous temperature readings indicated that the culture temperatures were slightly higher than the setpoints, 32°C for the 30°C setting and 39°C for the 38°C setting. The lower range light intensities used in this experiment (60 μmol photons m⁻² s⁻¹) may also influence the optimal growth temperature, and a more rigorous, multi-factorial investigation of Synechococcus sp. PCC 7002 growth is required to evaluate the optimal growth temperature of this strain. In addition to this unexpected growth response, there were significant changes in photosynthetic yields at 38°C in comparison to 30°C. At the higher temperature, the wild-type (7002) had reduced photosynthetic yields with values up to 50% lower compared to the 30°C temperature condition (Figure 2C). The FFA-producing strains showed an additional decrease in photosynthetic yield, with S02, the highest FFA producer, having the most reduced photosynthetic yields. To investigate whether these reduced photosynthetic yields were accompanied by changes in photosynthetic pigments, the absorbance spectra of the wild-type (7002) and engineered strains (S01 and S02) were measured for cultures at 38 and 30°C after significantly reduced photosynthetic yields were detected (day 16). While the absorbance spectrum for the wild-type (7002) did not show significant differences at the two temperatures, the spectrum of the S02 cultures showed reduced peaks for both chlorophyll-a (680 nm) and the phycobiliproteins (635 nm) at 38°C (Figure 3). The reduced growth, photosynthetic yields, and photosynthetic pigments that accompany FFA production at 38°C suggest that the FFA tolerance of Synechococcus sp. PCC 7002 is temperature-dependent.

At high temperatures, bacteria have been shown to change the degree of saturation of their cell membrane lipids, with increasing saturation accompanying increasing temperatures (Koga, 2012). The increased saturation of membrane FAs in Synechococcus sp. PCC 7002 at 38°C may affect the ability of FFAs to cross the cell membrane. Alternatively, the removal of saturated FAs by the recombinant thioesterase may compromise the cell’s ability to grow at this higher temperature. To investigate this hypothesis, the S02 strain was modified via gene knockout of the desB gene, encoding a desaturase involved in cold temperature tolerance in Synechococcus sp. PCC 7002 (Sakamoto et al., 1997). At 30°C, the desB mutant did not show any significant changes in FFA production, growth, or photosynthetic yield in comparison to S02 (data not shown). This suggests that DesB is not involved in FFA tolerance at this temperature, but it does not rule out a role for membrane saturation in the mechanism of temperature-dependent FFA tolerance in Synechococcus sp. PCC 7002.

To determine if higher yields of FFA can be produced in Synechococcus sp. PCC 7002 compared to S. elongatus PCC 7942, additional strategies for improving FFA biosynthesis were investigated. The acyl-ACP thioesterase from C. reinhardtii (fat1) was previously shown to have activity similar to ‘tesA in S. elongatus PCC 7942 (Ruffing, 2013a). Therefore, fat1 was expressed in Synechococcus sp. PCC 7002, along with fadD knockout, to form strain S03. Unexpectedly, S03 showed only a slight increase in excreted FFA concentration compared to S01, which has only the fadD knockout (Figure 4A). The fat1 gene was previously cloned from the associated C. reinhardtii mRNA transcript (Ruffing, 2013a), which likely includes a chloroplast-targeting signal for translocation of Fat1, a nuclear-encoded protein, into the chloroplast. Inclusion of the chloroplast-targeting signal may lead to either low enzyme activity or secretion of the enzyme, as the cyanobacterial membrane is evolutionarily similar to the chloroplast membrane (Giovannoni et al., 1988). To eliminate this potential source of low activity, a putative chloroplast-targeting signal was predicted using ChloroP 1.1 (Emanuelsson et al., 1999), and a new forward primer was designed to exclude this 5′ signal sequence, yielding a truncated fat1 (tfat1). Expression of tfat1 in S05 led to a nearly threefold increase FFA production and excretion compared to the fat1-expressing S03 strain (Figure 4A). However, the amount of FFA produced by S05 was still lower than that produced by the ‘tesA-expressing S02 strain. Additionally, S05 had lower growth and photosynthetic yield compared to S02 (Figures 4B,C); therefore, further strain development utilized the ‘tesA thioesterase rather than fat1 or tfat1 acyl-ACP thioesterases.

In addition to thioesterase expression, carbon fixation by RuBisCO is an important step for high carbon flux to support both growth and FFA biosynthesis. Similar to RuBisCO overexpression in S. elongatus PCC 7942, expression of the large and small subunits of RuBisCO from S. elongatus PCC 7942 (rbcLS) did not yield an improvement in FFA production in the Synechococcus sp. PCC 7002 strain S06 [(Ruffing, 2013a), Figure 4A]. Previous quantitative PCR studies in engineered S. elongatus PCC 7942 indicated that rbcLS expression is not significantly improved by integration of the P_trc-fat1-rbcLS synthetic operon (Ruffing, 2013a). Therefore, an additional promoter was inserted into the synthetic operon to drive rbcLS expression. The psbA1 promoter from S. elongatus PCC 7942 was cloned and inserted after ‘tesA to form the synthetic operon: P_trc-‘tesA-P_psbA1-rbcLS in S07, an engineered strain of Synechococcus sp. PCC 7002. Contrary to the results obtained in the S. elongatus PCC 7492 host (Ruffing, 2013a), rbcLS overexpression in S07 resulted in more than a threefold increase in excreted FFA concentrations (Figure 4A), suggesting that carbon fixation is in fact a rate-limiting step in FFA biosynthesis for Synechococcus sp. PCC 7002. The extracellular FFA concentrations in the S07 cultures had high variability, as illustrated by the error bars in Figure 4A, which represent the standard deviation of the three biological replicates. This measurement variability was due to FFA precipitation, illustrated by the inset image in Figure 4A, which made homogenous sampling difficult. The higher FFA production in S07 also resulted in reduced cell growth and photosynthetic yields (Figures 4B,C), indicating that high levels of FFA production may affect cellular physiology in Synechococcus sp. PCC 7002.