Our wet lab team worked to characterize parts that were successfully cloned. This includes carbonic anhydrase from N. vulgaris and pdc and adhB from Z. mobilis.
Carbonic anhydrase from N.vulgaris is known to be heat-resistant2. To test if our heat-resistant carbonic anhydrase would allow Synechocystis sp PCC 6803 to sequester a similar amount of CO2 compared to wildtype under room conditions, we used an airtight Tupperware container to test the decreasing concentration of CO2 over time. Unfortunately, the Tupperware we used wasn’t airtight even after we parafilmed the container. Therefore, we decided to take the hourly measure of CO2 concentration in an empty container. This would be our control to see if cyanobacteria would affect how fast the CO2 concentration dropped. Thus, we can assume the additional decrease in concentration would be from the cyanobacteria absorbing the CO2.
Pdc and adhB from Z.mobilis are known for their high efficiency. Pdc catalyzes the conversion of pyruvate to acetaldehyde. Acetaldehyde then is converted to ethanol by adhB. To test if PDC and adhB would allow Synechocystis sp PCC 6803 to produce ethanol we set up a titration curve using Schiff’s reagent. Schiff’s reagent is often used for ethanol detection in biological staining. When Schiff’s reagent reacts with ethanol it turns the sample a bright magenta color. We used this change of color to create a standard curve and to detect and quantify the ethanol produced by BL21 E.coli strain transformed with pdc and adhB.
We step up two controls of parafilm Tupperware with a CO2 monitor to test how fast the air leaves the box with no cyanobacteria. We also set up a box with wildtype Synechocystis sp. PCC 6803 cyanobacteria was taken from a seed culture originally inoculated from a colony on a BG-11 agar plate and grown at both 30°C with constant rotation at 200 rpm. The O.D.730 for the sample was 0.337. Our modified sample to test was SSynechocystis sp. PCC 6803 with our modified CA, “coordinated active site domain” of the enzyme carbonic anhydrase that promotes carbon sequestration. The O.D.730 for the sample was 0.400. All the samples were placed in identical Tupperware containers and parafilm in the same way with parafilm along the edge of the container. They were placed in a CO2 incubator brought up to 1%. Once the samples reached 1% CO2, we took them out of the incubator and placed them in room air still sealed. We waited up to 11 hours to see the drop in concentration of CO2.
After testing both the wildtype Synechocystis sp. PCC 6803 and our modified Synechocystis strain with the modified carbonic anhydrase from N. vulgaris (BBa_K5052911), we found that the wildtype cyanobacteria decreased carbon levels at a faster rate than the modified strain. The wildtype had a carbon reduction of -184 ppm/hour, while the modified strain's was -92.7 ppm/hour.
These results conclude that BBa_K5052911 does effect the carbon sequestration at room temperature since the rate of carbon capture was slower than wildtype.
EtOH Detection Experiments
Schiff’s Reagent Indicator Plates
Schiff’s reagent is a chemical solution that can be used to detect the presence of aldehydes, primary, and secondary alcohols. It is created by decolorizing a dye, typically fuchsin, with sulfur dioxide. When an aldehyde is present, the reagent restores the dye’s original magenta color, making it a useful test in biological staining1. Using the protocol by Jason Dexter and Pengcheng Fu, we attempted to make Schiff’s reagent indicator plates which could be used to test ethanol production in transformed BL21 strain and Synechocystis sp. PCC68032. Following the protocol, we attempted to make LB-agar indicator plates with 2% Schiff’s reagent. These plates would be used to test BL21 E.coli strain and Synechocystis sp. PCC 6803 was transformed with our ethanol cassette for ethanol production. A change in the color of the plates would indicate ethanol production2.
First, to confirm that Schiff’s reagent can detect alcohol we added 1% Schiff’s reagent to solutions of 100% ethanol, 70% ethanol, 70% isopropanol, and H2O (Figure 1). Like ethanol, isopropanol is an alcohol. Unlike ethanol, it is a secondary alcohol. Observing the intensity of magenta would also allow us to observe how much it reacts to different concentrations of ethanol as well as how differently it reacts between primary and secondary alcohols at the same concentration. The Schiff’s reagent turned all alcohol samples bright magenta confirming that it can be used to detect both primary and secondary alcohols in solution. Qualitatively, 100% ethanol showed the deepest shade of magenta. Between 70% ethanol and 70% isopropanol, it appeared that Schiff’s reagent reacts more with primary alcohols than secondary.
Figure 1: From left to right: tube with 100% ethanol, 70% ethanol, 70% isopropanol, H20
After demonstrating that our team was able to use Schiff's reagent to detect alcohols like ethanol, we attempted to make indicator plates by adding 4 mL of Schiff’s reagent to 200 mL LB-agar-kan (2% Schiff’s) to see if our colonies were producing ethanol. Upon adding Schiff’s reagent, the solution turned a bright red color (Figure 2). This indicated that Schiff’s reagent either reacted to something in LB-agar-kan or the LB-agar-kan was too hot which could affect the reactivity of Schiff’s reagent.
Figure 2: LB-agar-kan turned bright red after the addition of Schiff’s reagent.
We then attempted to overlay 200 uL of Schiff’s reagent onto premade LB-agar plates to see if Schiff’s reagent was going to react with cooled LB-agar. The plates were stored in the refrigerator overnight to make sure they were cooled enough. The LB-agar plates turned light pink upon adding the reagent, indicating a reaction with the LB media itself or with the agar (Figure 3).
Figure 3: The plate on the left had Schiff’s reagent added to it; the plate in the middle had Schiff’s reagent overlaid on it; the plate on the right is an LB-agar plate.
Following these results, we tested liquid LB media by adding 20 uL Schiff's to 1 mL LB to see if the Schiff’s reagent was reacting specifically with the agar or with LB media. LB turned a slight pink color. We then added 20 uL of 70% EtOH, but no visible difference was observed. After that, we tried adding 80 uL more of 70% EtOH to the same sample to get 100 uL of the final volume of Schiff’s reagent, but still, no visible difference was observed. Finally, we tried adding 80 uL more of Schiff's reagent for the final volume of 180 uL but still, no color difference was observed (Figure 4). These results led us to the conclusion that Schiff's reagent would not be a reliable metric for detecting ethanol in LB media. Considering that Schiff’s reagent was reacting to something in LB media, we decided to use the M9 minimal medium instead due to its ability to support cellular growth and the lack of components such as yeast extract and tryptone that are present in LB media.
Figure 4: The tube on the left contained LB+EtOH with Schiff’s; the tube in the middle contained LB with Schiff’s; the tube on the right contained LB.
To see if we can use Schiff’s reagent to detect and quantify our ethanol production in M9, a titration curve was produced. Schiff’s reagent was cooled in the fridge. Samples were vortexed after adding Schiff’s and let sit for an hour before reading the plate on the plate reader at 462 nm (K. Tennakone et al., 1987). The reading was performed again after 2 hrs and the curve was generated (Figure 5). The produced titration curve is linear which indicates a linear association between the percentage of ethanol and absorbance.
1mL of M9 + 100mL Schiff's (10%)
70% ethanol
50% ethanol
30% ethanol
20% ethanol
10% ethanol
M9 with Schiff's
M9
Figure 5: The titration curve
After performing the titration curve using the M9 medium, we saw that the reagent would be a reliable metric for ethanol detection because it showed increasing intensity of magenta with increasing concentrations of ethanol. Knowing this, we tested our E. coli transformed with the complete ethanol cassette to see if our parts worked. If our ethanol cassette is functional, we would be able to observe a difference in the intensity of magenta over time. BL21 + EtOH was grown at 37℃ in M9 minimal medium shaking at 100rpm. Since the ethanol cassette is under the control of the dark inducible promoter, PompC, the culture was covered with tin foil to limit the amount of light reaching the culture to allow the expression of AdhB and Pdc. The next day, we aliquoted 1 mL of just M9 medium as our negative control, BL21 strain with EtOH plasmid, and DH5a strain with the empty vector, pAM4891. We then added 100uL of Schiff's reagent and observed a color change (Figure 6).
Figure 6: From left to right: M9 + Schiff’s, DH5a + pAM54891 + Schiff’s, BL21 + EtOH + Schiff’s
The sample BL21 + EtOH + Schiff’s shows a significantly deeper shade of magenta than DH5a + pAM54891+ Schiff’s and M9 + Schiff’s alone. These results hint at the presence of ethanol as well as the functionality of AdhB and Pdc biobricks.
Our next goal was to quantify the ethanol produced by our transformed E. coli strains. This was done by setting up a 96-well plate similar to how we set up our titration curve.
Figure 7: The results of quantification
The results of a plate reader at A462 were promising, the difference between pAM4892 and the EtOH biobricks transformed E.coli was visible, but the data also indicated that the sample with E.coli transformed with pAM4891 contained 14% ethanol and the difference between the absorptions was not significant. This indicated a reaction of Schiff’s reagent with the components of M9 media.
Following these results, we decided to perform distillation. This served two purposes. Mainly to remove media that was affecting our results. The other purpose of performing distillation was to consider our overall design. The biophotovolteic cells were designed to be covered by hydrogel, so the ethanol secreted by cyanobacteria would go to the hydrogel, which would need to be distilled to extract the ethanol.
To see if our method for the distillation of ethanol from the media would work we had hydrogels soak up the media and ethanol produced by our E.coli, remove it from the culture, and heat it to distill the ethanol from the hydrogel. To achieve this we made 70% ethanol and 50% ethanol solutions with a total volume of 6 mL for each sample. We set aside 1 mL of each sample as our control and tested them with Schiff’s reagent to compare the color of the sample to the results we got out of the hydrogels. We let hydrogels soak for an hour in the samples. After the soaking, the hydrogel was put on a glass petri dish, covered with a 60 mm petri dish, and heated up to 78℃ until the droplets of liquid started to appear on the top of the petri dish. The 10 uL of Schiff’s reagent was added to the formed liquid and results were recorded.
Figure 8: Results after adding 10 uL of Schiff’s reagent.
The results showed that both control ethanol samples and ones distilled from hydrogen both turned the same shade of magenta indicating the presence of ethanol and proving that our distillation method worked (Figure 8). After getting the results, we tested just hydrogels. We soaked one hydrogel in the BG-11 media and another in water as our control. We used the same setup with the Petri dishes, unfortunately, Schiff's reagent reacted with the plate. After leaving it for a while all plates turned magenta. Following these results, we attempted distillation with a different collection vessel - a beaker. We soaked hydrogel in 50% EtOH, 70% EtOH, water and tested the beakers by adding 100 uL Schiff's and letting them sit for 10 min. The idea was to precoat the beakers in Schiff's which would allow us to see if there's ethanol in real time accurately. The beaker was also causing the Schiff's reagent to react but only slightly. Distillation was run for five minutes at 80℃ and 10 uL of Schiff's was added to the samples. The color change was observed.
Figure 9: Left to right: water-soaked hydrogel distillation, 50% ethanol-soaked hydrogel distillation, 70% ethanol-soaked hydrogel distillation.
The intensity of the magenta color was roughly the same between the two ethanol distillations, indicating the same concentration of ethanol distilled (Figure 9). These results proved that our distillation method. can be used to extract ethanol from hydrogels, so our next step was to perform distillation on the supernatants from our DH5a E. coli strain transformed with our ethanol plasmid and BL21 E. coli strain transformed with our ethanol plasmid to see if they produce ethanol. Four samples were set up to perform the distillation. Two were grown on the plates and two were grown in the liquid media. In both cases, E.coli were grown in the dark at 37℃. The plate samples were grown on LB-agar-km plates and liquid samples were grown in LB-km 5 mL cultures overnight. For plate samples, hydrogels were laid over the top of the lawn of E.coli. For liquid samples, hydrogels were soaked in the media extracted from liquid cultures after spinning them down and removing the pellet for 1hr. For the control, we soaked hydrogel in LB-km alone for 1hr. All samples were distilled and captured in 50 mL beakers. A 100 uL of each sample was collected in 1.5mL tubes and 10 uL of Schiff's reagent was added.
Figure 10: Results after 1 min after adding 10% Schiff’s. Left to right: DH5a + EtOH on solid, BL21 + EtOH on solid, DH5a + EtOH in liquid, BL21 + EtOH in liquid, LB.
The absence of color change in the LB sample indicated that distillation removed all possible components that could react with Schiff’s reagent, which means that the color change in other samples could be only due to the presence of ethanol. The BL21 sample on solid media showed a much darker shade of magenta than the DH5a sample on solid media, which makes sense considering that BL21 is an expression strain of E.coli. These results once again proved that our AdhB and Pdc biobricks work in E.coli (Figure 10).
To quantify these results Schiff’s reagent ImageJ titration curve was performed based on Figure 9. The following results were produced:
Figure 11: Schiff’s reagent ImageJ Titration Curve
Figure 12: Graph of ethanol concentration
Using the data from ImageJ Figure 10 was produced. The BL21 strain transformed with our EtOH biobrick and grown on solid media showed 75.02% ethanol per volume. The BL21 strain transformed with our EtOH biobrick and grown in liquid media showed 22.25% of ethanol per volume. Both results are significantly larger compared to the LB sample with 1.95% ethanol (Figure 11, 12).
Our team utilized two main cyanobacterial strains: Synechocystis sp. PCC 6803 and Synechococcus sp. UTEX 3154, in order to power our biophotovoltaic fuel cell. Throughout this process, we learned and documented methods for their cultivation by conducting several experiments and setting up our own assays to learn the best conditions for optimal cyanobacteria growth.
When reviewing existing research on cyanobacteria, we found that two types of media, BG-11 and Artificial Seawater, are commonly used for growing marine species such as Synechococcus sp. UTEX 3154. To determine which medium would support greater growth yield, we decided to compare the effectiveness of BG-11 and Artificial Seawater.
For our experiment, we used 1 mL samples of both BG-11 and Artificial Seawater inoculated with Synechococcus sp. UTEX 3154. Both samples were kept in a 38°C shaking incubator at 100 rpm with a 25% light intensity setting on our LED grow lights.
In the first 24 hours, Artificial Seawater appeared to promote better growth, with an O.D. (optical density) of 0.018 compared to BG-11’s 0.005. However, by day 5, BG-11 surpassed Artificial Seawater, reaching an O.D. of 0.033, while Artificial Seawater measured at only 0.022. This trend continued as the experiment progressed. After 11 days of growth, BG-11 showed exponential growth whereas Artificial Seawater plateaued.
Although Artificial Seawater initially promoted faster growth, BG-11 sustained a higher growth rate over time. Based on these results, we concluded that BG-11 is the more effective medium for cultivating cyanobacteria in this experiment, and we continued to use it as our primary growth medium.
In several studies on cyanobacteria growth, we observed that bicarbonate was added as an additional carbon source to support cyanobacterial growth1. To determine its potential benefits for our system, we conducted our own tests to evaluate its effectiveness.
To assess the potential advantages and disadvantages of bicarbonate flashing, the process of regularly adding bicarb to stimulate growth, we conducted several tests using 1 mL samples of Synechococcus sp. UTEX 3154 in different media. Samples were prepared by taking inoculant from the seed culture purchased from UTEX. We compared the effects of adding bicarbonate daily versus every other day in both BG-11 and Artificial Seawater (Artificial Seawater). Both samples were given an initial 24-hour growth periO.D. with no bicarbonate, and were kept in a 38°C shaking incubator at 100 rpm with a 25% light intensity setting on our LED grow lights.
By Day 4, all samples showed relatively similar O.D.s, ranging from 0.016 to 0.027 across the five test groups. However, significant changes emerged by Day 6. BG-11 liquid media with daily bicarbonate addition nearly doubled overnight, increasing from an O.D. of 0.065 to 0.102, while BG-11 liquid media with bicarbonate added every other day only increased from 0.040 to 0.057. In contrast, Artificial Seawater with bicarbonate every other day showed a decline from 0.048 to 0.026, consistent with our earlier findings that Artificial Seawater is less effective at sustaining cyanobacterial growth. Artificial Seawater with daily bicarbonate addition exhibited minimal growth, with an O.D. of just 0.006.
By Day 11, this trend persisted. The sample with the highest O.D. was BG-11 liquid media with daily bicarbonate addition at 0.347, followed by BG-11 with bicarbonate every other day at 0.285. The control sample, BG-11 with no bicarbonate added, had an O.D. of 0.109. Both Artificial Seawater samples, with and without bicarbonate, ended with O.D.s under 0.01, confirming that Artificial Seawater is not as effective for supporting sustained cyanobacterial growth.
While the samples that were flashed with bicarbonate were able to grow at a higher rate than those that were not, we observed that the bicarbonate was forming flakes in our media and was preventing our cyanobacteria from fully shaking in our incubator. Therefore, bicarbonate flashing was a useful way to grow up a culture with a very small starting O.D., but it was not useful when our samples reached higher O.D.s as sufficient gas-exchange was not efficient with the excess bicarbonate in the culture.
As Synechococcus sp. UTEX 3154 is an uncharacterized species, we wanted to investigate whether it would thrive better in more alkaline conditions, as marine environments tend to be slightly basic2>. We tested two different pH levels: 7 (the natural state of BG-11) and 8 (to simulate more basic, marine-like conditions).
In order to conduct this assay, we set up two cultures: one with Synechococcus sp. UTEX 3154 in BG-11 1% NaCl liquid media with at pH 7 (no additional base), and another in BG-11 1% NaCl liquid media adjusted to pH 8 using KOH and Tris HCl buffer. Both samples were kept in a 38°C shaking incubator at 100 rpm with a 25% light intensity setting on our LED grow lights.
Over 11 days of growth, the results were clear. The culture at pH 7 reached an O.D. of 0.127, showing consistent growth throughout the experiment. In contrast, the pH 8 culture only reached an O.D. of 0.006 and never entered the log growth phase. Based on these results, we concluded that UTEX 3154 grows best at pH 7 and decided to maintain the natural pH of BG-11 without further adjustment.
Since we were working with Synechococcus sp. UTEX 3154, a marine cyanobacterial species, we wanted to determine if salt concentration played a significant role in its growth.
We tested this with four different salt concentrations: 0% (no salt), 1%, 1.5%, and 2%. Four test tubes contained 2 mL of BG-11 media each with their varying levels of salt concentration and were inoculated with Synechococcus sp. UTEX 3154. Both samples were kept in a 38°C shaking incubator at 100 rpm with a 25% light intensity setting on our LED grow lights.
After 11 days of growth, the sample with 1% salt concentration had the highest O.D., followed by 0% salt, then 1.5%, and finally 2%, which had only reached an O.D. of 0.02.
To further investigate, we extended the assay until day 28 to observe the long-term trends. After several weeks, the sample with 0% salt (the control) showed the highest O.D., revealing it was the most effective for growth. This assay allowed us to determine that no added salt in the BG-11 media was the most optimal condition for growing our marine cyanobacterial species.
From our initial research, we believed that the optimal temperature for growing UTEX 3154 was 38°C with slight shaking, which is the protocol we followed for early experiments 3. However, after consulting with Dr. Alistar McCormick from the University of Edinburgh, a cyanobacteria expert, we learned that such a high temperature could lead to excessive media loss. Based on this advice, we decided to test our cyanobacteria cultures at 30°C and 38°C to determine the optimal temperature for growth.
We set up BG-11 liquid cultures inoculated with Synechococcus sp. UTEX 3154 at both 30°C and 38°C, maintaining a speed of 200 rpm and a 75% light intensity setting on our LED grow lights.
Over time, the Synechococcus sp. UTEX 3154 in 30°C had an exponential growth curve with a max O.D. of 1.146 after 16 days while the Synechococcus sp. UTEX 3154 in 38°C was only able to reach a max O.D. of 0.277 in 25 days. Based on these findings, we adjusted all future experiments to grow UTEX 3154 at 30°C for optimal growth.
In order to conduct successful homologous recombination to modify cyanobacterial genomes, a high concentration of antibiotics is needed to ensure that each copy of the chromosome is included. For example, when discussing with Dr. Toby Call from the University of Cambridge, we came across his PhD thesis regarding using cyanobacteria in biochemical systems. In his thesis, up to 500 ug/mL of kanamycin was used to isolate successful transformants 4. However, we decided to conduct our own assay to determine the optimal amount as the concentration of antibiotics needed when using a shuttle vector such as RSF1010 has not been extensively studied. We designed an assay to determine the maximum amount of antibiotic that our cyanobacterial strains could handle.
We began with 5 test tubes each with 2mL of Synechocystis sp. PCC 6803 and 5 test tubes with 2 mL of Synechococcus sp. UTEX 3154. Each culture of cyanobacteria started at an O.D. of 0.3 at a wavelength of 730 nm. 0 ug/mL, 25 ug/mL, 50 ug/mL, 100 ug/mL, or 200 ug/mL were added to each of the test tubes for each sample. The samples were kept with constant light and were shaken at 200 rpm. The O.D. at a wavelength of 730 nm was measured each day for a week of growth.
Our results showed that a minimum of 50 ug/mL was needed to successfully kill wild-type cyanobacteria While 25 ug/mL of kanamycin was sufficient in killing wild-type Synechococcus sp. UTEX 3154, we observed a dip followed by a sustained O.D., indicating that a spontaneous mutant may have arised. This assay allowed us to determine that 50 ug/mL of kanamycin is needed to successfully isolate transformed colonies.
We have identified seven parts that we hypothesized would improve the efficiency of our strains to sequester carbon, generate electricity, and produce ethanol. However, genetic manipulation of cyanobacteria is limited. Homologous recombination is the standard but has a lengthy selection process due to the need to ensure each copy of the chromosome in a cell contains the gene of interest, of which there could be hundreds1. The ploidy of cyanobacteria is also dependent on the growth stage which makes reliable modification via homologous recombination more difficult. To avoid these difficulties, our team opted to modify using a shuttle vector replicable in both E. coli and cyanobacteria, RSF1010. RSF1010-based plasmids are the only well-known shuttle vector between E. coli and cyanobacteria, consequently, we have developed novel solutions to ease the engineering process, including a promoter swap system as a way to efficiently switch out the promoter controlling our gene of interest, and an improved method of gene integration as a way to introduce two independent genes into the same cell without the risk of incompatibility of RSF1010-based plasmids. This allowed us to create cassettes of composite parts with all necessary components to address one of our three modules which we can modify our strains with.
This promoter swap system is based on traditional cut-and-paste cloning. Traditional cut-and-paste cloning was an attractive solution to us as the smallest of recognition sites can be easily added without disturbing the function of the full composite part. Two restriction sites are placed on either end of the promoter, DraIII cut site is placed upstream of the promoter and MluI cut site is in between the ribosome binding site and the start codon of the DNA coding sequence. Promoters can be exchanged via double digest with DraIII and MluI, performing a gel extraction of the vector, and inserting your new promoter of choice.
We have worked to modify three cyanobacterial promoters: Pcpc560–a constitutive promoter, PompC–a dark inducible promoter, and cLac145–an IPTG inducible promoter.
We characterized our promoter swap system and measured success by comparing EYFP expression in parts with promoters swapped to their native sequence. We took EYFP controlled by PompC (BBa_K5052902) and cLac145 (BBa_K5052900) and swapped out the promoters. For PompC + EYFP, we cut out PompC with DraIII and MluI to replace it with cLac145 and separately with Pcpc560. For cLac145 + EYFP, we cut out cLac145 with the same pair of restriction enzymes to replace it with PompC.
After performing traditional cut-and-paste to insert the respective promoters, the ligated products were used to transform DH5a E. coli. These transformants were grown on LB-agar-kan plates at 37C in the dark overnight. The next day, these plates were taken for fluorescent imaging.
This image was quantified for fluorescence on ImageJ by taking the area of the plate and calculating the intensity of the fluorescence. With this value, we can divide by the background values to find the corrected total cell fluorescence2.
When comparing swapped promoters to their native sequence, we see that the levels of fluorescence are vastly different. cLac145 + EYFP has a corrected total cell fluorescence of 11.93 compared to after we swapped it with PompC where the corrected total cell fluorescence increased to 16.321. When looking at PompC + EYFP, it shows a corrected total cell fluorescence of 18.875 which decreased greatly to 10.066 when swapped with Pcpc560 and to 11.354 when swapped with cLac145. Furthermore, comparing the strength of fluorescence between parts sharing the same promoter shows similar results, like between cLac145 + EYFP and PompC insert cLac145 which have a corrected total cell fluorescence of 11.93 and 11.354 respectively.
Plasmid incompatibility refers to the inability for two or more plasmids to coexist in one bacterial cell. This most often occurs due to both plasmids possessing the same replicon with specific controlling elements, or competition from similar partitioning elements1. To combat this issue, we developed a system of ‘super’ plasmids, as well as a plan to co-culture cyanobacteria such that each cell would only need to contain one plasmid of interest.
Each of the 3 modules of our project, carbon sequestration, electricity generation, and ethanol production had a planned super plasmid. This plasmid would contain all of the Biobricks that we planned to transform into cyanobacteria to get it to express the desired biological pathways. Additionally, the plasmid would contain the proper promoters for gene expression, and the kanamycin resistance genes necessary to test if our transformations were successful. In essence, each plasmid would be a self-contained system for the purposes of expressing multiple proteins of interest.
The carbon sequestration plasmid was planned to contain the SbtA and BicA genes, which both encode bicarbonate transport complexes. This plasmid being successfully incorporated into our cyanobacteria would result in a greatly increased uptake of carbon dioxide from the atmosphere.
Our electricity production plasmid was designed to carry the PsbE and PsbF genes that together express proteins for the cytochrome b559 protein complex. This complex is an essential part of photosystem 22, and should greatly increase the voltage produced by our cyanobacteria if properly formed.
As the final stage of our planned 3 modules, the ethanol plasmid carries the genes pdc and adhB from the Z. mobilis bacterium. These genes encode for the enzymes pyruvate decarboxylase and alcohol dehydrogenase respectively, which serve as critical components of the ethanol fermentation pathway. When expressed in cyanobacteria, the production of ethanol will be enabled, allowing us to collect a valuable byproduct from the cyanobacteria.
Creating the super plasmids in reality was conceptually very simple. Each biobrick had previously been ligated into the modified RSF1010-mobAY25F cloning vector through simple cut-and-paste. From there, based on the availability of restriction enzymes, one of the Biobricks would be cut out of its vector with a double digest, while the other vector would undergo a single digest to prepare it for ligation with the other gene of interest. Each super plasmid could be transformed into E. coli and grown in large amounts to prepare for eventual transformation into our cyanobacteria. However, this process was fraught with difficulty and setbacks.
Despite the simplicity of the planned experiments, we saw very little success with early attempts to digest our vectors and ligate both Biobricks together into one cloning vector. It is likely that the large plasmid resulting from having two genes present reduced the likelihood of proper ligations and transformations occurring. Most selection tests with bacteria indicated a failed transformation, and PCR analysis of the plasmids confirmed this negative result.
Ultimately it ended up being a matter of patience before we were able to confirm positive results in terms of a successful transformation. These results came about through a colony PCR analysis. Every time a transformation was performed, the resultant cells would be plated on a kanamycin selection plate. Any colonies that grew would be run through colony PCR using primers we designed specifically to multiply our genes of interest. When run against a positive control on an agarose gel, this approach enabled us to verify the presence of a successful super plasmid.
As seen in this comparison with an idealized version from SnapGene, the bands are exactly where we would expect them to be (accounting for different reactions being loaded into each well). This demonstrates that both the super electricity plasmid containing PsbE and PsbF, and the super ethanol plasmid containing pdc and adhB are present in separate E. coli cultures. Our carbon super plasmid was not able to be successfully ligated and transformed during the scope of this project.
After confirmation of the presence of our proper super plasmids, the colonies containing the successful result were grown quickly and allowed to multiply to a high degree. This was in preparation for the next step, natural transformation of cyanobacteria. The procedures and testing for this part of the project can be found in our Cyanobacteria section.
[1]Thomas, C. M. (2014) Plasmid incompatibility. Molecular Life Sciences 1–3.
[2]Pakrasi, H. B., Williams, J. G., and Arntzen, C. J. (1988) Targeted mutagenesis of the psbe and psbf genes blocks photosynthetic electron transport: Evidence for a functional role of cytochrome B559 in Photosystem ii. The EMBO Journal 7, 325–332.