Team:USTC/Results
Conduction system
Mtr
In our project, Mtr CAB is the most important and fundamental proteins as it plays the role to transfer extracellular electrons into the cytoplasm through the membrane. To examine whether the Mtr protein complex has the function as we expected, we used our engineered strain pMC( strain co-expressed Mtr CAB and Ccm A-H) to construct a bio-cathode. We monitored the current of the bio-cathode to see whether there would be a higher current in the experiment group than the WT strain.
Here, first we did an bacteria PCR to monitor the maintenance of the recombinant plasmids(pM28 contains the mtr CAB’s gene and the pTBC contains the ccm A-H’s gene). As we can see in figure 1, we could confirm that the strain is fine to use.
Figure 1.
So we started our bio-cathode assay to examine our theory. The protocol of the bio-cathode assay can be found in the notebook part in our wiki, look it up if you want to know more details!! In figure 2, we can easily confirm that the Mtr CAB protein complex was mature, as the pellets were red in pMC group, no matter the strain had been induced or not. When we did it the first time, there was no significant difference of the current of the bio-cathode between WT and our strain pMC(data not shown). We speculated that it was because we did NOT have the starvation step when we first did it, which is to cultivate the bacteria in a minimal salts medium for a certain time, like 4 to 6 hours. Because we did NOT have this starvation step, although we already used PBS to wash the bacteria 2 to 3 times, those nutritions still contained inside of the bacteria, providing another electron source when we were running the bio-cathode. So when the cathode was given a certain voltage, the bacteria still wouldn’t take up the electrons from the electrode.
Figure 2.
So we performed this bio-cathode assay for a second time, adding this starvation step into the protocol. In addition, after the starvation step, we used 1 mL of minimal salts medium to resuspend the bacteria and dropped it onto the graphite electrode to form a bio-film, which could help to make a better connection between the bacteria and the electrode, especially when we were using the Mtr pathway to transfer electrons. Here, in figure 3, you can see how we made this biofilm. 2 to 3 hours later, with a sufficient airflow in the laminar flow hood, the graphite electrode would dry up and form a great biofilm. With this biofilm, electrons could be transferred to the Mtr C protein directly from the electrode which can increase the efficiency of electron transferring. Then what we need to do was to construct this bio-cathode, put every part of this “toy” together and get the oxygen out of this container. Here in figure 4 is how we clear the oxygen out of the bio-cathode to create an anaerobic environment. Lastly, we connected the bio-cathode to the electric-chemical station to give a certain voltage to the cathode and monitor the current of the cathode as time went by as how figure 5 shows.
Figure 3.
Figure 4.
Figure 5.
Figure 6 is the result of this experiment. From the figure, we can easily notice that the red line, which is the Mtr-induced group, had a 50% higher current than the other two group after the bio-cathode turned into a stable state . This could strongly prove that the engineered strain pMC can transfer electrons into the cytoplasm, which led to the increasing of the cathode-current. But there would be a chance that this difference between these 3 groups was just the background noise between this three cathode, resulting from the hardware’s varieties. So we added fumarate into the system to see whether there would be a cathode catalyzed current happened in the pMC group. That’s why there was a sharp increasing in the figure. When we added fumarate into the system, the electrons on the electrode finally found a way to leak to—— the fumarate. So there would be a strong electron flow when we added fumarate into the system. But after a short time we introduced this sudden change into the system, the current will become stable again, slowly climbing back to the current it was. However, the time it took to get back to stable state can be a strong evident to prove our assumption——our engineered E.coli can transfer extracellular electrons into the cytoplasm!! The red line’s curve happened after we added fumarate into the system is kind of a typical curve of cathode-catalyze-current!! So, with this result, the cathode’s current to time under a certain voltage, we can confidently say that the Mtr CAB system work!!
Figure 6. The current result of the bio-cathode.
In conclusion, the Mtr CAB system can really function as an electron pathway to transfer extracellular electrons into the cytoplasm, even though it’s expressed in E.coli, but not it’s origin host Shewanella.!! In another word, our conduction system can function as we expected, transferring those electrons from the electrode into the cytoplasm, which means our E.coli can transform itself like transformer from a normal form to a special form that can “eat” electrons!
Ccm
1.Construction of ccm on pSB1C3 and co-transformation of ccm and mtr
We get ccm gene from the genome of E.coli by PCR and insert this sequence to pSB1C3 with pTet upstream successfully. The sequence of our ccm is validated by DNA sequencing from SangoTech. We co-transform the plasmids containing ccm and the plasmids containing mtr in BL21. Then we pick some colonies for cultivation and confirm the co-transformation of two plasmids (shown in Figure 1). We inoculate confirmed colony to 2x YT media and grown with 250 rpm shaking for 12 hours at 30˚C. 5 mL of overnight culture is used to inoculate 1L 2xYT media and were grown for 16 hours at 30 ˚C. After cultivation, we confirm the existence of our two plasmids in BL21 by bacteria PCR (shown in Figure 2).
2.We express mature MtrA and MtrC successfully
After cultivation, we collect our bacteria from 1 mL media by centrifugation. Obviously, our bacteria with ccm become red compared with wide type which shows our Ccm is expressed successfully because heme are attached to MtrA&C expressed on the outer membrane (shown in Figure 3).
We lyse the bacteria and extract the membrane and periplasmic fractions, respectively. Then we run SDS-PAGE of sample of each fraction. The molecular weight of MtrC, MtrB and MtrA is 72kDa, 77kDa and 36kDa respectively. We can confirm the expression of MtrABC from the bond of approximate molecular weight, but the expression of CcmA-H is not sure (shown in Figure 4). We attach a His-tag to MtrC so the expression of MtrC is confirmed from the result of Western blot (shown in Figure 5).
To insure the function of Ccm directly, we employ the method of TMBZ stain which is a common chemical analysis method for heme covalently bond to peptides. According to the theory of TMBZ stain, if Ccm A-H have catalyzed the attachment of heme to MtrA&C, there will be visible blue bond at corresponding position. By comparing the position of blue bond with protein marker, we make sure that our MtrA and MtrC are mature. These results prove that our Ccm functions well directly and our Ccm is expressed successfully indirectly (shown in Figure 6).
Besides, we design an experiment as a negative control. We transform the plasmids containing mtr (shown in Figure 7). Then we induce the expression of mtr without ccm under aerobic condition. We run SDS-PAGE and western blot of our samples (shown in Figure 8, Figure 9) and detect the heme via TMBZ stain (shown in Figure 10). It’s obvious that our MtrCAB is expressed compared with wide type from SDS-PAGE result. But there is no blue bond after TMBZ stain so we conclude that our Mtr is immature. These results also reveal the fact that Ccm A-H have no impact on the expression of MtrCAB but play a vital role in catalyzing the maturation of MtrA&C.
From these two experiments, we can reach the conclusion that MtrA&C get mature because of the function of CcmA-H which prove the successful expression of Ccm A-H. We construct mature MtrCBA system with the expression of CcmA-H and the first part of our project conduction system is fulfilled.
Reference:
[1] Thomas, P. E., Ryan, D., & Levin, W. (1976). An improved staining procedure for the detection of the peroxidase activity of cytochrome P-450 on sodium dodecyl sulfate polyacrylamide gels. Analytical biochemistry, 75(1), 168-176.[2] Jensen, H. M. (2013). Engineering Escherichia coli for molecularly defined electron transfer to metal oxides and electrodes. University of California, Berkeley
Photosynthesis system
1.cysdes-pLuxR-pSB1C3 construction and transformation
We obtain the sequence of cysdes gene from Genebank and synthesize this gene from IDT. We insert this sequence to pSB1C3 with promoter pLuxR which is provided by iGEM headquarters. The sequence of our ccm is validated by DNA sequencing from SangoTech. We transform the plasmids containing cysdes in BL21. Then we pick some colonies for cultivation and confirm the transformation of the plasmid by the methods of PCR (shown in Figure 1). From the result of electrophoresis, we confirm the transformation of cysdes plasmid with pLuxR.
2.Expression of CysDes
We inoculate 2 mL overnight culture to 200 mL LB media (1mM cysteine, 30mM glucose and 10mM HEPES are included) and cultivate for 2h at 37 ˚C. When OD600 reaches 0.4-0.6, add AHL to final concentration of 250nM. After 3h cultivation, collect the bacteria by centrifugation. Then extract the raw enzyme of CysDes by ultrasonication. We run SDS-PAGE of samples of raw enzyme, cell content obtained by 100 ˚C heating and wide type (shown in Figure 2). The protein CysDes is about 46kDa, we can find obvious bands at the about position of 45kDa which are unique to lanes of cell contents after induction and raw enzyme compared with the wide type. Although there are proteins of similar molecular weight in wide type, darker bands in experiment group meaning a high amount of proteins could prove the existence of high amount of CysDes. From the result of SDS-PAGE, we could confirm the expression of CysDes.
3.Growth curve of E.coli (BL21) under different concentrations of Cd2+
In our project, we add Cd2+, which is toxic to the growth of cell, to our media to synthesize CdS nanoparticles under the catalysis of our engineered E.coli. Considering that CysDes catalyzes the reduction of cysteine and Cd2+ is transformed to CdS precipitation to some extent, the existence of CysDes can strength bacterial resistance to Cd2+ toxicity. But the substrate of CysDes, cysteine, is a kind of necessary amino acid for bacterial growth. The adding of cysteine metabolic pathway may impede the growth of bacteria.
To figure out the impact of different concentration of Cd2+ and CysDes on the growth of E.coli and determine the appropriate concentration of Cd2+ for growth, we measure OD600 as a data for bacteria concentration at various conditions and time respectively. Then we draw the scatter graph and fit the growth curve with smooth line to show the tendency of growth (shown in Figure 3).
According to the graph of growth curve, we can reach these conclusions:(a) Adding of Cd2+ to media impedes the growth of wide type E.coli. But after 18h, wide type bacteria grow in low concentration of Cd2+ media (lower than 0.2mM) will reach the same platform stage as the group of media without Cd2+. Higher concentration of Cd2+(over 0.4mM) will limit the platform stage to a lower OD600.
(b) The metabolism of cysteine by CysDes slows down the growth of E.coli and delays the start of exponential stage compared to the wide type. But BL21 expressing CysDes can still reach the platform stage after 18h in the nearly same concentrations as the wide type.
(c) Adding of Cd2+ to media can also obstacle the growth of E.coli expressing CysDes and delay the start of exponential stage. But the change of Cd2+ concentration has no obvious effect on growth curve and cells expressing CysDes grow under 0.4mM Cd2+ can reach a higher concentration approximate to the group of 0.1mM and 0.2mM Cd2+ than the wide type.
(d) The expression of CysDes does strengthen the E.coli‘s resistance of Cd2+ toxicity, but also slow the growth of E.coli to some extent. We can supplement the media with appropriate amount of cysteine to reduce the negative impact caused by the expression of CysDes.
4.Enzyme activity analysis of CysDes in vitro
To analysis the enzyme activity of CysDes, we choose to detect the concentration of S2- which is reduced from cysteine under the catalysis of CysDes. Because of lacking appropriate purifying methods, we just analyze the activity of raw enzyme obtained via bacteria lysis. According to the method described in L.Chu et al. of hydrogen sulfide detection, we first cultivate the 1mL mixture of cysteine, PBS buffer and raw enzyme for 2h at 37 ˚C. Sulfide formation was determined by adding 0.1 ml of 0.02 M N,N-dimethyl-p-phenylenediamine sulfate in 7.2 N HCl and 0.1 ml of 0.3 M FeCl3 in 1.2 N HCl to the reaction tubes. The absorbance at 650 nm was determined after color development for 20 min at 20°C. Sulfide concentration is determined from the standard curve of Na2S.
From figure 4, the concentration of S2- in the group of CysDes is higher than the wide type which proves that CysDes promotes the reduction of cysteine to S2- with good enzymatic activity certainly.
Under the catalysis of same amount of raw enzyme, we measure the production of S2- with various concentration of cysteine after 2h at 37˚C (shown in Figure 5). The value of OD650 has approximate linear relationship with the concentration of cysteine which means CysDes almost catalyzes the reduction of all cysteine to S2- and CysDes functions well.
5.Transmission electron microscopy image of CdS nanoparticles and bacteria
6.CdS nanoparticles' effect to cathode-current
Reference:
[1] Chu, L., Ebersole, J. L., Kurzban, G. P., & Holt, S. C. (1997). Cystalysin, a 46-kilodalton cysteine desulfhydrase from Treponema denticola, with hemolytic and hemoxidative activities. Infection and immunity, 65(8), 3231-3238.[2] Wang, C., Lum, A., Ozuna, S., Clark, D., & Keasling, J. (2001). Aerobic sulfide production and cadmium precipitation by Escherichia coli expressing the Treponema denticola cysteine desulfhydrase gene. Applied microbiology and biotechnology, 56(3-4), 425-430.
Harvest system
1.Transformation and Expression
We transformed a plasmid PET22b containing KmAdh into E.coli successfully. We use KmAdh’s specific primers to do PCR to verify this achievement.
Figure 1.
We can see that the experimental group and the positive control have the same band but WT does not. This shows the transformation is successful.
Then we induced the expression of this enzyme. We use 200 mL LB to cultivate our bacteria in 37℃，250 rpm. When its OD600 reached 0.5-0.8 we added 20μL 1M IPTG in it to induce KmAdh expression.
Figure 2.
We can see from the SDS page that KmAdh was successfully expressed.
Then we centrifuge the cells at 8000 rpm, 4°C, for 10 min and then remove the supernatant . Resuspend the bacteria with 50 mL PBS; Centrifuge at 8000 rpm, 4°C, for 10 min and then remove the supernatant. Resuspend the bacteria with 15 mL PBS. The cells were disrupted via ultrasonication (Power 30%, 30 min. Total duty in cycles of 1s on, 2s off). Finally we centrifuge it at 14000 rpm, 4°C, for 20 min and retrieve the supernatant for future purification. The crude enzyme is purified by nickel column to get the pure enzyme.
Figure 3.
2.Enzyme activity test
NADH, as a necessary cofactor of KmAdh, has a significant absorption in 340nm. Along the process of the reduction reaction, the consuming of NADH will lead to a decrease of absorption in 340nm which allows us to test the activity of KmAdh by the spectrophotometer.
Figure 4.
After adding, we scan the 340nm UV absorption value over time. Because NADH is easy to be oxidized, we set a blank control. The system is the same as the above system, but with the same amount of PBS instead of KmAdh.
We can obviously see the rapid increase in the absorption value after adding enzyme, indicating that NADH is drastically consumed. This shows the purified enzyme function is normal and the KmAdh is successfully expressed in E. coli.
Figure 5.
3.Toxicity test
Considering that acetaldehyde and ethanol, the substrate and product of KmAdh, may do harm to the cell, we first made the growth curve of E.coli at different concentrations of acetaldehyde and ethanol to figure out a proper experimental condition. For acetaldehyde and ethanol, we both set four concentrations: 0%, 0.1%, 0.2% and 0.3%, and the results are shown in the following figures.
As the concentration of ethanol in the system increases, the growth of KMADH and WT is inhibited but KMADH’s growth is clearly better than WT’s at the same concentration. The reason is that KmAdh also has the effect of helping to break down ethanol. For acetaldehyde, the growth of KMADH and WT are both inhibited when the acetaldehyde concentration increases and KMADH’s growth is significantly better than WT’s when the acetaldehyde concentration reaches 0.3% (the highest concentration we set). This result is a rough proof of our KmAdh’s function is normal and the enzyme can be relatively high toxic acetaldehyde into less toxic ethanol to improve cell viability. When the acetaldehyde concentration and ethanol concentration in the system are the same, not only WT’s growth but also KMADH’s growth is inhibited. This indicates that the toxic effects of acetaldehyde on cells are stronger than ethanol.
According to the results, we decided to use 0.1% acetaldehyde as the substrate, for E.coli can live well.
Figure 6.
Figure 7.
