Our system can be divided into three biochemical steps: 1) transcription of the target sequence into mRNA, 2) reverse transcription of the target sequence into double-stranded cDNA, 3) recombination for the cDNA to replace the target sequence. We set out to separately verify each step before integrating together. Our team logo shows the mutants accumulated by varying blue colors.
Transcription
Figure 1. The flanking of reverse transcription initiation and loxP sequences does not disrupt Chl functionality.
Wildtype Chl gene continues to express and colonies are grown on the plate containing chloramphenicol (plate on the right). The L158X mutant cannot express functional resistance gene and thus the plate is devoid of cell colony (plate on the right).
We put the target sequence and its flanking elements together under a T7 stable promoter for high expression level of target RNA. To verify our R-Evolution system, we constructed 8 nonsense mutant of chloramphenicol resistance gene (Chl), bearing the 8 base pair substitution from sense codon to nonsense mutant. We verified this construct through culturing bacteria carrying the original version or mutant on plates containing chloramphenicol. We found that bacteria carrying the original Chl grow naturally, while no colony was formed on the plates of mChl (Table. 1). After adding the flanking sequences on both ends, we used the C158X mutant and showed that the gene’s function has not been changed by our conduct (Figure 1).
Table 1. Nonsense mutation disrupts the chloramphenicol resistance ability of Chl.
Amp stands for ampicillin, Chl stands for chloramphenicol, the left column means that they are added to the plate. Chl acts as the positive control, cells grow naturally on both plates, while cells transformed with the negative control plasmid can only grow on plates containing ampicillin. Chl gene carrying nonsense mutation at different sites does not affect its ampicillin resistance, but chloramphenicol resistance is lost as no colony is formed. Single colony was picked and shook in liquid culture overnight, the plate was coated with 100 μL culture after 104 dilution.
| Control | Chl | Chl-Y33X | Chl-W85X | Chl-E97X | Chl-S121X | Chl-L158X | Chl-K182X | Chl-Q190X | |
| Amp | 19 | 192 | 291 | 128 | 236 | 138 | 255 | 175 | 276 |
| Amp & Chl | 0 | 30 | 0 | 0 | 0 | 0 | 0 | 0 | 0 |
In addition, we also constructed and verified nonsense mutants of fluorescent protein EGFP and mCherry at the 158th and 159th amino acid (Figure 2).
Figure 2. mCherry cannot emit fluorescence.
The vertical axis shows the quantified level of EGFP expression. Mutants of the fluorescence proteins cannot emit fluorescence while the wildtype protein functions naturally. The fluorescence level (EGFP: excitation wavelength 485 nm, detection wavelength 528 nm; mCherry: excitation 550 nm, detection 590 nm) is quantified by the concentration of fluorescein, and normalized by the measured OD600 equivalent to the number of beads in the system. The fluorescein for EGFP and silica beads are from the iGEM distributed measurement kit, red fluorescence is quantified by rhodamine B (Sigma-Aldrich, #S1402-1G). Error bar in the two graphs on the first row indicates the SEM of three replicates.
Reverse transcription (RT)
The successful expression of RT
RT is expressed under an IPTG controlled promoter, we constructed a series of promoters by placing a LacO fragment under a stable promoter, and hopes to determine under which we could achieve most stringent control. The constructs we tested are: T5, T5-LacO, T7-LacO, LacUV5-LacO, J23119-LacO.
We initially attempted to verify RT’s expression through an EGFP fusion protein. We fused the EGFP to the C’ end of pol protein, linked by a GS tag. But this construct proved unsuccessful (data not shown), possibly due to the length and complexity of the gag-pol polyprotein. Then we turned to directly expressing EGFP in the place of RT under the control of IPTG. If EGFP can be successfully expressed, so should RT. And results are obtained for each induction promoter construct (figure would be supplemented in presentation and poster).
We found through careful examination that this failure is due to problems with our plasmid construct, so we moved the RT to another tested plasmid, and through SDS-PAGE, verified its successful expression (Figure 3). Gag-pol polyprotein is expressed as a whole with its stop codon mutated into its readthrough product, glutamine. The polyprotein has three functional parts, capsid protein, protease and reverse transcriptase. We made Y586F mutation on reverse transcriptase to increase its mutation rate. From the PAGE gel, we can see that all three bands could be seen when induced.
Figure 3. gag-pol polyprotein is successfully expressed and underwent excision in the cell.
SDS-PAGE is performed on whole cell-lysis. The gag-pol polyprotein is split into three pieces, capsid protein (60.4 kDa), protease (13.5 kDa) and reverse transcriptase (69.1 kDa). Both versions of reverse transcriptase, one wildtype, the other Y586F mutant, are tested. ‘-’ stands for uninduced sample, while ‘+’ stands for sample after induction. From the gel we could see that all three bands are brighter in the induced sample.
Recombination
Cre expression is controlled by Tet operon
We placed Cre under the promoter ptetR, whose expression is controlled by its inhibitor TetR. Regarding where we should place the inhibitor gene to maximize its expression, we opted between 2 options, one is placing it downstream of the LacI inhibitor, the other is to place it downstream of the kanamycin resistance gene (KanR). We tested both construct by placing an EGFP in these two places and measuring its fluorescence emission. Results show that the expression level is almost the same in both construct.
Cre initiates excision between two homologous loxP site
Placing 2 wild-type loxP on both ends of the target sequence (mCherry) in the same direction, and expressing it under a stable promoter (J23101). By co-transforming the target plasmid with another plasmid carrying Cre recombinase, we verified that our Cre protein functions accordingly by excising the mCherry sequence from the promoter (Figure 4).
Figure 4. lox511 remains compatible with wildtype loxP, though at a lower excision rate.
Wildtype loxP and lox511-mCherry-loxP are analyzed on two different gels, their marker bands are indicated. Wildtype loxP only has an excision band. lox511 has a slight full-length mCherry band slightly longer than 1000 bp, which correlates with the full length between two loxP, but excision band is still visible and brighter than that of full-length mCherry. This result suggests that lox511 still interacts with wildtype loxP and go through excision, but at a lower efficiency.
Through PCR amplification with the primers annealing to sequences outside the target, and subsequent electrophoresis, we found that the band from bacteria co-transforming Cre corresponds to the excision of mCherry.
lox5171 is most incompatible with wildtype loxP (wtlox)
When we were carrying out integrated human practice, we were warned by Prof. Wang that two homologous loxP would be excised at a much higher efficiency than performing recombination as we wished, so we searched the literature and selected 3 mutants that are said to be incompatible with wtlox but are compatible with themselves, they are lox511, lox2272 and lox5171.
We tested their incompatibility with wtlox by replacing one of wtlox into the mutant at the ends of mCherry, and co-transformed the plasmid with Cre (Figure 5 & 6). The result we obtained showed that lox5171-mCherry-wtlox performs best, and used it in further analysis (Figure 6).
Figure 5. Schematic diagram of loxP mutant incompatibility test.
Figure 6. Cre excises sequences flanked by homologous loxP sites, but are incompatible with its mutant version.
The above column shows which plasmids are transformed. The 3 middle lanes stand for Cre co-transforming with mCherry flanked on both ends by wildtype loxP (Lane 3), or with wildtype loxP on only one end, the other end being lox2272 (Lane 4) or lox5171 (Lane 5). mCherry flanked with lox2272 or lox5171 on one end does not go through excision so a full-length band was detectable, while mCherry flanked with wildtype loxP on both ends are excised and only a shorter band was seen.
Cre with degradation tags
When our modeling demonstrated to us that the expression level of Cre needs to be much lower than that of RT, we introduced degradation tags. By attaching them to the C terminal of Cre recombinase, the protein would be rapidly recognized and degraded by the E. coli’s native SsrA-SmpB degradation system. This construct could also solve the problem of basal leakage and continued existence after inducer removal.
Figure 7. Degradation tag greatly reduces the protein level at stable state.
WT represents the positive control of EGFP without any tag attachment. The five degradation tags are represented by their last five amino acid sequence. The vertical axis shows the quantitative analysis of EGFP fluorescence (excitation wavelength: 485 nm; detection wavelength: 528 nm), normalized by cell amount (OD600). The fluorescence is quantified by the concentration of green fluorescein, cell number is quantified by the number of silicon beads, both are from the distributed measurement kit. Fluorescence below detection level are eliminated. Error bar stands for the SEM of 3 replicates. t-test is performed between WT and each degradation tag, P<0.0001 (****).
Apart from the native AANDENYALAA tag, we also modified its last three or five amino acids into YALAV, YALVA, YALVV and WVLAA. We tested the stable expression level, as well as the degradation dynamic of each tag by attaching them to the C terminal of EGFP protein and measuring the change in fluorescence level (Figure 7). The stable state expression increases as the number of mutated amino acids increase, or the mutated site nears the N’ of the tag. Supported by our modeling result, we deemed that the XXX tag performs best and chose to use it in further experiments.
Modeling
Figure 8. Recombined Ptarget would occur when RT and Cre is expressed at a proper level.
Dynamics of the percentage of un-recombined / recombined Ptarget among all Ptargets is shown in the upper panel. The distribution of the percentage of substances at the steady-state is shown in the lower panel. Ps: un-recombined Ptarget. Pp: recombined Ptarget. The result that intermediate formed by un-recombined Ptarget and T7RNA polymerase shows that mutation on Parget can accumulate.
Our modelling successfully demonstrated that our system could function and mutation could accumulate along bacteria growth (Figure 8). For detailed explanation of our system, please visit our Modeling page.