We had 3 genomic variants that we named Y1, Y2 and Y3. The samples were normalized to similar concentrations and the PCR was run with 20ng of DNA material.
We ran a 2% agarose gel electrophoresis to check our PCR product size. We prepared two sets of tubes, one destined to the gel electrophoresis and one to be cleaned up as we do not have a UV “table” in our lab.
Results
Figure 1Genomic CUP1 PCR amplification.On the right is the low range ladder, the three lanes correspond to the three genomic variants amplified. All our amplified CUP1s have a size of around 200 base pairs (bp) which matches the theoretical size of CUP1: 183 bp.1
Conclusion We successfully amplified the CUP1 gene using PCR. We could move on and do a PCR clean up on our samples to purify the DNA.
PCR clean-up
2021-07-12
Aim Purify DNA fragments from the PCR mix to remove excess nucleotides and primers.
Results To verify our digestion worked properly, we ran a 1% agarose gel electrophoresis of our plasmid digested with different combinations of our restriction enzymes: BamHI alone, NheI alone, BamHI and NheI together or no enzymes.
Figure 2Gel picture of the digested vector backbone.Lane 1: GeneRuler 1 kb DNA Ladder. Lane 2: Plasmid digested with BamHI HF. Lane 3: Plasmid digested with NheI HF. Lane 4: Plasmid digested with both BamHI HF and NheI HF. Lane 5: Undigested plasmid. The bands in lane 2, 3 and 4 are slightly bigger than 6000 bp while the band in lane 5 is above the 10,000 band of the ladder. pCTcon2-V5 is 6468 bp.
The size of our plasmid, pCTcon2-V52, is 6468 bp. We should observe bands of this size when cutting the plasmid with either BamHI or NheI.
Linear DNA encounters less friction while running through the gel than circular DNA. Thus, on our gel, we expect to see the plasmid cut with either BamHI or NheI run faster than the undigested plasmid, as the plasmid would have been linearized.
The size of the insert between BamHI and NheI is 189 bp while the size of the plasmid without the insert is 6279 bp. In lane 4 where our plasmid was cut with both restriction enzymes, we thus expect a band near 6000 bp. Since the size difference is so small with the plasmid that was cut with only one restriction enzyme, we do not expect a difference on the gel. Furthermore, the insert band of 189 bp is too small to be seen on this gel thus we do not expect to see it.
Conclusion We were able to digest the pCTcon2V5 plasmid with BamHI and NheI and it worked properly. Thus we could continue with the next experiment.
Gibson assembly
2021-07-13
Aim Gibson assembly will allow to anneal our CUP1 gene to the now linearized vector backbone. The recomposed gene-containing plasmid will be ready to be transformed into bacteria.
Conclusion We moved on to bacteria transformation and this would confirm that the Gibson Assembly worked well.
Bacteria transformation and plating
2021-07-14
Aim Bacteria transformation with the newly assembled plasmid and selection on ampicillin plates. These plates allow for positive selection of the transformed bacteria as the plasmid contains an ampicillin resistance gene.
Plates were put in the inoculator at 37°C overnight. The next morning, we observed some colonies on our plates.
Results
Figure 3Picture of ampicillin plates A, B, C and D.A: pCTcon2-CUP1_Y1-V5 transformed bacteria. B: variant Y2. C: variant Y3. D: negative control, linearized pCTcon2-V5 transformed bacteria.
The colonies observed have the AmpR gene, that gives resistance to the bacteria against ampicillin.
Conclusion Our transformed bacteria had successfully taken up the plasmid containing the ampicillin resistance gene. Moreover, as we had asserted that the digestion was successful and as we know that bacteria can only function properly with circular DNA we could conclude that our Gibson assembly was successful.
Colony picking and culture inoculation
2021-07-15
Aim pCTcon2-CUP1-V5 amplification
Protocol Under the flame, with a sterile pipette, scoop up a colony and release the pipette tip into a 5 ml tube. Add 4 ml of sterile LB medium and place it in the shaker at 37°C.
Conclusion We could move on to the next step: Miniprep.
Miniprep
2021-07-16
Aim Purify the plasmid DNA from our overnight culture of bacteria transformed with the pCTcon2-CUP1-V5 plasmid.
Once the Miniprep performed, we measured our DNA concentration as well as purity using Nanodrop.
Results
Table 1Quantification of digested samples
Sample
Concentration
A260/A280
Sample 1 (Y1)
446.0 ng/μl
1.88
Sample 2 (Y2)
350.4 ng/μl
1.88
Sample 3 (Y3)
330.0 ng/μl
1.98
The last column is the absorbance ratio A260/A280 and represents the quality of the DNA. We consider a sample to be of good quality if the ratio lies between 1.8 and 2.
Conclusion These samples had a high concentration of DNA and appeared to be of good quality.
Sequencing
2021-07-16
Aim Compare our now amplified plasmid with the theoretical sequence to observe potential mutations that may have occurred during amplification in bacteria.
Protocol Send samples to a DNA sequencing company (Mycrosynth in our case).
We sent 2 clones for each transformant variant: YXCX where X stands for the number of the variant and clone respectively.
Results The alignment of the sequencing results are on this document, where the sequences number 1 to 12 correspond to, respectively Y1C1 forward, Y1C1 reverse and so on until Y3C2 reverse.
Y2C1 has a mutation (G to C) in both forward and reverse strands. We thus choose to ignore this one. It also seems like the forward primer of Y2C2 did not anneal, therefore we do not have a full strand. To avoid errors, we decide to only keep Y1 and Y3 variants.
Conclusion We decided to proceed with only Y1 and Y3 genomic variants and had two clones for each of them.
Yeast growth
2021-07-19
Aim Monitor the growth of yeast culture to identify the time window by which they are competent for transformation, that is a measured OD600 of 0.8 to 1.
Protocol In a 15 ml falcon tube, incubate 12 μl of wildtype yeast in 4 ml of YPD in a shaker at 30°C and 220 rpm. Measure the absorbance at 600 nm (OD600) every hour using a spectrophotometer. A calibration curve may be needed before the actual measurements.
We made 3 technical replicates and 2 biological replicates, but limited by the space on the shaker we could only do one.
Results On average between 2 experiments, OD600 of 1 was reached within approximately 12 hours.
Yeast transformation
2021-07-20
Aim Transform the yeast with our pCTcon2-CUP1-V5 plasmid and plate our transformants on a selective medium.
We prepared 3 samples: a negative control (no plasmid), a positive control (our plasmid backbone without the CUP1 insert) and the plasmid containing the CUP1 gene.
Our yeast strain, EBY100, is incapable of synthesizing tryptophan on its own and needs it from an external source, thus when plated onto tryptophan-less medium plates, they cannot grow. Our plasmid backbone encodes a Trp+ gene allowing the yeast to grow without tryptophan in its medium, allowing for positive selection of our yeast transformants.
We picked clones from each plate and amplified them in liquid SDCAA medium. In total we picked 6 clones from the yeast transformed with the recombinant plasmid : A, B, C, D, E and F.
Results
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Figure 4Plates A, B and C with transformed yeast.(a) Plate A: yeast transformed with pCTcon2-CUP1-V5 plasmid, colonies observed. (b) Plate B: positive control, yeast transformed with pCTcon2-V5 plasmid, colonies observed. (c) Plate C: negative control, wild type yeast (transformation with only water and no DNA). One colony was observed, probably a mutant. Small white dots are yeast colonies.
Conclusion We could conclude that the yeasts had been transformed successfully. We moved on to the next step: checking protein expression.
Western blot 1
2021-07-30
Aim Verify the expression of our protein of interest in yeast.
We used two mouse anti-V5 and anti-Tubulin as primary antibodies and a goat anti-mouse as a secondary.
The first Western blot was performed on clone A and B and had for negative controls, the wild type yeast as well as the yeast transformed with the plasmid backbone cultured in SDCAA (without galactose). For positive controls, the yeast transformed with the plasmid backbone cultured in SGCAA (with galactose) as well as a fly tubulin were runned onto the gel.
Figure 5Western blot 1.Figure 6Analysis of western blot 1.Aga2-V5 has a molecular weight of 21.3 kDa.
We loaded the following samples onto our gel: pCTcon2-V5 in SDCAA, pCTcon2-V5 in SGCAA, EBY100 (WT) in YPD, and two pCTcon2-CUP1-V5 clones (A and B) in SGCAA and SDCAA each. We obtained a band of ~ 50 kDa in the WT and plasmids with and without galactose, showing the presence of α-tubulin (49.8 kDa3) in our samples. We also have a band of ~ 20 kDa in plasmid + galactose showing that our Aga2-V5 protein was expressed (figure 6a). Figure 6b is showing the fly control expressing tubulin as well as Relish-V5, a protein harboring the V5 tag. Figure 6c shows the ratio of V5 signal versus tubulin.
However we didn’t obtain any signal, neither for the tubulin nor the protein of interest in the clone A and B lines.
Conclusion The clones were probably picked prematurely, and thus didn’t show anything on the gel. That’s why a second Western Blot was performed in order to check protein expression in our 4 other clones that were picked later: C, D, E and F.
Immunostaining 1
2021-08-09
Aim Check the expression of our CUP1 protein on the surface of our yeast transformants, clones C, D, E and F.
We used a mouse anti-V5 primary antibody and a goat Alexa Fluor 488 conjugated anti-mouse as a secondary antibody.
The results were observed under a confocal microscope. For each clone, two samples were analyzed, one where the cells’ membrane were permeabilized in order to see if the protein was expressed intracellularly and one where the cells’ membrane were not permeabilized.
Figure 7Clone C.Nuclei are stained with DAPI in blue and the V5 tag is stained with a green fluorophore.Figure 8Clone D.Figure 9Clone E.Figure 10Clone F.
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Figure 11Positive control: yeast transformed with the plasmid backbone (no insert).(a) DAPI channel (b) green fluorescence (c) merged channel.
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Figure 12Negative control: wild type yeast.(a) DAPI channel (b) green fluorescence (c) merged channel.
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Figure 13Negative control: yeast transformed with the plasmid backbone (no insert), membrane permeabilized.(a) DAPI channel (b) green fluorescence (c) merged channel.
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Figure 14Negative control: yeast transformed with the plasmid backbone (no insert), without antibody 2.(a) DAPI channel (b) green fluorescence (c) merged channel.
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Figure 15Negative control: yeast transformed with the plasmid backbone (no insert), without antibody 1.(a) DAPI channel (b) green fluorescence (c) merged channel.Figure 16Clone F, higher magnification.
All 4 clones showed expression of CUP1 on their surface. We assumed the expression is at the surface because: first, the antibody does not enter the cell, second, we can see a darker circle around the cell on figure 16 (higher magnification) representing the membrane and third, we did some controls with membrane permeabilized samples (figure 13). The membrane permalized samples show little signal that corresponds to the signal in negative controls WT (figure 12), backbone without antibody 1 (figure 14) and backbone without antibody 2 (figure 15), which we can assume is noise or unspecific binding of antibodies.
Unfortunately, not all cells were positive in one sample, which means that either our subsets of cells were not monoclonal and that not all the cells had the plasmid or that they all had the plasmid but weren't all expressing it.
Conclusion In order to see if the expression level differences were due to polyclonality of the yeast population we streaked the cells on a plate and picked a single colony in order to perform an immunostaining on a monoclonal population of cells. This resulted in no different results than before, we still had an heterogeneous population with cells displaying signal and cells with no signal. We concluded that the induction process was not 100% efficient and this was supported by the results of the paper from Chao et al.2
Western blot 2
2021-08-11
Aim Check the overall expression of our protein of interest in our yeast transformant clones C, D, E and F.
We obtained bands of approximatively 25 kDa in positive control as well as in clones C, E and F, showing that our protein of interest was present in these samples. Unfortunately we didn’t have any signal for the α-tubulin so we couldn’t normalize the signal and compare the expression level of the protein. As expected, in the negative control pCTcon2-V5 backbone plasmid uninduced, no band is visible.
We also noticed two bands on the gel, and verified the DNA sequence and saw that a second methionine, a start codon, is present downstream of the first one and in frame with the promoter, resulting in a second expressed protein, probably truncated. Unfortunately this shorter protein is the most expressed one, corresponding to the signal intensity.
Conclusion We’ve decided to continue the experiments with our 3 clones C, E and F, since D didn’t show any expression.
Western blot 3
2021-10-14
Aim Check the overall expression of our protein of interest in our yeast transformant clone F and normalize its signal relatively to α-tubulin.
Since we did not get any signal for tubulin in Western Blot 2, we decided to re-do the western with clone F to have a nice protein quantification plot.
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Figure 18Western blot 3.(a) Image of the nitrocellulose membrane with yeast transformed with plasmid backbone pCTcon2-V5 without insert, Clone F, i.e. yeast transformed with pCTcon2-CUP1-V5, both cultured with galactose in their media and wild type yeast (cultured in YPD). (b) Protein quantification plot, the black bars represent the signal for the backbone plasmid and the grey ones are for the Clone F. On the left is represented the signal of V5 tag and on the right, the tubulin signal. (c) is the normalized V5 signal for tubulin (V5 in % of tubulin).
We obtained a tubulin signal in both plasmid backbone pCTcon2-V5 and clone F (pCTcon2-CUP1-V5) but not in wild type yeast. This was probably due to the fact that a smaller quantity of cells were loaded on the gel, since the pellet was smaller in the WT sample. We could normalize the V5 signal relatively to tubulin and the result is shown on figure 18c. We can see that clone F expresses the protein a bit more than the backbone, which is consistent with what we observed in immunostaining.
Conclusion We were able to show that our membrane protein fusion was expressed in our engineered yeast and could quantify its relative expression in our clone F as well as in the positive control, the backbone plasmid transformed yeast.
Yeast growth curves
Aim Check for the yeast growth differences between the strains. This will allow us to see whether engineering the yeast would modify its ability to grow or survive.
Protocol
Prepare yeast cultures of known initial concentration for each of the strain to be tested (3 replicates per sample).
Load a 96 wells-plate with your samples (2 or 3 technical replicates for each sample).
Put in an overnight shaking microplate reader at 30°C, measuring the OD600 every hour.
If you don’t have this equipment at your disposal, place your yeast cultures in a 30°C shaker and collect samples directly from it every hour to measure the OD600 in a spectrophotometer or microplate reader.
We measured the growth of our transformed yeast (pCTcon2-CUP1-V5) in SGCAA, and we compared them to the growth curves of plasmid backbone (pCTcon2-V5) in SDCAA and in SGCAA and the wild type yeast (EBY100).
Results
Figure 19Yeast growth curves of different strains.
Copper absorption assay
Aim Measure the yeast ability to remove copper in solution for the different strains.
All measurements were performed with three biological replicates and three technical replicates.
The concentration of copper in the media was set at 4 mg/l for all copper absorption assay. We tested different conditions to identify the cause of bad copper absorption with our engineered yeasts (see the results below and the Results page).
Results
Figure (x)Measuring copper concentration over time with varying WT (EBY100) yeast concentration in the media.The yeast density was determined as a function of OD600 (OD600 of 1 corresponds to approximately 3 x 107 cells/ml).Figure (x)Measuring copper concentration over time in the media with WT (EBY100) yeast or yeast expressing CUP1.OD600 of the yeast culture was 4.0.
Results Wild type yeasts absorbed copper well, and we were not able to make our engineered yeast with the CUP1 monomer work better. However, the higher wildtype yeast concentration, the faster the copper could be absorbed.
Cloning the dimers
Plasmid digestion
2021-08-25
Aim Digest the backbone plasmid with restriction enzymes in order to linearize it for cloning by Gibson assembly.
To check that digestion worked properly, we ran an agarose gel electrophoresis of our plasmid digested with different combinations of our restriction enzymes: BamHI alone, NheI alone, BamHI and NheI and none.
Results To verify our digestion worked properly, we ran a 1% agarose gel electrophoresis of our plasmid digested with different combinations of our restriction enzymes: BamHI alone, NheI alone, BamHI and NheI together or no enzymes.
Figure 20Result of the backbone digestion on a 0.7% agarose gel.Lane 1: GeneRuler 1 kb DNA Ladder. Lane 2: Plasmid digested with BamHI HF. Lane 3: Plasmid digested with NheI HF. Lane 4: Plasmid digested with both BamHI HF and NheI HF. Lane 5: Undigested plasmid. The bands in lane 2, 3 and 4 have an approximate size of 6000 bp while the band in lane 5 is around the 8,000-10,000 bands of the ladder.
The size of our plasmid, pCTcon2-V52, is 6468 bp. We should observe bands of this size when cutting the plasmid with either BamHI or NheI.
Linear DNA encounters less friction while running through the gel than circular DNA. Thus, on our gel, we expect to see the plasmid cut with either BamHI or NheI run faster than the undigested plasmid, as the plasmid would have been linearized.
The size of the insert between BamHI and NheI is 189 bp while the size of the plasmid without the insert is 6279 bp. In lane 4 where our plasmid was cut with both restriction enzymes, we thus expect a band near 6000 bp. Since the size difference is so small with the plasmid that was cut with only one restriction enzyme, we do not expect a difference on the gel. Furthermore, the insert band of 189 bp is too small to be seen on this gel thus we do not expect to see it.
Conclusion pCTcon2V5 plasmid was successfully linearized by BamHI and NheI digestion.
Gene PCR
2021-08-26
Aim Amplify our CUP1 dimers sequences by Polymerase Chain Reaction (PCR) and add Gibson assembly overlap sequences at both extremities.
These dimer sequences are made of two copies of CUP1 fused by a linker that we designed (see the Design page for information). We ordered the sequences from Twist and went through a PCR to amplify and add the overlap sequences we needed for the Gibson assembly. We have 7 different dimer sequences, each discriminated by a unique linker. The samples were normalized to similar concentrations and the PCR was run with 20 ng of DNA material.
We then ran a 2% agarose gel electrophoresis to check the size of our 7 samples after PCR.
Results
Table 3Information about the dimers and the resulting transformed yeast subclones.
As we were only interested in the first bands of 420bp that correspond to the full dimers sequence, we directly cut out the DNA from the gel using a UV transilluminator. This would then be followed by a PCR clean up.
Conclusion We successfully amplified our CUP1 dimers by PCR and extracted the DNA of interest from the agarose gel.
PCR and digestion clean-up
2021-08-26
Aim Purify DNA fragments and digested backbone samples.
Results Samples are ready for the following Gibson Assembly with our digested backbone.
Gibson assembly
2021-08-26
Aim Gibson assembly will allow us to fuse our CUP1-dimers to the now linearized vector backbone. The recomposed gene-containing plasmid will be ready to be transformed into bacteria.
Conclusion We moved on to bacteria transformation and this would confirm that the Gibson Assembly worked well.
Bacteria transformation and plating
2021-08-26
Aim Bacteria transformation with the newly assembled plasmids and selection on ampicillin plates. These plates allow for positive selection of the transformed bacteria as the plasmid encodes an ampicillin resistance gene.
Plates were put in the inoculator at 37°C overnight. The next morning, we observed some colonies on our plates.
Results We observed colonies on every plate with our dimers transformants and on the positive control provided with the NEB Gibson Kit. We did not observe any colonies on the plate with the bacteria transformed with water (negative control).
Conclusion We could therefore assert that our Gibson assembly and transformations were successful. Our positive selection with ampicillin allowed us to be sure that the bacteria on the plates had successfully taken up our recombinant plasmids.
Colony picking and culture inoculation
2021-08-27
Aim Pick 3 colonies per plate that have survived Ampicillin treatment and grow them in liquid medium in order to amplify our plasmids DNA.
Protocol Under the flame, with a sterile pipette, scoop up a colony and release the pipette tip into a 5ml tube. Add 4ml of sterile LB media and place in the shaker at 30°C.
Results We had turbid tubes, indicating that the bacteria had grown.
Conclusion We have sufficient bacterial liquid culture for plasmid purification by Miniprep.
Miniprep
2021-08-30
Aim Purify the plasmid DNA from our overnight culture of bacteria transformed with the pCTcon2-CUP1-dimer-V5 plasmids.
Once the Miniprep performed, we measured our DNA concentration as well as purity using Nanodrop.
Results
Table 2Quantification of digested samples
Sample
Concentration
A260/A280
2A
143.7 ng/μl
1.83
2B
111.7 ng/μl
1.78
2C
102.3 ng/μl
1.69
2D
127.9 ng/μl
1.61
2E
77.8 ng/μl
1.86
2F
99.8 ng/μl
1.79
2G
89.5 ng/μl
1.89
We had three genomic variants per construct, hence we use the number 2 for dimer (version 2 of our yeast expressing CUP1 copies at their surface), then the letter A to G to identify the linker. We measure their concentration using Nanodrop and send them to be sequenced.
Sequencing
2021-08-31
Aim Compare our now amplified plasmid with the theoretical sequence to observe potential mutations that may have occurred during amplification in bacteria.
Protocol Send samples to a DNA sequencing company (Mycrosynth in our case).
Results Using SnapGene we selected, for each dimer, a variant that had no mutation in the reading frame.
Conclusion We kept only the clones without mutations for the next steps. We could move on to transform our engineered plasmids in the yeast cells.
Yeast transformation
2021-09-01
Aim Transform the yeast with our pCTcon2-dimer-V5 plasmids and plate our transformants on a selective medium.
According to the Yeast Growth Curve previously made during the version 1 (CUP1 monomer) experiments, we let yeast grow in YPD for approximately 10 hours in order for them to reach an appropriate OD600 = 8 concentration. We prepared 9 samples: a negative control (no DNA, just water), a positive control (our plasmid backbone without insert) and each 7 plasmids containing a dimer. After transformation, we plated our transformants on plates that do not contain tryptophan (SGCAA agar plates), for positive selection as the plasmid encodes a Trp+ gene, this allows the transformed yeast to survive on such a medium.
Results The negative control (water) did not show any colonies and the positive control (plasmid backbone) had many colonies. We could also observe colonies on our transformants plates.
Conclusion We picked clones from each transformed plate (Table 1). Each time a new clone is picked, it is amplified in liquid media (SDCAA at first but then SGCAA in order to induce the protein expression).
Western blot 2
2021-09-14
Aim Verify the expression of our protein of interest in yeast.
The first Western Blot was performed on all our dimers. The V5 tag is part of the membrane display system which itself is under the Gal promoter. Thus expression (and detection) should only occur in the presence of galactose.
Results The dimers have a molecular weight of 28.6 kDa, whereas the α-tubulin has a molecular weight of 49.8 kDa.
Figure 21Image of Western Blot of gel 1.Figure 22Information about the dimers and the resulting transformed yeast subclones.Figure 23Protein quantification.Figure 24Protein quantification: normalized V5.
We loaded the following bands onto our Western blot: samples from yeast subclones A1 to A6 and B1 to B3 in SGCAA. We obtained a band of ~ 30 kDa (figure 22), this size corresponds to the dimers theoretical size. Figure 23 shows the signal analysis of the bands done on the ImageJ software and figure 24 shows the ratio of V5 signal versus tubulin.
Figure 25Image of Western Blot of gel 2.Figure 26Figure 27Protein quantification.Figure 28Protein quantification: normalized V5.
We loaded the following samples onto our Western blot: sample from yeast subclones B4 to B6 and F1 to F6 in SGCAA. We obtained a band of ~30kDa (9a) corresponding to the dimer’s theoretical size. Figure 9b shows the signal analysis of the bands done on the ImageJ software and figure 9c shows the ratio of V5 signal versus tubulin.
Figure 29Image of Western Blot of gel 3.Figure 30Figure 31Protein quantification.Figure 32Protein quantification: normalized V5.
We loaded on this gel two yeast subclones for dimers 3, 4 and 5, and as controls, the plasmid backbone and the wild type.
Figure 33Image of Western Blot of gel 4.Figure 34Protein quantification.Figure 35Protein quantification: normalized V5.
We were able to see a band of approximately 30 kDa in each dimer sample, however other smaller bands were also visible. These are probably due to the second reading frame starting at the downstream second methionine.
Conclusion Our yeasts cells express two different proteins, one whose size matches the theoretical size of our dimers, and another corresponding to a second reading frame in our backbone. For the following experiments, we kept the subclones who have the best tub/V5 ratio taking into account the initial V5 quantification : A4, B1, C2, D2, E2, F3 and G2.
Immunostaining 2
2021-09-25
Aim Check the extracellular membrane expression of our dimers.
We used a mouse anti-V5 as primary antibody and a goat anti-mouse coupled with the Alexa Fluor Plus 488 as secondary. The results were observed under a confocal microscope.
Results
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Figure 36Nuclei are stained with DAPI in blue and the V5 tag is stained with a green fluorophore.(a) Yeast subclone A4, dimer 1 (b) Yeast subclone B1, dimer 2 (c) Yeast subclone C2, dimer 3 (d) Yeast subclone D2, dimer 4 (e) Yeast subclone E2, dimer 5 (f) Yeast subclone F3, dimer 6 (g) Yeast subclone G2, dimer 7. All our yeast clones are expressing their corresponding CUP1 dimer at their surface, although in one sample not all the cells express it. This is due to the fact that the expression yield is not 100%. We can see that some dimers are better than others, for example dimers 6 and 7 have good expression efficiency while dimer 1 have lower expression yield. We also don’t have the same amount of cells in each sample, due to experiment manipulations and not reliable OD measurements. Fortunately we can see that our protein of interest is expressed at the surface of the cell because of the darker green circle representing the cell membrane. This can be better seen on a higher magnification image.
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Figure 37Picture of the immunostaining of wild type yeast.(a) DAPI channel (b) green fluorescence channel (c) merge channel.
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Figure 38Picture of the immunostaining of plasmid backbone uninduced (- galactose) in yeast.(a) DAPI channel (b) green fluorescence channel (c) merge channel.
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Figure 39Picture of the immunostaining of plasmid backbone induced (+ galactose) in yeast, without the secondary antibody.(a) DAPI channel (b) green fluorescence channel (c) merge channel.
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Figure 40Picture of the immunostaining of plasmid backbone induced (+ galactose) in yeast, without the primary antibody.(a) DAPI channel (b) green fluorescence channel (c) merge channel.
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Figure 41Positive control : Picture of the immunostaining of plasmid backbone induced (+ galactose) in yeast.(a) DAPI channel (b) green fluorescence channel (c) merge channel. Our positive plasmid backbone control is expressing the yeast surface display at its membrane.Figure 42Immunostaining of Clone F (CUP1 monomer - pCTcon2-CUP1-V5).40x, green channel.
Conclusion All of our transformed yeast expressed the dimer at their surface, although the expression efficiency is not 100%. Dimer B and D seemed to be contaminated and had a lot of noise in the GFP signal. The results of dimer A, C, E, F and G are good.
Yeast growth curves
Aim Check for the yeast growth differences between the strains. This will allow us to see whether engineering the yeast would modify its ability to grow or survive.
Protocol
Prepare yeast cultures of known initial concentration for each of the strain to be tested (3 replicates per sample).
Load a 96 wells-plate with your samples (2 or 3 technical replicates for each sample).
Put in an overnight shaking microplate reader at 30°C, measuring the OD600 every hour.
If you don’t have this equipment at your disposal, place your yeast cultures in a 30°C shaker and collect samples directly from it every hour to measure the OD600 in a spectrophotometer or microplate reader.
We measured the growth of our 3 transformed yeast clones C, D and E as well as the plasmid backbone in SDCAA and in SGCAA. We also loaded the remaining wells with medium alone, without any yeast in it.
Results
Figure 43Yeast growth curves of different strains in SGCAA with different copper copper concentrations.
Copper absorption assay
Aim Measure the yeast ability to remove copper in solution for the different strains.
Protocol We grew our various engineered yeast strains in liquid media containing copper and we collected 2 ml aliquots of the culture at different time points after addition of yeasts in the media (t0, 15 min, 60 min, 90 min, 120 min). The yeast cultures were prepared in 500 ml Erlenmeyer flasks, and incubated at 30°C under 200 rpm shaking. Each of the collected samples were then slowly centrifuged (200g for 10 minutes) to separate the yeast cell from the media and copper sulfate solution. Note that the centrifugation step was performed at a low speed to avoid any cell lysis, hence preventing the presence of yeast components, free or bound to copper ions, in the supernatant. This is done mainly for two reasons. First, the absorbance wavelengths of yeast and copper are in the same spectrum range. Thud, any contaminations of yeast in the supernatant could lead to alteration and artefact values in the OD measurements, resulting in data with a higher concentration than reality. Second, if any pieces of yeast bound to copper ions would end up in the supernatant, these ions would also be detected by the spectrophotometer, also leading to values with a higher concentration than the one that would have been actually removed from our yeast. Once the centrifugation step was finished, the pellet, mainly characterized by yeast cells, and the supernatant, mainly characterized by media and copper sulfate, are isolated from each other and collected for different chemical downstream processes.
Copper concentration measurement (see also Cuprizone Solution Preparation and Copper Concentration Measurement protocol) In this assay, the aforementioned supernatant is mixed with cuprizone and brought to a slightly basic (pH ~ 8) condition. Cuprizone [oxalic acid bis(cyclohexylidene hydrazide)] is a selective and sensitive copper-chelating agent, typically used in spectrophotometric determination of copper(II). Indeed, this chemical compound is highly sensitive to copper and, once it interacts with a copper solution, it forms a complex that absorbs at wavelengths of 580-600 nm. The absorbance at 580-600 nm of the solution is therefore proportional to the amount of copper. However, according to Messori et al., the complex with Cu(II) and cuprizone is unstable because of the fast decomposition of the reagent, also affecting the stability of the absorbance over time4. Therefore, given the strong instability of the CuCPZ complex, we analyzed its absorbance kinetics to find a stable plateau, hence an optimal time to perform the measurements, avoiding any substantial variance in the measurements. We used a spectrophotometer UV-1600PC (#634-6001) with a filter allowing the measurement of the absorbance at 600 nm for that purpose.
All measurements were performed with three biological replicates and three technical replicates.
The concentration of copper in the media was set at 4 mg/l for all copper absorption assay. We tested different conditions to identify the cause of bad copper absorption with our engineered yeasts (see the results below and the Results page).
Results
Figure 44CUP1 surface expression in SGCAA-cultured yeast interferes with endogenous bio-accumulation The (YPD) and (SGCAA) in brackets depict the medium in which the yeast grew and are staying in during the experiment.
Figure 45Interference with endogenous bio-accumulation is not related to CUP1 expression nor the surface system itself.
Figure 46Rich medium is needed for bio-accumulation to occur.
Conclusion We were not able to make our engineered yeast work better than the wild type one, but we found out that the medium in which the yeast grows is an important factor to take into account when performing this kind of bioremediation experiment.
