<!DOCTYPE html> Results



During spring season, fruit cultivators throughout Switzerland incur enormous losses due to frost each year. A widespread and renowned crop in the Suisse Romande region are apricots, which were particularly affected by the cold this year. The aim of our project is to develop three different approaches to tackle this problem. We focused on different aspects of this issue and either inhibited the formation of ice crystals directly by using antifreeze proteins (AFP) or found a way to inhibit the production of ice nucleation proteins by Pseudomonas syringae syringae, which will reduce the amount of ice crystals that are formed during the late spring freezes.

Key Achievements

  • We successfully constructed 6 new composite parts containing the various AFP genes.
  • We expressed and purified RiAFP and DcAFP.
  • We were able to determine the thermal hysteresis of RiAFP and DcAFP, and therefore proved that they are functional.
  • We successfully demonstrated that the purified AFPs have a strong protective effect against frost damage on plants.
  • We managed to extract tailocins from the killer strain and visualised them using electron microscopy.
  • We confirmed the ability of our tailocins to kill our target strain thanks to an overlay assay.
  • We were able to determine the effect of P. syringae syringae on freezing time.
  • We proved that our tailocins affect freezing time of a solution containing P. syringae syringae.
  • We successfully inserted all three sgRNAs into three different plasmids.
  • We successfully inserted the donor sequence into two of the three plasmids carrying sgRNA.

Antifreeze Proteins

The aim of this part of the project was to produce an antifreeze protein solution, which can bind to ice crystals during late spring freeze and therefore protect the crops from frost damage.

Plasmid Cloning

Initially, we first amplified the various AFP genes by PCR and linearized the plasmids by restriction enzyme digestion. Then we assembled the various constructs by Gibson Assembly, transformed E. coli BL21 (DE3) cells by electroporation, and performed colony PCR to check if the genes were correctly assembled in the various constructs (Fig.1). We obtained the expected band sizes and confirmed their sequence by sanger sequencing. We therefore concluded that we managed to successfully clone FfIBP, DcAFP, and RiAFP in E. coli BL21 (DE3) cells.

Figure 1 | Colony PCR check. Left to right: L – 1 kb DNA Ladder, 1 – pCold-I-DcAFP (1179 bp), 2 – pET-17b-DcAFP (1231 bp), 3 – pCold-I-FfIBP (927 bp) 4 – pET-17b-FfIBP (979 bp), 5 – pCold-I-RiAFP (588), 6 – pET-17b-RiAFP (640 bp)

Expression and Purification

After this, we induced expression of the various AFPs with IPTG and ran a total cell SDS-PAGE (15%) of the E. coli BL21 (DE3) cultures stained with Coomassie blue. We used two different vectors, pET-17b and pCold-I, to compare how the promoters influenced expression efficiency for both AFPs. The expression was successful for FfIBP and RiAFP in both pET-17b and pCold-I plasmids. We observed thicker bands that matched the weights of the different antifreeze proteins (FfIBP 28.4kDa, RiAFP 12.8 kDa). The pCold-I vector worked better for both RiAFP and FfIBP, and the bands on the gels appeared to be thicker for this plasmid (Fig.2 and Fig.3). This can be explained by the fact that the natural expression of AFPs happens at cold temperatures, and any expression system adapted to these conditions will be more efficient. We induced expression at 15°C. At this temperature the 5’UTR of the mRNA from the pCold-I plasmid adopts a highly stable secondary structure and the TEE sequence then improves translation initiation of our AFP genes through a ribosome trapping process. The pET-17b plasmid does not have these features and is therefore less efficient. We can conclude that the pCold-I plasmid is a better system for expression of cold-stable proteins like AFPs.

Figure 2 | SDS-PAGE electrophoresis before and after FfIBP purification. RiAFP was expressed in E. coli BL21 (DE3) using both pCold-I and pET-17b vectors. A) pCold-I-FfIBP: L – 10 to 250 kDa Protein Ladder, 1 – Bacterial lysate, 2 – Purified FfIBP. B) pET-17b-FfIBP: L – 10 to 250 kDa Protein Ladder, 3 – Bacterial lysate, 4 – Purified FfIBP.
Figure 3 | SDS-PAGE electrophoresis before and after RiAFP purification. RiAFP was expressed in E. coli BL21 (DE3) using both pCold-I and pET-17b vectors. Left to right: L - 10 to 250 kDa Protein Ladder, Bacterial Lysate: 1 - pET-17b-RiAFP, 2 - pCold-I-RiAFP. Purified Protein: 3 - pET-17b-RiAFP, 4 - pCold-I-RiAFP.

From these initial test purifications with magnetic beads we were able to conclude that the expression and purification of FfIBP and RiAFP worked. Therefore, we upscaled the protein purification and induced expression in a 1-liter culture. After that, we performed two purification steps: the first using the His-tag affinity column and the second a gel filtration. Then, we eluted the protein in various fractions, ran a SDS-PAGE (15%) of the fractions containing the purified protein stained with Coomassie blue, and finally pulled the fractions containing FfIBP (Fig.4). Finally, we measured the final concentration of our protein with the nanodrop and obtained 178µM or 4.53mg/mL.

Figure 4 | SDS-PAGE electrophoresis after gel filtration purification of FfIBP. FfIBP was eluted into multiple fractions. Left to right: L - Protein Ladder, 10 to 250 kDa, fractions B3 to B12.

The upscaling for RiAFP did not work at first and we did not observe any bands that matched the weight of the protein on the total cell SDS-PAGE (15%) of E. coli cultures stained with Coomassie blue (data not shown). For the FfIBP purification, we used a baffled flask to grow the 1-liter culture for E. coli BL21 (DE3) expressing the construct. The edges of the flask maximize oxygen transfer to the culture media, but in some cases, they can also interfere with protein expression. Since we could not upscale the RiAFP expression using this method, we used a flask without baffles instead and succeeded that way. Then, we performed the same purification steps as we did for FfIBP purification. After gel filtration, we eluted RiAFP in various fractions, ran an SDS-PAGE (15%) of the fractions containing the purified protein stained with Coomassie blue, and finally pulled the fractions containing RiAFP (Fig.5). RiAFP does not contain any Tryptophan and Tyrosine residues in its sequence, hence, we could not measure its final concentration using a nanodrop and used the Bradford method to estimate it instead. In a standard reaction, 25µL of BSA concentration and 5µL of RiAFP concentration were mixed with 200µL of Bradford reagent. Absorbance was then measured at 595 nm. The calibration curve (Fig.6) is needed to calculate sample concentration. We obtained 0.75mg/mL or 58.94µM.

Figure 5 | SDS-PAGE electrophoresis after gel filtration purification of RiAFP. RiAFP was eluted into multiple fractions. Left to right: L - Protein Ladder, 10 to 250 kDa; fractions B2 to B8.
Figure 6 | Bradford assay standard curve of concentration versus absorbance. The concentration of RiAFP (in mg/mL) was determined using the equation y=0.0729x-0.015 with an R2 value of 0.9988, where y is absorbance and x is concentration.

The expression didn’t work for DcAFP in pET-17b. When we ran a total cell SDS-PAGE (15%) of E. coli cultures stained with Coomassie blue, we didn’t observe any thicker bands that would have corresponded to the protein. The root of the problem might be that the protein aggregate to form inclusion bodies. To remedy this problem, we attempted to optimize expression by using various concentrations of IPTG. Once more, we ran a total cell SDS-PAGE (15%) of E. coli cultures stained with Coomassie blue but did not observe thicker bands at the desired protein weight (Fig.7).

Figure 7 | SDS-PAGE electrophoresis of bacterial lysate of DcAFP expressed using the pET-17b plasmid. Expression was induced for 24 hours at 15°C with various concentrations of IPTG. L –Protein Ladder, 10 to 250 kDa, 1 –0mM , 2 –0.01mM, 3 - 0.1mM , 4 - 0.2mM, 5 - 0.5mM

Since we couldn’t optimize expression of DcAFP by adapting the concentration of IPTG, the next step would be to purify the protein from inclusion bodies. We adapted a protocol [1] to our DcAFP construct, which could be used in the future.

Results of AFP TH Values

The measurements we performed on our AFP solutions were done in part with the machine we designed, FROZONE. This device, coupled with a software we created, allows the precise control of the temperature of a copper plate placed in a vacuum chamber. We placed a drop of our AFP solutions on the copper plates and visualised their freezing and melting temperatures using a microscope.
This allowed us to precisely determine the freezing and melting temperatures of our AFP solutions, allowing us in turn to determine the TH values of our proteins at specific concentrations. The values we obtained can be visualised in the following (Fig. 24.).

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Figure 24 | Thermal hysteresis of RiAFP and FfIBP increase with their concentration.Connected scatter plot graph showing the mean of biological triplicate +/- standard deviation. Thermal hysteresis is measured with two different AFPs, RiAFP and FfIBP, at different concentrations (µM).

The data we obtained show that the thermal hysteresis of our proteins increases as their concentration in the solution increases.
From these results, we can conclude that our antifreeze proteins are both functional, as we have successfully measured their respective TH values at two distinct concentrations. With these results, we can also conclude that RiAFP is more effective than FfIBP at a concentration of 50µM.

Result of Protective Effect of AFPs on Plants

The following graph shows the percentage of damage measured on the leaves of Arabidopsis plants that were incubated overnight at -5 °C in various solutions of AFPs or in our buffer solution. Our results show that plants incubated in the buffer alone are significantly more damaged than those incubated in the AFP solutions, therefore confirming the protective effect of our AFPs. We were not able to detect, however, a significant difference between the protection level provided by FfIBP, RiAFP or a mix of both AFPs.

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Figure 25 | Our AFP solutions protect plants from frost damage.The graph shows mean +/- standard deviation. Percentage of Damage on Arabidopsis thaliana leaves measured after overnight incubation in a climatic chamber at -5ºC. Plants were immersed in phosphate-buffered saline (PBS) liquid solution supplemented with two different AFPs: FfIBP [200 µM] (n=13) and RiAFP [50 µM] (n=17) and a mix of them (RiAFP + FfIBP) (n=15) at a concentration of 25 µM and 100 µM, respectively. These solutions were compared with the control in which the plants were immersed in the PBS buffer without any AFP (buffer) (n=17). The damage ratio is significantly different between the buffer and all AFPs. ANOVA, * significant at p<0.05.Damage ratio between the different AFPs are not significant.


In order to kill Pseudomonas syringae pv. syringae B301D, our target strain similar to the frost-damage enhancing Pseudomonas syringae pv. syringae found on apricot trees, we are using tailocins naturally produced by Pseudomonas syrinage pv. aptata DSM50252.

Tailocins Extraction

We successfully extracted tailocins from our killer strain, P. syringae DSM50252.
We first induced the production of tailocins in our killer strain using different concentrations of mitomycin C, i.e. 0.5 or 3 µg ml -1 (cf. tailocin extraction protocol (link)). Tailocin production can be indirectly monitored via the death of the producer bacteria, as tailocins synthesis results in their lysis. We could induce production of tailocins with as little as 0.5 µgml -1 mitomycin C, as both concentration tested led to cell lysis compared to our control flask without addition of mitomycin C (Fig. 1).

Figure 8 | Mitomycin C induces tailocin production in P. syringae DSM50252. Liquid cultures of P. syringae DSM50212 (killer strain) exposed to 0 (A), 0.5 (B) or 3 (C) µgml -1 mitomycin C.

Having induced the production of tailocins by our killer strain, we then set out to test their efficacy in killing our target strain P. syringae syringae B301D. For this, we first proceeded to purify the tailocins from both the mitomycin-induced cell lysates and the non-induced culture as a negative control. We then used a soft agar overlay assay method (as described by Baltrus and Hockett) to assess the lysis of the target cells exposed to our purified tailocins.

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Figure 9 | Purified tailocins lyse P. syringae B301D.(A)Photographs showing P. syringae B301D cell lawns exposed to serially diluted tailocins extracted from P. syringae DSM5012 cultures exposed to 0, 0.5 or 3 µgmL-1 mitomycin C or purification buffer alone (Tn50). The black dot indicates where the sample was dropped. The buffer has no killing activity, as we can see that it does not produce a plaque when we spot it on the plate. (B) Schematic representation of expected morphologies of lysis plaques induced by a serial dilution of tailocins (top) or phages (bottom).

Our results indicate that we successfully purified tailocins from our killer strain and that these tailocins were able to kill our target strain (Fig. 2, A). Indeed, we observed clear concentration-dependent lysis of cells exposed to our diluted purified extract. The control solution, however, which we obtained by following the same extraction protocol but without inducing tailocin production with mitomycin C, showed some mild killing activity(Fig. 2, A ). This could indicate that some tailocins were produced even without mitomycin or that our killer strain produces another bacteriocin or antibiotic efficient against our target. Nevertheless, mitomycin C-induced lysates exhibit greater killing activities compared to the uninduced cultures, confirming that mitomycin C is essential to obtain significant tailocin yield. Additionally, we observed a higher killing activity in the 0.5 µg mL-1 mitomycin-induced extract than in the 3 µgmL -1 (Fig. 2, A), which suggests that the lower concentration led to a better yield of tailocins production.
As explained on the design page (link), our killer strain’s genome contains both a tailocin gene cluster and a prophage. As both can be induced by mitomycin C, we cannot exclude a priori that the killing activity is caused by the phage rather than the tailocins. Nevertheless, phages produce small individual plaques instead of big ones when they are diluted enough, whereas plaques produced by tailocins become gradually smaller and fainter as they are diluted (Fig. 2, B). As we do not observe individual plaques on our plates, it is most likely that the killing activity seen in our experiment is caused by tailocins. However, we cannot exclude that a small amount of phages is present and contributes to cell lysis. To prove that our tailocins only killed our target, further experiments should aim at deleting some of the phage’s genes from our strain’s genome.
As additional proof of the success of our tailocin purification, we performed negative staining of our tailocin extracts and visualized our samples by electron microscopy. This confirmed the presence of tailocins in our mitomycin C-induced extracts (Fig. 3), while no tailocins could be observed in our control sample (non-induced cells). Also, no phages could be seen in our samples, although more concentrated extracts could be required to observe them. By measuring the height and width of our tailocins, and obtained the following results: 168 nm long and 25 nm wide.

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Figure 10 | Electron microscopy images of tailocin extracts. From left to right : The first two images show tailocins extracted from P. syringae DSM50252 when induced with 0.5 µg mL-1 of mitomycin C. Image on the right shows a tailocin found in the sample obtained from cultures with 3 µgmL-1 of mitomycin C.

In parallel to obtaining tailocins directly from P. syringae , we aimed at cloning the tailocoin gene cluster in a plasmid to express it heterologously in Escherichia coli . As the cluster is relatively big (i.e. 14 kb), we first amplified it from P. syringae genomic DNA as four distinct fragments (Fig 4).

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Figure 11 | PCR amplification of the tailocin gene cluster as different fragments.(A) Schematic representation of the tailocin gene cluster from P. syringae B301D. Position of our primers on the gene cluster (in grey) and expected amplicons (in colors) are depicted. (B) PCR amplification of the tailocin gene cluster fragments. PCR products were run on a 1% agarose gel with a 1kb DNA ladder (NEB).

Plasmid Cloning

Having obtained our different fragments, we then attempted to assemble them using Gibson assembly but we were unfortunately unsuccessful. After transformation of E. coli cells with our Gibson products, we obtained some colonies growing on selective media and none in the control (in which we performed a Gibson without one of the fragments). The absence of colonies on our control plate seemed to indicate that the assembly had worked. However, the colony PCRs we later performed revealed that our transformed bacteria carried only the empty backbone ( Fig. 5 ). Due to lack of time, we were unfortunately not able to proceed further.

Figure 12 | Colony PCR of bacteria transformed with our Gibson product. (A) Schematic representation of our backbone with the tailocin gene cluster cloned (left) and empty backbone (right). The primers used for the colony PCR are represented by black arrows. (B) Gel electrophoresis showing amplicons obtained after colony PCR, as depicted in (A) above. Two sets of primers were used: Ta 09 and Ta 11 or Ta 09 and Ta 26 for two isolated transformants (labelled 1 and 2) and the empty backbone (P). In both colonies, the results observed showed that the plasmid did not contain our desired insert. PCR products were run on a 1% agarose gel with a 1kb DNA ladder (NEB)


Our Versus assay was designed to test if the tailocins we produced successfully killed and unarmed Pseudomonas syringae syringae. For this assay, we tested a P.syringae solution at OD1 and a P.syringae solution at OD1, which we had previously incubated with our tailocin solution for an hour. We placed a drop of each of these solutions on our FROZONE machine, set at -15°C. We then measured the time it took for each of the drops to freeze. Figure 27 illustrates our results, obtained using a t-test.

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Figure 27 | Pseudomonas death induced by tailocins increases time before freezing.Graph shows mean of biological triplicate +/- standard deviation. Time measured before observed freezing of NYB liquid solution supplemented with Pseudomonas Syringae Syringae (P. syringae syringae) or P. syringae syringae and tailocin proteins (Tailocin + P. syringae syringae). Times before freezing are significantly different. T-test, * significant at p<0.05, p=0.0114.

We can see that there is a significant difference between the time it took for the P.syringae solution to freeze versus the time it took for the same solution but previously incubated with tailocins to freeze. As the tailocins incubation is the only variable between the two solutions, we can conclude that these protein complexes allow the reduction of P.syringae’s ice nucleation activity.

Measuring the Time Before Freezing - P. syringae Solution

Pseudomonas syringae syringae (PSS) accelerates and facilitates the formation of ice crystals. Theoretically, adding P.syringae to a solution at a sub-zero temperature should shorten the time it takes for this solution to freeze.
To test this hypothesis, we used our FROZONE machine to perform the TTF assay. This assay measures the time it takes for a drop of solution to freeze at -15°C. The results of this assay, illustrated in Figure 26, show that it takes an average of 300 seconds for a 10µL drop of buffer to freeze at -15°C whereas it takes an average of 102 seconds for a solution containing an OD of 0.7 of P. syringae to freeze in these conditions. We also compared this data to the freezing time of a solution containing an OD 0.7 of E. coli, which takes an average of 1500 seconds to freeze. We tested two other concentrations of P. syringae to try and determine when the presence of the bacteria started to have an effect on the time before freezing. From these measurements, we established that even at very low concentrations, the bacteria have a drastic effect on the time it takes for a drop of solution to freeze.
This assay allowed us to confirm that P. syringae drastically reduces the Time before freezing, regardless of the concentrations tested.

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Figure 26 | Pseudomonas accelerates the freezing of liquid solutions.The graph shows a barplot with the mean of biological triplicate +/- standard deviation. Time measured before observed freezing of NYB liquid solution supplemented with five different samples: P. syringae syringae with an OD of 0.7 (P. syringae syringae 1); P. syringae syringae diluted twice with an OD of 0.39 (P. syringae syringae 0.5); P. syringae syringae diluted four time with an OD of 0.155 (P. syringae syringae 0.25); NYB buffer (Buffer); Escherichie coli with and OD of 0.72.


In this part of the project we primarily focused on engineering a plasmid responsible for delivering a CRISPR-Cas9 construct with a sgRNA targeting the genomic site of InaZ, the gene coding for the Ice Nucleation Protein. We tested three different sgRNAs in parallel (each sgRNA on a different plasmid) targeting different positions of InaZ.
In addition, as our purpose was to take advantage of the innate homologous recombination process of Pseudomonas syringae, we also wanted to insert a Donor Sequence into our plasmid, consisting of a 2 kb homology region allowing such mechanism to take place. These two elements would allow us to prove the concept that such an intervention could trigger the deletion of InaZ. In a second step, we would use the same techniques used to engineer the plasmid in order to engineer the final carrier consisting of a phagemid, allowing us to generate an engineered defective phage capable of infecting Pseudomonas syringae, eventually causing the deletion of InaZ.

All the protocols that we used to get our results can be found here.

Golden Gate Assembly (for sgRNA insertion)

We used a Golden Gate Assembly to clone the three different sgRNAs into pJH1 (i.e. our vector).
We first annealed the three pairs of complementary oligos which gave us the three different sgRNAs ready to be cloned by Golden Gate Assembly into pJH1 opened by BsaI-HFv2.

The actual Golden Gate Assembly reaction was hosted by a thermal cycler. We then transformed electrocompetent NEB5alpha with the Golden Gate reaction product, giving us the results shown in figure 13:

Figure 13 | Transformed E. coli NEB5alpha. On the right, the plate containing NEB5alpha bacteria transformed with pJH1 carrying sgRNA 1 is shown, one half (upper one) undergoing initial dilution in HEPES and the other (bottom one) without undergoing initial dilution. In the center (bottom plate) the plate containing bacteria transformed with pJH1 carrying sgRNA 2 is shown, following the same disposition as the plate on the right. The upper plate contains bacteria transformed with pJH1 carrying sgRNA 3, with the same disposition as the other two plates. On the left, the plate containing bacteria that did not undergo transformation is shown, representing a control

From here we decided to adhere to the following workflow:

Figure 14 | Representation of the followed workflow. Starting from plated transformed bacteria to extraction and amplification of the specific fragment (the right branch) as well as the plasmid extraction (the left branch)

First, we performed a colony PCR followed by a gel electrophoresis to check the presence and size of the amplified fragment. One of the two primers used during colony PCR is priming in the sgRNA insertion, meaning that if we observe a band, the presence of the guide RNA is confirmed, while its absence would be the result of a lack of sgRNA insertion. The electrophoresis gel is shown in figure 15:

Figure 15 | Electrophoresis results of the sgRNA insertion. Colony PCR after Golden Gate Assembly. The first lane on the left named “Ladder” corresponds to the molecular-weight size marker. Lanes numbered 1-14 correspond to single colonies tested. Wells 1-10 and 13-14 are from the insertion of the sgRNAs, while wells 11-12 were from the insertion of the donor sequence (not presented here)

All 12 colonies tested show a bright band at the expected molecular size of approximately 900 bp, which means that in those cases the sgRNA insert is probably present. We purified the PCR product and sent it for sequencing.

We also made a miniprep to extract the engineered pJH1. The sequencing analysis results are shown in figure 16:
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Figure 16 | Sequencing results aligned with pJH1 sequence in Snapgene. On the left pJH1 carrying sgRNA 1, sgRNA 2 in the middle and sgRNA 3 on the right

These results confirm that all three sgRNAs had been efficiently integrated into pJH1 (each sgRNA in a different plasmid entity).

Gibson Assembly (for Donor Sequence insertion):

We also used Gibson Assembly to clone into pJH1 (i.e. our vector) the donor sequence, carrier of the homology regions responsible for the homologous recombination event occurring in Pseudomonas syringae. We firstly ordered the donor sequence from IDT (Integrated DNA Technologies company) and dissolved it upon arrival.

Once the donor sequence was prepared for further manipulations, the first procedure that took place in this part of the project was the amplification of such sequence via PCR in order to have a sufficient concentration of it to perform a more efficient cloning procedure.
A verification of the amplified fragment was provided by a gel electrophoresis test, where the results are presented in figure 17:

Figure 17 | Electrophoresis results of the donor sequence amplification. The first lane on the left named “Ladder” corresponds to the molecular-weight size marker. The right lane, named “PCR prod.”, hosts the PCR amplification product. The red circle defines the molecular size of interest (i.e. 2 kb)

The length of the donor sequence is 2 kb, which is the molecular size where the gel in figure 17 shows the brightest and thickest band, confirming the correct amplification of this sequence.
We digested the backbone into which an insert sequence is supposed to be cloned. In our case, the backbone opening is achieved by a double digestion, where we used two different restriction enzymes (NheI-HF and SphI-HF) to avoid relegation of the resulting sticky ends.

To confirm the efficiency of the digestion we loaded the digested samples on a gel.
Figure 18 | Electrophoresis results of the pJH1 digestion. The molecular-weight size marker corresponds to the left well, named “Ladder”. The well named “pJH1” hosts the original plasmid (without undergoing digestion), the same as the one used as backbone during the double digestion. The wells 1-3 correspond to three digested pJH1, all of them carrying a different sgRNA (either 1, 2 or 3)

From figure 18, it is easily noticeable that the results are the expected ones, namely three bands at a molecular size near 8 kb appeared, which correspond to the linearised open plasmid. In addition, we observed three fainter bands at the bottom of the gel, which reflect the 337 bp sequences cut out.
After having efficiently amplified the donor sequence and properly digested the backbone, the Gibson Assembly was performed in order to insert the 2 kb (i.e. the donor sequence) into the 8 kb sequence (i.e. the open pJH1).

We then transformed the Gibson Assembly products into electrocompetent cells (E. coli NEB5alpha).
The plating results are displayed in figure 19:

Figure 19 | Plate results. On the left, one half (upper one) of the plate containing NEB5alpha bacteria transformed with pJH1 carrying sgRNA 1 and the donor sequence, while the bottom half hosts bacteria transformed with pJH1 carrying sgRNA 3 and the donor sequence. On the right, the upper half of the plate contains NEB5alpha transformed with pJH1 carrying sgRNA 2 and the donor sequence, while the bottom half contains the control, NEB5alpha that did not undergo transformation with pJH1

Next, we tested 12 clones by colony PCR and ran a gel to check the presence and characteristics of the amplified fragment, directly suggesting which colony presented the donor sequence and which did not.
The electrophoresis gel is shown in figure 20.
Figure 20 | Electrophoresis results of the donor sequence insertion. Colony PCR to confirm insertion of the donor sequence. The molecular-weight size marker corresponds to the left well, named “Ladder”. Wells 1-12 correspond to each single colony out of all that grew onto the LBA+Kan50 plates. All wells host colonies that underwent Gibson Assembly in order to insert the donor sequence

If the donor sequence is integrated, a band at a molecular size of 2’105 bp is expected, whereas if a band at 442 bp molecular size is visible, it means that the insertion did not occur properly and that the Gibson Assembly did not work for those colonies. As it is noticeable in figure 20, we obtained six colonies (i.e. 2-5, 8 and probably 9) with the correct insert. These PCR products were then sent for sequencing.

Figure 21 | Sequencing results aligned with pJH1 sequence carrying the donor sequence insertion in Snapgene.

As we can see in figure 21, the sequences confirmed that the team successfully inserted the donor sequence into pJH1. However, from the sequencing analysis results, as already suggested from the electrophoresis results, it turned out that one out of the three different types of pJH1 (one per each sgRNA type) (i.g. pJH1 bearing sgRNA 3) did not integrate the donor sequence. Nevertheless, we pursued our assays with pJH1 bearing either sgRNA 1 or sgRNA 2.
Once we obtained our engineered plasmid bearing both sgRNA and the donor sequence, we properly prepared Pseudomonas syringae pv. syringae B301D for transformation via electroporation.
To test the efficacy of our construct in deleting InaZ, we transformed the bacteria with three lines of plasmids, two of which bore sgRNA 1 while the other carried sgRNA 2.

The bacteria were then plated onto LB agar plates supplied with Kanamycin 50. The resulting plates are shown in figure 22:

Figure 22 | Plate results. On the left the plate containing bacteria transformed with pJH1 bearing sgRNA 1.3 (upper half) and sgRNA 1.5 (bottom half), while on the right there is the plate containing bacteria transformed with pJH1 carrying sgRNA 2.1 (bottom half) and a control containing bacteria that underwent electroporation without any plasmid

We did colony PCRs of the grown colonies by using two primers specific to the bacterial genome to confirm the InaZ deletion followed by running a gel. From the size of the amplified fragment, we can directly say whether the deletion was efficiently induced or not.
The electrophoresis results are shown in figure 23:
Figure 23 | Electrophoresis results of the colony PCR to confirm the deletion of InaZ. The molecular-weight size marker corresponds to the left well, named “Lad.”. Wells 1-17 correspond to each single grown colony. Well 18 hosts the template control, where no DNA was added to the PCR reaction mix. Well 19 represents another control where the bacteria did not undergo transformation with our construct

Unfortunately, all grown colonies still carried InaZ in their genome. In fact, the expected PCR product confirming InaZ deletion would have shown a band at around 2 kb, whereas, as shown in figure 23, all our PCR products have a band at about 6 kb, which means that InaZ is not deleted.
As we did not manage to delete InaZ from the bacterial genome, additional attempts would have been made if there was more time. Besides, small changes to the protocol could be tried, for example by either shortening the recovery time after electroporation or changing the temperature at which this process occurs. However, the reason we did not achieve our goal could also come from a problem in our construct. First, we might need to test if Cas9 got expressed. We could perform a Western Blot to test Cas9 expression. Second, we could test the efficiency of the sgRNAs. We could transform another plasmid containing the target region of the sgRNAs and see if Cas9 induces a cut in this plasmid in Pseudomonas syringae. Third, the homologous recombination might have been low efficient in Pseudomonas syringae, especially when the designed homology arms are not adjacent to the cut site. We could design a donor sequence carrying homologies adjacent to the cut site, which will not completely delete InaZ but at least disrupt it. In addition, instead of one cut, we could cut InaZ twice by using two gRNAs targeting the start and the end of the gene. This might facilitate the integration of the donor sequence by the HR repair mechanism, as the sequences next to the cut are directly homologous to the ones provided by the donor sequence.

  1. [1] D. Q. Zhang et al., Expression, purification, and antifreeze activity of carrot antifreeze protein and its mutants, In: Protein Expression and Purification, Vol. 35, No. 2, pp. 257-263, June 2004

    Available from:
  2. [2] K. L. Hockett et al., Use of the Soft-agar Overlay Technique to Screen for Bacterially Produced Inhibitory Compounds, In: Journal of Visualized Experiments, Vol. 119, 2017 Available from:
  3. Tailocins : Figures 2,4,5 created with
  4. Phages : Figure 2 created with

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