<!DOCTYPE html> Template



During spring season, fruit cultivators throughout Switzerland incur enormous losses due to frost each year [1]. A widespread and renowned crop in the Swiss 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 (AFPs) or found a way to inhibit the production of ice nucleation proteins by Pseudomonas syringae pv. syringae, which will reduce the amount of ice crystals that are formed during the late spring freezes.

First, we engineered bacteria that produce three different AFPs, which can bind to ice crystals and inhibit their growth. Second, we engineered bacteria that produce tailocins, which can kill P. syringae syringae, and therefore inhibit the formation of ice nucleation proteins. Third, we designed a phage that can deliver CRISPR/Cas9 and a guide RNA into P. syringae syringae to delete the gene coding for the ice nucleation protein.

To find out more about the results we obtained click here.

Antifreeze Proteins

In order to reduce frost damage in crops due to late spring freezes, we developed a treatment consisting of purified AFPs. AFPs bind to ice crystals and thereby inhibit any further ice growth. Their activity can be characterized using either thermal hysteresis (TH) or ice recrystallization inhibition (IRI). TH describes the difference between the non-equilibrium freezing and melting temperatures [2]. IRI activity prevents the growth of larger ice crystals, which could damage plant tissues, at the expense of smaller ones [3]. Not all AFPs have the same activity. Bacterial AFPs for example have moderate TH and IRI activities, whereas AFPs from fish and insects have high TH, but low IRI activity, and AFPs from plants have high IRI, but low TH activity [4].

For this project, we therefore chose three AFPs, each one with a different activity (Table 1 & Fig.1). This allowed us to make comparisons and find the most efficient AFP for our purpose. We chose to work with AFPs produced by Rhagium inquisitor (RiAFP), by Flavobacterium frigoris PS1 (FfIBP), and by Daucus carota (DcAFP).

Figure 1 | Activities of antifreeze proteins (AFPs). AFPs’ activity can be characterized by thermal hysteresis (TH) and ice recrystallization inhibition (IRI). RiAFP has high TH, but low IRI activity. FfIBP has moderate TH and IRI activity. DcAFP has high IRI, but low TH activity.

Antifreeze Protein Species of Origin Molecular Weight TH Activity
RiAFP Rhagium inquisitor 12.8kDa [5] 6K [5]
FfIBP Flavobacterium frigoris PS1 28.4kDa [6] 2.5K at 50µM [6]
DcAFP Daucus carota 33.9kDa [8] 0.35K at 1.0mg/mL [7]
Table 1. Sources, Molecular Weights and TH activities of RiAFP, FfIBP, and DcAFP.

The main approach we used to produce these proteins was to clone them into E. coli BL21 (DE3) using two different vectors, pET-17b and pCold-I, and then to induce expression with IPTG. Finally we extracted and purified the proteins using magnetic beads or a His-tag affinity column and gel filtration.

Plasmid Cloning

A list of all the primers used to clone the various constructs can be found here.

pET-17b contains a T7 promoter which allows regulated high-level expression of recombinant proteins. On the other hand, pCold-I includes a cold-shock protein A (cspA) promoter which allows expression of proteins that are more difficult and cannot be expressed with the T7 system. Therefore we compared expression of AFPs using each vector with the aim to optimize their expression and synthesis.

Figure 2 | Plasmid maps of RiAFP, DcAFP, and FfIBP inserted into the pET-17b and pCold-I vectors.

The Korea Polar Research Institute kindly sent us the pCold-I-FfIBP construct [5], which we transformed into E. coli BL21 (DE3). To insert FfIBP into the pET-17b plasmid, we amplified the gene by PCR using primers with the correct overlapping ends and linearized pET-17b using XhoI and HindIII restriction enzymes. We cloned the amplified FfIBP gene into the plasmid by Gibson Assembly and transformed the pET-17b-FfIBP construct into E. coli NEB5alpha. We extracted the construct from the cells, and finally transformed it into E. coli BL21 (DE3). RiAFP and DcAFP were synthesized by IDT. We amplified both RiAFP and DcAFP by PCR using primers with the correct overlapping ends. pET-17b was linearized using XhoI and HindIII restriction enzymes, and pCold-I was linearized using NdeI and XhoI restriction enzymes. We cloned both AFPs into the plasmids by Gibson Assembly and transformed the constructs into E. coli NEB5alpha. pET-17b-RiAFP, pCold-I-RiAFP, pET-17b-DcAFP, pCold-I-DcAFP (Fig. 2) were extracted and transformed into E. coli BL21 (DE3).

The various protocols used for the plasmid cloning can be found here.

Expression and Purification

Figure 3 | Representation of the various steps followed to express and purify RiAFP, DcAFP, and FfIBP.

Before inducing expression in large volumes of liquid cultures, we first did small-scale tests: an induction with 1 mM IPTG (we also did an expression optimization for DcAFP using an IPTG gradient concentration [0mM, 0.01mM, 0.1mM, 0.2mM, 0.5mM]) and purification with magnetic beads to verify if the AFPs could be correctly purified. The general steps we followed are described in Figure 3. Next, we grew 1-liter liquid cultures at 37 °C until an OD600 of 0.6 was reached. We then placed the culture in a cooling incubator at 15 °C for 30 minutes. We added 1mM IPTG and induced expression at 15 °C for 24 hours. Afterwards, we removed a small volume of culture to run a total cell SDS-PAGE (15%) of E. coli cultures stained with Coomassie blue and to verify that the AFP was correctly expressed. The remaining culture was centrifuged, and the cell pellet was resuspended in Cold Lysis Buffer. We sonicated the cells and centrifuged the cells once more to collect the supernatant, which we loaded into the His-tag affinity column. We eluted the AFPs bound to the column and concentrated the solution to a smaller volume. We performed a gel filtration to further purify the protein and finally stored aliquots at -80 °C.

A detailed protocol for AFP expression and purification can be found here.


To verify if our purified AFPs were functional and to compare their efficiencies, we performed various assays. We measured the TH activity of the purified recombinant proteins and assessed their effects on frost damage on plants. The details on these measurements can be found here.

Lastly, we wanted to see, if combining three different AFPs would allow us to exploit their distinct activities, which can be explained by differences in ice adsorption behavior [9], and design our final solution to be more efficient in protecting crops against frost damage.


Plant Frost Damage is not Only a Physical Phenomenon

Some bacterial species, which live on plant surfaces, were shown to promote ice crystal formation and thus to increase frost damage on plants [10]. Indeed, water does not only need temperatures below the freezing point to freeze; ice crystal formation also requires some kind of catalyst, in this case an ice nucleus, on which the crystal starts to grow. Mineral and dust particles are active ice nuclei, and therefore promote plant damage, but only at temperatures as low as around –10 °C. Moreover, plant tissues themselves are also relatively bad ice nuclei: when frost-sensitive plants are grown in sterile conditions, most of them can resist temperatures as low as –5 °C to –10 °C without frost damage. However, in the environment, frost-sensitive plants suffer at temperatures only slightly below 0 °C from frost damage which are caused by bacteria colonizing their surface, especially the species P. syringae syringae. P. syringae syringae produces ice nucleation proteins, called InaZ [11], on their membranes which catalyze the formation of ice crystals and thus cause frost damage on the plant at higher temperatures. The damage helps the bacteria to get into the plant as P. syringae syringae are also plant pathogens. For this reason, P. syringae syringae are a substantial problem for apricot producers [12].

This subproject therefore aims at eliminating P. syringae syringae with a phage derived bacteriocin with a narrow host range called tailocins. They are produced by several species of bacteria including P. syringae syringae in order to compete against closely related bacterial strains (often even of the same species) but do not harm cells from their producer’s strain [15]. They structurally look like phage tails and are encoded by phage genes repurposed by bacteria [13]. In the environment, Pseudomonas bacteria produce tailocins under stressed conditions (Fig. 4): some cells of the population will start expressing tailocins and lyse to release them, thus protecting the rest of the colony. When tailocins encounter their target bacteria, they bind to their cell wall and contract, thus perforating the wall. This creates a pore, which dissipates the proton gradient and kill the target cell. This mechanism is extremely efficient, as only one or few particles are sufficient to kill the cell [14].

Figure 4 | Production and killing mechanism of tailocins. Most Pseudomonas strains possess tailocins gene clusters in their genomes (1). Under specific conditions, they express their tailocins (2) and lyse to release them (3). When these tailocins recognize their target strain, they contract and perforate the bacterium’s cell wall, thus killing the competitor.

Tailocins have lots of advantages. Their narrow host range allows to target specifically a pathogenic species without harming potentially beneficial ones [16]. It was also shown that they can be easily modified to change their specificity by swapping the genes responsible for target recognition with the ones of another tailocin or even of a phage. They can thus be adapted to new target strains [17,18]. Contrary to phages, they are not able to replicate autonomously. Their release in the environment, therefore, raises less ethical and safety concerns as the one of phages [16]. It was also shown recently that the application of tailocins on a plant can efficiently prevent infections caused by P. syringae syringae [16].

Tailocin expression can be induced by exposing P. syringae syringae to mitomycin C, an antibiotic which causes a double-stranded break in the DNA [17]. The induction is strong enough so that the whole culture will start producing tailocins and lyse. Tailocins can then be extracted easily from this lysate. Mitomycin C, however, is both very expensive and toxic, which limits the possibility to produce tailocins at a large scale. We therefore aim at cloning the tailocin cluster of a suitable P. syringae syringae strain in E. coli and express it under the control of an inducible promoter in order to avoid the use of mitomycin C. This technique was already used in the past to produce engineered tailocins from Pseudomonas aeruginosa[19] and gave good results.

Choice of strains and identification of tailocins genes

The species P. syringae syringae includes a wide number of strains. The ones found on apricot trees are related to the the pathovar Pseudomonas syringae syringae and, to a lesser extent, Pseudomonas syringae morsprunorum [12]. We were not able to obtain the exact strains found on apricot trees, nor to find articles which had shown to which tailocins they were sensitive. As a proof of concept, we therefore produced tailocins from the strain Pseudomonas syringae pv. aptata DSM50252 which was shown to target Pseudomonas syringae pv. syringae B301D [17]. Both of these strains were easily available as they could be ordered from the DSMZ [20,21]; genome sequences are also available for both strains. Pseudomonas syringae pv. syringae B301D are part of the pathovar syringae like the Pseudomonas found on apricot trees, and were first isolated from a pear tree [20]. Even though it is not exactly the strain found on apricot trees, they are closely related. We, therefore, consider the strain to be a reasonable model.
The tailocin genes were identified using PHASTER [22], a tool aimed at identifying phage sequences in bacterial genomes. Tailocins are identified by the fact that they contain most elements of a prophage, but lack integrase and capsid genes [14]. All genes are present in a single cluster, which contains both structural genes and enzymes needed for the lysis of the producer cell. The results showed the presence of both a prophage and a tailocin in our killer strain’s genome [23]. The tailocin cluster is 13.8 kb long and contains 17 genes (Fig 5). There are two lytic enzymes: the holin and the lysozyme. The holin is the first gene of the cluster, as it is often the case in tailocins. Several genes are identified as structural tail genes by PHASTER; one of them is identified as a tail fiber, i. e. the protein responsible for target recognition and specificity. A chaperone (tail fiber assembly protein) is also present next to the tail fiber gene and helps the tail fiber to fold correctly [17]. Finally, there are some other phage-like genes whose function is not clearly identified by PHASTER (hypothetical proteins).

Figure 5 | Tailocin gene cluster of Pseudomonas syringae DSM50252. Gene annotation as identified by PHASTER

Tailocins extraction and characterisation

As a proof of concept, we want to first show that the tailocins of the producer strain we obtained, Pseudomonas syringae pv. aptata DSM50252, can indeed kill our target strain, Pseudomonas syringae pv. syringae B301D before engineering E. coli to produce tailocins. To do so, we will induce tailocins production in our killer strain using mitomycin C. We will then purify tailocins from the lysate using a precipitation method. In order to show tailocin killing activity, we will apply them on soft agar plates inoculated with our target strain and observe the formation of lysis plaques. As an additional proof, we will use electron microscopy to verify the presence of tailocins in the sample. In order to prove that tailocins would indeed cause the reduction of Pseudomonas syringae’s ability to promote ice crystal formation, we will measure the freezing time of water drops containing Pseudomonas syringae syringae B301D treated or untreated with tailocins, by using our custom-made device FROZONE. As a further characterisation step, we intend to apply tailocins on Arabidopsis thaliana containing Pseudomonas syringae syringae B301D to check that they can protect them from frost damage.

Tailocins production in E. coli

In order to make tailocins production cheaper and avoid the use of the toxic and expensive mitomycin C for safety reasons, we will aim to clone the gene cluster containing all the necessary genes to produce tailocins in E. coli and express them under the control of an inducible promoter.
As a backbone, we will use the plasmid pSG3651pLIBT7A, which was kindly given to us by Dr Michael Taschner, from the Department of Fundamental Microbiology of the University of Lausanne (DMF). This plasmid contains a lac inducible T7 promoter, which allows a strong protein expression and a tight control of it. Indeed, as our gene cluster contains lytic genes, we would need to lower their basal expression level as much as possible to avoid killing our producer cells. For the same reason, we will choose to transform our construct in BL21 pLys, which allows a tighter expression and was given to us by the Collier lab from the DMF.
As the gene cluster is quite big (13.8 kb), we decided to amplify it in four fragments of respectively 3 kb, 3 kb, 5 kb and 2.5 kb (Fig. 6). In order to assemble these fragments together with our backbone, we will use Gibson assembly. We will therefore design our primers in such a way that the fragments have suitable homology regions to assemble them.

Figure 6 | Tailocin genes cluster cloning process.

To express the tailocins, we will induce E. coli transformed with our construct with IPTG, which then leads to tailocins production and the lysis of the producer E. coli. The lysate will then be purified and the tailocins will be characterized as explained above.


The main idea behind the implementation of a phage in our project was to exploit its ability to infect bacteria to provide us with a specific and precise tool to target P. syringae syringae. In fact, bacteriophages are natural predators of bacteria and are highly specific to a limited range of prey, which is the reason why they are usually employed as a biocontrol against microbial pathogens [24]. Introducing a phage into our Aprifreeze product will allow us to specifically target P. syringae syringae, leaving the other bacterial species untouched by our intervention.

The final goal of this part of the project is to delete InaZ, the gene coding for the Ice Nucleation Protein (INP). This protein is harmful to fruit trees [10] as it promotes the formation of ice crystals [25]. This damages the plant cells and allows P. syringae syringae to obtain important nutrients [26].

In this part of the project, we decided to focus our efforts on designing a strategy to achieve the deletion of InaZ without the need of killing P. syringae syringae, in order to disturb the plant microbiota as little as possible. This is why we planned to engineer a viral-derived construct called phagemid. It is a plasmid-like DNA that can be encapsulated in a defective P1 bacteriophage as its unique genetic material [27]. If this defective phage is released onto a Pseudomonas species [28], it will infect its prey by injecting our phagemid that we engineered to trigger the deletion of InaZ without killing the bacterium.

P1 Phage Generation

In order to encapsulate the phagemid into the defective bacteriophage, we need a helper phage, which consists of another plasmid-like entity, carrier of all the genes necessary for the construction of a mature and infection-capable virus, when expressed in an E. coli species. In particular, these genes take care of the redirection of the bacterial molecular machinery to replicate the viral genome and to produce the capsid as well as inducing the viral lytic cycle [28][29].

However, this helper phage lacks the packaging signal called pac, which is a specific sequence that is recognized during the packaging of the viral genome into the capsid of the mature viral particles [30]. The beauty of this approach is that this packaging signal is only found in our engineered phagemid and will thus serve as a recognition site, allowing it to enter the viral capsid of the virions [28]. Therefore, we will produce P1 viruses uniquely carrying our phagemid and not the helper phage, as the latter lacks the packaging signal.

After harvesting the newly produced P1 virions, either by killing the bacterial host or by waiting for lysis plaques [31], these entities are ready to be delivered on P. syringae syringae via our Aprifreeze spray. Once sprayed onto P. syringae syringae, the phage will infect it and inject our engineered phagemid. We referred to it as a defective phage since it will not be able to reproduce at the expense of the bacterium [28]. This way, we prevent the spread of our virus in the environment and reduce its impact on other microbial populations.

Figure 7 | Representation of the mechanism occurring in E. coli once being transformed with the helper phage and the engineered phagemid, allowing the production of defective P1 phages. The mechanism is divided into five parts, defined by a number. Step 1: insertion of the engineered phagemid and the helper phage into E. coli, step 2 consists of the bacterial expression of the genes carried by the helper phage allowing the construction of phage parts and the replication of the phagemid by the host. In step 3 there is the encapsulation of the phagemid into the mature viral particles via the recognition of the pac signal, while step 4 shows the fully formed virions. Step 5 represents the phase where the host is lysed releasing the virions carrying the phagemid as their unique DNA material

InaZ deletion

To achieve our final goal of deleting InaZ from the genome of P. syringae syringae, we needed to engineer the phagemid in a way that it could trigger such a reaction. The strategy we adopted consists of cutting InaZ using CRISPR/Cas9 and repairing the double-strand DNA cut by exploiting the homologous recombination mechanism, where homologous sequences can be detected and exchanged with each other. The required recombineering proteins appear to be expressed in P. syringae syringae species [32], suggesting the presence of this mechanism and the feasibility of our approach. To corroborate this idea, it was also shown that in certain bacterial species, a higher expression of these proteins is triggered by DNA damage, and the homologous recombination can be used to cope with such a genetic accident [33]. We provide the template for homologous recombination on the phagemid and call this sequence the donor sequence. This sequence consists of two regions, one is homologous to the upstream sequence of InaZ, while the other shows homology to the downstream region of the same gene [34].

The CRISPR/Cas9 technology is derived from the adaptive immune system of many prokaryotes [35]. Once produced, our Cas9 will be associated with a single-guide RNA (sgRNA) [36], which is complementary to the DNA sequence we aim to cut [37]. If the sgRNA finds its complementary sequence, Cas9 will induce a double-strand break in proximity of it, opening the DNA double helix [38]. Thus, our phagemid also harbors Cas9 and a sgRNA targeting a particular sequence of InaZ.

Figure 8 | Schematic representation of the mechanism allowing to delete InaZ from P. syringae syringae genome. The properly engineered phagemid is first delivered to P. syringae pv. syringae B301D (the strain we used for the testing part). Once inserted into the bacteria, the CRISPR/Cas9 construct is expressed and the single-guide RNA will direct Cas9 towards InaZ, causing a double-strand break. The recombineering proteins are expressed and recognize the homologous sequences carried by the donor sequence, allowing the replacement of InaZ by the donor sequence, resulting in the deletion of InaZ from the bacterial genome

Phagemid engineering

To prove that this strategy is feasible, we decided to divide this part of the project into two distinct steps. Rather than starting with the direct manipulation of the phagemid, we chose to prove the efficacy of the strategy by engineering a simple plasmid that would correspond to our phagemid and show that we could efficiently delete InaZ by transforming Pseudomonas syringae pv. syringae B301D. The engineering process was carried out in the same way as we would have done with the phagemid. We started with an existing plasmid for carrying out CRISPR/Cas9 engineering in P. syringae syringae [35]. We only needed to change the sgRNA and insert the donor sequence, as the CRISPR/Cas9 cassette was already present in our customised plasmid. We first used Golden Gate Assembly to insert the sgRNA, and then used Gibson Assembly cloning to insert the donor sequence into our plasmid. Finally, we would transform P. syringae syringae with our engineered plasmid and analyse its chromosome in order to prove that we deleted InaZ.


To perform some of our experiments, a precise control of the temperature was needed. We therefore created FROZONE, a precise cooling device that fits under a microscope, acting similarly to a Nanoliter Osmometer. The design of our machine was inspired by the "MicroIce LTD". According to MicroIce, "A Nanoliter osmometer is a cooling stage mounted on an upright optical microscope. Cooling of the stage is achieved with the use of Peltier devices driven with a precision temperature controller."
Our nanoliter osmometer, FROZONE, was designed and created by us. The cooling of the device is achieved with a thermoelectric cooler controlled by a custom Python software. We also designed a vacuum chamber to minimize the crystallization of ambient humidity on our samples. Put simply, we created a machine that precisely controls the temperature of a copper plate in order to measure various aspects of the solutions we produced.
More information, please visit our Hardware and Software pages.


  1. [1] O. Lhotka et al., Possible increase of vegetation exposure to spring frost under climate change in Switzerland, In: Atmosphere (Basel), Vol. 11, No. 4, 1 April 2020
  2. [2] A. K. Gruneberg et al., Ice recrystallization inhibition activity varies with ice-binding protein type and does not correlate with thermal hysteresis, In: Cryobiology, Vol. 99, pp. 28-39, April 2021 Available from:
  3. [3] M. Chow‐Shi‐Yée et al., Inhibition of ice recrystallization and cryoprotective activity of wheat proteins in liver and pancreatic cells, In: Protein Sci., Vol. 25, No. 5, pp. 974-986, May 2016 Available from: /pmc/articles/PMC4838640/
  4. [4] A. Białkowska et al., Ice binding proteins: Diverse biological roles and applications in different types of industry, In: Biomolecules, Vol. 10, No. 2, February 2020 Available from: /pmc/articles/PMC7072191/
  5. [5] A. Hakim et al., Expression, purification, crystallization and preliminary crystallographic studies of Rhagium inquisitor antifreeze protein, In: Acta Crystallogr Sect F Struct Biol Cryst Commun, Vol. 68, Pt 5, pp. 547-550, May 2012 Available from: /pmc/articles/PMC3374510/
  6. [6] H. Do et al., Crystallization and preliminary X-ray crystallographic analysis of an ice-binding protein (FfIBP) from Flavobacterium frigoris PS1, In: Acta Crystallogr Sect F Struct Biol Cryst Commun, Vol. 68, Pt 7, pp. 806-809, July 2012 Available from: /pmc/articles/PMC3388927/
  7. [7] D. Worrall et al., A carrot leucine-rich-repeat protein that inhibits ice recrystallization, In: Science, Vol. 282, No. 5386, pp. 115-117, October 1998 Available from:
  8. [8] 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. 3, pp. 257-263, June 2004
  9. [9] L. L. C. Olijve et al., Blocking rapid ice crystal growth through nonbasal plane adsorption of antifreeze proteins, In: Proceedings of the National Academy of Sciences, Vol. 114, No. 14, pp. 3740-3745, April 2016 Available from:
  10. [10] S. E. Lindow, The role of bacterial ice nucleation in frost injury to plants, In: Annual Review Phytopathol, Vol. 21, pp. 363-384, 1983 Available from:
  11. [11] InaZ - Ice nucleation protein - Pseudomonas syringae pv. syringae - InaZ gene & protein. [cited 2021 Aug 19]. Available from:
  12. [12] D. Giovanardi et al., Characterisation of Pseudomonas syringae isolates from apricot orchards in north-eastern Italy, In: European Journal of Plant Pathology, Vol. 151, pp. 901-917, August 2018 Available from:
  13. [13] M. G. K. Ghequire et al.,The Tailocin Tale: Peeling off Phage Tails In: Trends in Microbiology, Vol. 23, No. 10, October 2015 Available from:
  14. [14] S. Carim et al., Systematic discovery of pseudomonad genetic factors involved in sensitivity to tailocins, In: The ISME Journal, Vol. 15, pp. 2289-2305, 2021 Available from:
  15. [15] S. Patz et al., Phage tail-like particles are versatile bacterial nanomachines – A mini-review, In: Journal of Advanced Research, Vol. 19, pp. 75-84, 2019
  16. [16] A. David et al., Prophylactic Application of Tailocins Prevents Infection by Pseudomonas syringae, In: Phytopathology, July 2021 Available from:
  17. [17] D. A. Baltrus et al., Localized recombination drives diversification of killing spectra for phage-derived syringacins, In: The ISME Journal, Vol. 13, pp. 237-249, February 2019 Available from:
  18. [18] D. Scholl et al., Genome Sequence of E. coli O104:H4 Leads to Rapid Development of a Targeted Antimicrobial Agent against This Emerging Pathogen, In: Plos One, March 2012 Available from:
  19. [19] J. M. Ritchie et al., An Escherichia coli O157-specific engineered pyocin prevents and ameliorates infection by E. coli O157:H7 in an animal model of diarrheal disease, In: Antimicrobial Agents and Chemotherapy, Vol. 55, No. 12, pp. 5469–5474, December 2011 Available from:
  20. [20] German Collection of Microorganisms and Cell Cultures GmbH: Details, Available from:
  21. [21] German Collection of Microorganisms and Cell Cultures GmbH: Details Available from:
  22. [22] PHASTER, Available from:
  23. [23] PHASTER, Available from:
  24. [24] M. Rabiey et al, Phage biocontrol to combat Pseudomonas syringae pathogens causing disease in cherry, In: Microbial Biotechnology, Vol. 13, No. 5, pp. 1428-1445, 08 May 2020 Available from:
  25. [25] D. Gurian-Sherman et al., Bacterial ice nucleation: significance and molecular basis, In: The Faseb Journal, Vol. 13, No. 14, pp. 1338-1343, November 1993
  26. Available from:
  27. [26] S. J. Roeters et al., Ice-nucleating proteins are activated by low temperatures to control the structure of interfacial water, In: Nature Communications, Vol. 12, No. 1, pp. 1183-1192, 19 February 2021
  28. [27] J. T. Kittleson et al., Scalable Plasmid Transfer using Engineered P1-based Phagemids, In: ACS Synthetic Biology, Vol. 1, No. 12, pp. 583-589, 21 December 2012 Available from:
  29. [28] C. Westwater et al., Development of a P1 phagemid system for the delivery of DNA into Gram-negative bacteria, In: Microbiology Society, Vol. 148, Pt. 4, pp. 943-950, April 2002 Available from:
  30. [29] H. Qi et al., Phagemid Vectors for Phage Display: Properties, Characteristics and Construction, In: Journal of Molecular Biology, Vol. 417, No. 3, pp. 129-143, 30 March 2012 Available from:
  31. [30] M. Russel, Filamentous phage assembly, In: Molecular Microbiology, Vol. 5, No. 7, pp. 1607-1613, July 1991 Available from:
  32. [31] W.M.A. Mullan, Isolation and purification of bacteriophages, In: Dairy Science food technology, 2001 Available from:
  33. [32] B. Swingle et al., Recombineering Using RecTE from Pseudomonas syringae, In: Applied and Environmental Microbiology, Vol. 76, No. 15, 11 July 2010 Available from:
  34. [33] P. J. G. Rauch et al., The expression of the Acinetobacter calcoaceticus recA gene increases in response to DNA damage independently of RecA and of development of competence for natural transformation, In: Microbiology, Vol. 142, Pt. 4, pp. 1025-1032, 1996 Available from:
  35. [34] K. Sakaguchi et al., A targeted gene knockout method using a newly constructed temperature-sensitive plasmid mediated homologous recombination in Bifidobacterium longum, In: Applied Microbiology and Biotechnology, Vol. 95, pp. 499-509, 2012 Available from:
  36. [35] P. Mali et al., Cas9 as a versatile tool for engineering biology, In: Nature Methods, Vol. 10, pp. 957-963, 27 September 2013 Available from:
  37. [36] J. Ho et al., The application of the CRISPR-Cas9 system in Pseudomonas syringae pv. actinidiae, In: Journal of Medical Microbiology, Vol. 69, No. 3, pp. 478-486, March 2020 Available from:
  38. [37] J. Doudna et al., The new frontier of genome engineering with CRISPR-Cas9, In: Science, Vol. 346, No. 6213, 28 November 2014 Available from:
  39. [38] J. M. Peters et al., Bacterial CRISPR: accomplishments and prospects, In: Current Opinion in Microbiology, Vol. 27, pp. 121-126, October 2015 Available from:
  40. AFP Figure 3 created with
  41. Tailocins Figures 4,5,6 created with
  42. Phages Figures 1,2 created with

Follow Us

Instagram logo Twitter logo linkedin logo


logo mail