Team:TUDelft/Description

PHOCUS

Project Description

A problem of biblical proportions

Locust plagues have afflicted human civilisations throughout history. Drawings in ancient Egyptian pyramids already display the devastation caused by the locusts' appetite and the ensuing famine [1]. Unfortunately, this problem of biblical proportion has not been left in the past. Recently, humongous swarms of the desert locust, Schistocerca gregaria, have threatened the livelihood of communities, endangering food security and economic stability [2]. The Food and Agriculture Organization (FAO) of the United Nations predicts that a staggering 10% of the world population will be affected by the current plague [3]. Although there are 19 species classified as locusts [4], our target is the desert locust because this is the species that inflicts the most damage [5].

Normally, desert locusts live solitary lives, but under the right environmental conditions they come together. When a critical locust density is reached, the phenotype of these insects changes from a solitary to a gregarious phase [4]. This phenomenon is referred to as phase polyphenism (Figure 1), and is key to the swarming behaviour observed during plagues. The switch to the gregarious phase is triggered when the locusts touch each other’s hind legs, or when they see or smell each other. They change size and colour, have a less restricted diet and become voracious [6], eating their bodyweight (2 grams) every day. Although this may seem insignificant, a swarm of one square kilometre consumes the same amount of food as 35,000 people a day [3]. Recently, a swarm in Kenya has been spotted with a size of 2,400 square kilometres, containing between 4 and 8 billion locusts. This swarm consumes the daily equivalent of 3.5 million people [2].

Phage polyphenism
Figure 1. Phase polyphenism of the desert locust (Schistocerca gregaria). Alterations of the desert locusts from a solitary phage to a gregarious phase.

The FAO and national governments are responsible for locust control. These organisations have two control methods available: chemicals and biopesticides. The chemical pesticides, although effective, are not specific to the desert locust. Therefore, they are also harmful to other animals, humans and the environment [7]. The alternative, a bio-pesticide called Metarhizium anisopliae, is a fungus that is specific to locusts. However, it works rather slow, with a killing time of 7 to 14 days [2]. Because of this lack of speed, it cannot be used in emergency responses during sudden outbreaks.
In essence, the people fighting the desert locusts are in need of a safe and fast solution to end these troubles once and for all. We as a team were shocked about these events and discovered an underlying passion in all of us to helping fight the desert locusts.

“Our mission is to tackle the locust crisis by developing a fast and safe biopesticide through responsible innovation and collaboration.”

PHOCUS; Target locusts from within

PHOCUS is a targeted bacteriophage-based biopesticide used to control the desert locust crisis. Our aim is to kill the locusts by producing toxic molecules in their gut. These molecules will be produced by the gut bacteria after infection with our engineered bacteriophage. The toxins we want to produce are the locust specific Cry7Ca1 toxin and short-hairpin RNAs (shRNAs) that target essential locust genes. This process is illustrated in Figure 2.

Project Overview
Figure 2. Project graphical overview. 1. Bacteriophages are engineered to contain DNA coding for our toxic molecules. 2. Engineered bacteriophages will be sprayed on vegetation in affected areas. 3. Locusts will eat the crops and ingest the bacteriophages. 4. Bacteriophages infect host bacteria and produce Cry7Ca1 and shRNA. Upon lysis, the cell bursts and the toxins and shRNAs are released. 5. Toxin and shRNA release causes locusts to die, swarms disappear.

Delivery: Engineered bacteriophages

Our delivery method consists of bacteriophages, as they are viruses that specifically infect bacteria [8]. PHOCUS targets the bacteria in the locust gut. Although the microbiota of different sections of the locust gut vary among individuals and species, the Enterobacter genus has been reported to be the most abundant [9]. To account for any variety in the core Enterobacter population in the locust gut, our biopesticide consists of a cocktail of phages targeting a variety of bacteria from the Enterobacter genus. To reach the locust gut target bacteria, our engineered bacteriophages will be sprayed on vegetation that will be eaten by gregarious locusts from the swarms. The relatively mild pH (7-8) [10] of their gut allows the bacteriophages to survive there [11].

We have used Escherichia coli (E. coli) BL21 (DE3), a bacteria from the genus Enterobacter, as our model organism, together with the bacteriophage T7. We have developed a proof of concept to choose the position where we would insert our desired gene in the engineered bacteriophage by using GFP as a reporter. We successfully engineered T7 phages to contain GFP using the Bacteriophage Recombineering of Electroporated DNA (BRED) method [12] (Figure 3).

Proposed implementation
Figure 3. The BRED method was used to engineer T7 bacteriophages to replace a non-essential gene with GFP. Linear double-stranded DNA (dsDNA) and T7 phages were co-electroporated in BL21 (DE3) cells. Recombination of the dsDNA and the non-essential gene resulted in engineered T7 phages containing GFP.

Mode of action: killing the gregarious locusts

Our goal is to kill the gregarious locusts present in the swarms by using a specific and unique complementary approach that combines the insecticidal activity of the crystal toxin Cry7Ca1 and the gene silencing effect of RNA interference (RNAi). The bacteriophage will carry the DNA coding for the Cry7Ca1 toxin from Bacillus thuringiensis (Bt), as it has been shown to be effective against locusts [13, 14], as well as shRNA that specifically targets essential locust genes. We successfully expressed the Cry7Ca1 toxin under a T7 promoter in E. coli BL21 (DE3) and purified it (see Results). Moreover, we designed and transcribed shRNA in vitro with T7 polymerase. As the shRNA needs to be transported from the locust gut to the hemolymph to be effective (see RNA interference (RNAi)), we designed a cargo consisting of different proteins. We validated the binding of a key protein of this proposed delivery cargo to the designed shRNA. In order to be converted into short silencing RNA (siRNA), the shRNA has to be cleaved by Dicer, an essential enzyme of the RNAi maturation pathway (see RNA interference (RNAi)). We have demonstrated that the designed and in vitro transcribed shRNA can be cleaved by Dicer (Results).

Cry7Ca1

Bacillus thuringiensis (Bt) is a ubiquitous spore-forming and gram-positive bacterium that forms parasporal crystalline protein inclusions at the onset of sporulation and during the stationary phase. These crystals contain δ-endotoxins that can be either insect-specific insecticidal crystal (Cry) proteins or cytolytic (Cyt) proteins. They have attracted worldwide interest as toxins for pest management because of their specificity [15, 16]. The toxin we intend to produce inside the locust gut is the Bt insecticidal crystal protein Cry7Ca1, which has been shown to be active against the locust species Locusta migratoria manilensis. The detailed mode of action of Cry proteins is still unclear, but it is believed that they act by forming pores in the insect gut epithelium cells (Figure 4).


Cry protein mechanism
Figure 4. Cry protein mechanism. Cry proteins are naturally produced by B. thuringiensis as a relatively inert protoxin ingested by insect larvae. The protoxin is then solubilised in the insect gut and processed by gut proteases, leading to their activation. The activated Cry toxin subsequently binds to specific receptors located on the brush border membrane vesicles of the insect gut epithelium. Oligomerization of its subunits leads to the formation of a pore structure in the cell membrane. This leads to cell lysis, gut damage and eventually larvae death [13, 17].
RNA interference (RNAi)

RNA interference (RNAi), also known as post-transcriptional gene silencing, is a biological response against double-stranded RNA (dsRNA). This response can be against endogenous parasitic and exogenous pathogenic nucleic acids, or act as a regulation method for the expression of protein-coding genes [18]. The presence of dsRNA forms in the locust hemolymph has been shown to lead to robust RNAi-mediated gene silencing in most locust tissues, reaching maximal effect at very low doses (30 ng) [19, 20]. We use the dsRNA approach to silence desert locusts genes specifically. For our project we produce shRNA as their processing by RNAi machinery leads to predictable products, minimising possible off targets. In addition, shRNAs are simple to transcribe and self-assemble, and they are specific and stable.

Proposed implementation
Figure 5. Schematic of the RNAi approach. When shRNA enters the cell it breaks down mRNA, leading to gene silencing. Schematic overview of the gene silencing process by siRNA. dsRNA or shRNA (shRNA) is processed by Dicer into short interference RNA (siRNA). RNA-induced silencing complex (RISC) is formed, hybridising with the target messenger RNA (mRNA), producing its cleavage [21].

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Our approach consists of producing multiple shRNA, dsRNA molecules that contain a hairpin turn and a dinucleotide 3' overhang, by transcribing them in a single tandem transcript (tshRNA). We aim to produce shRNAs that specifically target mRNA unique to desert locusts. Example targets include housekeeping and other essential genes, whose silencing will directly kill the locusts, or fertility-related genes to prevent breeding. The efficacy of gene silencing has been studied already in locusts [19] and other insects [23]. For this purpose, we intend to find any off-targets in other organisms by performing Basic Local Alignment Search Tool (BLAST) searches against databases such as the 1000 Insect Transcriptome Evolution (1KITE) dataset or off-target finder [24, 25].

A challenge when producing dsRNA is that it is produced inside the locust gut, but it has to reach the locust hemolymph to be active [19]. To do so, the shRNA has to cross the midgut epithelium and avoid being degraded by RNAses in the locust gut. To ensure that the produced shRNAs reach the hemolymph successfully, several protecting proteins can be co-expressed with the shRNA. An overview of this theory can be found in Design.

Complementary approach

Upon lysis, the Cry7Ca1 toxin punctures the gut lining, which is lethal to the locust [13,14]. This further facilitates the in situ produced shRNA to reach the hemolymph, where shRNA has been shown to work best [11]. When shRNA is produced and further processed into a mature RNAi, essential genes of the locust will be silenced, causing the locust to die [19, 20].


Mathematical models of phage infection and toxin production

Once a bacteriophage finds its bacterium, it injects its DNA into it. Upon infection, the bacteriophage hijacks the cellular machinery of the bacterium and turns it into a factory expressing and amplifying the injected DNA. The Cry7Ca1 and shRNA will be produced inside these bacteria together with new engineered bacteriophages. We decided to use lytic bacteriophages in our project to ensure enough production of our toxin. Lytic bacteriophages rapidly take over the host, make the host produce phages and toxins and cause release of toxins after lysis occurs [8]. When lysis happens, the bacteria burst and the produced compounds and bacteriophages are released inside the locust gut. Those bacteriophages then begin a new infective cycle (Figure 6).

Proposed implementation
Figure 6. Schematic of the bacteriophage-host interaction. Lytic bacteriophage infects its bacterium, the cellular machinery is taken over and new phages and toxins are produced. Consecutively the cell lyses, the toxins and phages are released. The newly made phages infect bacteria; the cycle begins anew.

To gain a better insight into this infection process, as well as feasibility of our biopesticide, we modelled the evolution over time of the number of bacteria, phages, and produced toxin. Furthermore, since speed is one of the design requirements of PHOCUS, we used modelling to gain insight in the speed at which the toxins are produced inside the locust gut. To determine the speed of our biopesticide we developed a model of phage infection and toxin production based on the work of Beretta and Kuang [26]. With this model we demonstrated the capability of our biopesticide to produce lethal quantities of toxins to kill the locust within 7 days. Furthermore, we demonstrated which design strategies are most effective in improving the effectiveness of our biopesticide. At last, we concluded that the rate at which bacteria acquire resistance does not directly pose a threat to our biopesticide.


However, in the desert locust gut, bacteria most likely reside in micro-colonies [Dr. O. Lavy, personal interview, 27, 28]. To understand how the spatial distribution of the bacteria in the gut influences phage propagation and toxin production, we modelled the bacterial population as a 2D biofilm with an agent-based description of phages based on the work of Simmons et al. [29]. This model allowed us to study the influence of the spatial organisation of bacteria on the effectiveness of our biopesticide. With this model we demonstrated the importance of phage mobility inside the biofilm for effective toxin production. Additionally, we showed how the spatial organization of bacteria in such biofilms promotes toxin production. At last, we demonstrated how biofilm heterogeneity (fraction of target and non-target bacteria) must be taken into account in the design of our biopesticide.


Summary

In short, we designed PHOCUS, a specific and safe biopesticide targeting the desert locust. The biopesticide consists of a cocktail of engineered bacteriophages targeting the Enterobacter genus in the gut of the locusts. The engineered bacteriophages contain the DNA coding for the locust specific Cry7Ca1 toxin and shRNAs. The feasibility of our biopesticide against locusts has been analysed through mathematical models. We showed that the toxic molecules are produced fast. Therefore, we conclude that our biopesticide has the potential to kill the locusts fast.

References
  1. Nevo, D. (1996). The desert locust,Schistocerca gregaria, and its control in the land of israel and the near east in antiquity, with some reflections on its appearance in Israel in modern times. Phytoparasitica, 24(1), 7–32.
  2. Roussi, A. (2020). Why gigantic locust swarms are challenging governments and researchers. Nature 2020 579:7799.
  3. FAQs | FAO | Food and Agriculture Organization of the United Nations. (n.d.). http://www.fao.org/locusts/faqs/en/
  4. Cullen, D. A., Cease, A. J., Latchininsky, A. V., Ayali, A., Berry, K., Buhl, J., De Keyser, R., Foquet, B., Hadrich, J. C., Matheson, T., Ott, S. R., Poot-Pech, M. A., Robinson, B. E., Smith, J. M., Song, H., Sword, G. A., Vanden Broeck, J., Verdonck, R., Verlinden, H., & Rogers, S. M. (2017). From Molecules to Management: Mechanisms and Consequences of Locust Phase Polyphenism. In Advances in Insect Physiology (pp. 167–285). Elsevier.
  5. The desert locust Plague: The World Bank Group Response. (n.d.). https://www.worldbank.org/en/topic/the-world-bank-group-and-the-desert-locust-outbreak
  6. Latchininsky, A. V. (2017). Locusts. In Encyclopedia of Animal Behavior (pp. 115–127). Elsevier.
  7. Zhang, L., Lecoq, M., Latchininsky, A., & Hunter, D. (2019). Locust and Grasshopper Management. Annual Review of Entomology, 64(1), 15–34.
  8. Clokie, M. R. J., Millard, A. D., Letarov, A. V, & Heaphy, S. (2011). Phages in nature. Bacteriophage, 1(1), 31–45.
  9. Lavy, O., Gophna, U., Gefen, E., & Ayali, A. (2019). The Effect of Density-Dependent Phase on the Locust Gut Bacterial Composition. Frontiers in Microbiology, 9(JAN), 3020.
  10. Kerby, G. P., Gowdy, R. A., Dillon, E. S., Dillon, M. L., & Sharp, T. Z. (n.d.). PURIFICATION, PH STABILITY AND SEDIMENTATION PROPERTIES OF THE T7 BACTERIOPHAGE OF ESCHERICHIA COLI I Downloaded from.
  11. Marinelli, L. J., Piuri, M., Swigoňová, Z., Balachandran, A., Oldfield, L. M., van Kessel, J. C., & Hatfull, G. F. (2008). BRED: A Simple and Powerful Tool for Constructing Mutant and Recombinant Bacteriophage Genomes. PLoS ONE, 3(12), e3957.
  12. Jing, X. et al. Crystal structure of Bacillus thuringiensis Cry7Ca1 toxin active against Locusta migratoria manilensis. Protein Sci. 28, 609–619 (2019).
  13. Wu, Y., Lei, C. F., Yi, D., Liu, P. M. & Gao, M. Y. Novel Bacillus thuringiensis δ-endotoxin active against Locusta migratoria manilensis. Appl. Environ. Microbiol. 77, 3227–3233 (2011).
  14. Palma, L. et al. Bacillus thuringiensis toxins: An overview of their biocidal activity. Toxins (Basel). 6, 3296–3325 (2014).
  15. Schnepf, E. et al. Bacillus thuringiensis and Its Pesticidal Crystal Proteins. Microbiol. Mol. Biol. Rev. 62, 775–806 (1998).
  16. Pigott, C. R. & Ellar, D. J. Role of Receptors in Bacillus thuringiensis Crystal Toxin Activity. Microbiol. Mol. Biol. Rev. 71, 255–281 (2007).
  17. Hannon, G. J. (2002). RNA interference. In Nature (Vol. 418, Issue 6894, pp. 244–251). Nature Publishing Group.
  18. Wynant, N., Verlinden, H., Breugelmans, B., Simonet, G., & Broeck, J. Vanden. (2012). Tissue-dependence and sensitivity of the systemic RNA interference response in the desert locust, Schistocerca gregaria. Insect Biochemistry and Molecular Biology, 42(12), 911–917.
  19. Santos, D., Broeck, J. Vanden, & Wynant, N. (2014). Systemic RNA interference in locusts: Reverse genetics and possibilities for locust pest control. In Current Opinion in Insect Science (Vol. 6, pp. 9–14). Elsevier Inc.
  20. RNAfold web server. (n.d.). http://rna.tbi.univie.ac.at//cgi-bin/RNAWebSuite/RNAfold.cgi?PAGE=3&ID=MLjhUGXlaY
  21. Robinson, R. (2004). RNAi Therapeutics: How Likely, How Soon? PLoS Biology, 2(1), e28.
  22. Lenaerts, C., Palmans, J., Marchal, E., Verdonck, R., & Vanden Broeck, J. (2017). Role of the venus kinase receptor in the female reproductive physiology of the desert locust, Schistocerca gregaria. Scientific Reports, 7(1), 1–12.
  23. Grobe, P. (2017). 1KITE - 1K Insect Transcriptome Evolution.https://www.1kite.org/
  24. Good, R. T., Varghese, T., Golz, J. F., Russell, D. A., Papanicolaou, A., Edwards, O., & Robin, C. (2016). OfftargetFinder: A web tool for species-specific RNAi design. Bioinformatics, 32(8), 1232–1234
  25. Beretta, E., & Kuang, Y. (1998). Modeling and analysis of a marine bacteriophage infection. Mathematical Biosciences, 149(1), 57–76.
  26. Dillon, R., & Charnley, K. (2002). Mutualism between the desert locust Schistocerca gregaria and its gut microbiota. In Research in Microbiology (Vol. 153, Issue 8, pp. 503–509). Elsevier Masson. https://doi.org/10.1016/S0923-2508(02)01361-X
  27. Hunt, J., & Charnley, A. K. (1981). Abundance and distribution of the gut flora of the desert locust, Schistocerca gregaria. Journal of Invertebrate Pathology, 38(3), 378–385.
  28. Simmons, M., Drescher, K., Nadell, C. D., & Bucci, V. (2018). Phage mobility is a core determinant of phage-bacteria coexistence in biofilms. ISME Journal, 12(2), 532–543.