In Vitro Assay

A High Throughput Method to Test Cry Toxin Activity

Within toxins families, Cry toxins are found previously to have an activity towards insects of different orders. In short, a functional Cry protein is only active when specific binding to the gut membrane of the target insect occurs. In more detail, these receptors have been found in the gut epithelium cells, where this proteins polymerize binding to its membrane receptor forming pores (holes). After the formation of this ion channels, the osmotic pressure is disturbed resulting in cell death of the target host¹².
Still a lot is unknown about currently characterized Cry toxins, where one might be effective against the Varroa destructor. In this section, an in vitro toxicity test is developed in order to be able to efficiently screen Cry toxins on their activity against Varroa destructor. From the Varroa destructor brush border membrane vesicles were made and loaded with 6-carboxyfluorescein to test the pore formation ability of Cry proteins, see figure 1a. As a proof of principle, brush border membrane vesicles (BBMVs) from the midgut of Tenebrio molitor were made and loaded with 6-carboxyfluorescein to test the pore formation ability of Cry3Aa. Cry3Aa is toxic to Tenebrio molitor larvae³⁴.

Brush border membrane vesicles are composed of phospholipid double layers originating from a midgut membrane. (link to protocol) This membrane therefore should also contain the membrane proteins of the midgut membrane. After incorporation of 6-carboxyfluorescein into the brush border membrane vesicles (link to protocol), the presence of an active Cry toxin can hypothetically be detected as followed: the Cry toxins will bind to their highly specific receptors⁵ and consequently create holes into the phospholipid double membrane layer, see figure 1b. After this event, the fluorophores leak out of the vesicles due to diffusion, which can be measured as an increase in fluorescence. This method has been proved to work for the activity of Cry3Aa on Leptinotarsa decemlineata⁶. Rausell et al. only demonstrated how the magnitude of the fluorescence increase links to toxicity, but this method has its limitations. Brush border membrane vesicles have the disadvantage of being unstable and undergoing degradation⁷. In other words, measurements on BBMVs can be influenced by spontaneous breaking of the vesicles. In our project, we took this into account by relating toxicity activity to a kinetic value.

The increase of fluorescence due to fluorophore leaking can be dedicated to dequenching. 6-Carboxyfluorescein is a self-quenching fluorophore. At a high concentration 6-carboxyfluorescein forms dimers. Because of dimer-dimer and dimer-monomer interactions, the fluorescence of a carboxyfluorescein molecule decreases drastically⁸. Within a vesicle used in this research, the concentration of the fluorophore will be high, which results in self-quenching behaviour. When the fluorophores leak out of the vesicles, the local concentration of fluorophores decreases. Hence, dequenching occurs, the fluorescence increases, see figure 1c.

Analysis of Brush Border Membrane Vesicles

To visualize the presence of brush border membrane vesicles, images were made with the use of TEM. This was done in collaboration with the iGEM team from Delft (link to protocol).

In here you can see vesicles - round shapes with sizes between the 100 nm and 300 nm. To gain more information about the composition of the samples that are visualized above, dynamic light scattering was performed (link to protocol). The dynamic light scattering results give information about the size distribution of the particles that are present in the solution.

Looking at both transmission electron microscopy results and dynamic light scattering results, the T.molitor samples have a large population of vesicles that vary their size around 200 nm. The population around this size is smaller, which might be due to the fact that the whole organism is used instead of only the gut for preparation of the latter sample.
To visualize the proteins in the brush border membrane vesicles, SDS-PAGE was performed. The results, shown in figure 4, show the presence of a variety of proteins. From the SDS-PAGE can not concluded which proteins are membrane proteins.

Fluorophore Leaking Experiments

The hypothesis is that due to addition of a detergent, the phospholipid bilayer will rearrange. This process will subsequently give the fluorophores the chance to escape their enclosed space and disperse in the solution. The dispersion of the fluorophores into the solution would then cause an increase in fluorescence as described previously.

To determine whether these brush border membrane vesicles were capable of releasing fluorophores and whether this release could be measured, the following was done: to brush border membrane vesicles, already incorporated with carboxyfluorescein, 1 volume percent of sodium dodecyl sulfate was added to disturb the their stability. Meanwhile the fluorescence was measured over time. (link to protocol)

Looking at figure 5, after addition of SDS (detergent), an increase in fluorescence is indeed visible. However, the same increase in fluorescence can be observed over time with vesicles that are in an environment without sodium dodecyl sulfate. Two explanations for this phenonem are that the brush border membrane vesicles are already porous or are not stable under the circumstances during measurement. Both processes would result in an increase in fluorescence over time. In order to compare induced fluorophore leaking from standard fluorophore leaking, the speed of the carboxyfluorescein leaking should be calculated and compared.

To determine whether a specific Cry protein can induce fluorophore release. Cry3Aa was isolated from Bacillus thuringiensis (link to protocol) and tested on brush border membrane vesicles obtained from Tenebrio molitor. Activated Cry3Aa gives in the method of Rausell et all.⁶ a clear band on a gel of a 68 kDa protein after SDS-PAGE was performed. The SDS-PAGE gel of the protein extract from Bacillus thuringiensis, shown in figure 7, shows a similar band around this size.

Tenebrio molitor BBMV were incubated with and without Cry3Aa and the fluorescence was measured overtime (link to protocol), see figure 8. From this graph it can be concluded that the presence of Cry3Aa results in a faster increase in fluorescent and therefore in a faster fluorophore release. To determine how large the role of induced leaking is, the kinetic properties of both measurements have to be compared. In order to compare these properties, it should be known which kinetic rules the fluorophore leaking follows. Plotting the natural logarithm of the concentration of fluorophores entrapped in vesicles against time, gives a straight line, see figure 9. The concentration fluorophores entrapped in vesicles is expressed as the difference of the maximum fluorescence and the fluorescence at a certain time point. The observed straight line in this graph tells us that first order reaction rules can be applied on the measurement ⁹. This means that the following formula can be applied to the obtained data:The reaction rate constant, kf, determines how fast the reaction goes. This constant can be used to compare one reaction with another. To find this constant the following formula was plotted through directly obtained data:
. The plotted formulas are visible in figure 8 and 9 as well.

Both samples were measured 6 times, however, to keep figure 8 and 9 clear, only one measurement per sample is shown. All data can be found here (link to raw data). For all measurements the reaction rate constants were determined as explained above and are shown in figure 9. The figure shows that in the presence of Cry3Aa, the kf values are higher and thus the fluorophores ooze faster. Overall, fluorophore leaking is accelerated with a factor of 1.59. Unless one of the other few proteins still present in the sample would unexpectedly interfere with the experiment, Cry3Aa induces leakage of carboxyfluorescein from Tenebrio molitor brush border membrane vesicles.

Toxin Engineering

Binding site mutagenesis

Design

In 1996, Rajamohan et al. demonstrated that mutations in the binding sites of Cry toxins can both decrease and enhance the specificity of a toxin towards its target organism. Knowing this, the 3D structure of the Cry3Aa toxin has been analysed. Three putative binding sites have been identified. These can be seen in the following table.

To mutate these, the plasmid pSB1A3_Cry3Aa_TEV_HIS has been used as a template. This plasmid contains the wildtype Cry3Aa toxin (Bacillus thuringiensis var. tenebrionis, Taxonomy ID: 1444). Its expression is regulated by the arabinose inducible promotor pBAD/araC (Bb_I0500). From this, three different plasmids have been created with randomized sequences within the binding sites (see protocols). By this we expect that each individual colony harbouring this plasmid containing an individual variants, in other words, a library of Cry3Aa proteins with randomly binding sites to be screened.

Screening

The mutants have been screened following the hightroughput procedure developed by Jacco (see above). In short, random colonies were picked from each of the binding mutants (n=144) for toxin expression and from these (n=24) were selected for activity testing (see protocols).As a negative control, cell free extract from empty expression strain, BL21(DE3) has been used, see Figure X... .

From these results, it can be concluded that the third binding site (amino acids 410-416) seems to be a good candidate for future engineering and specificity adaptation of this particular Cry toxin. Due to the relatively high deviation in reaction speed for the toxins 3.3.3 and 3.3.7, these should not be taken into account, as they are rather inconclusive. This leaves us with one proper candidate – the toxin mutant Cry3.3.5. This one has both a higher mean reaction velocity than the reference value and a relatively narrow range of measured relative k-values obtained for the 9 tested samples. Furthermore, the negative control shows that other proteins present in the cell free extract or cell debris (e.g membrane residue) interfere with our reading. Nevertheless, our preliminary results show that the mutant revokes a stronger response than the negative control, suggesting a specific binding to the Varroa vesicles.

Library sequencing

To find an indication what future engineering steps could be of use, the sequences of the altered binding sites have been obtained and looked at.

Phage display

Due to difficulty that is to obtain mites, the phage display experiments were set up in a model organism where a Cry toxin as already described, Tenebrio molitor (Cry3Aa). Furthermore, mealworms can be purchased in a lot of places and they are easy to maintain. In here, we expected to proof the applicability of this technique for specific protein engineering towards Varroa destructor. M13 was used and two different libraries were used; 7-mer flanked by a pair of cysteine residues (The Ph.D.™-C7C Phage Display Peptide Library) and the 12-mer (The Ph.D.™-12 Phage Display Peptide Library).

Display of M13KE-PIII libraries

Two different strategies for phage display were used, in vivo (conventional phage display) and in vitro (panning). For the in vitro display, a modified biopanning protocol, the Biopanning and Rapid Analysis of Selective Interactive Ligands (BRASIL) method, was used.

in vivo

First as a proof of concept feeding experiments were performed with fluorescine on mealworms and mites (see protocol). In short, the insects were fed with either a piece of carrot soaked with fluorescine or a bee larvae soaked with fluorescine. After washing with PBS buffer, the insect were ground up and fluorescence was measured. After seeing that the mites and mealworms could take up fluorescine, the phage libraries were fed to T. molitor and V. destructor and the bound phages recovered for further use(protocol). Due to a lack of mites, only two rounds of mite in vivo display could be performed. For the mealworms, the titre decreased drastically per round, see Figure X. Thus, a second run with propagation of phages between the round has been performed. During this second round, an unexpected incident caused us to stop the in vivo phage display. While tittering a phage library obtained from a mealworm, no plaques with the expected morphology (blue, lysogenic) showed up on the plate. Instead, a M13 plaques, several lytic virus were recovered (see XXX for different morphologies). The experiments have thus been stopped due to a fear of contamination in the samples. To investigate the source of the contamination, all possible sources have been looked at: used buffer, used top-agar, and the plates. The source of the unexpected plaques turned out to be the mealworm gut. XXX et al. found that phages can be found in mealworms naturally. When using guts of mealworms that have not been fed any phages prior to the process, still a lot of plaques of different kinds could be seen on the titre plates. From this, we could conclude that there are naturally some bacteriophages that E. coli can be a host for in the gut of T. molitor. While those phages could be of interest for general bacteriophage studies, or further specific binding studies, they do interfere with our phage display experiments in two ways: 1) Knowing that phages are already inside of the gut (in relatively high amounts) they might block the receptors for our phage libraries 2) As the phages and their properties are not known, working with them might not be safe As there was a shortage of living mites for the in vivo display anyway, we decided to switch to an in vivo phage display method.  Sequencing data

in vitro

For an in vitro assay, vesicles of the midgut of T. molitor and V. destructor have been produced, according to the general protocol. Briefly, cells were lysed, lipids were separated and then left for reassembly. These vesicles should have no phages attached on the outside, due to the way they have been prepared. The only chance of having phages that are naturally inhabitating mealworms or Varroa mites was thus that they have been trapped inside of the vesicle during the preparation steps and so relatively low. Unfortunately, the titre of phages bound to the vesicles prepared from Varroa destructor decreased dramatically after the first round of display and dropped to 0 after the second round. This indicates that the phages did not bind to the vesicles in a way that is strong enough to keep them attached during the separation step of vesicles with attached phages from phages. The reasons for this might be investigated and the protocol used for this phage display might still be optimized. However, the sequences of the phages left after one round have been obtained.

Varroa Isolates

Staining Isolates

To provide an alternative to currently available pesticides, we wanted to adopt Cry toxins to kill Varroa destructor. Cry toxins, also known as Bt toxins, have been used commercially for decades and can limit the environmental impact of pesticide use ¹. Cry toxins are produced by the insect pathogen Bacillus thuringiensis. They are activated by the alkaline conditions of the insect gut, after which the protoxin is cleaved off. The toxin can then recognize receptors on the gut membrane and bind to them. It will form oligomers with other toxin molecules and make a pore in the gut membrane. This is what causes the insect’s death ². Their mode of action can make them highly specific to certain species of insects.

While over 800 Cry toxins are known, research on the workings of these toxins mainly focuses on conventional pests such as the Colorado potato beetle, cabbage moths and mosquitoes. Field experiments on non-target invertebrates show that Acari (mites and ticks) are not significantly affected by Bt crops³ (transgenic crops which express Cry toxins, such as Bt cotton or Bt maize), but these use only a few known Cry toxins. Other studies have shown that there are strains of B. thuringiensis which are harmful to Acari. While most of these strains or toxins were tested against herbivorous species of mites^4,5,6,7, some are known to kill species of ticks^8,9,10,11. In fact, two strains have been discovered which show toxicity to V. destructor^12,13! Unfortunately, they did not identify a particular Cry toxin.

To find our own Cry toxin, we adapted a procedure from Rampersad et al.¹⁴ and Alquisira-Ramírez et al¹². We gathered over 800 dead mites from the Dutch foundation “De Duurzame Bij”¹⁵, Bennekom, The Netherlands.

Our method of sample preparation selected for spore formation, but not the B. thuringiensis rod-shaped morphology. Therefore, 106 colonies from the mites were investigated with brightfield microscopy. They were stained with Coomassie Brilliant Blue to make Cry toxins more visible. Five isolates were identified as Bacillus-like species. Some of them, including Bacillus thuringiensis HD-350, are shown in Figure 2.

In Vivo Toxicity

We wanted to test the strains we identified for their toxicity to Varroa destructor. Protein extracts and SDS-PAGE were performed. The toxicity assay was based on the assay used by Alquisira-Ramírez et al.¹², but we faced one major problem: the assay, including the negative control, killed all of the Varroa mites. Therefore, it was not a reliable indicator of the isolates’ toxicity to Varroa. An alternative would be to use an assay where we dip honeybee larvae in protein extracts from the strains; a Varroa mite which feeds on the larvae should then ingest any toxins. Unfortunately, Varroa mites tend to share feeding sites and use them repeatedly¹⁶, which means that not all mites would consume the protein extract.

A sample-size calculation was performed in R with the following code to assess the feasibility of such an assay.
> p=0.66
> p0=0.5
> alpha=0.05
> beta=0.1
> (n=p*(1-p)*((qnorm(1-alpha)+qnorm(1-beta))/(p-p0))^2)

Here, p is the alternative hypothesis, p0 the null hypothesis, alpha the type I error and beta the type II error. The values of these variables were based on the following assumptions:

• The null hypothesis is 50% mite mortality. Natural mite mortality is very high in the laboratory.
• Linea Muhsal estimated during feeding experiments that approximately 16% of mites ingested the fluorophores she had dipped honeybee larvae in.
• If an effective Cry toxin is ingested, we expect to see 100% mite mortality. This is an optimistic assumption.
• 16% of mites eat a toxin which causes 100% mite mortality, so if there is a toxin, we should see 66% mite mortality.

With these assumptions, a sample size of 75 was calculated. We were generally able to gather a maximum of 40 mites per week. Gathering enough mites to test all our samples was unfeasible. Therefore, pursuit of this type of toxicity assay was cancelled in favour of further development of an in vitro assay.

16s rRNA sequencing and SDS-PAGE results

All five isolates with Bacillus-like morphology had their 16s rRNA sequenced. A search of NCBI’s 16S ribosomal RNA sequences database resulted in the following alignments, each with approximately 99% identity of their best match. The isolates and their matches are displayed in Table 1.

It can be difficult to tell Bacillus species apart with just a 16s rRNA sequence¹⁷, so an SDS-PAGE gel of the protein extracts was prepared to further analyse them. This gel is shown in Figure 3.

Proteomics analysis of isolate V82

Based on the previous results, our Lysinibacillus isolate V82 had the most potential as a putative mite pathogen. It is relatively closely related to a known insect pathogen, Lysinibacillus sphaericus, a species which is known to make a 100 kDa mosquiticidal toxin ¹⁸. Our isolate also showed a strong band at the 100 kDa region (Figure 3). A new SDS-PAGE gel was prepared with independently grown V82 cultures and bands were cut out for peptide extraction, so the 100 kDa protein could be analysed with LC-MS/MS. Negative control bands were cut out approximately ~1 cM above the 100 kDa bands. MaxQuant was used to analyze the data.

Analysis of the LC-MS/MS results resulted in Figure 5. Figure 5 is a vulcano plot; the x-axis plots the protein abundance ratio between the 100 kDa band and its respective control band, while the y-axis plots the p-value of this difference in protein abundance. Therefore, a protein in the left half of the plot is more abundant in the control bands, while the opposite goes for proteins in the right half of the plot. The higher it is on the y-axis, the more significant this difference is.

Contaminations such as keratins and the trypsin used to digest the samples should not differ significantly between the samples. This is also shown in Figure 5 as most contaminants (the orange data points) are present in the bottom half of the vulcano plot. The relative label-free quantification (LFQ) intensity of the significant proteins shown in Figure 4 was very low, in all cases lower than the LFQ intensity of trypsin. From a biological perspective, this makes sense; there is no reason for an organism to produce mass amounts of alanine tRNA ligase. However, this and the fact that only 3% of peptides could be matched indicates that the 100 kDa protein was not present in the prepared Lysinibacillus sequence database. Therefore, the peptides were also matched against a Bacillus database, but this yielded similar results. Genomic DNA was extracted from V82 and sent for sequencing so the LC-MS/MS data could be run against the isolate's own DNA. This genomic DNA was also analyzed with the Toxin Scanner.

References

References for in vitro essay

1. Alejandra Bravo, Sarjeet S. Gill, Mario Soberón. Mode of action of Bacillus thuringiensis Cry and Cyt toxins and their potential for insect control. Toxicon. 2007 Mar 15; 49(4): 423–435. ↩

2. Hideo Ohkawa,Hisashi Miyagawa,Phillip W. Lee. Pesticide Chemistry: Crop Protection, Public Health, Environmental Safety. Wiley-VHC Verlag GmbH & Co KGaA Weinheim. 2007. p.193↩

 

References for Varroa isolates

1. Lacey, L. A., Grzywacz, D., Shapiro-Ilan, D. I., Frutos, R., Brownbridge, M., & Goettel, M. S. (2015). Insect pathogens as biological control agents: back to the future. Journal of invertebrate pathology, 132, 1-41. ↩

2. de Maagd, R. A., Bravo, A., & Crickmore, N. (2001). How Bacillus thuringiensis has evolved specific toxins to colonize the insect world.TRENDS in Genetics, 17(4), 193-199. ↩

3. Marvier, M., McCreedy, C., Regetz, J., & Kareiva, P. (2007). A meta-analysis of effects of Bt cotton and maize on nontarget invertebrates.science, 316(5830), 1475-1477. ↩

4. Ahmed, N., Wang, M., & Shu, S. (2016). Effect of commercial Bacillus thuringiensis toxins on Tyrophagus putrescentiae (Schrank) fed on wolfberry (Lycium barbarum L.). International Journal of Acarology, 42(1), 1-6. ↩

5. Chapman, M. H., & Hoy, M. A. (1991). Relative toxicity of Bacillus thuringiensis var. tenebrionis to the two‐spotted spider mite (Tetranychus urticae Koch) and its predator Metaseiulus occidentalis (Nesbitt)(Acari, Tetranychidae and Phytoseiidae). Journal of Applied Entomology, 111(1‐5), 147-154. ↩

6. Dunstand-Guzmán, E., Peña-Chora, G., Hallal-Calleros, C., Pérez-Martínez, M., Hernández-Velazquez, V. M., Morales-Montor, J., & Flores-Pérez, F. I. (2015). Acaricidal effect and histological damage induced by Bacillus thuringiensis protein extracts on the mite Psoroptes cuniculi. Parasites & vectors, 8(1), 1. ↩

7. Erban, T., Nesvorna, M., Erbanova, M., & Hubert, J. (2009). Bacillus thuringiensis var. tenebrionis control of synanthropic mites (Acari: Acaridida) under laboratory conditions. Experimental and Applied Acarology, 49(4), 339-346. ↩

8. Fernández-Ruvalcaba, M., Peña-Chora, G., Romo-Martínez, A., Hernández-Velázquez, V., de la Parra, A. B., & De La Rosa, D. P. (2010). Evaluation of Bacillus thuringiensis pathogenicity for a strain of the tick, Rhipicephalus microplus, resistant to chemical pesticides. Journal of Insect Science, 10(1), 186. ↩

9. Hassanain, M. A., Garhy, M. E., Abdel-Ghaffar, F. A., El-Sharaby, A., & Megeed, K. N. A. (1997). Biological control studies of soft and hard ticks in Egypt. Parasitology research, 83(3), 209-213. ↩

10. Neethu, K. B., Priji, P., Unni, K. N., Sajith, S., Sreedevi, S., Ramani, N., ... & Benjamin, S. (2015). New Bacillus thuringiensis strain isolated from the gut of Malabari goat is effective against Tetranychus macfarlanei. Journal of Applied Entomology. ↩

11. Zhioua, E., Heyer, K., Browning, M., Ginsberg, H. S., & LeBrun, R. A. (1999). Pathogenicity of Bacillus thuringiensis variety kurstaki to Ixodes scapularis (Acari: Ixodidae). Journal of medical entomology, 36(6), 900-902. ↩

12. Neethu, K. Alquisira-Ramírez, E. V., Paredes-Gonzalez, J. R., Hernández-Velázquez, V. M., Ramírez-Trujillo, J. A., & Peña-Chora, G. (2014). In vitro susceptibility of varroa destructor and Apis mellifera to native strains of Bacillus thuringiensis. Apidologie, 45(6), 707-718. ↩

13. Tsagou, V., Lianou, A., Lazarakis, D., Emmanouel, N., & Aggelis, G. (2004). Newly isolated bacterial strains belonging to Bacillaceae (Bacillus sp.) and Micrococcaceae accelerate death of the honey bee mite, varroa destructor (V. jacobsoni), in laboratory assays. Biotechnology letters, 26(6), 529-532. ↩

14. Rampersad, J., & Ammons, D. (2005). A Bacillus thuringiensis isolation method utilizing a novel stain, low selection and high throughput produced atypical results. BMC microbiology, 5(1), 1. ↩

15. This is the homepage of “De Duurzame Bij”. They want to stop treating Varroa with pesticides. ↩

16. Kanbar, G., & Engels, W. (2005). Communal use of integumental wounds in honey bee (Apis mellifera) pupae multiply infested by the ectoparasitic mite Varroa destructor. Genetics and Molecular Research, 4(3), 465-472. ↩

17. Bavykin, S. G., Lysov, Y. P., Zakhariev, V., Kelly, J. J., Jackman, J., Stahl, D. A., & Cherni, A. (2004). Use of 16S rRNA, 23S rRNA, and gyrB gene sequence analysis to determine phylogenetic relationships of Bacillus cereus group microorganisms. Journal of clinical microbiology, 42(8), 3711-3730. ↩

18. Berry, C. (2012). The bacterium, Lysinibacillus sphaericus, as an insect pathogen. Journal of invertebrate pathology, 109(1), 1-10. ↩