Team:Brno Czech Republic/Experiments
Experiments
Here, you can find more information about the experiments we've done. You can read about our reasons for performing the experiment, its theoretical background and a short description of our workflow. You will find the result of each experiment in the section Results & Conclusion. If you are interested in more specific information about the methodist we used, you can take a look at our Protocols here or through the hypertext link mentioned with specific experiments.
At Masaryk university, we were allowed to work in a laboratory, which is usually used for teaching practical courses during the semester. There we had everything we needed - flow boxes, cyclers, centrifuges, pipettes and so on. We also had the support of doctoral students and technical assistants of the Experimental biology department. Unfortunately, the coronavirus situation in our country worsened at the end of the summer, so we had to stop with our experiments earlier than planned.
We have however already planned out what to improve in the future. We have also already brainstormed how to characterize the rest of our synthetic constructs which we will hopefully share with you next year.
Basic experiments
Preparation of growth media
The protocols for media and agar preparation can be found here: PROTOCOL: Preparation of growth media.
Preparation of synthetic constructs and primers
Preparation of synthetic constructs
Motivation:
For the transformation of organism B. subtilis we had 6 synthetic constructs prepared by company IDT within the offer mediated by iGEM contest.
These synthetic modules are IM, ScafL, LysBL-DocCt, LysGG-DocCt, ScafD, MlrB-DocBc, MlrC-DocAc and are further described in the section Project design.
Short description:
- Genes were delivered in a powder form and they had to be resuspended according to the instructions provided by the producer.
- Synthetic genes were shortly centrifuged at Short spin mode and then resuspended in 50 µl nuclease free H2O in the flowbox.
- Resuspended genes are less stable, therefore it was necessary to keep them on ice or put them in a freezer at -20 °C.
- We worked with them only in Flowbox to minimize risk of DNA contamination.
- It is better to use pipette tips with filters for the gene stocks, because pipettes might contain aerosols from previous pipetting. Possible contamination could devalue the whole stock.
Preparation of primers
Motivation:
Within the CYANOTRAP project we worked with a lot of primers. Their detailed description is here.
Short description:
Our primers were also synthesised by the IDT company. They were also delivered in powder form and it was necessary to resuspend them in nuclease free H2O, according to provided instructions.When working with primers, we always used 10 nM solution, which we prepared from 100 nM stock solution. The stock solution was created by resuspending delivered powder. We assigned each primer a number to be able to distinguish them and talk about them easily.
Long-Range PCR of synthetic constructs
Protocol is available here: PROTOCOL: Long-Range PCR
Motivation:
We routinely used long-Range PCR in our experiment for the amplification of our DNA. Another reason was our issues with colony PCR. Amplified DNA was used for many purposes, for example as gel electrophoresis, sequencing or storage.
Short description:
When performing Long-Range PCR, we followed a protocol that came with the GoTaq® Long PCR Master Mix by the company PROMEGA here. While working, it is necessary to keep everything sterile. Positive and negative controls are also necessary.
Preparation of glycerol stocks and vectors for cloning
Protocols are available here (PROTOCOL: Preparation of glycerol stocks) and here (PROTOCOL: Isolation of plasmids).
Motivation:
We received our strain of B. subtilis and the vectors for our project from Dr. Libor Krásný. He is the head of the Laboratory of Microbial Genetics and Gene Expression at the Academy of Sciences of Czech Republic. After the plates with strains of B. subtilis and E. coli cells containing our vectors were successfully delivered, we needed to make glycerol stocks from them and isolate our vectors from E. coli.
Short description:
In total we got 5 plates with following strains:
You can read more about our bacterial strains and vectors in Project design
In the end, we've worked mainly with vector pDG3661 and a little bit with pDG1664. Other strains carrying our vectors were stored at - 80 °C.
One colony from each plate was then streaked on a fresh plate with Lysogeny broth (LB) agar. For E. coli strains, we used a medium with ampicillin (concentration: 100 µg/ml). Due to initial miscommunication, we thought that the ideal temperature for the growth of B. subtilis is 28 °C. We used this temperature in this experiment and in the following weeks. After realising our mistake, we proceeded to use the correct temperature of 37 °C. Lower temperature caused delayed growth. The plates with E. coli were incubated at 37 °C.
Next day one larger colony from the plate with B. subtilis was chosen and it was densely spread on 6 different plates. One colony was chosen from each plate with E. coli and each colony was densely spread on 2 different plates. We've inoculated two plates with each colony, to have a backup in case of contamination.
Following day the plates were harvested and the bacteria were transported into 2 ml of liquid LB medium with 20% glycerol. These glycerol stocks were then stored at -80 °C.
Plasmid were isolated from a liquid overnight culture of corresponding E. coli strain, we used PureYield™ Plasmid Miniprep System by Promega Corporation.
Preparation of genetic modules in Escherichia coli cloning host
Test of vector background of Escherichia coli
Protocol is available here: PROTOCOL: Test of vector background of Escherichia coli
Motivation:
After transforming E. coli with our vectors after restriction cloning, we got only a small number of transformants. Therefore, we tested the transformation background to determine if the growth we are observing is just the background. Background refers to the fact that in some cases even a vector which is cleaved with two different restriction enzymes and dephosphorylated with antarctic phosphatase, could reconnect.
Short description:
One batch of competent cells was transformed with supercoiled plasmid. Another batch of cells was transformed (see transformation experiment: Cloning and preparation of E. coli below) with a vector cleaved with two restriction enzymes - in our case HindIII and EcoRI - and also dephosphorylated with Antarctic phosphatase. Both batches were inoculated at the LB agar plate with appropriate antibiotics. Plates were incubated overnight in 37°C.
Preparation of genetic modules in Escherichia coli cloning host
Protocol is available here: PROTOCOL: Preparation of genetic modules in Escherichia coli cloning host
Motivation:
We transformed our constructs into E. coli Dh6α NEB® 5-alpha Competent (High Efficiency) in order to multiply our vectors with inserts and to make glycerol stocks.
Short description:
First, the plasmid was isolated from a well grown overnight culture of E. coli. Then the empty vector pDG3661 was cleaved by two different restriction enzymes - EcoRI and HindIII. Antarctic phosphatase was added to keep the plasmid from self-ligating. As for our insert, we used amplified PCR products of synthetized genes by IDT. Subsequently, they were digested with the same restriction enzymes under the same conditions (with no Antarctic phosphatase added).
We used different ligation strategies during preparation of each of these plates. T4 ligase from Promega was used for the first ligation reaction. This reaction ran overnight at room temperature. The cells transformed with this reaction mix were inoculated on plate 2 (see Results). The ligation reaction, with which we transformed the cells plated on plate 3, ran for 15 min at room temperature. This reaction mix contained T4 ligase from NEB. Our last ligation reaction was used to transform cells plated on plate 4. Unlike in previous reactions, the products of restriction digestion were first separated on agarose gel and then extracted using the kit mentioned above. The ligation reaction ran for 15 min at room temperature. Ligation mixes were added to E. coli cells, which then underwent heat-shock and were incubated for 75 min before being inoculated on a LB plate containing ampicillin. In this experiment we also used positive control. Our positive control of competence ability of competent cells contained empty supercoiled plasmid pUC19. The cells grew overnight at 37 °C. Next day we evaluated the results.
Proof of transformation of Escherichia coli
Protocol is available here: PROTOCOL: Proof of transformation of Escherichia coli
Motivation:
We needed to determine the success of our transformation and the presence of our vector. We've had some issues with colonyPCR, so we've had to use different methods of proof - restriction digestion and sequencing.
Short description:
Firstly, we purified our plasmid from overnight culture. Isolated plasmid was then cleaved by two different restriction enzymes - EcoRI and HindIII. The restriction reaction lasted 2 hours at 37°C. Afterwards, we used agarose electrophoresis to show the size of the fragments.
Preparation of vector pDG3661 for sequencing
Motivation:
The aim of this experiment was to verify the sequence of isolated plasmid pDG3661, specifically the sequence of our insert - the Immobilization module (IM). This experiment further demonstrates the success of the cloning and E. coli Dh6α transformation.
Short description:
We followed the protocol for sequencing of plasmid templates, provided by the SEQme company. We used isolated plasmid pDG3661_IM (See PROTOCOL: Isolation of plasmids). We added forward and reverse primers to the samples of isolated plasmid. Both primers were added separately, so there were two sequencing mixes for each sample and the sequence was ready to be read from both strands. We send these samples to the company SEQme based in Czech republic.
Integration of the exprssion casette with IM into the chromosome of Bacillus subtilis
Preparation of competent cells of Bacillus subtilis
Protocol is available here: PROTOCOL: Preparation of competent cells of Bacillus subtilis
Original publications:
Spizizen J. (1958). TRANSFORMATION OF BIOCHEMICALLY DEFICIENT STRAINS OF BACILLUS SUBTILIS BY DEOXYRIBONUCLEATE. Proceedings of the National Academy of Sciences of the United States of America, 44(10), 1072–1078. https://doi.org/10.1073/pnas.44.10.1072
- modified by dr. Krásný - Laboratory of Microbial Genetics and Gene Expression
Dubnau, D., & Davidoff-Abelson, R. (1971). Fate of transforming DNA following uptake by competent Bacillus subtilis. I. Formation and properties of the donor-recipient complex. Journal of molecular biology, 56(2), 209–221. https://doi.org/10.1016/0022-2836(71)90460-8
- modified by dr. Krásný - Laboratory of Microbial Genetics and Gene Expression
Motivation:
We needed to prepare natural competent cells of B. subtilis. The Laboratory of Microbial Genetics and Gene Expression and dr. Krásný from Prague really helped us at the beginning of our experiment. He also recommended this type of competence indication - as he has very good experience with this protocol.
Theory:
The preparation of naturally competent cells is necessary for successful transformation of B. subtilis. It is also possible to prepare electrocompetent cells from B. subtilis, but that wasn't necessary in our case. This method can be done only in certain media and in the post-exponential phase of bacterial culture growth. These competent cells are able to efficiently take in exogenous high-molecular-weight DNA. In the competent culture, not every cell is in its competent state. It is however enough to generate a sufficient amount of transformants. Their preparation includes a two-step process using glucose minimal salt-based media with Spizizen salts.
Short description:
This experiment requires the preparation of two different media - SPI & SPII and an overnight culture of B. subtilis. The next day, overnight culture has to be inoculated into the SPI medium in duplicate - one for measurement and one for later use. After inoculation, the OD600 of the culture must be measured. The cultivation must continue until the values indicate the departure from the exponential rate (this is very important). This usually takes around 5 hours and 30 min. As the values deviate from the exponential curve, culture is inoculated into the SPII medium and cultivated under the same conditions for 90 minutes. Afterwards, the culture is placed on ice to stop its growth and centrifuged. Pellet is resuspended in a fraction of the supernatant and glycerol. Aliquots can then be made and stored at -80°C.
Testing competent cells of Bacillus subtilis
Protocol is available here: PROTOCOL: Transformation of Bacillus subtilis
Motivation:
The ability of our natural competent cells to accept plasmid DNA needed to be verified. We performed this experiment many times before the synthetic genes arrived to optimize the process. In the end we determined the best conditions for our transformation.
Theory:
This experiment primarily verifies the competency of the cells and the efficiency of their transformation. The use of controls is essential for this experiment. B. subtilis cells without plasmid pDG3661 carrying a resistance to chloramphenicol were used as a negative control. These bacteria should not be able to grow in the presence of this antibiotic. It is also necessary to confirm that our competent cells are alive by growing them in LB agar with no antibiotics, where colonies should appear regardless of the success of our transformation.
Short description:
Our natural competent cells were transformed with plasmids pDG3661_IM. We then followed this protocol (time of shaking was 60 min). Afterwards, we inoculated the bacteria onto LB plates with chloramphenicol. Cells transformed with empty plasmid pDG3661 were used as positive control. The cells were cultivated overnight at 37 °C.
Transformation of Bacillus subtilis
Protocol is available here: PROTOCOL: Transformation of Bacillus subtilis
Motivation:
We transformed naturally competent B. subtilis 168 in order to produce our proteins - specifically the IM.
Short experimental description:
We first had to isolate pDG3661_IM from E. coli Dh6αfrom previous experiments. The naturally competent B. subtilis cells were preserved in the form of glycerol stocks. We added 1 µg of plasmid to 100 µl of natural competent cells. We also prepared positive and negative controls. The positive control contained empty supercoiled plasmid pDG3661 carrying antibiotic resistance. The competent cells without plasmid were used as the negative control. These controls confirmed that the transformation was performed well and that the antibiotic was working on non-resistant cells. We incubated the tubes containing competent cells and plasmids on a shaker at 200 rpm and 37 °C for 1 hour. Afterwards, we planted the competent cells on agar plates with selection medium containing chloramphenicol. The cells grew overnight at 37 °C. Next day we evaluated the results.
TOPO cloning
Protocol is available here: PROTOCOL: TOPO cloning
Motivation:
Beside restriction cloning we also performed TOPO cloning, in case the restriction cloning wouldn't be successful.
Theory:
TOPO cloning is a tool of synthetic biology, which enables cloning of an insert into a vector without using a ligase. It utilizes the ability of Taq polymerase to add a single deoxyadenosine to the 3'-end of the PCR products. Taq polymerase thus creates a sticky end of the insert, which can then be used to ligate it to our vector. Important role also plays DNA topoisomerase I., which recognises a specific DNA sequence and catalyzes its digestion and the subsequent re-ligation. Under optimal conditions, the insert can be ligated to the specific vector thanks to the activity of topoisomerase I.
Short description:
First we amplified our synthetic gene using a Long-Range PCR. We've then added this insert to a plasmid specifically designed for topo cloning - Topoisomerase I-activated pCR™4-TOPO® vector
To transform the competent E. coli cells with this mix, we followed the variant 2 in Protocol. Transformants were then streaked on Xgal plates with ampicillin. The plates were incubated overnight at 37 °C in the dark.
As we were trying out the TOPO cloning, we've decided to redo restriction cloning as well. In the end, we've gotten colonies of transformed cells in both cases. We've opted to continue with the transformants from restriction cloning because in that case, our insert was already in the correct plasmid for the chromosomal integration in B. subtilis.
Proof of the successful transformation of Bacillus subtilis
Protocol is available here: PROTOCOL: Proof of the successful transformation of Bacillus subtilis
Motivation:
After transforming B. subtilis with our vectors, we needed to confirm that the integration of our IM gene and the lacZ gene into the chromosome of B. subtilis was successful. As we had some issues with colony PCR, we had to use different methods of proof. Therefore we decided to isolate genomic DNA of B. subtilis and use standard long-Range PCR for its amplification. We then used sequencing and also the AmyE test to confirm the integration of our gene.
Theory:
We transformed B. subtilis with the vector pDG3661_IM containing also the lacZ gene. The sequence is designed for integration into the amyE region of the chromosome of our bacteria. Our primers were designed to be complementary to the chromosomal sequence in the proximity of the integration site. PCR amplicons should thus contain a part of the B. subtilis chromosome, our synthetic, the lacZ gene and a component of vector pDG3661 for ectopic integration. The size of these amplicons should be around 8074 bp.
There are many different methods of isolating the genomic DNA of the B. subtilis. We tried two and tested their effectiveness. The first method - we called it the physical method - used extreme temperature changes. The second - chemical - method utilized a lysis buffer. We also tested the impact of the age of our cultures, as B. subtilis forms endospores which are not suitable for DNA isolation. As we expected, fresh overnight culture gave better results than a culture which was stored for a couple of days.
Short description:
We used fresh overnight cultures and older cultures grown on plates. Both methods of chromosomal isolation are described in detail in the protocol. The physical method resulted in higher yield of DNA (about 150 ng/µl). As for purity, there was no difference between the two methods.
We used the isolated DNA as a template for long PCR. The amplicons were then visualised on agarose gel.
AmyE test (Starch hydrolysis test)
Protocol is available here: PROTOCOL: AmyE test (Starch hydrolysis test)
Original publication:
Guérout-Fleury, A. M., Frandsen, N., & Stragier, P. (1996). Plasmids for ectopic integration in Bacillus subtilis. Gene, 180(1-2), 57–61. https://doi.org/10.1016/s0378-1119(96)00404-0
Motivation:
AmyE test is a simple method of demonstrating ectopic integration into the chromosome of B. subtilis. The integration site in the chromosome encodes š¼-amylase, which is able to degrade starch in the media. If the integration site is intact, the š¼-amylase is produced and a clear halo around the colonies can be observed after iodine treatment as the starch gets digested. If the integration into chromosome is successful and the sequence encoding š¼-amylase is split by the inset, the enzyme is not produced and starch is not degraded. In our experiment we tried using an AmyE test to confirm the transformation of our B. subtilis with oue vectors. We compared our samples with B. subtilis 168, which was not transformed with any DNA.
Short description:
Two layer agar is used for this method - the first layer is a regular agar containing corresponding antibiotic and the second layer is a thin agar with starch supplement. After the addition of a few crystals of iodine, the starch will turn dark blue. Do not forget to work in the laboratory hood as iodine fumes are harmful.
Modified Blue-White selection
Protocol is available here: PROTOCOL: Modified Blue-White selection
Motivation:
We wanted to also use a modified Blue-White selection in order to verify chromosomal integration. Unfortunately, after a few confusing results, we realized that our construct, which was cloned upstream of the spoVG-lacZ gene, has a STOP codon at the end and spoVG-lacZ does not have its own RBS site. Due to this flaw in our design, we could not use this selection method. If we decided to perform this test in the future, we would also place an RBS at the end of our synthetic construct. We realized that iGEM is also (and maybe mainly) about learning from our mistakes.
Theory:
The vectors which we chose for our project include spoVG-lacZ gene. This gene was created by fuzing the spoVG gene from B. subtilis and lacZ gene which encodes ß-galactosidase. This enzyme is able to degrade the Xgal substrate to gain a colorful product. Colonies with active ß-galactosidase turn blue and colonies without it stay colorless. In our case, colonies containing our insert would thus turn blue, which is not how the Blue-white method is usually done. In our project the transcription of the spoVG-lacZ gene is controlled by constitutive promoter Pveg, so we did not use IPTG to induce the protein expression.
Short description:
In this experiment LB agar with added Xgal is used. We prepared this agar and then we inoculated the bacteria onto the plates containing this medium.
Preparation of sample for sequenation - Bacillus subtilis
Motivation:
We needed to verify the sequence which was integrated into the chromosome of B. subtilis. This would demonstrate the success of our transformation and chromosomal integration. We tried to get a good sequencing result but unfortunately, we were not able to prepare the sample properly.
Short description:
We used the services of the company SEQme, based in Czech republic. The sequencing was performed three times. In the first attempt we used the whole chromosomal DNA of B. subtilis 168, which was transformed with vector pDG3661_IM. We used primer 1 and 29, which allowed us to obtain a sequence of the main part of our synthetic gene – the immobilisation module. More information can be found in the section Primers. However, the results weren't optimal, so we performed this experiment once again with optimised sample preparation.
When repeating the sequencing, we prepared chromosomal DNA isolated with the same method. We then amplified desired DNA sequence, using the same primers as were being used for sequencing, specifically primer 1 and 13. The PCR product was then separated from chromosomal DNA on 0,6 % agarose gel. Afterwards, we performed gel extraction and prepared the samples for sequencing according to this protocol and sent them for sequencing. In the third attempt, we used the same method. Despite our efforts, the results were not optimal.
Characterization of immobilization module
Calibration of the device with iGEM Measurement kit should precede this experiment. We however did not receive it this year and, due to a lack of time and the coronavirus situation, we also werenāt able to handover this material to another iGEM team. We have preformed the experiment in the final days of our lab work and we consider its result as unclear and inconclusive. Next year we will try to get the iGEM Measurement kit for calibration of the device in order to measure OD600 in time.
Cultivation experiment
Protocol is available here: PROTOCOL: Cultivation experiment
Original publication:
You C., Zhang X. Z., Sathitsuksanoh N., Lynd L. R. and Zhang Y. H. P. 2012. Enhanced microbial utilization of recalcitrant cellulose by an ex vivo cellulosome-microbe complex. Appl. Environ. Microb. 78 (5): 1437–1444. DOI: 10.1128/AEM.07138-11
Motivation:
For optimal extracellular production of heterologous proteins, B. subtilis should be cultivated in 2x Mal medium. This experiment was inspired by the work of You et al. in 2012.
Short description:
Expression cultures after an overnight cultivation and the control strains were inoculated into 10 ml of liquid LB medium with and without chloramphenicol, respectively. This selection strategy was kept the same for the whole experiment. After 16 hours of growth at 37 °C and 200 rpm, 20 ml of fresh LB medium were inoculated with the cultures so that the starting OD600 would be 0.05. OD600 was then measured every 30 minutes. When the culture reached OD600 of 1.2, 50 ml of modified 2x Mal medium were inoculated with these cultures to get the starting OD600 of 0.0048. OD600 was then measured every 30 minutes. After the culture reached OD600 of 1.5, fresh 50 ml of modified 2x Mal medium were inoculated with these cultures to get the starting OD600 of 0.0025. The culture was cultivated for 14 hours and then used for further experiments. The protocol contains the preparation of the modified 2x Mal medium as well as the cultivation procedure.
Collection of samples and their SDS-PAGE and Western blot analysis
Protocol is available here: PROTOCOL: Bradford protein assay , SDS-PAGE, Coomassie staining, Western blot, Proccesing of samples for SDS-PAGE
Motivation:
We used SDS-page and Western blotting to prove the expression of the IM in B. subtilis. We tried to gather the samples from the cultures with different OD600 and several different methods of sample processing to obtain the best results.
Theory:
SDS-page is a method used to separate proteins based on their size, using a highly linked gel matrix and direct electrical current. The effect of different charges is suppressed as the proteins are denatured and encapsulated by negatively charged SDS molecules.
Western blot is used to verify the presence of a protein through its interaction with specific antibodies, which are then detected by chemiluminescence. From the SDS gel, the proteins are transferred to the PVDF membrane. Primary antibodies can then interact and bind to the Tag on our protein. Afterwards, secondary antibodies carrying chemiluminescent enzymes bind to primary antibodies.
Short description:
The first round of sample collection was performed thusly. At first the samples were collected from an overnight culture in LB, then from 2 x Mal medium at the OD600 of 1.2 and finally after 24 h cultivation in LB. These samples were processed with B-PER reagent and 2 M urea. SDS-page and Western blot was carried out using c-Myc antibody. We tried antibodies from mouse and rabbit. Primary mouse antibody was monoclonal c-Myc (9E10) from Franz Klein lab. Secondary was anti-mouse IgG + IgM, (H+L), conjugated to HRP, from Pierce # 31444. Primary rabbit antibody was polyclonal c-Myc (A-14) from Santa Cruz Biotechnology, INC. Secondary rabbit antibody was anti-rabbit IgG, (H+L), conjugated to HRP, from Pierce # 31460. The first concentrations of antibodies were 1:1000 of the primary antibodies and 1:15000 of the secondary antibodies.
Unfortunately, the results yielded from negative control were rather similar to the expression strains and there is no clear proof of the presence of the IM. The gel was also stained with coomassie blue, but also there was no visible difference between the control and expression samples.
For our second attempt, we tried to make some adjustments. The first incubation was accidentally done at 30 °C. We used the correct temperature of 37 °C for the second run. This time we also gathered whole cell lysates and whole cells. We also adjusted the concentrations of primary and secondary antibodies to 1:500. The first samples were collected from overnight cultures growing in LB medium, another batch of samples was obtained from 2x Mal medium at the OD600 of 1.5. Final samples were collected after the cultures were incubated in 2x Mal medium for 24 hours.
Preparation of cellulose microbeads
Protocol is available here: PROTOCOL: Preparation of cellulose microbeads
Motivation:
We needed to clean the cellulose microbeads from the 20% ethanol in which they were stored.
Short description:
Approximately 0,5 ml of microbeads were transferred from the storage container to Eppendorf tubes and left to sediment. The ethanol was then removed and the rest of it was left to evaporate in a thermoblock at 50 °C. After that, the microbeads were resuspended in 0,5 ml of PBS buffer. This suspension was left to sediment and the PBS buffer was discarded. The washing was repeated three times and at the end the microbeads were resuspended in a small amount of PBS. You can find the detailed protocols here.
Cellulose binding experiment
Protocol is available here: PROTOCOL: Cellulose binding experiment
Motivation:
We wanted to find out if the IM enhances the ability of B. subtilis to bind to cellulose and to simulate the conditions in the CYANOTRAP device.
Short description:
The experiment was done using two expression strains and a control strain of unmodified B. subtilis 168 (C). The cellulose microbeads mixed only with the PBS buffer were also used to test the sterility of our microbeads (M). The cells were cultivated as described in the Cultivation experiment. After reaching the OD600 of 1.6 (approximately), two sets of samples were collected. In the first set, 0.75 ml of each strain in the PBS buffer at the OD600 of 1; 0.1 and 0.01 were prepared. In the second set, 5 ml of each strain in the PBS buffer at the OD600 of 1 were prepared.
The first set of samples was mixed with approximately 0,75 ml of cellulose microbeads that were prepared following this protocol. The second set was mixed with approximately 5 ml of the same material. The cells were then incubated with the microbeads for 1 hour at room temperature with rotation.
After that, the microbeads were left to sediment for 15 minutes. The supernatant was separated from the pellet and 100 µl of the supernatant of each sample (11.1 of the IM strain; 11.5 of the IM strain; control, M) from the first set of samples were applied onto LB agar plates. The pellets of the same set were washed twice with the PBS buffer to wash away any unbound cells and then the 100 µl of each type of sample were applied onto LB agar plates as well. These plates were incubated overnight at 37 °C and photographed the next day.
The pellet of the second set of samples was also washed twice with PBS for the same reason and then 2 ml of the pellet,resuspended in a small amount of the PBS buffer, were transferred to a plastic gravity column with the cutoff limit of 20 µm. This cutoff ensures that the cells bound to cellulose microbeads will stay above the filter while the unbound cells will appear in the flowthrough. 20 ml of fresh PBS was added to the microbeads with (presumably) bound cells and it was left to drop to a collection tube. After the entirety of the PBS buffer passed through the column, 100 µl of each sample was applied to LB agar plates. The plates were then incubated overnight at 37 °C.
