Difference between revisions of "Team:Nottingham/Lab"

Ideas

Immediately after meeting for the first time, we were tasked with finding a project that we would be working on over the summer. At first, the idea seemed daunting but before we knew it, amazing (sometimes outrageous) ideas were being suggested.

Almost a month later, after numerous presentations, we narrowed down our potential projects to five:

Ideonella sakaiensis

Ideonella sakaiensisis a bacterium that can degrade plastic (PET) using two enzymes. It was discovered outside a plastic recycling facility in Japan and subsequently isolated by research teams at Kyoto Institute of Technology and Keio University(Yoshida et al., 2016). The idea of being able to tackle the plastic crisis affecting the oceans and marine life was a particularly appealing one. However, we quickly realised that we didn’t know how to transform the organism and the time-scale for how long it would take. Also, our project was very similar to other research being conducted and so it would be difficult to come up with a novel way to address the crisis.

Water bottle biosensor

According to the House of Commons Environmental Audit Committee, in their first report of the season (2017-19), in the UK alone we use ‘13 billion plastic bottles every year’ and ‘only 7.5 billion are recycled’. This prompted the team to suggest a biosensor that detects harmful substance in plastic bottles. As a result, the average consumer would buy fewer plastic bottles and reuse their bottle. However, many solutions immediately were proposed that counteracted the need for the biosensor-this included the fact that many people buy permanent water bottles that they wash and reuse; it would be more cost effective to campaign recycling efforts than to market a biosensor and it is cheaper to just recycle a plastic bottle and buy a new one than to buy a biosensor. Indeed in the report by the Environmental Audit Committee (potential stakeholders), they suggest increasing the number of water fountains in open spaces and improving recycling through a number of methods including Deposit Return Schemes (which incentivise consumers).

Bdellovibrio bacteriovorus

Bdellovibrio bacteriovorus is a predatory bacterium that preys on other (Gram-negative) bacterial species (Rendulic et al., 2004). Its enzymes and their mechanisms are being studied in order to better understand the bacterium for future use as a possible therapeutic agent or as a method to penetrate biofilms (Kadouri and O'Toole, 2005; Sockett and Lambert, 2004). Our team was interested in engineering the bacterium to use it (and its enzymes) as a ‘pathogen eating machine’in food processes-for example targeting Clostridium botulinum, which releases the botulinum toxin. On top of it being an interesting species to work on, we had a leading expert on the Bdellovibrio species at our university. However, the main bacterium we were targeting (Clostridium botulinum) is a Gram-positive bacterium so is not recognised by Bdellovibrio. Also, it would take many weeks to engineer Bdellovibrio for single gene mutations, which is difficult to transform using random transposon mutagenesis. This was a project that required many years which our team did not have!

mRNA interference of Streptococcus mutans

Streptococcus mutansis one of a number of bacteria involved in tooth decay and is the most prevalent. According to Public Health England, in the UK, ‘almost a quarter (24.7%) of 5 year olds have tooth decay’ of which 3 or 4 teeth are affected. Dental health problems also have a heavy financial burden on the NHS-it spends around £3.4 billion per year on dental care. What makes S. mutans particularly hard to deal wit his its ability to form biofilms regulated by glucosyltransferases which catalyse sucrose to adhesive glucan. In particular, GtfB and GtfB seem to be the most important in biofilm production-mutations in the gtfB and gtfC genes disrupted microcolony formation on saliva coated surfaces(Koo et al., 2010). Our idea was to use a bacteriophage (a virus that only infects bacteria) to deliver micro RNAs or small interfering RNAs to silence those genes. One of our supervisors works with phage so she would be able to guide the wet lab team. We opted to silence the toxins rather than kill the bacteria because we wanted a way of preventing glucan formation without disturbing the balance of the oral microbiome

mRNA interference of Clostridium difficile

Clostridium difficile is an anaerobic bacterium capable of forming spores (meaning it persists in the environment). Clostridium difficil einfection(CDI) is a major infection which causes hospital and community acquired-diarrhoea. It particularly affects those who have long-term hospital stays (especially the elderly), are immunocompromised/immunodeficient (for example due to chemotherapy) and/or are on broad-spectrum antibiotics. CDI has a heavy financial burden-according to Zhang et al. (2017), the annual costs due to C. difficile infections in the US alone are an estimated $6.3 billion with almost 2.4 million days spent in hospitals. According to the Centers for Disease Control and Prevention, between 1999 and 2007 there was an increase in the estimated number of deaths due to CDI from 3,000 to 14,000 which was seen across Europe and Canada as well (McDonaldet al., 2012; Lessa et al., 2012). This has been linked to a hypervirulent, resistance strain of C. difficile.

Within the SBRC, our supervisors work with a wide range of Clostridial species in the Clostridia Research Group. Unlike with S. mutans, there was more experience within the team and so the supervisors would be able to work with us more closely and advise us on this project.

It was hard to decide between working with Ideonella sakaiensis, Streptococcus mutansor and C. difficile. But in the end, the team voted and chose mRNA interference of C. difficile which became our project; Clostridium dTox.

Project design

C. difficile & phage characterisation

C. difficile strain SBRC 078 was isolated previously in the SBRC from clinical faecal samples and belongs to the hypervirulent PCR ribotype 078. The strain contains the genes tcdA and tcdB encoding for both toxins. Phage phiSBRC was previously isolated in the SBRC from an environmental sample and can infect and form plaques on C. difficile SBRC 078. A lysogenic version of C. difficile SBRC 078, which contains phage phiSBRC integrated into the bacterial chromosome, was created previously in the SBRC.

C. difficile growth analysis

The growth profile of the wildtype version of C. difficile SBRC 078 was compared to the growth profile of the lysogenic version of this strain. To assess this the growth of both strains was monitored for 24 hours and the OD at 600 nm was measured and the maximum growth rate was calculated using the equation

where t1 is the OD at the start of exponential phase and t2 is the OD at the end of exponential phase. This data was used to inform the model parameters and was required to ensure that in the human gut the lysogenic bacterial strains created in this project would grow in the same manner as the wild-type cells and therefore would outcompete them.

Phage burst size

Phage burst size was assessed to determine the number of infectious phage particles produced per bacterial cell during one infection cycle. This was determined by measuring the number of infectious phage particles (in plaque forming units per ml) produced over a time-course after infection of C. difficile SBRC 078 with phiSBRC. The titre of free phage at each time point was determined by enumeration of plaques using a plaque assay. C. difficile SBRC 078 was infected to a multiplicity of infection (MOI) of 1. The burst size was calculated as the Final Phage Titre/(Infection Phage Titre – Titre of Unbound Phages). This data was used to inform the model parameters.

Promoter library

Our project aims to supress toxin production in C. difficile and we chose two different strategies to pursue this aim. Briefly, these strategies involve either dead-Cas9 (dCas9) or antisense RNA (asRNA) to inhibit toxin production. Both strategies will require a careful consideration of the genetic parts involved in the device. Of particular importance is the choice of promoter employed to control expression of the dCas9, guide RNA or antisense RNA. Tailoring expression to an appropriate level is often an important design consideration in genetic engineering. In our case it was thought that the use of a strong promoter would be of greatest benefit for both strategies we pursued in order to maximise either the amount of guide RNA and dCas9 or the amount asRNA. Concentration of these components within the cell was expected to correlate with the degree of toxin suppression and since the objective was to supress toxin to the greatest extent possible, we aimed to find and characterise strong promoters within C. difficile.

The promoters we chose to characterise were as follows:

BBa_J23106 (from iGEM registry)

BBa_J23114 (from iGEM registry)

BBa_J23119 (from iGEM registry)

PCsp_fdx (from Clostridium sporogenes)

PCac_thl (from Clostridium acetobutylicum)

PCdi_TcdA (from C. difficile)

PCdi¬_TcdB (from C. difficile)

All seven promoters were intended to be assessed in both E. coli and C. difficile. PCsp_fdx and PCac_thl were chosen since they have been used extensively in studies on C. difficile as well as related organisms and both are considered to be strong promoters (Heap, 2018, Heap et al., 2009). A comparison of the two suspected strong promoters was made with the native promoters controlling toxin expression in C. difficile PCdi_TcdA and PCdi¬_TcdB. It was thought to be interesting and potentially useful to discover the strength of the toxin promoters and potentially their variance in their expression in different conditions. Three existing iGEM registry promoters were also chosen to be assessed in C. difficile. This served two functions, firstly it improved the registry in terms of part characterisation as there is currently no data on their use in Gram-positive organisms. Secondly, since these promoters have been well documented in E. coli they could give a good indication of the strength of the Clostridial promoters when used in E. coli.

Occasionally cloning dCas9 in E. coli can be problematic, potentially due to unwanted off-target effects of the protein, the DNA binding nature of the enzyme or due to the size of the gene itself. To facilitate cloning and yet maximise dCas9 activity in C. difficile the ideal promoter would have low expression in E. coli and yet high expression in C. difficile. The choice of promoters and decision to assay them in both E. coli and C. difficile was designed to help us choose the optimal promoters for the toxin suppression projects, characterise existing iGEM registry parts in novel contexts and add potentially valuable Clostridial/Gram-positive promoters to the registry. Two different assays were chosen to assess the promoters described above. In E. coli, a GFP assay was chosen due to its widespread use, ease, cost, precision and reliability. However, GFP assays have not been successfully used in Clostridia, due to the requirement of oxygen for GFP, and as such other reporter assays are commonly used. One such reporter assay is the GusA assay in which the expression of the reporter gene gusA can be accurately measured via the eventual release of a fluorescent compound 4-methylumberlliferone (4-MU). The assay relies on the fact that the protein encoded by gusA is a glucuronidase which converts the non-fluorescent 4-methylumberlliferyl glucuronide (4-MUG) into the fluorescent (4-MU).

GusA assays can be performed in E. coli as well as Clostridia and so both GFP and GusA assays were used in E. coli. Our GFP assay was inspired by our InterLab experience as we thought it would be useful to use the protocols and calibration curves we obtained from the study to standardise our data. This would help us give context to the strength of the promoters by comparing them to the InterLab positive and negative controls, using the calibrations curves generated through our InterLab study to ensure that the results would be reproducible by any other laboratory using different equipment.

Heap, J. T. (2018). Stringency of Synthetic Promoter Sequences in Clostridium Revealed and Circumvented by Tuning Promoter Library Mutation Rates. https://doi.org/10.1021/acssynbio.7b00398

Heap, J. T., Pennington, O. J., Cartman, S. T., & Minton, N. P. (2009). A modular system for Clostridium shuttle plasmids. Journal of Microbiological Methods, 78(1), 79–85. https://doi.org/10.1016/j.mimet.2009.05.004

CRISPRi

The origin of the CRISPR/Cas9 based genome editing tool

The CRISPR/Cas system is a naturally occurring defence mechanism in bacteria. It confers adaptive immunity against mobile genetic elements (MGEs), like phages. CRISPR-Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)- are short segments of repetitive DNA that are identical to each other. Due to the palindromic nature of the sequence, RNA transcribed from these repeats forms hairpin turns. These small CRISPR RNAs (crRNAs) are interspaced with unique DNA sequences known as spacer DNA. These DNA regions encode sequences complementary to MGEs introduced to the cell previously by phages. Upstream of the CRISPR array one can usually find the cas genes. These genes encode the Cas proteins that play an essential role during the different stages of bacterial immunity against intruding DNA. When bacteria are infected by a virus or phage, the CRISPR array is transcribed and the product, crRNA, forms a complex with (multiple) Cas protein(s), called the Cas-crRNA ribonucleoprotein complex. Depending on the organism different proteins are involved in this process. The crRNA guides the Cas protein(s) to the invading DNA which leads to degradation of dangerous virus or phage DNA before the infection has started.

The CRISPR ‘revolution’

To date the CRISPR system has been adapted to form a new revolutionary gene editing tool, mostly known under the name CRISPR/Cas9. This technique relies on two main components: the short guide RNA (sgRNA) and the Cas9 protein. The sgRNAs can be split into two parts: the “seed” region, which is a 20 bp sequence that can be easily modified to be complementary to target a gene of interest, and a “handle” region which facilitates the binding of Cas9 to the sgRNA. The sgRNA functions as a ‘guide’ to ensure Cas9 creates double-strand breakage at a specific place in the DNA. In eukaryotes these double-strand breaks are glued back together via non-homologous end-joining (NHEJ) During NHEJ, a few nucleotides are removed from the site of cleavage which leads to the disruption of the gene. This disruption might affect or even completely abolish the activity of the protein it encodes which could lead to phenotypic changes. On the other hand, most bacteria cannot perform NHEJ and therefore die when Cas9 cuts the DNA as their genome cannot be replicated. However, if the target region has been successfully modified, the sgRNA will no longer guide the Cas9 protein to this region of the genome thus preventing cleavage. This makes the CRISPR/Cas9 system an ideal selection tool when editing the genome of bacteria as wild type cells will be killed whereas mutants will survive.

Diagram explaining transcription and translation in bacteria.

Diagram explaining CRISPRi.

Initially, the CRISPR/Cas9 tool was used to generate knock-out and knock-in genes in various bacterial species. It was quickly realised that this system could easily be altered and so can be used for a much wider range of applications. Genetic modifications of the Cas proteins has allowed scientists to use this system to purify specific DNA regions, image DNA in living cells, introduce specific mutations in the DNA code, and activate or repress target genes, the latter being the most interesting in light of this project (Clostridium dTOX). In order to use the CRIPSR/Cas9 tool for the repression of protein expression, the RuvC and HNH nuclease domains were inactivated in the Cas9 protein resulting in a catalytically inactive protein, nuclease dead-Cas9 (dCas9). Instead of creating double-strand breaks, dCas9 (temporary) binds to a sgRNA complementary DNA sequence thus preventing the mRNA polymerase from binding/continuing transcription and in doing so preventing transcription. This process of reducing protein expression levels with dCas9 is called CRISPRi.

Why using CRISPRi for Clostridium dTOX

Ultimately the success of Clostridium dTOX depends on how efficiently the C. difficile toxins are repressed thereby turning C. difficile from a pathogenic (toxic) strain into a non-toxigenic strain. Rather than killing the C. difficile population in the gut completely, the creation of competition between non-toxic and toxic >em>C. difficile can stop the infection as well as preserve natural gut microbiota to prevent other opportunistic bacteria from populating the gut. CRISPRi has been shown to effectively decrease the production of proteins in cells and can be easily engineered to target different DNA regions making it a good candidate to target the promoter region of TcdA (PtcdA) and the promoter region of TcdB (PtcdB) with the aim of reducing toxin production. Moreover, multiple sgRNAs can be expressed and directed to each promoter thereby improving the tightness of the system.

Initial planning/preparation

C. difficile infection (CDI) symptoms range from mild diarrhoea to life-threatening pseudo-membranous colitis and toxic megacolon. They are essentially caused by the production of two toxins: TcdA and TcdB. The two toxin genes are part of the PaLoc gene cluster encoding three accessory proteins: TcdR, TcdC and TcdE. TcdR is a sigma factor essential for the initiating the translation of tcdA and tcdB. Besides controlling toxin expression, TcdR is also involved in the regulation of many other genes in C. difficile. Repressing the expression of TcdR might have an effect on the general cell fitness, therefore it was decided to target the promoter regions upstream of tcdA and tcdB instead thus minimizing the effect on other cell processes. Moreover, it has been shown that Cas9 binding to the promoter to inhibit the initiation of translation results in stronger repression of expression when compared to prematurely ending translation by targeting Cas9 to the gene sequence. Therefore, in this project, the sgRNAs are designed so that they direct Cas9 to the promoter region to inhibit the initiation of translation.

Potential dCas9 targets within the PtcdA and PtcB promoter sequence were selected using the Benchling platform and converted into sgRNA sequences. For PtcdA, six sgRNAs were designed and, for PtcdB, five sgRNAs were selected. In order to treat CDI, both toxins should be repressed simultaneously because each toxin individually can cause CDI symptoms. Therefore it was decided to first test the sgRNAs for toxin A and toxin B separately in E. coli after which the best repressing sgRNAs for each toxin would be taken forward and combined in C. difficile. In C. difficile only the expression of sgRNAs and dCas9 is required as all other components necessary for the system to work are already present. The overall toxin repression can then be measured with a cytotoxicity assay.

Vector design details

After thorough consideration, plasmid pMTL84121 was chosen to express our future construct as it contained all the necessary parts for successful cloning: - pCD6 Gram-positive replicon – for stable maintenance of the vector in C. difficile - p15a+tra Gram-negative replicon – for stable maintenance of the vector in E.coli - catP gene – gives chloramphenicol resistance to bacteria and is used as a selection marker - Multiple cloning sites (MCS) - provides cloning sites for inserts To test our experiment in E. coli and C. difficile, we constructed the following vector: pMTL84121-PthI-dCas9-Pfdx-tcdR-T10-PtcdA/PtcdB-gusA-T14 + gRNA. What Why pthI Promoter for dCas9. dCas9 Cas protein that binds to DNA and blocks transcription due to steric hindrance Pfdx Promoter for tcdR. Our team results showed this promoter had the strongest activity in C.difficile than any other chosen promoter aimed at E. coli (link to asRNA) tcdR An important regulatory gene in C.difficile, which affects synthesis of toxins A and B as well as other important chemical reactions T10 Terminator for tcdR. It produces termination of tcdR transcription PtcdA/PtcdB Promoters for toxin A and B respectively GusA Reporter gene T14 Terminator for reporter gene gusA. It produces termination of gusA transcription sgRNA Constructed sgRNA complementary to specific region of PtcdA/PtcdB Experimental plan: After carrying our initial planning of the components for our experiment, we came up with a following experimental proposal: 1. Cloning of PthI-dCas9 insert into pMTL84121: pMTLdCas9 2. Cloning of ptcdA/ptcdB-GusA-TB0014 into pMTL84121-PthI-dCas9: pMTSdCas9-GusA 3. Cloning of pfdx-tcdR-TB0010 into pMTL84121-PthI-dCas9-ptcdA/ptcdB-GusA-TB0014 4. Cloning of sgRNA (A1-A7, B1-B5) into pMTL84121-PthI-dCas9-Pfdx-tcdR-T10-PtcdA/PtcdB-gusA-T14: pMTLdCas9-GusA-sgRNA 5. Transformation of full vector pMTL84121-PthI-dCas9-Pfdx-tcdR-T10-PtcdA/PtcdB-gusA-T14 + gRNA into E.coli 6. Checking the repression of GusA by targeting ptcdA/ptcdB with respective sgRNA via GusA assay. 7. Transform potentially suppressing sgRNAs into C. difficile. 8. Test sgRNA with signs of repression in C. difficile via cytotoxicity assay.

Antisense RNA

Antisense RNA (asRNA) is another means of reducing gene function. In this strategy a piece of RNA is transcribed which is complementary to the coding strand of a target gene in the reverse orientation, in other words it is the antisense to it. This has the effect of sequestering the coding mRNA in an RNA-RNA duplex which is unable to effectively bind the ribosome meaning protein synthesis of that gene is inhibited. The RNA-RNA duplex molecule may also be targeted by RNases specific to double stranded RNAs meaning the target mRNA is degraded faster due to the presence of asRNA.

Many studies have utilised an asRNA approach to genetic studies. Typically asRNA will not be capable of completely eliminating gene function since target mRNAs may not encounter the asRNA molecule within the cell before being translated. For this reason, asRNA has primarily been used when gene ‘knockdowns’ are desired rather than total gene knockouts. In knockdown strains the expression of the target gene is reduced rather than entirely eliminated. Non-model organisms which are less genetically accessible frequently do not have established methods for creating gene knockouts. In this case asRNA is often used to gain initial insights into gene function since all that is required for asRNA studies is knowledge of the target gene sequence, a plasmid capable of replication in the organism and a means of transformation.

Diagram explaining transcription and translation in bacteria.

Diagram explaining antisense RNA.

Our project aims to create a phage which will suppress toxin production in C. difficile once integrated into its genome. To do this we will genetically modify a phage known to infect strains of C. difficile named phiSBRC. The phage will be modified to include constructs to suppress toxin production either via a CRISPRi (dead-Cas9) approach or via asRNA. There were several design considerations when approaching this problem. Firstly the construct should be capable of significantly suppressing toxin production, it was not known whether dead-Cas9 or asRNA would be superior in this respect. Secondly, the eventual genetic construct we choose should be sufficiently small that we can alter the phage genome without adversely affecting its normal function. One potential limitation was thought to be the total amount of DNA which the phage could package into its head. In this respect asRNA could have a significant advantage over a dead-Cas9 approach since the total size of the genetic construct can be much smaller. Antisense RNA constructs can simply consist of a promoter and a short asRNA region of less than 100bp while the cas9 gene alone is more than 4kb long. However, since it was not known which approach would produce more effective toxin suppression both approaches were pursued.

C. difficile has two well characterised toxins which cause epithelial cells to undergo apoptosis these are TcdA and TcdB. It was thought that each construct we create should aim to supress both of these toxins simultaneously since research has concluded that each can operate independently from the other (Kuehne et al., 2010). The general form of our constructs therefore is to have asRNA parts downstream of promoters with a transcriptional terminator between these promoter-asRNA pairs. It was thought to use two different promoters to as to avoid unwanted recombination events within our constructs. As discussed previously, the optimal promoter for expressing the asRNA parts was thought to be the strongest promoter possible. We were therefore looking for the two strongest promoters in C. difficile we could find. Our results from the promoter assays we performed indicate that PCsp_fdx¬ is the strongest promoter assayed in C. difficile. Unfortunately we were unable to clone the GusA reporter plasmid for PCac_thl which was the other Clostridial promoter expected to give strong expression. However, the PCac_thl GFP reporter plasmid was created and assayed in E. coli where it gave the highest expression of any promoter tested including the positive control used in the iGEM InterLab studies. This result was consistent in our laboratory as well as that of our two collaborating teams representing Imperial College London and University of Warwick. The Clostridial promoter P¬Cac_thl has been routinely used in Clostridial research and outperforms every other promoter in E. coli, therefore it can be hypothesised that P¬Cac_thl would exhibit the strongest levels of expression in the GusA promoter assay in C. difficile, had this plasmid been constructed. As such PCac_thl and PCsp_fdx were chosen as the two strong promoters from which to express our asRNA parts.

When choosing the length of the antisense RNA we consulted the scientific literature. There is some contradicting advice on this topic with E. coli asRNA parts seeming to be significantly shorter than those used in the few asRNA studies we found performed in clostridia. There are important design considerations here since there is a compromise to be made. Longer asRNA parts seem to generally give a greater degree of suppression but are also more likely to give unwanted off-target effects. This is probably because they can bind the target mRNA more tightly but are also more likely to have regions of short similarity with other non-target mRNAs within the cell. With this in mind we chose to try two different lengths of asRNA binding to the coding region of the target gene as well as the entire region upstream of the gene expected to include the ribosome binding site. ‘Construct One’ has a binding region of 24bp, this is the length suggested by a recent review paper on this topic (Hoynes-O’Connor & Moon, 2016). ‘Construct Two’ has a binding region of 50bp, this is much longer though still significantly shorter than the hundreds of base pairs previously used in clostridial studies (Desai & Papoutsakis, 1999; Fagan & Fairweather, 2011). Both of these constructs target both of the toxin genes we are interested in.

Construct One diagram

Construct Two diagram

Desai, R. P., & Papoutsakis, E. T. (1999). Antisense RNA strategies for metabolic engineering of Clostridium acetobutylicum. Applied and Environmental Microbiology, 65(3), 936–945.

Fagan, R. P., & Fairweather, N. F. (2011). Clostridium difficile has two parallel and essential sec secretion systems. Journal of Biological Chemistry, 286(31), 27483–27493. https://doi.org/10.1074/jbc.M111.263889

Hoynes-O’Connor, A., & Moon, T. S. (2016). Development of Design Rules for Reliable Antisense RNA Behavior in E. coli. ACS Synthetic Biology, 5(12), 1441–1454. https://doi.org/10.1021/acssynbio.6b00036

Kuehne, S. A., Cartman, S. T., Heap, J. T., Kelly, M. L., Cockayne, A., & Minton, N. P. (2010). The role of toxin A and toxin B in Clostridium difficile infection. Nature, 467(7316), 711–713. https://doi.org/10.1038/nature09397