Overall Project

Our project is about trying to reduce the waste going to landfill by engineering a bacteria to degrade polystyrene. Some bacteria have been found to form biofilms on polystyrene, indicating that polystyrene may be being degraded. We have several different parts to our project, including a citizen science experiment (CSE).

It is not clear how the bacteria actually degrade the polystyrene, as no pathway for polystyrene degradation has been found so far. It is possible that the reason why degradation appears to be so slow is because no degradation pathway has evolved specifically for polystyrene yet, so some enzymes in other pathways may be doing any degradation (though as they are not adapted for polystyrene, the rate will be very slow). There are a couple of pathways we have been looking into that could cause this: the styrene degradation pathway (as styrene is the monomer of polystyrene, so has similar structure to one repeating unit), and the toluene degradation pathway (as this would break open the bulky, unreactive phenyl side chain, making it easier for aliphatic degrading enzymes to degrade the main chain). We've done some modeling of the first enzyme in the toluene degrading pathway, and future Leicester iGEM teams may be able to artificially mutate the first enzyme to see whether our potential modification could work (see the modeling page).

We intend to extract the genes involved in any pathway degrading Expanded Polystyrene (EPS), and/or develop a new pathway involving modifications to existing enzymes able to degrade aromatic and aliphatic hydrocarbons to fit polystyrene and its derivatives into the active sites. The bacteria strain that we insert these new genes into should then be able to degrade polystyrene at a higher rate than natural bacteria, and future iGEM projects at the University of Leicester could harness this bacterial strain to produce useful products.

Project Details

Making Polystyrene selection media

The first stage involved making a selection media to isolate bacterial colonies that could survive with polystyrene as effectively the only carbon source available, making it difficult for bacteria unable to degrade polystyrene to survive. However, salts containing the other minerals needed for survival must be present. We found a minimal salts media recipe (Atiq et al.,2010) that avoided other carbon sources while still making other essential elements available. We spent several weeks attempting to dissolve polystyrene into the agar, using various solvents such as acetone (which just took the air out of expanded polystyrene, rather than dissolving the polymer, and dissolved petri dishes), toluene (which dissolved polystyrene 'sugar' (pre-expanded polystyrene beads donated to us by Styropack ltd.)but also dissolved the plates) and various alcohols (which didn't dissolve polystyrene). In the end, we decided not to use a solvent, and instead sprinkled the polystyrene 'sugar' on top of the minimal media once poured, so that the 'sugar' was available to bacteria spread on the surface.

The Protocol

Polystyrene selection media (or polystyrene minimal media) is made first by making up a minimal broth using the recipe from (Atiq et al.,2010) by dissolving 1g K₂HPO₄, 0.2g KH₂PO₄, 1g NaCl, 0.002g CaCl₂.2H₂O, 0.005g Boric Acid, 1g (NH₄)2SO₄, 0.5g MgSO₄.7H₂O, 0.001g CuSO₄.5H₂O, 0.001g ZnSO₄.7H₂O, 0.001 g MnSO₄.H₂O and 0.01g FeSO₄.7H₂O, per litre H₂O. Melt 2% electrophoresis grade agarose to the broth to solidify it after cooling using a microwave, and heat until the agarose is completely melted/dissolved. Allow media to cool by keeping it at 60 degrees C until needed. To make a selection plate, use 95% media and 5% polystyrene 'sugar' (polystyrene beads before they've been expanded in the manufacturing process). We suggest 10 grams as a total mass of minimal media and solution. You should first pour out the media, and before solidifying, sprinkle the polystyrene on top, to prevent the 'sugar' from sinking to the bottom thus allowing bacteria colonies to access the polystyrene as a carbon source. The agarose is the only other carbon source in the plate (except obviously the plate itself), so any colonies that grow should be utilising polystyrene as a carbon source, or at least have evolved pathways to deal with the toxic effects that a long chain hydrocarbon could cause bacteria (such as interfering with the cell membrane, due to the hydrophobic nature of parts of both).

The Citizen Science Experiment

The CSE was used to get the public involved in trying to find bacteria that could live on, and degrade expanded polystyrene (EPS). We asked them to bury a strip of EPS in the ground and leave it for a length of time (around 2 months), to find out whether bacteria might establish colonies, thus indicating that a colony could at the very least bind to, and possibly degrade EPS. The kits were Risk Assessed, and a copy of the assessment was included in the kit, along with instructions, a strip of EPS, and a self seal bag to put the EPS into. This was all contained in a stamped, addressed envelope at a cost to the public of £2 (+50p postage if bought online via our blog to cover the cost of sending the kit to the person). We recently had our first batch of kits back, ready to analyse.

The hope is that we'll find a bacteria that has been using polystyrene as a carbon source, so we can extract the genes responsible and attach them to high expression promoters to increase the amount of protein each bacterium produces. We will also be trying mutagenesis on these bacteria to increase the rate of reaction of the enzymes involved in the pathway.

The Protocol

The first part involves halving the expanded polystyrene (EPS) lengthways with sterilized scissors in our returned citizen science experiment (CSE) kits so that we'll still have some of each sample left should we need it. First we need to do a practice with unused EPS strips (control and test). Pour/pipette 20ml LB agar into a petri dish and allow to set. Put line down middle in permanent marker USING A RULER! Then label one side swab (for the swabbing/scraping material) and the other side strip for the EPS strip (can use other names if you prefer). Add 50 micro litres of PBS (phosphate buffered saline) to each side of the half strip of EPS and scrape with spreader, before spreading onto the left half of the agar (or side labeled as 'swab'). Try to swab any mud off the strip if possible. Then place the half strip in the centre of the other half labeled 'strip'. Leave for a week at room temperature on the lab bench and examine agar for colonies. The other half of the strip should be put back into cold storage for future use.

After letting the organisms grow for a few weeks, select some vigourously growing colonies and spread them out onto a polystyrene minimal media plate divided into 6 sections, where each selected colony is allocated its own segment. Leave at room temperature or at 37^oC, checking each day. The bacteria that grow are likely to only grow very slowly, so it could be a month or more until you see a visible colony, due to the low polystyrene degrading rates of bacterial enzymes. If a colony grows, extract some, add the sample to luria broth, and grow overnight on a shaker at room temperature or 37 ^oC. Boilate the colony by boiling the bacteria in distilled water for 10 minutes, before centrifuging at 13,000 RPM for 5 minutes to remove cell debris- the supernatent should contain DNA, which you can then add to a PCR mix and amplify overnight

DNA extraction

DNA extraction is of course vital when trying to make biobricks and to extract genes of interest, and we've had varying success with several methods. These have included QUIGEN's Blood & Cell Culture DNA Maxi kit, CTAB and Maxwell DNA extraction.

QUIGEN Blood & Cell Culture DNA Maxi kit

The protocol was as in instruction booklet supplied. Unfortunately, this method didn't really work, despite 4-5 retries with E. coli as a test.

CTAB

This protocol was given to us by Miranda Johnson of the level 2 lab on our floor to use after the QUIGEN kit didn't work. It is designed for Staphylococcal chromosomal DNA extraction, but we had some success with the extraction regardless of the species of bacteria used:

Using an overnight culture, spin down 1.5ml of culture in an 1.5ml Eppendorf tube for 10 minutes at 13,000 RPM. remove the supernatant, then respin down so as to be able to remove any more supernatant. Resuspend the cells in 250µl P1 buffer (QUIGEN). Add 15µl 5M NaCl to the Eppendorf tube before adding 2µl 100mg/ml lysozyme and mix. Incubate the tube in a 37^oC waterbath untill the solution become clear. This is when the cells have lysed. Add 2µl 25mg/ml proteinase K (QUIGEN) and the same amount of RNase. Add 27µl 10% SDS, invert the tube and incubate at 37^oC for 20-30 minutes. If you are using a class 2 pathogen, but have your main work area in a class 1 lab, then after decontaminating the outside of the Eppendorf tube, you can now bring it into a level 1 lab. Add 97µl 5M NaCl and mix well. Wearing goggles, add 81µl CTAB (caution: corrosive, Toxic, Irritant so wear eye protection). Incubate at 65^oC for 20 minutes. Add an equal volume (500µl) of 24:1 chloroform:isoamylalcohol mix (suspected carcinogen, toxic, irritant, so only use in a fume cupboard). Mix thoroughly but gently. Spin the tube at 13,000 RPM for 10 minutes, and transfer the upper layer formed to another tube, avoiding any precipitate. Add an equal amount of isopropanol to this upper layer and mix thoroughly but gently. Spin for 10 minutes at 13,000 RPM. Remove all alcohol avoiding the pellet (use a vac line if possible), but don't allow to completely dry or else you may have difficulty resuspending the pellet. Resuspend the pellet in 100µl of TE buffer (more if the pellet is large).

Maxwell prep

To prepare your bacteria for this DNA extraction, you follow the CTAB protocol untill the first incubation step (when the solution goes clear after cell lysis). Because, as students, we're not actually allowed to use the machine, so our lysed cells were put into the machine by Dr Badge. This got us ourt most reliable and best extraction.

Sau3A1 partial digest

A Sau3A1 partial digest was originally intended, after DNA extraction, to be a step towards making a DNA library, which we could then have grown on our polystyrene minimal media to find genome fragments that could degrade polystyrene. However, we ran out of time, and ended up attempting to PCR out Tod operon genes that we thought could be partially responsible for polystyrene degradation. We set up DNA digestion assays to find out the time taken to get fragments of a particular length (from 1Kbp to 10 Kbp). We started doing the digests at 37^oC, but we realised that this caused Sau3A1 to digest DNA too quickly, even at very low concentrations.

The protocol

First off, a reaction mixture was prepared using: 4µg of DNA (in our case, we needed 53.7µl of DNA in buffer), 7.5µl 10x NE buffer 1, 0.75µl 100x BSA, 0.5µl Sau3A1 (to be added last), and topped up to a final total of 75µl with distilled H₂O. 1µl Sau3A1 is 4 units of enzyme (1 unit can digest 1µg λ DNA in 1 hour @ 37^oC in 50µl total reaction volume). The reaction we found, was easier to control and get better gel photos at room temperature. We made 8 stopping aliquots to stop samples of the reaction at time=0, 5, 10, 15, 20, 25, 30 and one taken before enzyme was added. Each stopping aliquot was made up of 4µl Loading dye, 2µl 0.5M EDTA and 6 µl H₂O put in iced water. At each time, 8µl of reaction mixture was removed from the reaction mixture and added to a stop aliquot (EDTA binds to and removes Mg²⁺ ions from Sau3A1, inhibiting the enzyme, and the iced water stops the enzyme from working). These were then run on a 0.7% agarose gel and transilluminated to reveal a clear curve in DNA fragment length as time increased:

Primers and PCR

When we realised that it would be quicker to try to extract genes via PCR rather than via a DNA library, we BLAST searched for genes present in the Tod operon, and found the whole operon was remarkably conserved, with only a couple of the genes needing any kind of degeneracy in their primers. We concentrated on genes within the operon that didn't have any restriction sites incompatible with the basic Biobrick standard:

Unfortunately, with the 2 strains of P. putida, none of the primers amplified the target genes, meaning that the strains do not encode any of the Tod operon genes. Because of this, it meant we couldn't make a biobrick.

The next step

Results