Team:Sydney Australia/Experiments
Experiments
Key Experiments
Aim
To confirm the production and activity of the PSIK, PSID and PSIM in individual constructs prior to combination in single inducible vector.
Aim
To confirm the production and activity of the PSIK, PSID and PSIM in individual constructs prior to combination in single inducible vector.
Aim
This part of the project aimed to express and characterise the activity of the PsiH, a P450 mono-oxygenase enzyme that converts tryptamine to 4 hydroxytryptamine in vivo before putting it into our single-cell system.
Aim
To transform our two plasmids into the same cell so the entire pathway from the mushroom can be recreated functionally in E.coli.
Aim
Aim
The aim of this experiment was to determine which type of codon harmonisation/optimisation (rank-order method, absolute frequency method, relative frequency method and IDT codon optimisation) would improve the expression of the VVD protein compared to the wild-type codon usage. The results of this experiment would then be used to codon harmonise our psi genes to improve their expression.
Aim
To increase the fluorescence of the IDT-codon optimised VVD to further increase its utility as an iLOV-based blue fluorescent protein.
Protocols
Innoculating or Transferring Cultures
- Wipe down the bench with ethanol or F10FC before starting.
- Work close to the Bunsen burner.
- Use the hot ‘blue flame’ to get a strong updraft, flame the necks of bottles, and to flame your loop (if using a metal loop, do not flame plastic loops, you will have a bad time).
- The safety flame (yellow) is only good for seeing when you have lit your bunsen burner and for lighting the ethanol on your glass spreader, if using. Again, do not light plastic spreaders on fire - you will have a bad time.
- Keep the work area clear of clutter, ensure everything you need is within easy reach such that no part of your body will need to cross the flame. You do not want to set on fire either.
- Don’t do culture work on top of your lab book or other paper notes. These should be kept in green spaces. Paper near aseptic work increases the risk of fire, and your beautiful notes going into the autoclave.
- Flame the neck of glass bottles and tubes when opening and closing them
- Pass through flame back and forth briefly three or four times over three seconds.
- Be mindful of where you put things, act as though the bench is lava.
- i.e. once you remove a sterile plastic loop from the packet, or flame your metal loop, do not put it back down on the counter or it will have to be re-flamed or disposed of.
- Do not contaminate your workspace by putting loops with culture on them
- If your lab coat sleeves hang from your wrists (i.e. are not elasticated so they fit snugly on your arm), do not allow them to flap around. Tuck them into your gloves if possible, tape them or elastic band them to fit snugly, or roll them up if not working with corrosive or toxic reagents.
- Fun fact: loose sleeves + bacteria + flame = contamination or fire.
Sterile Bottles, Solutions, and Gear
- Label EVERYTHING carefully! Write chemicals used, plasmid names, media type, date, and your name.
- Make sure any hazardous reagents are clearly labelled with the chemical name, the concentration, and the nature of the hazard (e.g. flammable/corrosive/toxic). Store all hazardous reagents in the appropriate area. DO NOT leave unlabelled hazardous chemicals on the bench. You never know if any of your labmates are feeling thirsty and decide to ignore the ‘no drinking rule’. Do you want their dumb deaths on your hands?
- When you take lids off, do not put them down on the bench unless necessary. If you need to put the lid down (weakling), put them upside down, so the screw thread is facing upwards. Surfaces are a huge source of bacterial contamination.
- Don’t leave lids of bottles for any longer than necessary. Turns out the air is lava too.
- Getting sterile Eppendorf tubes out of the autoclave beaker:
- OPTION 1: take off foil top, carefully pluck out the tube by its base or edge, place carefully in a tube rack, repeat if necessary, replace foil.
- OPTION 2: take off foil top, shake out tubes into the foil, carefully place in the tube rack - being careful not to touch the inside of the tube, replace foil.
- DO NOT : rip off the foil, plunge your hand into the beaker, rummage around like it’s a bargain bin, emerge triumphantly with a tube, then leave the foil in disarray until some poor sad soul realises they need to send the whole batch down for autoclaving again because of your inconsiderate behaviour.
Pipetting with Gibsons
- Wipe down your pipettes with ethanol every. single. day. If you do not do this, I really don’t think you can call yourself a Molecular biologist! (luckily for me I identify as a microbiologist so I’m safe)
- Also do this when changing from microbial to DNA work, or if you’re concerned you (or someone you know) have contaminated your pipette somehow.
- If you’re having persistent issues with contamination, do not just try to dump your pipette on someone else’s bench and take theirs. There is another way!
- Dissemble the pipette according to the manufacturer’s instructions.
- Soak in bleach (0.5% hypochlorite = a 1/10 dilution of household bleach, freshly prepared), so that all parts of the pipette are covered.
- Leave to soak for 30min. Do not leave for longer or you are at risk of the pipette parts rusting.
- Rinse extensively with water, then with 80% ethanol.
- Allow to dry.
- Remember that just the tip is sterile! No matter how recently you have swabbed your pipette with alcohol, the main body of the pipette is still contaminated in some way (DNA, microbes, enzymes, they’re everywhere! Remember - the air is lava).
- The body of the pipette should not come into contact with anything that needs to remain sterile (e.g. the inner walls of bottles or falcon tubes).
- Hold the pipette vertical! Or as close to it as you can get. If you hold it sideways the liquids may run up the barrel of the pipette, damaging the inner workings, or worse - contaminating your culture or PCR with whatever gunk got up there before!
- As long as you are careful with your pipetting, most work will not require plugged pipette tips.
- They are only really required for RNA work and 16S RNA PCRs as these procedures are incredibly sensitive to DNA/RNA contamination.
- Be careful when getting tips out of the box, we don’t want any tip casualties. Open the box the whole way, put the tip on firmly, move the pipette away, then close the box.
- Don’t hit the tip on the lid of the box, don’t leave the box open long after you’ve got the tip out, and don’t go around knocking other tips on your way out like some sort of pipetting Godzilla!
- All the pipette tip wants is to be part of an experiment, don’t let them die before they get their time in the spotlight.
- Preparation prevents poor performance! Swab down your bench and pipettes with ethanol, label your tubes, and most importantly - read the protocol carefully!
- Keep all enzymes in the freezer or on ice at all times. No exceptions! (except maybe like three exceptions) Restriction enzymes, ligase, polymerases, reverse transcriptase, phosphotase, kinase, Gibson assembly mix, etc.
- Keep all DNAs (plasmids, PCR products, genomic DNAs) either in the fridge or freezer when they are not actively being used. DNA can sit on the bench for a little while (few minutes up to an hour), but don’t abandon it in the harsh outdoors.
- Double check all buffer and incubation requirements. It may help to write things such as times and temperatures on a separate piece of paper as you read through the protocol. That way you can make a list and check it twice!
- Make up the bufer first upto to 1x concentration or what is needed for your experiment, THEN add your enzymes and/or DNA. Least expensive to most expensive as I say *wink*.
- Handle your tubes carefully.
- Don’t touch any part of the inside of your tubes, including the inside part of the lids with your gloves or gross human fingers. Eww.
- Be careful when opening the tubes especially as your thumb can easily slip onto the inside of the lid without realising! Try using your thumbnail, not your whole thumb, to carefully flick open the lid.
- Be extra careful when putting the tubes on ice - the ice must not get inside your tubes! Don’t push the tubes too far down into the ice, handle them carefully. It can be helpful to use a ‘floaty’ (thin foam tube holder) to keep your tubes from going too far into the ice.
Setting up PCRs and Other Enzyme Reactions
When doing molecular biology experiments using bacteria, the ability to properly make antibiotic stocks is life and death. Too weak, and you risk mutating untransformed cells into antibiotic-resistant mutants that could accidentally be unleashed on the world and kill us all… too strong… and you ruin your experiment by killing everything and have a really bad day.
To make your time and effort worthwhile, try to make 50 or 100mL of stock at a time to be frozen in 1mL aliquots. These steps will reference quantities for a 50mL batch, but this can be scaled up or down depending on your available antibiotic-stock-making equipment and your antibiotic needs.
Typical Antibiotic Concentrations: Stock & Usage
- Ampicillin: 100 µg/ml in broth —> 100mg/mL in stock
- Chloramphenicol: 25 µg/ml (usually - 12.5 µg/ml for fosmids) —> 25mg/mL in stock
- Kanamycin: 50 µg/ml —> 50mg/mL in stock
- Tetracycline: 10 µg/ml (light-sensitive! wrap plates/broths/stocks in foil) —> 10mg/mL in stock (Foil wrapped or in black-opaque tubes)
- Streptomycin: 200 µg/ml —> 200mg/mL in stock
- Gentamicin: 10 µg/ml —> 10mg/mL in stock
Protocol
- Grab your supplies. For a 50mL batch you will need:
- One falcon (50mL) full of ultra-pure water (i.e. MilliQ)
- Two empty falcon tubes (50mL)
- One 50mL syringe - plastic tip, not metal as we don’t want needle stick injuries
- One filter steriliser tip to fit your 50mL syringe
- Your antibiotic powder of choice
- A teeny tiny spoon
- A falcon tube holding rack
- Place your falcon tube holding rack and one empty falcon on your scales, set scale to zero.
- Weigh your dried antibiotic to make up desired concentration (e.g. 250mg Kanamycin, 5000mg Ampicillin), using the all-important teeny tiny spoon to measure into the empty falcon.
- Using a measuring cylinder, measure out 50mL of your ultra-pure water. Pour measured water into the falcon with the antibiotic powder.
- Seal the falcon tube and shake it like a polaroid picture until the powder is dissolved. Top tip: make sure that the tube is sealed or you will get antibiotic water everywhere!
- Now we need to filter sterilise the stock, so prepare your other empty falcon tube by placing it into the rack. Open your filter steriliser tip, affix it to the end of your syringe, remove the plunger so you can pour your stock into the syringe.
- Don’t leave the plunger on the bench, keep it in your hand - remember, the bench is lava!
- Once the syringe is full, you can put the plunger back into the syringe and push your stock through your filter steriliser.
- You have now made a stock! Congratulations! It’s time to make your aliquots!
- Get out 1.5 mL tubes and label them with the antibiotic name/abbreviation.
- Pipette 1mL of antibiotic into each tube. You may end up not having enough antibiotic to put into all 50 of the tubes, sometimes you lose an mL or so along the way. This is normal. Take deep breaths and just be proud you managed to make at least 45 tubes.
- Place into a labelled ziplock bag and freeze at -20C until required.
- Stocks can be defrosted by warming in hands as required.
If you're going to do any Synbio, you're bound to need a TONNE of plates. Here's the best method we've found for making them!
- Figure out the total number of plates you will need, then multiply x 25 to get the approx. amount of agar you will need (in ml). It's better to make more than you think you need! Alternatively, just make up the agar in multiples of 400 ml, this a standard amount that will make ~16 plates (this is about right for one ‘sleeve’ of sterile Petri dishes, which have 20 dishes in them).
- Add all the required media ingredients (eg. for LB agar, this is 10 g tryptone, 5 g yeast extract and 5 g of NaCl per litre) – make sure you correct these ingredient weights for the amount of agar you are making (e.g. multiply by 0.4 if you are only making 400 ml).
- Dispense the media into multiple lots of 400 ml in 500 ml media bottles. Make sure that the media bottles have a plastic pouring ‘lip’ on them, not just a naked glass top; the lip is important to be able to pour the agar neatly. You could also make multiples of 200 ml in 250 ml bottles, but don’t make >400 ml in a 1 L bottle – this is too heavy to easily pour plates with.
- Once you have dispensed the media into the bottles, now you can add the appropriate amount of agar to each bottle. This is 17 grams per litre (=6.8 grams per 400 ml).
- Add bottle caps. Don’t screw these on all the way, leave them a bit loose. Hold the caps in place with a small piece of autoclave tape.
- Autoclave the agar. If the agar media will not be autoclaved on the same day, keep it in the cold room until the next day (microbes will start to grow within a few hours, especially in rich media like LB).
- When the agar comes out of the autoclave, if it is still molten and very hot, put it in the hot water bath (~60°C) until it cools down to pouring temperature (~15 min). If you don’t want to use it immediately, leave on the bench until it solidifies (if it hasn’t already), then tighten the cap and it can be stored indefinitely at room temp. If agar has solidified and you want to use it straight away, microwave it (lid loose!) for approx 10 min on 50% power (these settings are for 400 ml of agar at room temp, adjust accordingly!), then put in hot water bath for ~15 min.
- Prepare laminar flow hood for plate pouring as follows: give it a blast of UV for ~15 min then turn off UV light,), take out the doors, then turn on the light and fan. Swab down with 80% ethanol or F10SC.
- If you don’t have access to the laminar flow hood, you can make plates on your regular lab bench. This is usually fine, but swab down the bench with ethanol first, and work close to the Bunsen flame. (if you swab your gloves with ethanol, be careful to let this evaporate before turning on the Bunsen burner!)
- Collect one bottle of molten agar from the waterbath, wipe down the outside with paper towel, and take to your bench. Add any “after-autoclaving” additions at this stage, from filter-sterilised stock solutions (this may be antibiotics, or trace metals solution, or Tween etc). Mix by swirling for 10-20 seconds, don’t shake the bottle as this will cause persistent bubbles. Its good practice to write all the additions on the bottle, then tick them off after each addition. Forgetting these is bad!
- Take the final agar to the laminar flow hood. Open up one sleeve of plates (at the ‘base’ end), and slide them out. Keep the sleeve! Label all the plates with the type of medium (e.g LB-Cm25). Pour the agar into the first plate; use a single smooth pouring motion. Stop when it reaches all the edges. Put the lid on. Do the next plate. Repeat. Don’t stack up the plates in big towers, they will take longer to cool.
- Leave the plates for about 10 min to allow the agar to solidify. Then invert the plates with the lids off to allow them to dry for 20-30 min.
- Pack the plates back into the sleeve, then label the sleeve with the type of agar and the date of manufacture and your name.
- Store plates in the cold room. They should stay good for >6 months if you have been careful!
DON’T ADD THE AGAR YET! (this doesn’t dissolve, it settles out)
DON’T ADD ANTIBIOTICS, TRACE METALS, or TWEEN yet, if these ingredients are needed.
If you don’t dry the plates, you won’t get nice spread-plates, and the risk of contamination is greater from other microbes ‘climbing’ into the plate over the edge.
This protocol is based on the Coleman Lab protocol.
The solutions needed for the plasmid prep are listed at the end of the protocol. These should work with any kind of silica-based purification column and could also be adapted to purification using free silica particles (e.g. diatomaceous earth).
Wear appropriate PE including lab coat gloves and safety glasses. Hazards include NaOH, SDS and guanidine.
Method for 5mL Culture
- Inoculate 5 ml of LB broth containing the appropriate antibiotic with a loopful of E.coli culture from a fresh plate of the same medium (‘fresh’ here means less than ~2 weeks old). Grow overnight (16-24 h) at 37°C with shaking.
- Ampicillin: 100 μg/ml
- Kanamycin: 50 μg/ml
- Streptomycin: 200 μg/ml
- Chloramphenicol: 25 μg/ml (usually) or 12.5 μg/ml (for fosmids)
- Tetracycline: 10 μg/ml (light-sensitive! wrap plates/broths in foil)
- Gentamicin: 10 μg/ml
- Centrifuge the culture at 3000 rpm for 10 minutes.
- Pour off supernatant into culture waste and resuspend the cells in 1 ml of buffer EB by vortexing. Transfer to 1.5 ml Eppendorf tube. Centrifuge 1 min at ~10,000 g. Pour off supernatant, keep pellet.
- Resuspend the cell pellet in 200 μl buffer P1 by vortexing.
- Add 300 μl buffer P2. Mix by rapidly inverting the tube 10 times. Incubate 10 min at room temp. (No longer than 10 min!) The mixture should go clear and become viscous as the cells break open.
- Add 400 μl buffer N3. Mix by rapidly inverting the tube 10 times. It is essential that the N3 buffer is thoroughly mixed in. Viscosity should disappear, and a white precipitate should appear. Incubate for 5 minutes at room temp. Centrifuge for 5 min at ~10,000 g.
- Place a spin column (silica-based e.g. ‘Econospin’ brand) into its collection tube (if it isn’t already set up that way), and carefully pipette 750μl of the plasmid extract supernatant onto the column. Avoid the white pellet and any white junk that may be floating on top. Centrifuge 30 seconds at ~10,000 g. Discard flow-through into culture waste.
- Add 700 μl buffer PB to the column, centrifuge 30 sec at ~10,000 x g. Discard flow-through.
- Add 700μl buffer PE to the column, centrifuge 30 sec at ~10,000 x g. Discard flow-through.
- Repeat step 9.
- Spin the ‘empty’ column one more time for 2 min at ~10,000 x g to remove the last traces of PE.
- Discard collection tube, and place the column part with the lid open on a clean ‘Kimwipe’. Place on a 60 C° heatblock for ~10 min to evaporate residual ethanol. Avoid touching the ‘nipple’ at the bottom of the column with your fingers.
- Place column into an fresh Eppi tube, and add 30 μl of buffer EB to each column. (ensure EB goes onto the membrane at the bottom, not the wall). Allow to sit for 2 min. Centrifuge 1 min at ~10,000 g.
- Digest 2 μl of the plasmid preparation with an appropriate restriction enzyme (eg. an enzyme that cuts once or twice) (see restriction digest protocol), and run on an agarose gel to evaluate yield. Note that the Nanodrop may not be accurate for plasmid preps.
It is critical to remember to add the antibiotics to the LB broth. Some common antibiotics and their typical concentrations are shown below:
Adaption for 50ml culture
- Inoculate 50 ml of LB broth containing the appropriate antibiotic with a large loopful of your E.coli culture from a fresh plate of the same medium.Grow overnight (16-24 h) at 37°C with shaking.
- Aseptically transfer (pour) the cells into a 50 ml Falcon tube. Find or make a balance tube with the same volume of water. Estimating equal volumes by eye (+/- 1 ml) is OK for low speed centrifuges, but a balance should be used for more sensitive machines or when spinning at higher speeds (>3000g) (for these, need accuracy +/- 0.1 mg).
- Centrifuge at 4000 rpm for 10 minutes. Pour of supernatant and keep pellet
- Resuspend the cell pellet in 800 μl buffer P1 by vortexing. Transfer 200μl of the resuspended cells into 4 fresh Eppi tubes. Continue as for 5mL protocol from step 5.
Plasmid Prep Solutions
- Buffer P1: 5 mM EDTA, pH 8. Sterilise by autoclaving. Then add RNAse to 300 μg/ml.
- Buffer P2: 1% SDS, 0.2 M NaOH. Not sterilised.
- Buffer N3: 4.2 M guanidine HCl, 0.9 M K-acetate, pH 4.8. Not sterilised. Adjust the pH using glacial acetic acid
- Buffer PB: 5 M guanidine HCl, 30% isopropanol. Not sterilised. Prepare in a sterile bottle or tube containing the correct amount of autoclaved RO water.
- Buffer PE: 10 mM Tris-HCl, pH 8, 80% ethanol. Not sterilised. Prepare in a sterile bottle or tube containing the correct amount of autoclaved RO water.
- Buffer EB: 10 mM Tris-HCl, pH 8. Autoclaved. Usually this would be prepared by diluting a 1M Tris-HCL pH 8 stock.
Usually this would be prepared by starting with a 0.5 M EDTA, pH 8 stock, and a 10 mg/ml aliquot of boiled RNAse.
Prepare from stocks of 10% and 2 M NaOH (these don’t need to be sterilised, but it is not a bad idea to prepare these and the final buffer in sterile bottles or tubes containing the correct amount of autoclaved RO water). Keep in a tube or bottle with minimal headspace and tightly closed. Will eventually develop precipitate of Na2CO3 – remake the solution if you see this. Recommend to make fresh approx. monthly.
Notes: As with all molecular biology and microbiology work, aseptic technique is crucial. Clean your bench, tube racks, and pipettes with 80% ethanol before starting, and work carefully to ensure you don’t contaminate the PCR reagents or mixtures with microbes, DNA, or enzymes from your hands or the bench or the ice bucket etc etc. Ensure pipette tips, Eppi tubes and PCR tubes are sterile.
This protocol can be used to add restriction sites to ends of DNA sequences. When making primer, add the restriction site sequence to the 5’ end of the primer.
Different PCRs behave differently. Some PCRs are very straightforward while others are very challenging. The factors impacting the difficulty of PCR include the following:
- Type of DNA Template: the easiest template is a plasmid or PCR product containing your sequence of interest. These very clean and have a very high ratio of target sequence to non-target sequence. A medium difficulty template would be genomic DNA from a single microbe.
- Purity of DNA template: highly purified DNA will amplify much better than crude DNA (e.g. boiled cells), which will amplify better than whole cells, which will amplify better than a complex and dirty template (e.g whole soil) (this is unlikely to work at all!). That said, you can certainly do PCR on whole cells of most gram-negative bacteria, including E. coli (this is known as ‘colony PCR’), and this is very useful for rapidly screening clones.
- Type of primers: specific primers work well, degenerate primers work less well, and the higher the degeneracy, the worse they perform. Primers that are designed well (no dimers/hairpins or only weak dimers/hairpins) will work better than primers that are poorly designed (strong dimers/hairpins). Primers that have lots of junk at the 5’ end like long non-target sequences or fluorochromes will perform worse than primers that are exactly the same as the template. Primers with mismatches to the template may still work, but the more mismatches present, the worse they will work, and the closer the mismatches are to the 3’ end, the more serious the problems will be.
- Type of target gene, and its copy number: If your target gene is on a plasmid, it will amplify more easily than a chromosomal gene. If your gene has many copies in the template DNA, it will amplify better than a gene which is very rare. If your target gene is 16S rDNA or some other highly-conserved gene, and your primers are ‘universal’ primers, you might expect problems with contamination, since this gene is *everywhere*, including on your hands, on the bench, in your pipette etc. (the negative control with no DNA added is critical in this case!). On the other hand, if your target gene is only found in your particular favourite and unusual organism, then contamination will be less of a concern.
- Size of PCR product: The smaller the product, the easier the PCR. Product size should not cause problems up to ~ 1 kb, but as you go larger than this, the PCR will become increasingly challenging. This also depends a lot on the type of polymerase used (Phusion is better than Pfu, which is better than Taq, in terms of getting large products). Anything larger than ~5 kb is going to be difficult to amplify, even for an experienced user – success here requires using a fancy polymerase, *excellent* primers (well-designed, no dimers/hairpins), clean and high-quality template with high signal:noise ratio, fresh reagents, and excellent hands-on technique. In theory, PCR products up to ~20 kb are possible, but these are extremely difficult to obtain in practice.
Protocol for setup of PCRs
Notes: The usual PCR is 25 μl volume, this is appropriate for “screening” purposes. However, if you are trying to make a lot of PCR product for cloning, then scale reactions up to 50 μl per tube. A PCR that is working well should give you about 50 ng of product per μl of reaction, thus 2.5 μg per 50 μl. For a cloning experiment, if you prepare 8 x 50 μl PCRs, this should give you plenty of insert DNA (10-20 μg), even if your PCR is not working at high efficiency. This is way in excess of the theoretical requirement, but allows for losses due to subsequent purification steps, and the likelihood that you may need to repeat the experiment a few times
Example Reaction Mix (per 25μL)
- 10xbuffer: 2.5μl (->1x)
- sterile milliQ water: 20 μl
- dNTPs (10 mM): 0.5 μl
- primer #1 (50 μM): 0.25 μl
- primer #2 (50 μM): 0.25 μl
- polymerase* (5 U/μl): 0.25 μl
- DNA template: 1uL
For 8 Reactions** (one strip of tubes)
- 10xbuffer: 22.5μl
- sterile milliQ water: 182 μl
- dNTPs (10 mM): 4.5 μl
- primer #1 (50 μM): 2.3 μl
- primer #2 (50 μM): 2.3 μl
- polymerase* (5 U/μl): 2.3μl
- DNA template : 8 x 1uL
* This means thermostable DNA pol e.g. Taq, Pfu, Phusion, Q5, etc. but NOT Klenow or T4 pol.
** Calculations done as if 9 x 25 μl reactions were being prepared, to ensure mix doesn’t run out.
Template considerations: The correct amount of DNA template depends on the type of sample. If you are using a plasmid, you only need a tiny bit (~1 ng) because the ratio of target:non-target sequences is very high. For genomic DNA of a single microbe, a more usual amount would be 10-20 ng per reaction. For metagenomic DNA, you may need to go even higher (100 ng per reaction), although note that there are diminishing returns here due to the presence of PCR inhibitors in many templates that will interfere with PCR. For complex templates like soil DNA, the optimum concentration of template often needs to be determined experimentally. More is not always better; in some cases, diluting the template (e.g. 1/10) will yield a band when undiluted template gives no band at all.
Set up of Mastermix and Reactions
- Exact setup of PCR depends on which element is the variable in the reaction. Usually this is the DNA template, but sometimes it is the primer. The ‘Master Mix’ should be made to include everything except the variable being tested. The below protocol and above recipes are written assuming that this variable is the DNA; you need to modify if the same template is being tested with multiple primers.
- Calculate how many PCRs you need altogether, and thus the total volume of master mix required. It’s important to make more than you need (~10% more), as pipetting errors will always change the expected volumes a little bit. Write up the exact recipe that you need in your lab book.
- Label your strip tubes on the side, near the top. If you label the lids or the bottom of the tube, the labels tend to come off during thermocycling. Place the tubes in a tube-rack (a 96-well micro-titre plate works well). Leave the lids off the tubes – you are more likely to contaminate the reactions via excessive opening and closing of tubes than from stuff falling in from the air.
- Retrieve the PCR reagents from the -20°C freezer and thaw them (except polymerase, this remains liquid even at -20°C due to glycerol in buffer). It’s important that the 10x buffer, dNTPs, and primers are thoroughly thawed out before use. Give them a brief vortex or flick to mix. These reagents don’t need to be kept on ice during the time needed to set up the PCR, but don’t leave them on the bench longer than needed.
- Prepare the master mix in a sterile Eppi tube, containing everything except the variable being tested. Add ingredients in the order listed in the recipes above. Some folks will insist that this must be done on ice, opinions differ. It is true that you will minimise premature polymerase activity by setting up on ice, but whether this really makes a difference for most PCRs is arguable. Setup on ice is supposedly more important for Pfu and Phusion which contain 3’-5’ exonuclease activity which can ‘eat’ your primers during setup. If you set up on ice, risks of contamination from the ice itself must be managed.
- Aliquot 25uL of the master mix into each of the PCR tubes.
- Adding the DNA
- (DNA template in solution) Add DNA template (1 μl) to all the tubes, being very careful to match the sample #’s to the labels on the tubes. You need to concentrate your attention here and really focus in order to avoid missing a tube or putting two templates in the same tube. Be careful to change tips between every DNA template.
- (colony PCR: DNA in cells). Touch a white tip onto a colony or patch of growth and pick up a *small* amount of cells. It should be just enough so that you can see there are cells there, not much more than this. Then dip the tip in and out of the master mix in the first PCR tube approx 5 times and use the same tip to streak out a patch on a patch plate. You don’t need to wipe the whole chunk of growth on the inside of the tube, just the dipping in and out action will dislodge enough cells to give you enough template DNA. Less is more in this case! Repeat for other colonies/patches.
- Put lids on tubes, ensure they are snapped on tight, place immediately in thermocycler. Double check your program parameters before starting. See below for detailed thermocycling instructions.
- Return all reagents to the freezer.
- After thermocycling is completed, run out your PCRs on an agarose gel to check their size and yield. Typically, we would load 5 μl of PCR mixture. This should give a very strong band if the PCR has been successful.
- If you are intending to use the PCR product for cloning, check for the presence of non-specific bands (usually fainter, smaller bands than the expected one) – if these are numerous and/or strong, you may need to try a higher annealing temp, or revisit primer design, before the product can be cloned. Alternatively, you can cut out the desired band from the agarose (see our Purification of DNA via spin column protocol).
Thermocycling
A standard thermocycling protocol is given below (for Taq polymerase)
- Initial denaturation: 95°C, 5 min
- Denaturation: 95°C, 30 sec
- Annealing: 5°C lower than the melting temperature of the primers, 30 sec
- Extension: 72°C, 1 min per 1000kb product (differs between polymerases)
- 35 cycles of these steps
- Final extension: 72°C, 10 min
- Hold: 15°C
The numbers that are in bold indicate variables that need to be optimised for every individual PCR. Sometimes, other variables may also need to be modified too, as below Annealing temperature: This is typically set at 5°C lower than the melting temperature (Tm) of the primers. If these Tm’s are different, use the lower one for this calculation. This calculation is very crude, and it is best to optimise annealing temperature experimentally by doing a ‘gradient PCR’, e.g. testing an annealing temp range from 5° lower to 5° higher than the predicted best temp. Aim to make primers with Tm’s around 65°C, which gives a predicted annealing temp around 60°C. The usable range for annealing temp’s is approx 50°C – 70°C.
Extension time: This is set primarily by the length of the desired PCR product. For Taq polymerase, this is 1 minute of extension per kb of product. However, note that different polymerases require different extension times: Pfu needs 1.5 min per kb, and Phusion needs 0.5 min per kb. If you use a longer extension time than necessary, you risk increasing the yield of non-specific products; if you use a shorter extension time than necessary, you risk a low yield of the desired product. Denaturation: The optimal denaturation conditions for Phusion pol are different than for Taq or Pfu. Change the initial denaturation to 98°C - 30 sec, and the denaturation in each cycle to 98°C - 10 sec.
Number of cycles: This can be increased to 35 or even 40 cycles to give a higher yield of product, but this risks introducing more mutations into the PCR amplicons, and also gives a higher chance of secondary non-specific products being formed. Conversely, the # of cycles can be reduced to 25, which will minimise mutations and non-specific products, but will also lower yields.
Hold temperature: It’s OK to leave your PCRs in the machine overnight. While some protocols recommend fridge temp (4°C) for this ‘Hold’ step, this is actually bad for the machine – it will accumulate condensation on the block, which will degrade the block over time. A good compromise is to set the hold temp to 15°C, which keeps the samples cool(-ish), maintaining them in good condition, but is not cool enough to cause condensation to form.
PCR Enhancer Additions: Two enhancers that might be worth testing for problematic PCRs are Bovine Serum Albumin (BSA) and dimethylsulfoxide (DMSO). BSA is good for binding inhibitors and is typically added at 0.5 mg/ml final conc. from a 10 mg/ml stock (=1.25 μl per 25 μl reaction). DMSO is good for enabling amplification of GC-rich templates and is typically added at 1 μl of neat DMSO per 25 μl reaction (=4%).
Prepping your Primers
This diagram shows some of the basic PCRs that can be used to determine whether a plasmid clone is:
- The desired recombinant
- Some other kind of recombinant
- Vector only
PCR using primers F1-R1 is often the starting point for screening. This is known as left junction PCR. Getting a product of the expected size in this PCR is a good indication you have the right recombinants.
PCR using primers F2-R2 (right junction PCR) can then be done on colonies positive in left junction PCR. If the clones are also positive in this PCR, it is further evidence that they are the construct you’re after.
PCR using the primers F1-R2 is known as a spanning PCR or an insert spanning PCR. This PCR will amplify the entire insert region. This PCR can be used to determine if the insert is the correct size, but bear in mind that this is not an all or nothing PCR like the junction PCRs. In spanning PCR the clones will give a small product if they have no insert, and a large product if there is an insert. However, if you have a mixed population of cells you may get a false negative result as the small product will be amplified preferentially, even if there are positive clones in there too. This is why pure colony isolates are so important. Spanning PCR is most often used to prepare the insert region for sequencing rather than initial clone screening as longer PCR products take longer to thermocycle.
Primers used for the amplification of the original DNA insert (not shown in the diagram) can also be used for screening, but they will not tell you whether the insert is oriented correctly like the junction PCRs will.
Positioning of the primers is important, especially for later steps such as sequencing. Make sure that all primers (F1, F2, R1, R2) are at least 50bp away from the ligation join - as the first 20-30bp of sequence read are usually ‘junk’. However, if the primers are too far away from the ligation join you’re amplifying and sequencing things that you don’t need to, which increases time and expense. Placing the primers about 100-150bp away from the ligation join is good, and ought to give a positive PCR product of around 200-300bp.
Initial Patching and PCR Screening
- Choose 20 well-isolated colonies from your transformation plates. (if your construct has colour selection, ensure you pick colonies that are not connected to any negatively coloured colonies). Circle and number these on the base of the plate.
- Label three PCR strip tubes with numbers 1-20, leaving three blanks (one per strip) as negative controls.
- Using an agar plate containing the appropriate antibiotic, divide the plate into patches. On the bottom of the plate draw a 5x5 grid. Label 20 of the squares with numbers 1 to 20 (see diagram).
- Prepare a PCR master mix with appropriate left junction primers, enough for ~30 PCRs. Aliquot this into 3 x 8 lots of 25uL in the labelled PCR strip tubes. Taq polymerase is good to use for this job as it is very robust to any ‘mess’ in the PCR, but other polymerases can also be used.
- Using a white pipette tip (suitable for p10 or p2 pipettes), pick up some growth from one of your colonies of interest. Make sure you can see some cells on the tip, but you don’t need a lot.
- Dip the tip in and out of the first PCR tube five times. You don’t need to dislodge the whole chunk of growth, enough cells will fall into the mix to give you a product.
- With the same tip, transfer the remaining growth onto the first patch on your agar plate by scratching the tip across the surface of the agar to make a set of three close parallel lines (see diagram). Stay clear of the borders of each square, we don't want the colonies to get too friendly with each other!
- Repeat steps 5 - 7 for the remaining colonies, making sure to be careful that you match your tube number to the plate square number each time. Cross of circled colonies after you have sampled them - no need to test one colony twice!
- Put the PCR tubes in the thermocycler and incubate the agar plate overnight.
- Run the PCR products on an agarose gel and note which colony numbers gave a positive result.
- If none of your clones are positive then you may need to test another twenty from your transformation plate. However, if none of those are positive either you will need to do some troubleshooting as something is likely wrong - check your ligation and transformation controls, check your primer sequences for issues such as dimers, and make sure your thermocycler conditions are appropriate for your polymerase and your primers. You may end up needing to repeat the cloning process. If a second cloning still yields no positive colonies it is possible that your gene of interest is toxic to your host cells - you may need a diferent vector (e.g. inducible, lower copy number) or a diferent host or diferent incubation conditions.
Secondary PCR Screening
- After getting between three and seven positive colonies from your primary screening you may want to confirm these results with a right junction PCR.
- Set up a PCR master mix with appropriate right junction primers. Make enough master mix so there is at least 10% more than you need so you don’t run out.
- Make up another agar plate, but this time with just enough squares for the number of positive colonies. It may be helpful to you to label the squares with the numbers of the positive patches (i.e. if your positives were numbers 7, 12, and 19 you would label your three squares with those numbers rather than 1, 2, and 3.)
- Follow steps 5 - 8 from the above protocol, taking some colonies from the patch plate you made.
- Put the PCR tubes in the thermocycler and incubate the agar plate overnight.
- After thermocycling, run the products on an agarose gel. Note which colonies were positive.
- Choose one positive colony to inoculate a 100mL LB-antibiotic broth to prepare for a plasmid prep.
- Keep the patch plate! Wrap it with parafilm and keep it in the cool room. You may end up needing to use a diferent positive colony if there are problems with the first one.
Even though you’ve performed two junction PCRs you can’t be sure your construct is completely correct. It is advised that you validate your construct using a restriction digest or through sequencing . Small mutations cannot be easily detected by junction PCR, and they can completely change the functionality of your construct!
- Choose one or a combination of restriction enzymes that will cut the plasmid 1-3 times. Make sure that all the predicted fragment sizes are different enough to separate on a gel. If you’re using multiple restriction enzymes, make sure that they all work in the same buffer and at the same temperature.
- Get your buffer out of the freezer and ensure it is totally thawed and mixed before use.
- Get restriction enzymes out of freezer and put them on ice immediately. It is critical that the enzymes stay cold at all times.
- Set up the digest in a sterile Eppendorf tube in the following order:
- Incubate at the preferred temperature. If using HF enzymes, 15 minutes is sufficient, but can digest for up to 24hrs. 30 minutes – 1 hour is optimal.
- Run on agarose gel to check. Note: For Bam restriction enzymes, you must use SDS containing loading buffer or purify or heat kill the sample (15 min 80°C) before running the gel, as Bam binds tightly to the DNA and will slow the migration of the DNA and change the apparent size of the bands.
2uL 10x buffer
YuL MilliQ water
XuL plasmid, containing 250ng DNA
1uL each RE
----------------------------------------------
20uL total
Ensure you are changing tips between every addition to avoid contaminating the restriction enzymes.
Ideally, you would not use more than 5uL of plasmid. If you use more that 5uL, the digest may not work due to junk in the plasmid prep (e.g. salts) interfering – in this case, consider further purification and concentration of the plasmid e.g. by ethanol precipitation. Calculate the amount of MQW needed (Y) by subtracting the total volume, 20uL, from the amount of plasmid needed (X) and the number of restriction enzymes being used.
This protocol is to turn the sticky ends left over from a restriction digest into blunt ends. Mung bean nuclease has the ability to chew back the overhang.
Can be done in the same reaction mixture as the digest that creates sticky ends or can be done subsequently after heat-killing or spin column purifying DNA. Reaction can be scaled up or down as needed.
A reaction contains:
- 10x Cutsmart Buffer
- A certain number of ng DNA (make sure you know how many!)
- 1 Unit Mung Bean Nuclease per ng DNA. The concentration of NEB Mung Bean Nuclease is 10,000U/mL, which is the equivalent of 10U/uL. Therefore, the number of uL of nuclease is the number of ng of DNA divided by 10.
- Total to desired volume with MQW (30uL is a good size, depends on the amount of DNA)
Make up reaction in Eppendorf tube. Incubate at 30°C for 30 minutes.
After the 30 minutes is over, purify the DNA by spin column. DO NOT ATTEMPT TO HEAT KILL!
Making gels is one of the most crucial parts of any iGEM project. They can be used for gel purification, restriction digest diagnostics, and so much more. Here's our preferred method for making gels:
- Figure out how many samples you have and choose well-formers and gel volume accordingly: the smallest gel tank uses 50 ml agarose and takes up to 10 samples, the largest tank uses 150 ml agarose and takes up to ~40 samples (if two rows of well-formers used). Remember to leave a lane for the molecular weight marker!
- Decide what agarose concentration and what buffer are appropriate. For many gels, 1% agarose and 1x TAE will be fine. If you are trying to separate very small DNAs (<500 bp), you can increase up to 2% agarose, and if you are trying to separate large DNAs (>5 kb), you can decrease down to 0.7% agarose. If you want to run a gel very quickly (15mins), use 1 x LAB buffer. If you need to purify DNA from the gel for cloning or sequencing, use TAE (borate from TBE or LAB interferes with later enzyme reactions).
- Decide whether you need very accurate band-sizing or not. If you do not (e.g. just want to see band or no band when screening clones), you can put the detection dye (GelGreen or Thizole Orange) into the gel. If you do need high accuracy, instead use post-staining.
- Set up the gel-casting tray by masking-taping the ends of the tray. Place the well-former (comb) such that it is level and straight and sitting ~2 mm above the base of the gel forming unit. It is crucial that the well formers are NOT TOUCHING the casting tray – in that case, your wells will have holes in the bottom and you will lose your samples. 5. Using a small, clean, dry conical flask, weigh out the appropriate amount of agarose powder for the correct and % agarose gel that you intend to make, then pour in the appropriate volume of TBE or TAE buffer. Put in microwave, and heat on high for 1 minute. Put on heat resistant gloves, then take out to examine whether all the agarose is dissolved. If not, replace for another 30 sec on high power, and repeat examination and heating until all dissolved.
- Cool the conical flask down under a stream of cold water, while swirling. Be careful not to get water INTO the flask. After about 20 seconds, see if you can hold the flask comfortably in your hand (no glove) – if it is too hot to hold, continue cooling, re-check every 20 seconds or so. When the flask feels warm but not uncomfortably hot, it is ready to pour (this is approx. 45-50°C). If you cool it too much, it may set solid. In this case, microwave again (as above) in 30 sec increments to re-dissolve.
- If appropriate: add GelGreen or Thizole Orange dye to the gel. For gels with long run times, >40mins, do not use Thizole Orange, as it will run off the gel. Use 5uL per 50mL gel of stock for both dyes.
- Pour the molten agarose into the casting tray. Use a single smooth motion, don’t stop and start. Stop when the agarose is 3/4 of the height of the ‘teeth’ on the well-forming comb. Or alternatively, stop when you judge the wells are deep enough to hold the amount of sample that you need to load. What you DON’T want to do is over-fill the agarose so that it goes over the teeth – this will result in a channel that connects all the wells, and subsequent cross-contamination of the samples.
- Allow the gel to set. This takes about 15 minutes but will be faster if set up in the cold room.
- When gel is set, remove the masking tape, and put the gel in the electrophoresis tank. Fill the tank with enough of the appropriate buffer to fills both wells and covers the gel by 1-2mm, then carefully pull out the well formers (straight up, don’t yank them side to side or forwards and back). The reason for the buffer is to stop the wells collapsing on themselves (this can happen with thin wells at lower % agarose).
- Choose the appropriate molecular weight marker. For DNAs <1 kb, use the NEB 100 bp ladder. For DNAs from 1 kb – 10 kb use the NEB 1 kb ladder. You may need to load two different ladders, one on each side of the gel.
- Prepare a row of spots of purple loading dye on a Parafilm strip, of appropriate volume and number. This is a 6x buffer.
- List in your lab book a map of which samples will be loaded in which locations. Avoid the first and last wells if possible, these tend not to run as straight as the more central wells.
- Mix your first sample with the first spot of loading dye, and load into the first well. Steady the pipette tip with the finger of your non-pipetting hand to ensure accurate dispensing. The tip needs to be just inside the well, don’t push it all the way down in the well. Change tips, mix up the next sample with blue dye, and load again. Repeat for all samples, including the marker.
- Put the lid on the gel box, check that the terminals are connected correctly (negative terminal should be closest to the wells, positive terminal is far from the wells, ie. Run towards Red). Run the gel at ~150 volts (small gel box) up to ~250 V (large gel box). Check that you can see gas bubbles at the electrodes when you start running the gel. If not, check all the wire connections.
- Stop the gel when the fast-running blue dye (bromophenol blue) (= usually the only blue dye) is near the end of the gel (this may take 30-60 min). You may need to run longer to get good separation of large products (>5 kb). Run for shorter time for small products (<500 bp), or they may run off gel.
- If you added GelGreen or Thizole orange to gel, it is now ready to image on the transilluminator. Otherwise, post-stain in a solution of Thizole Orange (5 μl per 50ml water) for 30 min on a mechanical rocker, before transilluminating.
GENERAL NOTES
This protocol is good for restriction fragments, small digested plasmids, and PCR products. It is NOT good for genomic or chromosomal DNA. Use the FastPrep reagents or CTAB-phenol type prep instead for genomic DNA. Digested plasmids work well up to about 5 kb, but the yield quickly falls off past this point. Generally, plasmids don’t purify as well as PCR products on these columns.
- Figure out what volume of sample you have. If this is less than 100 µl, then make it up to 100 µl with TE. The volume you now have will be called “1 volume”. Add 5 volumes of buffer PB to the DNA solution. e.g. if you have a 100 µl restriction digest, add 500 µl PB.
- Place a silica-based spin column (e.g. ‘Econospin’) into its 2 ml catch tube (if it isn’t already set up that way). Load up to 750 µl of the DNA-PB mixture onto the column. Spin at ~10,000 g for 30 sec. Discard the flow-through into culture waste.
- If you still have more DNA-PB mixture left, repeat the previous step until all of the mixture has been put thru the column. The columns will hold a total of ~10 µg DNA, which is a lot!
- Add 750 µl of buffer PE to the column, spin ~10,000 g for 30 sec, discard flow-through.
- Repeat step 4.
- Spin again for 30 sec to remove all traces of PE from the column. Discard both the flow-through and catch tube, and transfer the spin column onto a clean Kimwipe. Leave the column lid open. Transfer Kimwipe to 60°C oven, and allow to dry for 10 min.
- Transfer spin column to a sterile 1.5 ml Eppi tube, and add 20-50 µl* of EB buffer to the centre of the spin column – ie on the membrane, not the walls of tube. Allow to sit for 2 min. Spin at ~10,000 g for 1 min, retain Eppi tube with DNA solution in EB, discard spin column.
* Usually we want high concentration rather than high yield, so use 20 µl. If max. yield is important, or if you have lots of DNA, use 50 µl. Note that you lose approx 5 µl EB during the procedure.
General Notes
This protocol is good for restriction fragments, small digested plasmids, and PCR products. It is NOT good for genomic or chromosomal DNA. Digested plasmids work well up to about 5 kb, but the yield quickly falls off past this point. Generally, plasmids don’t purify as well as PCR products on these columns.
Gel-purification: Don’t expose the DNA to any UV light and don’t use borate-containing buffers in the gel. Even minimal exposure to short-wave UV light will greatly damage DNA, and make it hard to ligate later. Long-wave UV (e.g. hand-held lamp) is less damaging than short-wave UV (transilluminator), but best to avoid UV altogether. Borate can cause problems for later enzyme steps.
Notes on Gel Extraction Using Gel-green (recommended)
- f you need to purify DNA from an agarose gel, use TAE-agarose (don’t use TBE!). Put Gel-Green (1 μl) in the gel. Load plenty of DNA so it is easy to visualize.
- Run gel as usual, then visualize under blue illumination (NOT UV!) in a dark room or box. There is a blue light in the lab for this purpose and orange-tinted glasses also help. Cut out your band(s) of interest with a clean, sharp scalpel blade, and put them into an Eppi tube.
- Weigh the tube containing gel slice and subtract weight of empty Eppi tube to figure out the weight of gel. Proceed to ‘main protocol’ below.
- Weigh the tube containing gel slice and subtract weight of empty Eppi tube to figure out the weight of gel. Proceed to ‘main protocol’ below.
Main Protocol
Be sure to have your gloves and safety glasses on. QG contains guanidine which is somewhat toxic.
- If your weight of agarose is less than 100 mg (=100 μl), then make it up to 100 μl with TE. The volume you now have will be called “1 volume”. Then add “3.5 volumes” of QG buffer to your gel slice. eg. if you have 100 mg of agarose, add 350 μl of QG.
- Slice up or mash the agarose in an Eppi (<300 mg agarose) or a McCartney bottle (>300 mg). This will speed dissolution. Melt the agarose with heating (60°C for 5 min is usually enough, mix occasionally), then when it’s all dissolved, allow to cool to room temp.
- Add “1.5 volumes” of isopropanol to the mixture. e.g. for 100 mg agarose, use 150 μl isopropanol.
- Place a silica-based spin column into its 2 ml catch tube (if it isn’t already set up that way). Load up to 750 μl of the DNA / QG / isopropanol mixture onto the column. Spin at ~10,000 g for 30 sec. Discard the flow-through into culture waste.
- If you still have more DNA /QG / isopropanol mixture left, repeat the previous step until all of the mixture has been put through the column. The columns will hold a total of ~10 μg DNA, which is a lot!
- Add 750 μl of buffer PE to the column, spin ~10,000 g for 30 sec, discard flow-through.
- Repeat step 6.
- Spin again for 30 sec to remove all traces of PE from the column. Discard both the flow-through and catch tube, and transfer the spin column onto a clean Kimwipe. Leave the column lid open. Transfer Kimwipe to 60°C oven, and allow to dry for 10 min.
- Transfer spin column to a sterile 1.5 ml Eppi tube, and add 20-50 μl* of EB buffer (5 mM Tris, pH 8) to the centre of the spin column – ie on the membrane, not the walls of tube. Allow to sit for 2 min. Spin at ~10,000 g for 1 min, retain Eppi tube with DNA solution in EB, discard spin column.
* Usually we want high concentration rather than high yield, so use 20 μl. If max. yield is important, or if you have lots of DNA, use 50 μl. Note that you lose approx 5 μl EB during the procedure.
DNA Purification Solutions (Spin Colummn)
- Buffer QG: 5M guanidine HCl. Not sterilized. Prepare by adding GuHCl to autoclaved RO water in a sterile bottle/tube.
- Buffer PE: 10 mM Tris-HCl, pH 8, 80% ethanol. Not sterilised.
- Buffer EB: 10 mM Tris-HCl, pH 8. Autoclaved. Add appropriate vol. of autoclaved 1 M Tris-HCl to the correct amount of autoclaved RO water in a sterile bottle or tube.
Add appropriate vol. of autoclaved 1 M Tris-HCl to the correct amount of autoclaved RO water in a sterile bottle or tube, then add 100% ethanol to give 80% final conc.
Use excellent aseptic technique at all times. All materials must be sterile. Protocol can be scaled up or scaled down as required. 100mL E. coli culture produces about 50 x 220uL aliquots of competent cells. Each aliquot is sufficient for 4 transformations, with a little bufer for safety. Competent cells can be used fresh, so feel free to make them fresh for your experiment but as this is quite an involved process, do your labmates a solid and make a big batch to put into the freezer for future use. Freezer stocks will not survive multiple freeze-thaw cycles so once you take an aliquot out of the freezer any leftovers must be disposed of.
After step 7 you will need to work on ice and in the cold room to increase the quality of the final cell prep. These steps will be written in blue for easy differentiation.
- Streak desired E. coli strain (e.g. TOP10, BL21(DE3), JM109) from glycerol stock or other source onto plain LB agar, no antibiotics (means you need to be extra careful!). Incubate overnight at 37C.
- Check that your culture from the night before looks pure. (hint: if it’s green and fuzzy, that aint E. coli). If it’s looking good, inoculate a plain 5mL LB broth with several of the colonies from your LB plate. Incubate overnight (16-24hr) at 37C with shaking.
- At the beginning of the day, place two centrifuge tubes in the freezer, ready to use.
- Aseptically inoculate 100mL plain LB broth (in a 500mL Schott bottle or Erlenmeyer flask) with 3 ml of overnight culture; this should give an initial OD600 of ~0.05. (check this to be sure).
- Grow cells at 37C with shaking until the culture reaches an OD600 of ~0.5 (if you’re short on time, above 0.3 is passable. If you overshot it, anything up to 0.7 is okay). This can take up to 3 hours, but some E. coli are over achievers so after the first hour check the OD every 15-30mins.
- Aseptically transfer (also known as carefully pour) the cells into the cold, sterile centrifuge tubes or bottles (one 250 ml Sorvall bottle or 2 x 50 ml Falcon tubes) you prepared earlier. Balance these, either by transferring culture aseptically between the tubes/bottles or by adding sterile water or LB to the lighter tube/ bottle.
- (Brrr… now’s the chilly part!) Centrifuge at 4000 rpm (~3000 g in Centaur/ Centurion machine) at 4°C for 10 minutes. Note: centrifuge needs to be cold. Turn on and set temperature beforehand. You can spin faster, up to say 7000 g, but above this point, faster is not better, and cell pellets will be hard to resuspend, but if you skipped arm day by all means give it your best shot.
- Working in the cold room with the cells on ice, pour of the supernatant into culture waste (don’t let the centrifuge tube/bottle actually touch the edge of the culture waste bottle). Try to remove as much of the liquid as possible – give it a shake / tap to assist this.
- Resuspend the pellet gently in 33mL RF1 solution by vortexing and/or shaking the tube/bottle. Its OK to be rough with the cells at this stage in the process, but you shouldn’t need to shake or vortex for more than 10 sec or so.
- Incubate on ice for 1 hour, then pellet the bacteria again at 4000rpm, 4°C for 10 minutes.
- Working in the cold room, pour of the supernatant into culture waste. As before, try to remove as much of the residual liquid as possible.
- Resuspend the pellet in 8mL RF2 solution by vortexing or shaking. At this stage, the cells have become more fragile due to the RF1 treatment, so its important not to shake/vortex any longer than ~10 sec (this shouldn’t be necessary).
- . Incubate on ice for 15 minutes. While the cells are incubating, set up all your Eppi tubes (~50) on ice with the lids open, so they are pre-chilled, and ready to receive cells. Label these tubes on top with the strain name before putting them on ice (labelling becomes difcult with cold and wet tubes!). Be careful to only push the tubes only about 2/3 of the way into the ice. If they are pushed in too far (right up to the lip of the tube), you risk getting ice or melted ice (not sterile!) into your cell aliquots
- Working quickly (but still carefully!), aliquot 220 µL of cell suspension into the prechilled Eppi tubes. Once dispensed, close tube lids tightly, and collect all tubes into a bag/box with a clear and prominent label and store it immediately in the -80°C freezer. It’s a good idea to label both the outside of the bag/box AND place a label written on paper inside. Label with the date you made the batch so they can be easily identified if anything goes wrong in testing.
- Test the transformation efciency of the freshly-prepared competent cells using a known amount of a plasmid standard (see protocol for heat shock transformation).
- Streak out a loopful of the cells onto plain LB medium (using either a sample of a frozen aliquot or some residual cells remaining in the large centrifuge tube/ bottle), and incubate at 30°C for three days to allow any of the common types of contaminants (e.g. Staphylococcus) to grow. This is to check the purity of the cell stock. The streak-plate should look completely uniform, with colonies of only one type (E.coli), and no heterogeneity in the initial patch or the streaklines which would indicate a mixture of bacteria is present.
- If either of your tests fail (e.g. you do not get transformed cells, you get heterogeneity on your streak plate) dispose of your batch of competent cells.
Competent Cell Solutions
RF1 Solution
- 500mL Reverse Osmosis Water (ROW)
- 6.06g RbCl (final conc. 100mM)
- 4.95g MnCl2 (final conc. 50mM)
- 1.47g K Acetate (final conc. 30mM)
- 0.74g CaCl2 (final conc. 10mM)
- 75g Glycerol (15% w/v)
RF2 Solution
- 500mL Reverse Osmosis Water (ROW)
- 1.06g MOPS (final conc. 10mM)
- 0.6g RbCl (final conc. 10mM)
- CaCl2 (final conc. 75mM)
- 75g Glycerol (15% w/v)
For both solutions: add all the ingredients to 300mL of the 500mL ROW, then make up the final volume using a measuring cylinder.
For RF1: After adding all the ingredients, adjust pH to 5.8 with conc. acetic acid. pH will change very quickly and only requires ~ 10µL of acetic acid. Sterilize by filtration into an autoclaved media bottle.
For RF2: After adding all the ingredients, adjust pH to 6.8 with NaOH or HCl, as appropriate. Sterilize by filtration into an autoclaved media bottle.
Reagents
- Suspension Buffer: 1x TE (10mM Tris, 1mM EDTA pH 8).
- Ni+ bead suspension in 20% EtOH.
- Buffer A: 20 mM Tris, pH 8
- Buffer B: 20 mM Tris, 500 mM NaCl, 5 mM imidazole, 0.1% Triton X-100, pH 8
- Buffer C: 20 mM Tris, 500 mM NaCl, 150 mM imidazole, 0.1% Triton X-100, pH 8
- Buffer D: 50 mM Tris-HCl (pH 7.5), 20% glycerol, 0.1 mM EDTA, 1 mM DTT (Add the DTT after autoclaving.)
Growing and collecting cell lysate
- Inoculate 500mL of LB containing the appropriate antibiotics with your cells contain your HIS tagged constructs. A large loopful of cells from a plate resuspended in 1mL of plain LB or an overnight 5mL seed culture of your cells should be enough of an inoculant.
- If you are using a T7 polymerase containing cell line (BL21 etc) and vector with a HIS tagged sequence under a T7 promoter incubate your cultures at 37oC with shaking (200rpm) until OD600 0.4-0.6 then induce with IPTG (1mM)
- Leave the cultures to shake (200rpm) at your desired protein expression temperature overnight. Deciding the ideal expression conditions may require sampling of cell lysates from smaller cultures under various conditions using SDS PAGE. Good working temperatures to test expression include 15oC, Room Temp (21oC -25oC), 30oC and 37oC. See ‘SDS PAGE of Cell Lysates’ protocol for lysate testing methods.
- Centrifuge the cultures (3900g, 15min, room temp) and pour off the supernatant.
- Resuspend cell pellet in 25mL of Buffer A by vortexing.
- Centrifuge once more (as above) and pour off the supernatant.
- Resuspend the pellet in 5mL of Suspension Buffer.
- Freeze suspended cells at -30 for 30 minutes.
- Thaw suspension and add 30uL of 1x protease inhibitor cocktail.
- Transfer 500uL aliquots of the suspension to sterile bead beating tubes (containing 2 large beads (5mm) two small scoops of medium beads (60 mesh) and two small scoops of small beads (75-150 microns).
- Bead beat tubes for 30 seconds then move tubes to ice for 2 minutes. if results suggest poor lysis repeat this step.
- Centrifuge bead beating tubes at 15,000g for 5 minutes (preferably at 4oC)
- Move tubes back to ice immediately
- Combine the supernatant from all bead beating tubes into one prechilled 15mL Falcon tube and keep on ice (be careful not to collect any glass beads)
- Collect 30uL of your now combined cell lysate in a 1.5mL eppendorf labelled ‘Cell Extract’ and freeze at -30oC for future SDS loading (see ‘SDS PAGE of Cell Lysate’ protocol). Proceed with HIS tag purification with the remainder of your cell lysate.
- Pack a 10mL syringe tightly with 1-1.5cm of non absorbent cotton wool (plunge hard to compact the cotton wool at the base of the tube.
- Suspend your column over ice and remove the plunger and add 2-3mL of your Ni+ bead slurry to the column
- Allow the beads to settle then slowly add 10mL of Buffer B to the column, allowing the solution to move through and out the bottom of the column into a waste beaker. Ensure the column does not dry out however.
- Add your cell lysate solution and allow it to move through the column. Collect the flow through in a 15mL falcon tube labelled ‘Flow sample’ on the ice below.
- Add 20mL of Buffer B to the column gently to flush out any unbound protein in the column. Collect the flow through in a 50mL falcon labelled ‘Wash sample’
- Add 5mL of Buffer C to the column and collect the flow through.
- Repeat step 6 twice more collecting the flow through in separate tubes each time.
- Pipette 30uL of each fraction to new individual 1.5mL eppendorf tubes for SDS PAGE analysis (See ‘SDS PAGE of Cell Lysates’) (These can be frozen at -30oC). If required you store your protein for immediate use at 4oC or freeze them at -30oC. Note that the activity of your enzymes may be affected by long term storage of your enzyme at these conditions.
- Prepare your ‘Eluted’ fractions for dialysis against 1L of Buffer D overnight at 4oC with a low speed magnetic stirrer.
Preparing Cell Lysates
- Prepare 1.5-2mL bead beating tubes using the appropriate amount of glass beads. Two 5mm beads (large) and two small scoops of both 60 mesh beads (medium) and 75-150 micron beads (small) should be enough for one tube.
- Sterilize by autoclaving.
- Pellet your cells from broth or scrape off a plate and resuspend in 500uL of 1xTE (10mM Tris, 1mM EDTA pH 8) and 5uL of a 1x protease inhibitor cocktail (This will only extract soluble protein from your cells). How much culture to use is tricky to know, more concentrated is usually better, but too many cells and suspension may be gloopy and hard to work with. Use more bead beating tubes if working with larger cultures to avoid this problem. The pellet from 2mL of broth culture or a large loopful from a plate is a good starting point for 1 bead beating tube.
- Take your tubes over to the bead beater and unscrew the tube-holder.
- Space your tubes around the disc (make sure the caps are on tight and the sides of the tubes are labelled!), replace the cover as tight as you can.
- Beat for 30 seconds. If results suggest poor lysis, or if you are working with Gram-positive bacteria or yeast etc, you will need to do multiple cycles of beating – in these cases, chill your tubes on ice for 1 min between beatings. You may need 5 x 30 sec beatings to fully lyse tough cells like mycobacteria.
- Centrifuge at ~10,000g for 5 minutes ideally at 4 °C to remove beads and any remaining whole cells. If you must centrifuge at room temperature keep lysates on ice for 5 minutes before centrifuging at ~15,000g for 2 minutes.
- Retain the supernatant and keep on ice.
- Check the absorbance of supernatants using a Nanodrop (Protein setting or 280nm wavelength) to determine mg/mL (Abs of 1 = 1mg/mL protein). Ideally we want around 5mg/mL for nice PAGE gels (between 2 and 10mg/mL may be OK). If It’s less than 2mg/mL either repeat with more cells or bead beat for more cycles. If it’s more than 10mg/mL dilute down to 5mg/mL. Keep in mind the Nanodrop reading is only an estimate for protein concentration, since there is other junk in the cell extract that will interfere. For some applications like enzyme assays, you may need to use a different protein quantitation methods.
- Using your nanodrop estimates calculate how much of each sample you’ll need to get a constant amount of protein in each lane on your PAGE gel. It’s very important to try and match the total protein in each lane so that we can compare e.g induced vs uninduced samples etc. between 25-50ug per lane usually works out well. For example if your sample is 5mg/mL that’s equal to 5mg/uL so for a 40ug lane you would need 8uL of your sample.
- Add an appropriate amount of SDS-PAGE sample buffer to your sample (final conc: 4% SDS, 20% glycerol, 0.002% Bromophenol Blue, 5% fresh B-mercaptoethanol, 62.5mM Tris-HCL pH6.8). i.e for the 8uL sample above you would need ~8uL of a 2x SDS-PAGE sample buffer stock solution. Make up 50% more sample + SDS sample buffer than you actually need to prevent loss in the next step.
- Heat samples at 100°C for 3min then allow them to cool down before loading them on a gel.
To examine the insoluble fraction of your cells resuspend the culture in 500uL of SDS sample buffer (4% SDS, 20% glycerol, 0.002% Bromophenol Blue, 5% fresh B-mercaptoethanol, 62.5mM Tris-HCL pH6.8).
Preparing your SDS PAGE gel.
- Prepare your 1x SDS running buffer first.
- SDS
- Tris base
- Glycine
- MilliQ
- 10g
- 30.3g
- 144g
- To 1L
- Take your prepared gel cassettes (precast or homecast) and remove the well comb, holders (home cast) and/or bottom tape (precast).
- Put them into the housing with the short plate sides facing inwards and then clip into place. It’s a good idea to mark the location of the wells with a permanent marker at this stage to make it easier to see where the wells are later. If running multiple gels it can also be good to mark the top of the longer plate in a way to help distinguish which gels contain what samples.
- Put the assembly into the clear tank and check that the top part makes contact with the terminals.
- Fill the space between the gels with 1x SDS PAGE running buffer and fill the tank up to the 2 or 4 gel line depending on how many gels you are running (if concerned about overheating lower weight proteins <25kDa, always fill the tank to the 4 gel line with 1x SDS running buffer). If you only have one gel, use the ‘buffer dam’ on the second side so you get a distinct pool of buffer at the top (an empty gel cassette, tape not removed works well here).
- Load 5uL of protein ladder standard to one lane (don’t need to boil or add buffer to the ladder). Add the appropriate amount of each of your samples in their respective lanes. Make sure to mark down your loading order! Ensure the gel is asymmetrical as it might get flipped over later during imaging.
- Run the gel at 200V for 40 minutes or until the dye front reaches the bottom of the gel. Running the gel at a lower voltage for longer may improve the resolution of your gel. Some housings leak so check the gel every now and again and ensure that there is enough buffer between the gels, it may need to be topped up.
- When the dye has reached the bottom of the gel, stop and disassemble the housing. Used buffer can go down the sink.
10x SDS running buffer.
Reagent
Amount
pH to 8.5
Visualising SDS PAGE
- Carefully pry open the plates using a wedge tool or flat head screwdriver. Carefully pry off the gel using gloved fingers or sterile tweezers. It will be reasonably tough but may still tear or break so be careful.
- Place it in a plastic dish and add enough distilled water to suspend the gel slightly.
- Next...
- If you are using a Biorad pre-cast gel with stain free imaging capability move directly to an imager and proceed with the stain-free UV imaging protocol for that machine, transferring the gel carefully to the imaging plate. If you are checking for a protein of interest ensure it contains tryptophan in order to visualize it using Biorad’s stain free technology.
- If you are not using a stain free gel proceed with coomassie staining by pour off the water into a waste container and adding coomassie blue solution to cover.
- Put some gladwrap over the top and put on a gentle rocker for half an hour.
- Pour the stain into a methanol waste bottle and cover with high-destain solution (40% MeOH 10% Acetic Acid) and return, covered, to the rocker for another half an hour.
- Pour off into the methanol waste and replace with low-destain solution (10% MeOH, 10% Acetic Acid) and rock, covered overnight. The high-destain step may be omitted and done the next morning if there is still a lot of background stain. Don’t worry about a bit of background stain, but less is better as it will enhance the contrast.
- Gels can be stored in the cold room between two sheets of clear plastic sealed together with tape for later reference or mass-spec band excision. You can annotate the plastic directly but be aware the gel can shift in the plastic over time if not kept flat. If you want to discard the gel, this must be treated as contaminated waste, due to possible acrylamide monomer still present... discard into your chemical waste bag.
Preparing dialysis tubing.
- Cut tubing into pieces of convenient length (10-20cm, this will depend on your sample volume).
- Boil the tubing in 10minutes in a large volume of 2% (w/v) sodium bicarbonate and 1mM EDTA (ph8.0)
- Rinse the tubing with distilled H20
- Boil the tubing for 10 minutes in 1mM EDTA (ph8.0)
- Allow the tubing to cool then store it in 4oC Be sure that the tubing is always submerged
Preparing your protein samples for dialysis.
- Handle with fresh gloves only.
- Wash the tubing inside and out with sterilized distilled water.
- Double over one end of the tubing and clip it.
- Pipette your protein sample in through the open end.
- Double over the open end and clip that.
- Submerge your tubing submerged in the solution you wish to dialyse against.
- Leave overnight at 4oC stirring slowly (magnetic stirrer).
General Notes
This protocol procceds after Digestion & Ligation protocol. This protocol must be done is asceptic condition and will test a users asceptic technique.
- Remove one or more aliquots (as required) of chemically competent cells of your E.coli strain from the -80°C freezer. Thaw the cells e.g. by rubbing them in your hands or put them briefly in a 37°C waterbath, but don’t let them stay warm! As soon as they are thawed, put them onto ice.
- Divide the cells into the appropriate number of 50 µl aliquots in separate Eppi tubes on ice. Add your DNA samples to each tube; you can use up to ~10 ul of ligation mixture or plasmid here, but note that typically 3 µl of ligation mix or 1 µl of plasmid would be standard.
- Make sure you include both a positive control and a negative control in the transformation experiment. The positive control should be 1 µl of a plasmid with the correct antibiotic resistance (same resistance as the plasmid used for the ligation), and should also be a plasmid stock that you know is in good condition (based on agarose gel).The negative control is simply no DNA added.
- Put the cells into a foam ‘floatie’ and put on ice. Ensure at least the bottom half of the tube (approx 2 cm) is embedded in the ice, don’t just rest them on top of the ice. Allow the cell/DNA mixtures to incubate on ice for 15-30 min.
- Take your esky of ice over to the 42°C waterbath or 42°C heat block. Put the floatie into the waterbath. Allow 45 seconds for heat shock. (Plus or minus 10 seconds, this needs to be exact!). Then transfer the floatie straight back onto ice (embed into ice, as above, don’t just rest on top).
- Allow transformation mixtures to sit for 2 min on ice, then add 1 ml sterile LB broth to each tube. You can also use more fancy media (e.g. SOC or SOB), but there is not that much difference.
- Incubate on 37°C shaker for 1 hour. Put the tubes horizontal so they get good shaking action. eg. put the tubes laying flat on the shaker platform and masking-tape into place. Make sure the lids are tight! You can incubate without shaking, and you can incubate for less time (30 min), but it won’t work as well in these cases.
- Label the LB-antibiotic plates before starting the next bit; you need two plates for each ligation condition or plasmid type, since we will plate out two different cell concentrations of each to ensure we get countable/pickable numbers of colonies. Double check the plates to ensure you are using the correct type of antibiotic(s) for the type of plasmid(s) you are using.
- Pipette 100 µl of the first cell suspension onto one LB-antibiotic plate (label ‘100 µl’ in addition to other info). Sterilise the glass spreader with ethanol and flame (CAUTION! READ THE SOP FIRST!), and spread the cells around the plate with the spreader. Do this by pushing the spreader with a backand-forth motion, while turning the plate around in a circular motion. Be careful not to touch the spreader on your fingers! Keep spreading for approx 10 seconds.
- Spread 100 µl of the remaining samples, each onto a separate, appropriately-labelled plate.
- Centrifuge all the tubes at ~15,000 rpm for 1 minute in a micro-centrifuge. Pour off most of the supernatant into culture waste (being careful not to touch the tubes on the edge of the culture waste bottle). Leave a little bit of liquid behind (about one or two drops).
- Vortex the cells in the remaining liquid for about 10 seconds, until they are not sticking to tube anymore, and you have a nice smooth, even, cell suspension.
- Pipette the cells from the first cell suspension onto the appropriate pre-labelled LB-antibiotic plate (label with ‘pellet’ in addition to other info), spread plate as above. Repeat for the remaining samples
- Incubate all plates at 37°C overnight. Note that for some plasmids and ligations, it may be beneficial to instead try room temp for 2-3 days – this lowers the copy number of pUC type plasmids, and is useful to allow retrieval of clones that might be toxic to the host.
- When examining your plates, first check your controls. The positive control should have thousands of colonies, perhaps even a confluent lawn of growth, especially on the ‘pellet’ plate. the negative control should have no colonies at all. If you don’t see these results with the controls, anything you see on your experimental plates is questionable. Common problems and their interpretation are summarised in the Table below. Detailed troubleshooting of the different ligation controls and their results is described in a different protocol. “Mix up of labelling” can cause MANY problems – be super careful with your labelling ! (do this BEFORE starting the hands-on bit of the procedure, and double-check everything)
If the plates are properly dried, you should feel the spreader start to ‘stick’ to the agar, this means the liquid has been drawn into the agar. If this doesn’t happen after ~20 seconds, stop spreading, but next time, dry plates for longer! If the plates are incubated with a lot of liquid still on them, you may not get nice discrete colonies (the cells will swim around in the liquid, making a mess).
| Problem | Interpretation/solution |
|---|---|
| Lots of growth of the negative control (thousands of colonies or lawn) |
|
| Some growth on the negative control (a few colonies) |
|
| No growth or very little growth on the positive control plate |
|
A resting cell suspension contains cells with induced enzymes capable of doing metabolic reactions, but which are suspended in non-growth medium so they can’t make new proteins or cells. Under these non-growth conditions, substrate depletion initially follows zero-order kinetics (linear), allowing simple calculation of apparent specific activities (k) from a straight line fit through the substrate depletion data. Resting cell suspensions are also useful because they allow concentration of cells from a large culture volume into a small volume, thus boosting the reaction rates, and sometimes making reactions measurable that can’t be seen in growing cultures.
Protocol
- Choose a growth medium that you know will give you the enzyme activity of interest.
- Determine what volume of culture you need to grow to see the activity. If the activity is weak, you may need to grow 500 ml of cells to make a 10 ml resting cell suspension (50-fold concentration factor), but a strong enzyme activity may be visualised in the same volume of culture as resting cell suspension. If the activity of your assay is too high, the cells will consume the substrate too quickly for you to get good quantitative data. If the activity is too low, a linear substrate depletion curve may not be apparent (or you may have to measure depletion over a very long period of time). If you incubate “resting cells” for too long, funny things can happen, for example the cells can uninduce, or other enzymes that you don’t want may switch on.
- Think about what controls you need. Will the negative control consist of the same strain grown in non-inducing medium? Or will it be an E. coli with vector-only, not the cloned gene of interest? Is there a positive control you can use?
- Think about replicates. Do you need to grow multiple cultures on the same day? Or should you stagger these over different days? Is it OK to make replicate cell suspensions from the same parent culture...is that still a true ‘replicate’?
- Think about the need for sterility. Usually, once the culture has grown, we are in the realm of biochemistry rather than microbiology, so it’s OK to use clean but non-sterile glassware and plasticware and solutions etc in a resting cell assay. However, if you want to keep the solutions for a long time, or if the assay runs for a long time, or uses very rich ingredients (for example glucose), you may need to be more careful.
- Plan the experimental protocol and timing. Make sure you have all the required media, consumables (especially sterile centrifuge tubes), and required solutions. Make sure you are familiar with the way the cultures grow and the analysis methods required before growing up lots of cultures or large volumes of cultures – a quick preliminary experiment is often invaluable to get the feel for an assay before committing a large amount of resources. How long do the cultures take to grow? How long does the assay take to run? Will you need to be in the lab at 3 am on Sunday morning? (if so, maybe rethink when you will inoculate the culture!)
- Inoculate the cultures from a fresh inoculum (either plate or broth) and grow to mid-log phase. “Mid-log” (or mid-exponential) means different things for different bacteria and different media, but this will typically be around OD600 = 0.5. If in doubt, let the cultures grow to full density, then assume that mid-exponential is about half of that OD600. In some unusual cases, you may need to grow cultures into stationary phase or death phase or even lag phase, if the enzyme you are interested in is induced at those times, but for most common cellular enzymes, you will get best activity when the cells are happy and healthy in mid-log.
- If you are using wild-type cells which are induced by the growth conditions themselves, skip ahead to the next step. If you are using recombinant cells, where the inducer is artificial (for example IPTG, arabinose, tetracycline), add the inducer now at mid-log phase. Allow to induce for 4- 48 hours depending on the strain, conditions, and enzyme of interest. For E. coli cells and standard induction systems, 4 hr is usually enough, but for mycobacteria or other slow-growers or for unusual enzymes or cold incubations, induction may require a lot longer. Monitoring OD600 during induction is a good idea to get a sense of how induction impacts on growth.
- Harvest the cultures by centrifugation. Bacteria can be pelleted at quite low g forces and times, e.g. 3000 g for 10 min is fine in most cases. Check the conversion from g to rpm if in doubt – these are usually in the same range (3000 g is about 4000 rpm in a ‘Centaur’ machine 50 ml rotor), but different rotors have different conversion factors. Some cells like mycobacteria may require additives (0.05% Tween-80) to centrifuge properly, if these aren’t already in the medium (and these additives need to be maintained in both wash buffers and the assay buffer). Cells pellet easier at higher salt concentrations; pure water is quite difficult to pellet cells in.
- WASH THE CELLS! This is important, and often overlooked. If there are significant amounts of rich growth medium (e.g LB) present in your resting cell assay the cells can grow, which will interfere with determining rates of reaction. Washing is done by centrifuging and then resuspending in a buffer like KP (10 mM K2HPO4, pH 7) or MOPS or PBS etc. To resuspend cells, its best to flick or vortex or shake the tube/bottle rather than using a pipette – clumpy cells in particular will stick to tips and be lost. You can shake vigorously, as this won’t hurt most bacteria.
- More cell washing hints: Don’t forget to add Tween to wash solutions for clumpy cells like mycobacteria (add Tween 80 after to wash solutions after autoclaving). Don’t use plain water as a wash buffer, and ideally use a wash buffer that is the same as the base components in your assay buffer in the later steps. Typically, we would do two wash steps in a convenient volume, for example 20 ml buffer in a 50 ml Falcon tube for culture pellets from a few hundred ml of culture, or 0.5 ml buffer in an Eppi tube for pellets from a few ml of culture.
- What temperature to use for centrifugation and wash steps? This is a bit controversial. As a starting point, it is best to use a cold centrifuge for both these steps, since this should “stop the cells in their tracks” and minimise any changes their physiology. However, in some cases, the cold treatment may shock the cells, and they may respond better to room temperature centrifugation and storage during setup. In all cases though, avoid them getting hot! (>30 C)
- After washing, resuspend the cells in a small volume of the same buffer and keep on ice while you are preparing the assays. Though see above, sometimes room temp might be better. By “small volume” here, we mean like 100 x less than the original culture volume. This gives us the greatest flexibility for assay setup. It’s easy to dilute cells further, but annoying to concentrate them. Measure the OD600 of the cell suspension, using an appropriate dilution if the spectro reading is more than 1. Calculate back to figure out the actual OD600 of the suspension.
- Set up the assay reactions in appropriate types and sizes of tubes. If the substrate is volatile, you will need gas-tight bottles (serum bottles), which are crimp-sealed. Otherwise, you can use McCartney bottles, Eppis or Falcon tubes. If using McCartney bottles or other recycled glassware, wash these with RO water before using, to remove traces of detergent. If the cells need aeration, ensure sufficient headspace (say 75% of sealed bottle volume), or incubate loosely capped. What is your target OD600? (how much activity are you expecting?) – this will determine the volume of the assay, or more specifically, the ratio of the assay volume to the original culture volume. How many samples do you need to take, and what volume are these? (this also impacts the assay volume)
- Set up a master mix, which contains all the ingredients for your assay (buffer and substrate, maybe cofactors or stabilisers). Aliquot this mix out into your individual tubes/bottles, leaving room to add cells to bring the assays to their full volume. Pre-incubate the master mix at the assay temperature for at least 10 min before starting, then add cells.
- Immediately take a time zero sample. If this is a liquid sample, centrifuge this immediately in a cold centrifuge, and transfer the supernatant to a fresh tube for immediate assay, or for freezing and later analysis. Label tubes first! Keep in mind that the sampling process itself can interfere with the assay procedure. For example, if you are too slow, the temperature of the incubating shaker won’t be maintained in the cultures during sampling. If you take too much of the initial liquid volume out via sampling, this will change the way volatiles partition in the bottle.
- Incubate the tubes with shaking at the required temperature. Keep in mind the need for good mixing! For smaller tubes or bottles, it is often better to incubate them horizontally to get good mixing action. This is even more important when assaying volatile substrates, where movement between liquid and gas phases happens. Sample at appropriate intervals – this may be between 1 minute to 1 hour, depending on activity levels.
- At end of assay, discard cell suspension to autoclave waste, and if using glassware, rinse this out with water straight away (also discard to autoclave waste), so that cells don’t dry onto the glass. If using volatile substrates, you can leave bottles open in fume hood overnight before discarding culture, to allow volatiles to disperse.
- After doing your assays, calculate the rate of substrate disappearance or product appearance; this will usually be in nanomoles per minute. This can then be converted into an “apparent specific activity” (k) by dividing by the protein content of the assay (mg). We can use previous standard curves to convert OD600 to amount of protein:
- For E.coli grown in LB: protein (g) = 0.21 x wet weight of biomass, and biomass (g/L)= OD600/0.534, therefore protein (g/L) = 0.393 x OD600 or [protein (μg/ml) = 393 x OD600] (Kangwa et al 2015)
- For Pseudomonas putida grown in LB: [protein (μg/ml) = 69 x OD600] (Mai Anh Ly, PhD thesis)
- For Mycobacterium smegmatis grown in MSM-glucose: [protein (μg/ml) = 99 x OD600] (Mai Anh Ly, PhD thesis)
- For Mycobacterium chubuense grown in MSM-acetate: [[protein (μg/ml) = 112 x OD600] (Laura Nolan, Hons thesis)
General Notes
One of, if not the most important protocols of synthetic biology. It’s an essential protocol to memorise and master as most if not all your experiments hinges on the success of this particular method.
A few notes to begin with:
- Quantify the DNA concentration of your chosen plasmid and insert DNA (usually a G-block or a PCR product). This can be done through nanodrop, however keep in mind that the nanodrop DNA concentration result of your plasmid is rough estimate and sometimes incorrect. Therefore you may have to run you’re a digest of your plasmid on gel and estimate relative DNA concentration to the DNA ladder standard.
- Check which restriction buffer is appropriate. For double digests, it’s OK to use a buffer which gives 100% activity for one enzyme, and 75% activity of the other, but lower than this is not good.
- Check the appropriate digest temperature, its usually 37°C, but not always. Search online or on the stock restriction enzyme tube for the proper temperature for digestion.
Digestion of Vector and DNA insert
THE GOLDEN RATIO FOR BETWEEN PLASMID AND DNA INSERT*
~3:1 ratio between Insert DNA Mass: Vector DNA Mass
*Of course this is influenced by the relative sizes of your vector and insert DNA and you may have to tune this ratio to suite your needs but you want to have AT LEAST a higher ratio of insert DNA to your Vector DNA e.g. ~100 ng of DNA insert: ~25 ng of plasmid = 4:1
Plasmid Digestion – Plasmid Master Mix
This particularly useful if you are cloning several genes into the same vector/plasmid.
- Retrieve 10 x restriction buffer from freezer, thaw completely, and vortex to mix. The same tube of buffer can be used many times, if you are careful with your aseptic technique.
- Retrieve the plasmid from the freezer, allow to thaw, (e.g. in 37°C waterbath, or rub in your hands, or on bench etc), then put it on ice when it is thawed. It’s not good to leave the plasmid stock at room temp or above for prolonged periods or it may degrade due to traces of nucleases.
- Retrieve the restriction enzyme(s) from the freezer, put IMMEDIATELY on ice. These are heat-sensitive and you need to look after them. Do not leave them at room temp. Keep on ice while setting up the reaction, then immediately put back in freezer. These don need to be thawed, they are in a glycerol solution which doesn’t freeze at -20°C.
- Calculate an appropriate mass of Vector DNA for the plasmid digest master mix. For the example below, lets calculate how much volume of plasmid you need to use to get 250 ng – call this ‘X’, and calculate how much water to add to the digest; this is (100 – 10 – 2 – X) – call this ‘Y’.
- Label your tube(s), then set up the digest(s) by adding the ingredients in the following order. Make sure you use excellent aseptic technique, and change tips every time. It’s OK to set this up at room temp, the reaction tube doesn’t have to be on ice. Mix restriction enzyme buffer + DNA + MQ water mixture by pipetting up and down and centrifuging down before addition of the RE. Enzymes like to be in their correct buffer conditions to function.
- 10 µl* of 10x restriction buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion).
- ‘Y’ µl of sterile MQ water
- ‘X’ µl of plasmid DNA
- 2 µl of (each) restriction enzyme
Total Volume: 100 µl
- Mix by pipetting up and down or by flicking, then tap on bench to get liquid to bottom of tube.
- Incubate at correct temperature for approx. 2 hours. If it’s a High fidelity RE, 1hr is more than enough. (1 - 4 hr is OK, but overnight digest is too long; this can lead to ‘raggedy ends’ of the plasmid even if it looks OK on a gel; this is due to non-specific nuclease activity).
NOTE: 100 µl seems like a large volume to use, but this helps dilute any impurities in the plasmid prep. Increasing the total volume of the digest and/or reducing the volume of plasmid added often help to improve the quality of a poor digest. If the digested plasmid is for the purpose of ligation, we can use a large volume, since it will be column-purified & concentrated anyway.
Digestion of Insert DNA
This will be most likely be G-block from oligonucleotide manufacturer or PCR product with the appropriate restriction enzyme sites at the ends. As said before, you want to have a higher ratio of insert DNA to vector DNA. This digestion can be concurrently performed with the digestion of the plasmid.
- Calculate an appropriate mass of Insert DNA. For the example below, lets calculate how much volume of Insert you need to use to get 100 ng – call this A, and calculate how much water to add to the digest; this is (15-1.5-1-A) – call this B.
- Retrieve 10 x restriction buffer from freezer, thaw completely, and vortex to mix. The same tube of buffer can be used many times, if you are careful with your aseptic technique.
- Retrieve the restriction enzyme(s) from the freezer, put IMMEDIATELY on ice. These are heat-sensitive and you need to look after them. Do not leave them at room temp. Keep on ice while setting up the reaction, then immediately put back in freezer. These don’t need to be thawed, they are in a glycerol solution which doesn’t freeze at -20°C.
- Retrieve the Insert DNA from the freezer, allow to thaw, (e.g. in 37°C waterbath, or rub in your hands, or on bench etc), then put it on ice when it is thawed. It’s not good to leave the Insert at room temp or above for prolonged periods or it may degrade due to traces of nucleases.
-
Label your tube(s). Set up digest by adding ingredients in the following order. Make sure you use excellent aseptic technique, and change tips every time. It’s OK to set this up at room temp. Mix restriction enzyme buffer + DNA + MQ water mixture by pipetting up and down and centrifuging down before addition of the RE. Enzymes like to be in their correct buffer conditions to function.
- 1.5 µL* of 10x restriction buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion).
- ‘B’ µl of sterile MQ water
- ‘A’ µl of insert DNA
- 1 µl of (each) restriction enzyme
Total Volume: 15 µl*
*This volume is quite flexible. Adjust if needed.
- Incubate at correct temperature for approx. 2 hours. If it’s a High fidelity RE, 1hr is more than enough. (1 - 4 hr is OK, but overnight digest is too long; this can lead to ‘raggedy ends’ of the plasmid even if it looks OK on a gel; this is due to non-specific nuclease activity)
- Purify both vector and insert DNA using Qiaquick columns (or similar), elute in 15 µl EB.
Ligation
- Retrieve one 10x ligase buffer aliquot from the freezer. This should be a small amount of buffer (e.g. 10 µl) in a small generic tube, not the large tube from the manufacturer with ~1 ml of buffer Ligase buffer (unlike restriction buffer) cannot be repeatedly frozen and thawed, it starts to ‘die’ after even one freeze/thaw cycle.
- Retrieve the T4 ligase enzyme from the -20°C freezer and put IMMEDIATELY on ice. This reagent is VERY heat sensitive, and must be handled with care.
- Put your labelled tube(s) on ice, then set up the ligase reaction in this tube on ice. Be careful not to get ice or melted ice in the tube - this is not sterile! Add reagents in this order, change tips each time. Mix restriction enzyme buffer + DNA + MQ water mixture by pipetting up and down and centrifuging down before addition of the RE. Enzymes like to be in their correct buffer conditions to function.
- 2 µl of 10 x ligase buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion)
- 8 µl purified insert DNA
- 8 µl purified plasmid DNA
- 2 µl of T4 DNA ligase enzyme Total Volume: 20 µL
- Mix by flicking briefly then incubate either 4°C overnight or room temp for 1 hour. Return the ligase enzyme immediately to -20°C freezer. Throw out any unused thawed ligase buffer.
If you want to don’t want to deal with column DNA purification you can use:
Cloning/ligation protocol using heat-killed digests
This protocol is especially useful for ligations involving very small or large fragments which do not get retained very well during column purifications. It’s also worth considering when you don’t have much DNA in your sample and want to minimise losing it. Note that some restriction enzymes are not heat-killable (e.g. PstI-HF and BamHI) so check this first! (NEB website).
The thermostat on a heat-block isn’t very accurate and usually the block is 5-10°C below the set point. Check the thermometer before you start. Putting the thermometer in an Eppi tube of water in the block will give you the closest idea of what your sample is actually being heated to.
A downside of this protocol is that it doesn’t remove small offcut bits of DNA like column purification does. If for example you are digesting a PCR product that isn’t too small, or a two sites in a plasmid with only a few bases between them, it would be better to column purify these digests to reduce the chance of the small fragment re-ligating back in.
- While waiting for vector and DNA insert digests, turn on the heating block and set to whichever is the highest heat kill temperature of the two restriction enzymes. When digest is complete, place the tube with DNA into the heating block for 20 min. Turn off heat block! Chill DNA on ice for 5 min.
- Retrieve one 10x ligase buffer aliquot from the freezer. This should be a small amount of buffer (e.g. 10 µl) in a small generic tube, not the large tube from the manufacturer with ~1 ml of buffer Ligase buffer (unlike restriction buffer) cannot be repeatedly frozen and thawed, it starts to ‘die’ after even one freeze/thaw cycle.
- Retrieve the T4 ligase enzyme from the -20°C freezer and put IMMEDIATELY on ice. This reagent is VERY heat sensitive, and must be handled with care.
- Put your labelled tube(s) on ice, then set up the ligase reaction in this tube on ice. Be careful not to get ice or melted ice in the tube - this is not sterile! Add reagents in this order, change tips each time:
- 2.3 µl of 10 x ligase buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion).
- 10 µl purified insert DNA
- 10 µl purified plasmid DNA
- 1 µl of T4 DNA ligase enzyme Total Volume: 23.3 µl
- Give the tube a quick spin to get all the liquid to the bottom. Ligation can be done at room temperature for 30 min on your bench or overnight in the cold room.
Proceed with Transformation Protocols!
Special Mentions
This protocol was provided to the iGem USYD team by Professor Elizabeth Gillam of the School of Chemistry and Molecular Biosciences at the University of Queensland. Thanks for sharing this protocol :D!
Day 1:
Transform DH5aF'IQTM cells with both the P450 expression plasmid and pGro7, using either electroporation or ultracompetent cells generated by the Inoue method (see separate protocol sheet). Alternatively, DH5aF'IQTM pre-transformed with pGro7 can be stored as a glycerol stock then transformed freshly with the relevant P450 expression plasmid. Select by growth overnight at 37°C on LBglu/amp/cam plates.
Day 2:
- Put on starter cultures: LBglu/amp/cam of appropriate volume (eg. ~3 ml for 0.5 ml to seed 50ml bulk culture) inoculated from frozen transformed glycerol stock or PREFERABLY a single colony from a LBglu/amp/cam plate of freshly transformed cells. Grow overnight at 37°C, max shaking speed, to saturation.
- Make up 1L TB base, aliquot and autoclave 50mL TB broths in 500mL flasks in which the bacteria will be cultured. P450 yields are better in small scale cultures than in larger batch cultures.
Day 3:
- Add first lot of additives (PREINDUCTION PHASE) to each culture flask.
- Inoculate these bulk cultures containing TB base and additives for preinduction phase as soon as possible in morning with 1/100 volume of starter cultures ie. 0.5 ml starter culture per 50 ml of expression medium.
- Incubate cultures for five hours at 25°C, shaking at ~ 160-180rpm. This is the time required for the pre-induction phase.
- At t = 5hrs, add second lot of additives (IPTG, ALA, and ARA) to initaite induction of protein expression. Continue to incubate at 25°C, ~160-180 rpm for a further 43 hours (i.e. harvest at t = 48 hours)
Day 5:
- Harvest cultures at t = 48 hours and prepare spheroplasts as normal (see other protocol sheets).
Specific Media
LBglu/amp/cam Media - per 1 litre
- 10g bactotryptone
- 5g Yeast Extract
- 10g NaCl
- 0.3 mL/L 10M NaoOH Autoclaved to sterilise. When cool, add 10ml/l of 20% w/v glucose (filter sterilised). Immediately before use add 1ml/l of 100mg/ml ampicillin and 1 ml/l 20mg/ml chloramphenicol (from methanolic stock).
LBglu/amp/cam agar plates
Prepare as above but add 15g/l bactoagar prior to autoclaving. When media has cooled to ~ 60°C or less, add glucose, amp and CAM and mix well before pouring plates.
TB (Terrific Broth) BASE MEDIA
Per litre
:- 12g bactotryptone
- 24g yeast extract
- 2g bactopeptone
- 1ml 1M NaCl
- 40ml 10% glycerol Make up to 900 ml with milliQ water and autoclave.
Notes
Try to make up TB as soon as possible before use and ALWAYS AUTOCLAVE IMMEDIATELY AFTER PREPARING BASE MEDIA! Unautoclaved TB goes off VERY rapidly! If you absolutely have to store it before sterilising, keep at 4°C for the minimum time possible and observe carefully before sterilising and using. Keep at RT after sterilising. Always aliquot base media into the flasks in which you will culture the bacteria prior to autoclaving. Don't make up a bulk mixture, autoclave, then subaliquot into sterile flasks, since this increases the chance of introducing contamination
ADDITIVES FOR PREINDUCTION PHASE
Immediately before starting induction, add (aseptically, per litre complete TB medium (i.e. adding to 900ml TB base)):
- 100 ml KPi for TB
- 1ml 1M Thiamine *
- 250µl trace element solution *
- 1ml 100mg/ml ampicillin
- 1ml 20mg/ml chloramphenicol
NOTE:
Medium with bactotryptone, yeast extract, bactopeptone, glycerol, KPi for TB, thiamine, trace elements, ampicillin and IPTG is the basic start point for optimising expression - make modifications on this as appropriate for any given trial.
KPi for TB:
- 23.1g KH2PO4
- 125.4g K2HPO4 Make up to 1 litre with milliQ water and autoclave separately. DON'T AUTOCLAVE KPi + TB base media, it will precipate into a crappy mess.
Trace element stock:
- 2.7g FeCl3.6H2O
- 0.2g ZnCl2.4H2O
- 0.2g CoCl2.6H2O
- 0.2g Na2MoO4.2H2O
- .1g CaCl2.2H2O
- 0.1g CuCl2
- 0.05g H3BO3
- 10ml conc. HCl Make up to 100ml with milliQ and autoclave.
These salts preciptate out of solution easily. Therefore needs to directly to medium and mix.
Antibiotics & ETC
Ampicillin: 100mg/ml sterile milliQ water make up immediately before use and filter to sterilise
Chloramphenicol: 20mg/ml in high quality methanol (CAM is very insoluble in water)
1M Thiamine: 337mg / ml sterile milliQ water make up immediately before use and filter to sterilise
Notes:
Thiamine, CAM and Amp can all be added to KPi; mixture then added in one step to medium. e.g. if have a total of 900 ml TB base, autoclave 100 ml of KPi for TB, then filter sterilise 1ml amp, 1 ml CAM and 1ml thiamine into the KPi, then add 5.15 ml of this to 45ml TB base). Add trace elements separately (12.5µl per 50 ml culture). If using previously-opened stocks (eg trace elements) it is safest to filter sterilise an aliquot again immediately before using. Avoid adding supplements to TB until just before inoculation with culture, since this medium is rich and will easily grow (other types microbes.
ADDITIVES FOR INDUCTION
Following the 5hr preincubation, cultures will be supplemented with (per litre)
- 1ml 1M IPTG (1000X stock: typically 50µl/ 50 ml culture)
- 1ml 500mM d-ALA (1000X stock: typically 50µl/50ml culture)
- 10ml 400 mg/ml arabinose (100X stock: typically 500µl/50 ml culture)
500mM d-ALA: 84 mg/ ml milliQ - make up immediately before use and filter to sterilise (d-ALA appears to be essential for certain forms so we add for all).
1M IPTG (Isopropylthiogalactoside) 238.2mg / ml sterile milliQ water make up immediately before use and filter to sterilise
400mg/ml arabinose (ARA) make up immediately before use and filter to sterilise Ara may require brief incubation at 37°C or under a hot water tap to dissolve.
These can be mixed together in the appropriate proportions (1:1:10 IPTG: ALA: Ara) and filter sterilised and added in one lot (e.g. 600µl / 50 ml culture) to each culture flask.
General Notes
Golden Gate Assembly (GGA) is used for cloning one or more inserts into a vector by designing overlapping ends which allow the combination of multiple inserts through the use of a non-palindromic restriction enzyme, BsaI, which is then followed by ligation and transformation.
Pre-cutting of inserts and vector
- Add 1µL 10x Cutsmart (NEB) buffer (1x buffer contains 50 mM Potassium Acetate, 20 mM Tris-acetate, 10 mM Magnesium Acetate, 100 µg/ml BSA, pH 7.9 at 25°C) to a 1.5 mL tube.
- Add 1µL BsaI-HF (200U)
- Combine Equimolar amounts of inserts and vector to the tube.
- Set thermocycler to 37°C and incubate BsaI-HF DNA mixture for 15 minutes
Ligation
- After the BsaI-HF DNA mixture or GGA (Golden Gate Assembled) sample was been incubated, calculate a volume of GGA sample (X) and a volume of MQ water (Y) to make up to a total volume 15µL with:
- 1µL 5x Cutsmart Buffer
- 1.5µL 10mM ATP
- 1µL T4 DNA ligase
- GGA sample 'X' µL
- MQ 'Y' µL Total volume = 15 µL
- Run samples on the following thermocycler program with the folowing program parameters:
- 37°C 2 minutes
- 16°C 2 minutes For 25 cycles
- Then 75°C for 15 minutes
- Then hold (stasis conditions) for 4°C
- After running your GGA sample in the PCR conditions above, add 1uL 5x Cutsmart Buffer, 1uL BsaI, and 3uL MQ water (Total Volume is now 20uL!)
- Set thermocycler to 37°C and incubate BsaI-HF DNA mixture for 15 minutes
- Transform 3-8 uL of GGA sample to a suitable competent cell line.
- And we are done... Why are you still here?
Clean up step - This step isn't neccassary but it's purpose is to digest un-ligated free floating inserts. for
There are many ways to go about this, but this is roughly how we did it. If you are using GFP-family protein or a protein with similar excitation and emission wavelengths, I suggest using the iGEM Measurement protocol. But without further adieu.
After screening your fluorescent colonies by PCR and checking correct sequence of the plasmid by Sanger sequencing. Select your purified colonies, and if your vector is inducible, induce them at time (t=0) or just before chuck at the bad boi (plate reader).
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Culture in 5ml LB broth overnight at 37C 200rpm shaking. Ensure you are also culturing a negative Control (Empty/parental vector) and a positive control such as known fluorescent culture expressing a protein with similar excitation and emission spectrums, or the wild-type version of your expressed fluorescent protein.
- Measure OD of overnight cultures dilute to OD600 0.02 using LB media with appropriate antibiotics and hold on ice (limits growth).
- IN VERY ASCEPTIC CONDITIONS e.g. A biosafety cabinet; In a 96 well plate, aliquot 100 uL of your chilled cultures into replicates for each of your selected test samples and controls. After aliquoting samples, immediately put back the 96 well plate lid and chill on ice to limit growth and prevent contamination.
- Set the plate reader, with at a constant temperature and shaking, to measure OD600 and Fluorescence (your specific excitation and emission wavelengths) every 15 minutes for 18-24hrs.
- Then process your resulting data using your favourite spreadsheet processor! HAVE FUN :D!
Notes: You can typically disregard the first reading as condensation impacts the the OD600 and fluorescence readings at t=0.