Bucci lab methods

Background

I notoriously tweak my protocols a little bit everytime I perform them. It is important to think critically about each step of a protocol as you are doing it. There are always ways to make improvements, whether from an experimental viewpoint or efficiency. A protocol should be a dynamic document. That said, here is a general outline of some protocols I've used in my labwork.

If this is your first time here, please read through the protocols first and make sure you have all of the reagents necessary in order to complete them.

Electrocompetent Cell Preparation

If, like most, you are a person who prepares your electrocompetent cells at cold temperatures because it "increases efficiency"... then stop that voodoo nonsense and read this paper ASAP!! I used to have to go up and down 2 flights of stairs to our cold centrifuges between each washing step, keep my reagents on ice, keep my containers on ice, keep my hands on ice (just kidding), keep the cells as cold as possible, because the cells are fragile and they might die! Or something like that. None of that appears to be necessary. I really applaud the group in the linked paper because this has saved me a lot of time and tedium. Did I mention you should read their paper ASAP!? It is a very robust method. I've performed it successfully with various strains of E. coli, Salmonella, Klebsiella, Proteus, and Pseudomonas.

1. Grow strain to be transformed in 1-5mL LB overnight @ 37C w/ shaking
2. Next day, subculture the O/N 1:100 into 3-5mL of fresh LB media.
3. Put into shaker at max speed (mine only goes to 300rpm) at 37C and wait (depends on growth rate of bug of interest. You want mid exponential phase, so like OD600 = 0.4-0.6. I call it 2-3 hours for E. coli)
4. Centrifuge at ~5000*g for 5 minutes (Quick aside: stop using RPM to denote speed of a centrifuge. Its an essentially a worthless unit for this purpose unless the whole world uses the same rotor. RCF or bust)
5. Discard supernatant (I just invert the tube and flick it a couple times). Resuspend in 1mL of UltraPure Distilled Water (or whatever pure water you trust).
6. Optional: Transfer to a microcentrifuge tube. I do this because I prefer working with these to working with a 15mL conical tube.
7. Repeat steps 4 and 5 a few more times. I usually do 3 resuspensions, using 1mL of water each. On the final supernatant removal step, you may want to try to decant with a pipette, but be careful because your cell pellet is going to be pretty soft/loose.
8. On the final resuspension, only resuspend with 30uL of your Ultrapure water.
9. Add ~1uL of your purified plasmid to the mixture
10. Electroporate (I have the BioRad Micropulser and I use the 0.1cm gap electrocuvettes).
11. Resuspend immediately with 37C SOC (or LB with 0.1% glucose).

Then just proceed as normal (incubate ~1hr with shaking at 37, then put on pre-warmed (37C) selection plates and incubate O/N). It works. It seems too easy. But I promise, it works. Read this if you don't believe me.

Note 1: The paper that you should totally read and cite states that maximum efficiency was achieved at 23C. For me, this is close enough to room temperature. Try the method at whatever ambient temperature you keep your lab and I'll bet you that it works.

Note 2: If I am transforming with something like a ligation mixture or a gibson assembly mixture (i.e., any non-purified plasmid) then I always use commercial electrocompetent cells from NEB.

Transformation selection plates

So, you prepared your room temperature competent cells, how efficient will they be?! Who knows! Follow this method and you'll recover isolated colonies. I always suggest to prepare your selection plates on the same day as you are doing your transformation, and have the plates poured prior to the electroporation step. This will allow enough time for the plates to solidify and be transferred to the 37C incubator, so they are at 37C when the plating takes place. One tip though: don't let your plates dry out completely. This works perfectly if there is a lot of moisture and/or condensation on top of your agar.

1. Prepare your LB agar selection plates with appropriate antibiotic and pour into petri dishes, so that you have 4 plates for every electroporation/transformation that you've done.
2. Once solidified, you can UV them, but I never do. Just transfer them to the 37C at least 45minutes before you plan to use them.
3. Once your transformations have incubated for 1 hour (the part under step 11 above), remove from the 37C and do the following:

~4log scale cfu/mL plating

1. Find your plates with the most moisture. If none have moisture, ok you can add ~25-50uL of pure water on top.
2. Transfer 1uL of the transformed culture onto the moist plate, transfer 10uL to the another plate (with some moisture), and 100uL to the 3rd. Then put the culture back in the 37C incubator.
3. Spread the liquid throughout the plates. I use a glass hockey stick and sterilize with 70% isopropanol (because I like a big orange flame). If you move from low concentration to high concentration, you don't need to sterilize the stick in between plates. Immediately put these plates back into the 37C incubator.
4. Repeat step 1 - 3 for every transformation you are plating.
5. Centrifuge the remaining ~850uL of culture for 5 minutes at 5000*g. Remove the supernatant by pouring it out, but retain ~100uL of liquid.
6. Resuspend the pellet in the leftover supernatant, and transfer to the 4th (final) plate and spread.

That's it. This allows you to plate the entire transformation reaction between 4 plates, where you essentially have a 4 log distribution in cfu/mL, so you can get isolated colonies regardless of your efficiency.

Edit: I find now 4 plates is overkill. If you're a first timer, do the 4plate method. Now I more frequently plate ~10uL on one plate, and centrifuge, discard, resuspend, and plate the remainder on a second plate. These are going to me more like 2 log different but you are likely to find isolated colonies on one or the other.

Microcin-related Research

The Delgado Assay

This assay was originally used in our lab to understand the ability for strains of E. coli to secrete inhibitory molecules (microcins, in our case), and to see if those molecules persist in an environment after the producing strain is killed. Originally adapted from this paper (See Fig 2), though I call it the "Delgado" assay just because I read this paper and the name stuck. I am certain that the assay was first developed by someone else in olden days.

You will notice the mention of 2,2'-dipyridyl (synonym bipyridyl, and referred to here as "DP"). Since my work is mostly with siderophore-microcins, iron limiting conditions appear to play a role in (potentially) both siderophore-microcin production as well as target strain susceptibility. This is a topic that could be further explored in the future.

Briefly, to perform the Delgado Assay:

Day 1:

• Streak fresh plates for isolation from frozen stocks with appropriate antibiotic selection of all microcin producing and target strains.

Day 2:

• Prepare assay LB plates with appropriate induction molecules (e.g. IPTG, L-rhamnose, L-arabinose, etc.), and with 0.2mM DP. I've anecdotally heard that microcin-based inhibition is more pronounced in minimal media compared to rich, but if using an overproduction plasmid, as done in this manuscript, then rich media is ok.
• Take single colonies from the Day 1 streak plates (using a sterile toothpick or a sterile pipette tip) and stab into the assay plates. I generally do this in at least biological duplicates (i.e. 2 separate colonies).
• Incubate overnight at appropriate temperature (37C if looking at enterobacteria like E. coli)
• Create a liquid overnight culture of the target bacterial strain you wish you assay against.

Day 3:

• Inactivate growth of the stab cultures with chloroform vapors. Use glass petri dishes and put them in a ventilation hood (no lids necessary). Place a single piece of filter paper in the center of the glass petri dish (type of filter paper doesn't really matter. Something cheap). Pipette 500uL of chloroform onto the filter paper, and then immediately take your overnight stab culture plate and place upside down over the filter paper, exposing the bacteria to the chloroform vapors, and allow to sit for at least 10 minutes. At this point, the filter paper I use is completely dry and the plastic petri dishes containing bacteria have begun to degrade. A more detailed explanation of this protocol can be found here.
• Transfer plates with lids slightly ajar to a 37C incubator for 10 minutes to allow all chloroform vapors to leave the plate.
• (Optional) Transfer plates to a biosafety hood and treat with UV light for 15-20 minutes (I do this, but I doubt it is necessary).
• Plates are now ready for overlays

To perform the overlay, you will need to create a "soft agar" mixture of agar, overnight target strain culture, fresh media, and DP, to a final volume sufficient for 3mL to be added to each inactivated stab plate:

• Melt 3% agar in the microwave.
• Subculture (1:100) the overnight target strain in a fresh tube of rich broth, and add DP to a final concentration of 0.2mM. Add the 3% agar to a final concentration of 0.65%, and mix briefly by pipetting up and down. You have to work quickly because the bacteria/LB/DP/agar mixture will begin to solidify in likely less than 30 seconds.
• Add 3mL of the soft agar mixture to each plate and allow to dry/solidify (it will only take a few seconds).
• Transfer the plates to the incubator and allow to incubate overnight. If you have strong microcin production, you may begin to see halos within ~6 hours)

Day 4:

• Observe. Take pictures if you see anything particularly interesting!

Microcin Production and Isolation from liquid culture

The Bucci lab has taken advantage of the Maltose binding protein (MBP) fusion method for overproduction and purification of microcins. The protocols and methods we've utilized are all derived from the NEB handbook on MBP fusion purification, linked here. One variation we've employed is 6xHis-tagging the MBP N-terminus, thus allowing us to easily remove it from the mixture after TEV protease digestion. We also purchase a His-tagged TEV protease, from NEB. We do not have our POI His-tagged. Edit: I realize if accessing the site internationally, the link above may not work. A simplified version of the manual is internationally accessible here. Sorry for any inconvenience.

After you're familiar with the standard methods of amylose resin chromatography, starting on page 9 of the NEB manual, here are some notes before starting:

• Adding the iron chelating agent at the moment of subculture often results in very poor growth, especially in BL21 strains. We often wait 1 hour after subculture before adding dipyridyl.
• For induction, we find that very early induction results in the highest yields. Often we wait just ~2 hours after subculture, when the OD600nm is approximately 0.2. Even waiting for higher OD and then having a long induction time seems to result in lower yields.
• It would be wise to continue to improve upon what we've established by developing your own experiments. However, we've settled on commonly using the following protocol:

Day 1:

1. Inoculate 100mL overnight culture for every flask dedicated to overexpression. We usually grow 2L batches, with 100mL overnight dedicated to each 2L, even if they are clones.
2. Keep the cultures cold until you are ready to leave for the day. Then shake them at room temperature, or 16C if you have a cooling shaker, at ~150 rpm. The point is to prevent cultures from being in late exponential phase by morning when you want to subculture.
3. Prepare the LB media, antibiotics, inducer, dipyridyl, and 20% D-glucose. We often do 8L at a time, so make sure you're LB is ready before morning arrives.

Day 2:

1. Pre-warm your LB broth to 37C, if possible.
2. Pour LB broth into the 2L flask, up to approximately 1.8L. Anticipating a final volume of 2L, add 0.1% D-glucose, antibiotics (we use Chloramphenicol @ 25ug/ml and Ampicillin @ 100 ug/ml), and 100mL of inoculum. Then fill to 2L with LB.
3. Shake flasks at 200rpm or higher, 37C.
4. At 1 hour, add dipyridyl to 0.2mM.
5. At 2-3 hours, add the inducer (we use IPTG at 0.4mM)
6. At 5-7 hours after induction, harvest the cells by spinning at 10,000rcf. Remove all supernatant and resuspend cells into ~25mL total volume using column buffer as described in the NEB manual. Store in a 50mL conical tube.
7. Freeze at -20C overnight (or up to a few weeks).

Day 3:

1. Thaw cells in a cold water bath. You want to keep the samples cold for the remainder of the protocol.
2. Think about which fractions (e.g. crude lysate, flow through, wash, pre/post protease elution) you want to save for PAGE analysis or other downstream uses and make sure to put a small amount on ice before moving to the next step.
3. Once thawed, sonicate using a probe sonicator. Try to minimize foaming and use a high amplitude with short bursts. We've settled on a protocol of 15seconds on, 10 seconds off, for 3 minutes, using 100% intensity.
4. Fill to 50mL with Column buffer
5. Centrifuge at 13,000rcf, at 4C, for ~45 minutes.
6. While the cells spin, prepare your columns according to the NEB manual.
7. Drink some coffee
8. Harvest the crude lysate (supernatant), dilute the crude lysate 1:12 using column buffer (ie dump your 50mL crude lysate into ~600mL column buffer). Adjust pH to 8.
9. Load the column, wash, and elute according to the manual. We usually discard the first 5mL of the elution, and then capture the next 30mL.
10. Using molecular weight cutoff filters (we use 15mL Amicon 10,000Da cut-offs), concentrate your entire elution down to ~ 200uL.
11. Digest the concentrated elution using TEV protease overnight at 4C.

Day 4:

1. Using Ni-NTA method of choice (either agarose or beads), remove the TEV and MBP. Do not use membrane filters if studying microcins (as we are), as they often bind to membranes unpredictably.
2. Analyze (PAGE, Bradford, Qubit, etc.)
3. Enjoy

Clostridium Research

I first want to acknowledge the Clostron Research Group, because almost all of my Clostridium-related work has been done based on methods from the lab of Nigel Minton at the University of Nottingham. The publications from this group are really top-notch, in my opinion. If you want to work with Clostridium (especially if you are doing genetics), please make yourself familiar with this team's body of work.

Clostridium difficile

Primarily, I work with C. difficile strain 630. I've sometimes seen it referred to as C. difficile strain CD630 in the literature, but I believe strain 630 and CD630 are the same thing. It has a relatively well-annotated genome sequence, it is non-hypervirulent, tcdA+, tcdB+, and has a relatively easily circumnavigable restriction barrier, making it more amenable to plasmid uptake than other strains of C. difficile or Clostridium in general. Purdy et al, 2002, (from the Minton Lab) were able to transform Cdif CD6 and CD3 by modifying plasmids to be void of the specific restriction sites recognized by the restriction/modification (RM) systems of those particular strains, however strain 630 did not demonstrate any site-specific nucleases, but instead just general mechanisms.

The pMTL80000-series of vectors was then generated by the Minton group towards many Clostridium-based applications. It is a great concept, where these vectors can be used by the entire Clostridium community, rather than having specialized vectors built by all labs that want to do Clostridium work. The paper is here, and in the Bucci lab I use pMTL82151 for C. diff 630 chromosomal modifications. To build homologous recombination regions, I use SOE-PCR, which you can find a guide to on my Cloning Tips page.

In order to grow C. diff, I use BHIS media, which is Brain Heart Infusion (BHI), with the Supplementation of 5 mg/mL Yeast Extract and 0.1% L-cysteine. I prepare the BHI and Yeast Extract and then autoclave to sterilize. The L-cysteine I prepare a 5% solution, filter sterilze, and immediately transfer to the anaerobic environment. Then I add to the BHI + Yeast Extract as needed. Chamber conditions for C. difficile 630 can vary. I have grown the strain at both 10% CO2, 10% H2, and 80% N2, as well as 5% CO2, 5% H2, 90% N2 with success.

gDNA isolation and purification

In order to isolate the genome DNA of C. difficile, you can essentially follow the procedure as given by the Promega Wizard gDNA Purification Kit, which a few exceptions. A general outline is given below:

1. After 1-2 days growth, pellet the culture by centrifugation, within the anaerobic chamber if possible.
2. Resuspend the pellet in 480uL of anaerobic EDTA solution (50mM; pH 8.0). I generally prepare the EDTA solution from powder, do the pH adjustment with HCl and NaOH, and then filter sterilize.
3. To weaken the cell membrane, add 120uL of 10mg/mL lysozyme (Lysozyme from Chicken Egg White, Sigma) or 60uL Lysozyme (10mg/mL) and 60uL of Lysostaphin (10mg/mL), I've had roughly equal success with either method. You can do this step in an aerobic environment, but I try to move quickly to limit C.diff's exposure to O2.
4. Incubate at 37C for 1hour.
5. Centrifuge @ 15k*g for 2 minutes and discard the supernatant
6. Follow the protocol from Promega, adding 600uL of Nuclei Lysis Solution as described in step 6 (of the Promega protocol)
7. Incubate at 80C for 5 minutes
8. Increase the temperature to 94C and incubate an additional 8 minutes. Be careful, sometimes our hot plate will start to boil the water at this setting. If the water is boiling, the eppendorff tubes are going to pop open (speaking from experience...), and you should remove the heat momentarily. I've tried this protocol just keeping the temperature at 80C and I did not recover any DNA. The near boiling water seems to aid in the degradation of the cell membrane.
9. Then just follow the Promega protocol as given. Yields achieved generally in the 150 ng/uL - 300 ng/uL range.