same weird weird staph transformation results - any idea what is going on? (Nov/25/2009 )

I've been trying to obtain an overexpressor in staph. I've done TONS of electroporations in gram positive and this is the very first time I get this strange results (in order of attempts):

1. Lawn of bacteria in EVERY plate. I increased the antibiotic concentration. Run an antibiotic test on broth first and the result was a 4X times more concentrated than the paper I am following stated. Anyway... proceded.

2. I am electroporated 2 different strains obtained from 2 different sources. both behave the same. No more lawn, but colonies everywhere. Including the negative control

3. Increased even more the antibiotic concentration. Grow in broth without antibiotic for 1.5 hours after electroporation and then added antibiotic and grow more for another 1.5 hours, then plated. Result: one strain had no growth whatsoever. But the other one (the one I really want to transform) had a few colonies in most plates, by the periphery, including negative control again).

A few notes:the strain dies at certain concentration of kanamycin (the marker I am using) in broth, but not in plates (but I used 3 different batch of plates and made sure they were not too hot before adding the antibiotic). This strain is a restriction minus and that's why I want to use it as opposed to the other one (which did not grow on any plate) because the other strain shreds the plasmid. However, I did the same test on the restriction minus strain and they both seem to shred the plasmid. But the point here is that I am getting colonies in the negative (no plasmid) plate, even using a s**tload of kanamycin on it!

Any suggestions of what my be going on?

I want to get a new strain, but my boss thinks we should try with a different plasmid instead.

planktonica on Nov 26 2009, 08:09 AM said:

OMG, I don't think there is a person out there is having exactly the same problem as mine . I have 2 Staph strains (mutant RN4220 and an isolated strain), a shuttle vector with Kanamycin resistance gene and pE194 plasmid of S.aureus (Erythromycin resistant). The shuttle vector we have was used for Bacillus and it has pUB110 origin, so it should work in Staph, but both the 2 strains I have resist to Kanamycin and I always get many colonies on both plates, even when I increase the Kanamycin up to 450 ug/ml. I though that the isolated strain has Kan resistance because of its native plasmids, so I tried to eliminate it using Ethidium Bromide and electroporation. However, both 2 methods weren't efficient, even when I extracted plasmid from the colony showing no Kanamycin resistance, I got the same band as the wild type strain. I used this colony to make competent cell again, but somehow it still showed the colonies on Kanamycin plate with such a high concentration. I found out that in some colonies they shred the plasmid as in your case.
At that time, I'm trying again and again the electroporation using the pE194 plasmid. Seem like the Erythromycin is a better choice. However, I don't success with the isolated strain, even using the same conditions as RN4220.
I'm making the E. coli - Staph shuttle vector based on pE194 with my genes to facilitate the colony screening. I'm thinking about inactivating the restriction system of Staph using heating condition as described elsewhere. I'll try that if I fail again.
Even we come up with the same problem and no solution, but I'm glad to know I'm not the only one ^^. Please keep following this topic, if any of us solve the problem, let share to help each other. Thank you.

I am using RN4220 too, that that is my main problematic strain! I added up to 850 ug/ml of kanamycin and although it won't grow on broth, it still grows on my negative control plates!!! Also, even when it claims to be restriction minus, I added plasmid to some lysate I had for other experiment (lysate from RN4220 strain and from ISP179, separately) and ran a gel. I've got a shredded plasmid in both (however, I am not sure about how reliable this test is).

The vector I am using is a shuttle vector (puc19 based) with kan resistance marker. I am sure I am getting the right transformants, but with a negative control full of colonies, screening for the real transformant in the transformation plate would be finding a needle in a hay stack!

I just need to make an overexpressor!!!! If I can get to transform RN4220 that would be a great advance for me.

How concentrated are you using your plasmid?
Is your RN4220 strain resistant to kan?

planktonica on Nov 27 2009, 04:50 AM said:

The same here . Not only the RN4220, but also my isolated strain, they can't grow in the broth with 100 ug/ml Kan but after transformation, the colonies on the negative plate always appear after around 20h. Recently, I increased the concentration of Kan to 600 ug/ml but it's still the same.
I heard that the plasmid concentration is very important so I tried with different amount. The last experiment I used ~ 3ug plasmid for 50ul competent cell. I harvested cells for competent cell preparation at OD600 of ~0.5-1.0 at RT. My electroporation procedure is: mix cells and plasmid for 5 min at RT before pulsing, then pulse with the voltage of 1.8kV (I tried 2 kV, 2.5 kV and 3kV before but much of cells died and using high plasmid concentration can lead to the "arc" phenomenon when pulsing). I use the 0.2 cm gap cuvette and the apparatus from Bio-rad. I incubate my cell in B2 medium for 1-1.5 h before spreading. It's always the equal number of colonies on both plates, so I suspected the transformation didn't succeed and this was confirmed by plasmid extraction --> no plasmid at all.
No problem with my second vector pE194. However when I conjugated it with pGEMT vector for shuttle vector construction, I can't introduce the plasmid to RN4220 and the shredding seemed to be happened when I got the colony with many plasmid band but the PCR was negative.

How about trying the heat-inactivation method described in this paper (J. Lofblom, 2006)? Regardless the unexpected Kan resistance of RN4220, if you can dramatically increase the transformation efficiency, it's much easier to screen the correct colony.

About the unexpected Kanamycin resistance of RN4220, I think there's might be a mechanism behind. Staphylococci grow weakly with the low supply of oxygen, so does the antibiotic resistance. On the plate, there is much more of oxygen compared with the medium, so it's easier to resist the Kanamycin. Do you agree with this explanation?
One more thing is that aminoglycosides are active against aerobic and facultative aerobic Gram-negative bacilli and some Gram-positive bacteria of which staphylococci but they are not active against anaerobes and rikettsia. Is that any connection?

Hi quasimondo,

Well, the aeration rate in tube vs plate sounds logical, I am not sure if that would be the explanation but sound reasonable.
I will spare you some kanamycin: I tried up to 850 ug/ml and still the same.

I will try at higher concentrations. But now I am suspicious that my plasmid amount is low (I have a midiprep with about 900 ng/ul). However, my boss wants me to try again with higher antibiotic concentration.

We will try to get a new plasmid.

By the way, my protocol uses ice incubation (30 min) after electroporation. And we don't incubate the competent cells/plasmid mix at all. Actually, my previous supervisor stressed the fact that the cuvette had to be ice cold for electroporation (not so sure if that makes any difference at all).

planktonica on Dec 1 2009, 06:09 AM said:

Hi planktonica,
I don't think the incubation on ice is important in this case. I've got the protocol from other lab and they also use RT. They emphasized that the plasmid concentration is the most important thing. One more thing, I noticed how we prepare the competent cell is also affects the antibiotic resistance. Harvest the cell at low OD (0.5-0.6) somehow reduce the resistance. It seems like the adaptation mechanism of Staph is really good, they always have new variations after division, I think. Actually, I streaked my competent cell on B2 plate, pick 50 colonies to new plate and screen for Kan resistance, they showed different level. I choose the worst one (almost can't grow) for competent cell preparation. The new control cell grew weakly on Kanamycin, but it's the same as the transformed cells .
I'm doing the heat inactivation method, hope that it can help.
Looking forward your good news ^^. Good luck.

Hi,

Yestereday I electroporated again. I used more plasmid and a higher concentration of Kan (1 mg/ml)
Also, as a control and test I plated cells before competent cell treatment (just the overnight culture) in a kan1000 plate, also plated the competent cells and the competent cells after I gr4ew them in media at 37C for 1.5 hrs after electroporation.

The plate with overnight culture was empty.
The plate with competent cells had a few colonies in the periphery.
The negative control after 1.5 hr incubation in media had a few cells.
The transformation plates were mostly empty, except for 3 lonely tiny colonies that may (or may not) be actual colonies.

My thoughts:
Is it possible that the 15% glycerol treatment and heat shock of the competent cell preparation stress them so much that triggers some strange mechanism that makes them survive the kan plate??
Are those tiny colonies in the transformation plates actual transformants that are just too weak and such a crazy amount of antibiotic that are taking even longer to grow?

I am incubating the plates longer, to see if that makes any difference in size of the colonies and then testing the colonies (if they grow) by PCR.

Regarding the plasmid check. I am not actually sure that it was a reliable assay because I got the same results with the restriction minus strain and the clinical isolate.

Hello planktonica,
My approach also didn't work. I did the same protocol as mentioned in the paper: thawing of competent cell (40ul) -> quick harvesting -> treat the cell at 56 degree for 2 minutes -> add 500 ul of electroporation buffer (500mM sucrose + 10% glycerol) -> centrifugation 5000xg for 15 min -> resuspend competent cell in 40 ul buffer -> perform the electroporation. I still use the Kan concentration of 600 ug/ml with the hope that the relative colony number should be higher in my transformation plate but at last the control still grew (grow directly the competent cells without electroporation) and ... nothing on my transformation plate. I did the normal electroporation at the same time (without heat inactivation) but they show no different with the control.
In my previous experiment, I suspect the cell after electroporation could increase the Kan tolerance, so I grew them directly and there was no problem. Also, the non-competent cell can't grow on the low plate --> we observe the same thing (both my RN4220 and isolated strain). Considering the washing and glycerol treatment, the cell would reduce the intracellular ionic content (reducing electric resistance of the cell, I think). The ionic balance directly affect the respiration of the cell, which is related to oxygen consumption --> maybe the same reason when we compare the cell growth in the medium and on the plate!
Right now I temporarily pause this work for writing the manuscript, so I'm thinking about that more thoroughly. If you have any further result or idea, please tell me, I'm following this topic daily. Maybe at least we could figure out what is the culprit behind this weird thing.
PS: I also thing something went wrong with your assay, but I need time to find out what you missed. About the colony PCR, I think it's not the good approach for Staph screening or we have to optimize the protocol for this target. Normally I just got very blur and smear band by colony PCR. The best way is plasmid extraction and compare with the wildtype one. The last thing is I haven't known that you use the heatshock protocol for your Staph. I didn't think about that before because what I've found are just related to electroporation. So, could you provide me your protocol? maybe I'll try that.
Thank you and good luck!