24 replies to this topic

[green_bear]

Hi everybody, I am new to this forum. Anyway, I would really appreciate if you could give me some suggestions on this.
Currently, we are trying to get a 3kb plasmid containing RFP constitutively expressed into P.a. We are following the protocol in this paper:
10 minutes preparation of electrocompetent P.a.

I will summarize the steps which we used as below:

Quote

A. Prepare electrocompetent P.a.:
1. Inoculate single colony in 6ml of LB overnight.
2. Distribute cell culture equally into 4 microcentrifuge tubes.
3. Centrifuge at 16000 x g and room temperature for 2min and discard supernatant.
4. Wash each tube of cell pellet with 1ml of sucrose solution and centrifuge. Repeat Step 4 twice.
5. Resuspend all the 4 cell pellets with a combined volume of 100ul of sucrose to produce 10^-9 viable cells.

B. Electroporation:
1. Transfer 3ul or 500ng of purified DNA into 100ul of electrocompetent cells.
3. Transfer the mixture into pre-chilled cuvette.
4. Slot cuvette into BioRad Gene Pulser and pulse shock (2.5kV, 200Ω and 25µF). Add 1ml of LB at room temperature immediately into the cuvette. Transfer mixture into 1.5ml microcentrifuge tube and shake for 1 hr at 37oC.
5. Centrifuge at 16 000 x g and discard 900ul of supernatant. Resuspend cells in 100ul of residual medium.
6. Plate entire cell culture onto LB plate with suitable antibiotic (Chloramphenicol 20ug/ml). Incubate at 37oC overnight.

The only 2 steps which are slightly different from the paper are in bold:
- After the pulse shock, it takes us about 30 seconds (not immediately as suggested) before we add LB, as we need to move the sample into fume hood.
- We incubate the cells for 1 hour in 1.5 ml microcentrifuge tube, not glass tube.

The problem we encounter is that, after this 1-hour incubation step, the cell solution develops a transparent glue-like matrix which attach to all the cell pellets. We suspect that this might be biofilm, but have no idea which leads to its formation. So we always end up plating both the cells and this matrix on the plate. The next morning, we have indistinct colonies growing all over in all the plates, including negative control (cells without plasmids added). So it might be that this biofilm protects the cells from the antibiotics.

Have anybody else encountered this problem before, or had  any suggestion on this?

Thank you very much in advance :-)

G_B

[HomeBrew]

Hello green_bear -- welcome to the BioForum!

What do you mean by "indistinct colonies"?  Do you mean a lawn of growth?  If so, try plating your cells at dilution, or reduce your non-selective growth period to 30 minutes.

The "transparent glue-like matrix" is probably a high MW polysaccharide -- most likely alginate.

Why do you need to move your cells into the fume hood before adding media?  If this is absolutely required, can you move your electroporation machine there as well?  I don't think this is having a significant effect on your experiment though...

Does your recipient strain grow on your chloramphenicol plates?  Maybe you need to go higher than 20ug/ml -- we usually use chloramphenicol at ~30 ug/ml.

[green_bear]

Hi HomeBrew,

Many thanks to your prompt reply and suggestions.
Yes, I should have referred it as lawn of growth rather than indistinct colonies. Next time I will try both of your suggestions.
Also, could you give some details on how alginate is produced? We've never had this problem before while growing P.a. So it could only be due to the electroporation. I think it's the core of the problem, as this matrix tends to stick to all the cell pellets, so we could not resuspend them, and ended up plating the matrix as well. I realize that reducing the non-selective growth (from 2 to 1 hour) could actually help reduce the amount of this matrix. I will try to reduce further, perhaps to 30 minutes. Besides this, would you have any other suggestion?
In fact our recipient strain could not survive in LB broth with 20ug/ml. But they did survive while being streaked on agar plate as we used them as the negative control for the above experiment. I will test with higher Cm concentration next time, but still think that the alginate that you mentioned has something to do with this.
About the fume hood, I was always told to do any bacterial work inside that to avoid contamination, and ensure safety. The electroporation machine belongs to another lab, so we can't move it. Anyway, I also think it would not matter much.

Thank you!
G_B

[phage434]

It sounds to me as if you might be simply killing the cells, and the "alginate" is cells lysed and releasing cytosol.  Is the electroporation cuvette arcing or popping during this process?  I would try lower voltages.  Also, you might try lower cell densities.  For reasons I don't understand, this can often improve efficiencies.  You might want to see whether you still have viable cells after electroporation, by serial dilution and colony count on non-selective plates.  Some strains may have restriction systems which will dramatically reduce transformation efficiency.

[HomeBrew]

I think phage434 has a point -- I immediately thought "alginate" in conjuction with P. aeruginosa, but as I think more carefully about it, this could be cell debris released by lysis, as I have seen this kind of junk before, even in E. coli.

I really would work out your selective conditions and plating dilutions -- it's going to be tough to draw any conclusions until you are sure your electroporants are selected for and you can get them as well-isolated colonies.

[fishdoc]

After reading the protocol, it seems to me you are plating an awful lot of bacteria onto 1 plate. We do a similar type of electro-transformation (mostly for E. coli). Following electroporation, we add the 1 ml of LB and incubate 1 hr to recover. However, following recovery, we do not pellet the cells and resupend in 100 ul, we just use that ~1 ml directly to plate. Usually I plate 10, 50, and 200 ul onto selective plates, so that's up to 1/5 of the total cells or so that get plated. Frequently, there is a lawn on the 200 ul plate, because (I think) the mass of cells plated overcomes the antibiotics present in the media. Using lower volumes, or even diluting 1/10 or 1/100 usually gives much better results.

Most of the time we are using ampicillin (200 ug/ml), so I don't know how that relates to chlor or the ability of bacteria to overcome the antibiotics, but since your contol cells (no plasmid) are also growing on the selective media, the first place I would start is to plate much less bacteria, and maybe also not even pellet/resuspend, which is concentrating the bacteria.

As for the matrix following recovery, I don't know what that would be.

[green_bear]

phage434 and Homebrew, I really appreciate all your thoughts.
In fact today I tested the same protocol with both P.a. and E.coli MG1655. For the latter, I did not see any of the glue-like thingy. So could it be due to both reasons: alginate produced by P.a. was released out due to cell lysis?
You guys are right, I will play around with different antibiotics concentration, as well as serial dilution first.

By the way, do you know whether Pseudomonas do produce AHLs while growing as planktonic cells in liquid medium? I know roughly that AHL concentration is higher at higher cell density, but I'm not sure whether this is only applicable to when they are growing on solid surface (and forming biofilm), or also in liquid medium.


Have a nice weekend!

G_B

[green_bear]

Hi fishdoc, thanks for your advice as well.
Indeed this is the first time we've ever done electro-transformation, so we just stick to a recent paper with high citation regarding the process for P.a. If tomorrow I still see a lawn of bacteria, I'll probably reduce the concentration for plating the next time.
The problem is still that most bacteria stick to that matrix, so it's really hard to adjust cell density before plating. As phage434 suggested, I'd lower the voltage to see whether I can get rid of that stuff.

G_B

[green_bear]

Hi guys,

I'd like to give some updates regarding my exp.
Well, previously I actually misunderstood the protocol somehow. The steps quoted are for transformation of chromosomal DNAs. So for replicative plasmids (as what I'm doing right now), the centrifugation step before plating is omitted. So doing that way I could get rid of the glue-like matrix.

I determined that 30ug/ml Cm in LB plate is indeed sufficient to kill the negative control PA.

I did as what fishdoc suggested, using 50ul and 200ul out of the 1ml culture (after recovery growth) to plate. With 200ul, I saw lawn of bacteria even for negative control (without plasmid). I guess it's due to too much cells plated. With 50ul, I did not see sign of bacteria after 16hours, but saw a thin layer after 24 hours. Could this be due to the breakdown of antibiotics in the plates?

So up to this point the transformation is still unsuccessful

Do you guys have any advice on this?
By the way, besides electro-transformation, has anyone tried the MgCl2 and heat-shock method for P.aeruginosa? I know the efficiency is much lower, but I still want to give it a try as my plasmid is only about 3kb...

Thank you!

G_B

[fishdoc]

green_bear, on Jan 27 2010, 01:30 PM, said:

Hi guys,

I'd like to give some updates regarding my exp.
Well, previously I actually misunderstood the protocol somehow. The steps quoted are for transformation of chromosomal DNAs. So for replicative plasmids (as what I'm doing right now), the centrifugation step before plating is omitted. So doing that way I could get rid of the glue-like matrix.

I determined that 30ug/ml Cm in LB plate is indeed sufficient to kill the negative control PA.

I did as what fishdoc suggested, using 50ul and 200ul out of the 1ml culture (after recovery growth) to plate. With 200ul, I saw lawn of bacteria even for negative control (without plasmid). I guess it's due to too much cells plated. With 50ul, I did not see sign of bacteria after 16hours, but saw a thin layer after 24 hours. Could this be due to the breakdown of antibiotics in the plates?

So up to this point the transformation is still unsuccessful

Do you guys have any advice on this?
By the way, besides electro-transformation, has anyone tried the MgCl2 and heat-shock method for P.aeruginosa? I know the efficiency is much lower, but I still want to give it a try as my plasmid is only about 3kb...

Thank you!

G_B

By "thin layer" do you mean the beginning of a lawn? Or do you mean small colonies forming? I'm not familiar with P. aeruginosa, all my work has been with E. coli. Within 24 hrs (and usually within 16), E. coli is normally growing fine. If not, it's usually a failed ligation or electroporation or the insert is toxic to the cell. I wouldn't think RFP would be toxic to the cell unless it was produced in really high amounts. The other thing would be to make sure the ori of the plasmid is compatible with replication in P. aeruginosa, or making sure P. aeruginosa doesn't carry a native plasmid that is incompatible with the plasmid you're trying to transform.

[phage434]

So, if the transformation protocol is under control, you might want to check a few other things.  A P. aeruginosa restriction system could be cutting your DNA.   You might want to check what restriction systems are typically present in your strains.  Do you have a control plasmid that is functional in Pseudomonas that you could use to verify transformation?  What origin of replication are you using in your plasmid?  Have you sequenced your plasmid to assure that it is what you think it is?

[HomeBrew]

When I worked with P. aeruginosa, we usually moved plasmids in via conjugation (see Goldberg and Ohman. (1984). J. of Bacteriol., 158:115-1121), but your vector must be mobilizable by a helper plasmid.  Recently, I successfully transformed PAO1 using cells made competent by the rubidium chloride method, so that works, too.  But you need to get your selection method tightened up so that you have no growth at all on your control plates, or you're going to have difficulties with any method.

[green_bear]

Hi guys, thanks so much for your replies!

@fishdoc: Yes, I meant beginning of a thin lawn.

fishdoc and phage434, you are both right, I should check with the person I got this plasmid from about the its compatibility with P.aeruginosa. But say, I know the sequence of my plasmid, is there any way I can do the check myself online? I also have the RFP insert in pUC19, a common E.coli vector. Does anyone know whether this vector is compatible in P.aeruginosa? There's also the 12kb RSF1010-derived plasmid which is P.a.-compatible, but the last time I tried extracting the plasmid, I could not see any band under gel electrophoresis.

As Homebrew mentioned, I am still not 100% sure about the selection method. Last time I tried streaking colonies directly from a P.aeruginosa plate into LB plates of 20,30 and 50 ug/ml Chloramphenicol. Cells did not grow in the first plate, while I could see lawns on the other two. But the fact that untransformed cells did grow when 200ul of recovery culture was plated (and even 50ul after 24h) makes me a bit doubtful of selection condition. I am not sure whether this is due to, as fishdoc suggested, "the mass of cells plated overcomes the antibiotics present in the media".

In addition, which OD of cells should I use to prepare electro-competent cells? I notice that the protocol mentioned overnight culture in LB was used. But this is kinda weird, cause after overnight incubation, normally the bacteria already reached stationary phase and might not be that active any more...Do you guys think I should re-inoculate this culture in the morning (about 100 times dilution), and grow into an OD of 0.5-0.7?

And any comment on the washing and electroporation medium? I used 300mM sucrose (dissolved in distilled water) for all the steps.


G_B

[fishdoc]

I don't have the protocol in front of me, but we make E. coli electrocompetent by growing an overnight culture, then using that to inoculate about 40 mls of LB (1:100 inoculum) and growing for a few hours to an OD of between 0.6 and 1.0. Cells (10 mls) are put into centrifuge tube and pelleted, resuspended in DI water, pelleted, resuspended in DI water, pelleted, resuspended in 10% glycerol, pelleted, resuspended in 2 ml 10% glycerol, transferred to 1.5 ml tubes, pelleted, resuspended in 200 ul 10% glycerol, put in 1.5 ml tubes (50-80 ul) and stored at -80C.

[green_bear]

Hi, just a quick update from today.
I stored recovery culture left over from last time electroporation in 4degC fridge. Yesterday I took 100ul of it to plate on LBC plates (30ug/ml). After 16 hours, I saw some colonies on the plate, but none on negative control plate. But as the colonies were still growing (I could not pick them from the plate cause they were not tightly attached to the agar surface), I let it incubated further more. About 1 hour later, I saw lawn of bacteria on both plates. So is this due to break-down of antibiotics?

G_B