You can try a gentle rinse with sterile water (or nutrient solution) prior to sonication.  This should leave the biofilm in tact. 

Dear Kedar,

Nice question and study.

Maybe you can provide some details on how the biofilm was grown on a silicone catheter and how the catheter was washed. E.g. was the catheter submerged in growth medium (causing growth on the inside and outside) or was the medium flown trough the catheter (growth on catheter inside during medium flow).

Under low shear conditions and/or high nutrient concentration usually a biofilm develops with a low cohesion strength. Assume that using a vortex will remove quite some biofilm (flocs may be visible with naked eye or stereo microscope). Most likely washing of the catheter will remove some biofilm as well. Agree with Teresa that a gentle washing may be most effective to separate planktonic from biofilm cells. Suggested is to use the same osmotic strength medium as used for the growth (not e.g. demi water).

Maybe some control measurements can be considered to quantify the extent of biofilm release to the medium during rinsing/washing/the vortex treatment. A consideration is to measure the concentration of adenosine tri phosphate (ATP, measure for active biomass) of both the (i) growth medium and the (ii) accumulated material in suspension obtained from the sonicated catheter after gently rinsing as pretreatment. The ratio of ATP will enable to distinguish the degree of growth in the medium and on the material (depending on the experimental conditions the growth in the media may be minute compared to the on the material). Also, part of the medium can be filtered over a polycarbonate membrane (accumulating cells and biofilm on the membrane) and using a dye like DAPI or acridine orange and a fluorescence microscope will enable to see to what extent flocs of biofilm are present in the medium after rinsing the catheter. This technique can be found in literature and guidelines under total direct cell count.

Success!

Best regards, Hans

Thank you very much, Teresa and Hans!

The current question is regarding the static model where I immerse 1cm catheter piece in E coli culture in LB medium for 2 days. For a control I use a 10 min immersion of a fresh catheter piece to determine the effectiveness of washes. I can still see a lot of planktonic count after gentle washes in PBS. That is why I tried the vortexing. It helped me get rid of the planktonic bacteria but the doubt nagging me is that I am getting an underestimation if biofilm is removed by vortexing and am getting an overestimation if gentle wash does not remove planktonic. And these over- or under- counts are not (naturally) consistent over experiments.

I want to apply the quantification to my flow model also but only after sorting this problem out. 

Thanks again!

Dear Kedar,

Thanks for the additional information. I understand that you want to study the biofilm growth. The batch incubation experimental approach causes growth occurring in the nutrient rich liquid and on the surfaces in contact with the liquid. The rapid and strong growth of bacteria in both the liquid and on the material are most likely difficult to discriminate.

Usually, exact reproducible biofilm development may be difficult to achieve.

Based on the information provided, maybe the following may give some ideas for follow up studies, with more reproducible biofilm growth.

- batch growth potential tests in which the nutrient rich liquid is periodically (daily) replaced and the catheters are maintained in the flask (or the catheter pieces are transferred to another flask with fresh liquid). During the study the flask content is mixed in an incubator. At the end of the study the liquid is replaced and the biomass on the catheters quantified.

- growth during defined liquid flow. E.g. a fixed flow rate of liquid through the catheter will cause biofilm growth without the need to discriminate the growth in the liquid and on the material.

Best regards, Hans

You will always have a third variable, ie those cell which are on the biofilm but not physically-attached to the substrate. Reproducible washing is difficult to do. Basically you have to decide how you will wash and show how the number of cell detached  reaches a plateau (cells on biofilm, but not attached) . You then continue to increase the sheer in the washing procedure until a second plateau in cells removed is reached. These are those in the biofilm, not just on the biofilm. Doing the whole experiment in a flow cell is essential if you need to quantify sheer of the wash liquid.and its ability to cause biofim detachment. The works of WG Characklis deal with flow cells and surface-associated sheer force. See Characklis and Cooksey 1984 Biofilms and microbial fouling. Adv. Appl. Microbiol. 29:93-138. and publications that quote this

Thank you very much! I will look into these suggestions 

You might want to consider using a live dead type stain where you just stain for the live cells.  You could then visualize the cell locations on a microscope.  This would allow you to look at the cells after each washing to see if you have washed away those on the surface or those pieces of biofilm.  Or you could just use a E. coli producing GFP in the study.  This would just be an addon suggestion to the previous replies.

Thanks a lot!

Reproducible washing can be achieved (I did) but it all depends on the biofilm forming ability of your strain and optimisation of your washing. Our biofilm was very strongly attached and I also had to scrape the surface to remove the biofilm. You will always have some bacteria that attaches from the medium and this would be removed with gentle washing. I doubt that a significant amount of the biofilm would be lost upon washing if the strain is a good biofilm formant. Are you using static or flow conditions? If you are placing your catheter pieces into bacterial culture then handling them during washing will remove some of the biofilm that forms on the outside, increasing bacterial count in the wash solution. Washing conditions will also be different on the inside and outside of the catheter. Inside the catheter it will be harder to remove even loosely attached bacteria especially if the lumen is very small. It would be better to grow the biofilm just inside the catheter. Sonication by itself did not work for me and that is why I had to scrape the inside of the catheter to remove the biofilm. You would have to look at the sonication process as well as the sonication efficiency of many sonic baths is not the same across the bath's surface. That is, it may perform better when the sample is placed in the middle of the bath and worse towards the sides.

I am not sure what your aim is but it is possibly inevitable that a very small fraction of the biofilm bacteria comes off. As long as your procedure is optimised it should not matter. Also consider the boundary layer when washing.

Thank you very much, Pajkos!

Copying my previous reply:

The current question is regarding the static model where I immerse 1cm catheter piece in E coli culture in LB medium for 2 days. For a control I use a 10 min immersion of a fresh catheter piece to determine the effectiveness of washes. I can still see a lot of planktonic count after gentle washes in PBS. That is why I tried the vortexing. It helped me get rid of the planktonic bacteria but the doubt nagging me is that I am getting an underestimation if biofilm is removed by vortexing and am getting an overestimation if gentle wash does not remove planktonic. And these over- or under- counts are not (naturally) consistent over experiments.

I want to apply the quantification to my flow model also but only after sorting this problem out. 

Thanks again!

Hello,

For my project I was growing biofilm in a micro titter plate and in order to count planktonic and biofilm phase cells, I was removing planktonic cells with Gilson pipette slowly trying not to disturb the biofilm. After that I would wash the wells twice with sterile distilled water also slowly and genly, and after that I would scrape of the biofilm for 20 seconds. I hope it helps. Good luck with your project!

You can rinse twice gently with PBS before sonication  but I would also perform the experiments at least 3  times  to calculate an avarage of  bacterial count in biofilm

Thanks Maria, I tried rinsing gently in PBS, but what I observe is that even planktonic bacteria, which are not a part of biofilm, are not effectively removed and hence give an over-estimation. I have done this with a non-biofilm mutant as well as a 10-min immersion control.

Dear Kedar,

We use for biofilm formation assay minimal media not enrich media like LB. The media that used in standard biofilm assay in O' Tool lab/ Giesel school of medicine is typically,  M63 (minimal salts medium) supplemented with MgSO4, glucose and casamino acids (CAA). This media gives strong attached biofilm and easily remove the unattached bacteria. The enrich media has many amino acids that lead to attache the bacteria to the surface and will not remove even with the twice washing.

Also that fact of biofilm formation on implemented devices like a catheter and other surfaces is in hard environment like no oxygen, no nutrient elements, ...... 

Thanks,

Hadeel

Thanks Hadeel!