Contrary to Dr. Shah, I want to state that the principle of methylene blue staining is not membrane porosity but the ability of living cells to reduce methylene blue stain to a colorless product. While living cells have an active dehydrogenase system which reduce methylene blue, dead cells are unable to reduce this stain. Thus viable cells are colorless under this stain while dead cells stain blue.

For Dr. Ceyhun, I think you should optimize the methylene blue concentrations that you are working with. Try different concentrations between 0.01% to 0.1%. Also, try to use cells that are in their exponential phase of growth. If methylene blue is allowed to stain  cells for a very long time, it may lead to false negative results (many blue cells showing non-viability). Hope my answer helps. Bye

Hi Ceyhun,

mix equal parts of your yeast solution with a 0.1% (w/v) methylene blue solution. This results in a final MB concentration of 0.05%. Mix the suspension well and let it react for one minute. Afterwards proceed to count the cells by use of a counting chamber.

Best regards, Christian

Hi Christian,

Thank you for your answer. I have tried the same of your advice but it gave very high dead cells percentage in the sample which I did not expect that. I don't know where I am wrong. Is it suitable to use less concentraion (for example 0,05% w/v methylene blue solution, not in final concentraion) or certainly the right concentraion is 0,1% (w/v)?

Best regards.

Do you use exponentially growing cells? Or do you have a stationary culture? If you use cells from the latter culture you will probably get a rather high percentage of dead cells (especially if you started your culture with 2% glucose in the medium). I have the feeling that not your MB staining causes the problems but the culture itself... What are your cultivation conditions?

I tried stainig both with exponentially growing cells and stationary phase. I had high percentage with both conditions, but higher in stationary phase as expected. Cultivation conditions are 30 celcius at 200 rpm with YPD broth (1% yeast extract, 2% peptone and 2% dextrose). Do you think that approximately 40% of dead cells are normal for exponential phase?

40% of dead cells seems not realistic, as you say this is much too high...(but we are talking about control cells, not cells treated with a toxic agent, right?) I would expect this value in later stationary phase but certainly not in the exponential growth phase.

I have some more suggestions for you:

1.) How many cells do you use for your staining? A highly concentrated sample may be very difficult to count and a sample that is too much diluted may give erroneous results. The best way is to have 100 (or fewer) yeast cells at 400 times magnification per microscopic field area.

2.) Approx. 1000 cells should be examined in total.  When you are counting cells, buds that are less in size than half the parent cell are ignored.

3.) Take your samples from a YPD culture that has been incubated for 16 hr at 30°C.

I hope some of these suggestions help you. Good luck!

Yes, 40% of dead cells for control cells.

For your suggestions: 

1) After dilutions, approximately 40-50 cells in each field at 400 times magnification of microscope. I will try not to dilute as much as this. I should try to keep near 100 cells.

2) So far, I've counted 300-400 cells in each time. I will try your suggestion.

3) I took samples which were incubated for 16 hours as you suggested.

Thank you for suggestions,

Best regards.

dear, find this protocol

Principle of Procedure used:

The dead cells have increased porosity of the membrane, which allows easy penetration of dyes, and thus they get intensely stained. On the other hand, live cells resist diffusion of the dye because of controlled porosity, integrity and permeability of the plasma membrane. This principle is used here to differentiate live cells from dead cells with the help of dye like Methylene blue or neutral red at a concentration that is non-toxic to the cells. Here, dead cells appear stained blue in color, whereas, live cells appear faintly stained or colorless.

Demonstration of viable cells.

Requirements:

yeast culture suspension

0.5 % Methylene blue solution

Glass slide

Cover slip

Sterile pipettes and test tubes

Procedure:

1. In a sterile tube, transfer 5.0 ml of yeast suspension aseptically.

2. To this, add one drop of 0.5 % Methylene blue solution under aseptic condition.

3. Set the tube aside for five minutes at room temperature.

4. Transfer 3 to 4 loopful of stained suspension from tube on a glass slide.

5. Place coverslip and observe under high power.

6. Record your results.

Contrary to Dr. Shah, I want to state that the principle of methylene blue staining is not membrane porosity but the ability of living cells to reduce methylene blue stain to a colorless product. While living cells have an active dehydrogenase system which reduce methylene blue, dead cells are unable to reduce this stain. Thus viable cells are colorless under this stain while dead cells stain blue.

For Dr. Ceyhun, I think you should optimize the methylene blue concentrations that you are working with. Try different concentrations between 0.01% to 0.1%. Also, try to use cells that are in their exponential phase of growth. If methylene blue is allowed to stain  cells for a very long time, it may lead to false negative results (many blue cells showing non-viability). Hope my answer helps. Bye

I think right with Eke.