Low conductivity media for yeast and E.coli - (Feb/06/2007 )
Hi,
I'm currently working on a project that is treating with yeast and E.coli cells. The cells should be exposed to an electric field while they are situated in a fluid. Because the electrodes get into direct contact with that fluid it should have a very low electrical conductivity.
At first some experiment conditions:
- whole duration 1/2 to 1 hour
- time the cells are in the field: about 10 to 20 seconds
- cells are supplied in a dry form (centrifugated, without fluid)
Unfortunately I'm not a microbiologist, so the answer to this question is not so easy for me 
I have already tried to get helpful information and I would like to tell you my present standard of knowledge:
- Media with salt in it (like 0.9% NaCl) have a conductivity that is too high
- Media with sugar in it have a low conductivity but an increased viscosity (cells should easily move in the fluid) and the cells will grow
- The osmolarity should suit the cells
My actual favourite would be a solution with sugar, but only enough so the cells live for the time of the experiments and do not grow too fast. Additionally the experiments would be conducted at room temperature (24° degrees Celcius) and not at the ideal temperature of 37° degrees for E.coli, so the growth rate would not be so high.
So, my actual questions are:
Is it right that an isotonic sugar solution (5g sugar in 100g water) has the correct osmotic pressure for cells? Could it be used or is there another problem like the pH value, or do I need normal water rather than deionized water for this solution?
Do you know a culture media or buffer solution that is suitable for yeast and for E.coli as well, that has a conductivity as low as possible and a very low viscosity (like water). By the way, the low conductivity is more important than the low viscosity! Additionally it would be great if the solution is as easy to fabricate on our own as possible (or cheap 
).
I really hope someone can help me!
Cheers, Juergen
Both e coli and nearly all yeast species have cell walls. Thus if said cell wall was intact, the cell is not terribly concerned about the tonicity of its' growth media. However growth rate is sensitive to pH.
E coli being a pig of a bug, thus isn't too bothered on the media it is eating. So your media should reflect the media that the yeast species likes. If both organisms are to be grown in the same flask, do note that e coli will probably out compete the yeast. Ie most of the biomass will be e coli cells. Also, some kind of positive selection would be nice if you intend to grow a mix culture. Antibiotic selection on both E coli and the yeast.
As for your media hunt, have you looked up the standard media for e coli and the yeast species? There are many standard preparations.
I am not sure if you are refering to Saccharomyces cerevisiae, Schizosaccharomyces pombe or another yeast species. So for no reason, I shall assume you are working on pombe! (the yeast species I work on)
Here is my attempt at formulating a high yeild low conductivity media.
For 1 Litre
5g yeast extract (what better food for yeast cell then other yeast cells)
5g tryptone (high growth for e coli.)
50mM NaCl (e coli needs Cl ions. below 40mM the growth rate is affected)
2mM NaOH (pH need to be around neutral for growth of e coli. E coli pretty much won't grow below pH 6. Pombe is a little more resistent to acid... this small amount of NaOH will give a tinny buffering capacity)
5g glucose (pombe needs glucose, however glucose cause e coli to rapidly acidify the media... and rapidly diminishes its growth. 0.5% is about the minimum glucose concentration for reasonable pombe growth and slighly higher then the maximum glucose concentration for e coli (assuming growth in a shut system.. a non continous system)
This is technically a rich undefine media.
You could look up minimal media formulation for your two microbe species. And design your own media from there.
But in anycase, the conductivity for any media that is decided upon, should be conducted.
If you want to use a buffer. I would suggest MOPS... (It has the draw back of not being autoclavable.) Another buffer possible is Tris, however E coli doesn't like high tris concentrations, (can't remember the concentration) so buffering capacity is limited.
Thank you very much for your detailed answer!
Do I get it right that the media you mentioned helps the cells to grow?
The point is that I do not want the cells to grow during the experiments!!
The yeast and E.coli cells are ready for testing, I only have to add some solution to get them into our testing device. They only have to live for about 1 hour in our media while the test runs and should not grow.
From that point of view a buffer solution would be the right thing. Unfortunately, all buffers have a high conductivity, as far as I know, because of the salt that they contain.
So, a solution with sugar would be an alternative, but the sugar let the cells grow. I thought that if I work at room temperature the growth rate would be reduced, maybe close to zero.
Do I understand it right that the most important factor for cells to live is the osmotic pressure and that the pH value is not so critical if the growth rate does not concern me?
How do I compose an isotonic sugar solution? With deionized water? What sugar should I use (glucose, ...)? Would that solution be applicable for my purpose?
By the way: I will work with Saccharomyces cerevisiae.
Regards, J.
quote name='celllover' date='Feb 7 2007, 12:04 AM' post='87051']
Thank you very much for your detailed answer!
Do I get it right that the media you mentioned helps the cells to grow?
The point is that I do not want the cells to grow during the experiments!!
The yeast and E.coli cells are ready for testing, I only have to add some solution to get them into our testing device. They only have to live for about 1 hour in our media while the test runs and should not grow.
From that point of view a buffer solution would be the right thing. Unfortunately, all buffers have a high conductivity, as far as I know, because of the salt that they contain.
So, a solution with sugar would be an alternative, but the sugar let the cells grow. I thought that if I work at room temperature the growth rate would be reduced, maybe close to zero.
Do I understand it right that the most important factor for cells to live is the osmotic pressure and that the pH value is not so critical if the growth rate does not concern me?
How do I compose an isotonic sugar solution? With deionized water? What sugar should I use (glucose, ...)? Would that solution be applicable for my purpose?
By the way: I will work with Saccharomyces cerevisiae.
Regards, J.
[/quote]
Ah, I see. My mistake. A holding solution rather then a growth media is desired.
That is correct, if growth is no concern, then pH isn't that critical. Just don't drop it below 6 or rise it above 9. However neither is osmotic pressure of extra importance either. The cell wall again comes to the rescue.
Nevertheless, if worries of damaging cell is of concern. I would suggest looking up spheroplast buffer solutions.
eg
0.4M mannitol (used to maintain e coli spheroplast. Also used in one paper to maintain cerevisiae spheroplast)
or
1M sorbitol (standard yeast spheroplast stabilising solution)
I would suggest that you use deionised water. Thus there is no need to worry if extra minerals from the tap have any compounding effect. The above solution is made in the way of any normal solution. However for autoclaving, if possible I would do a sugar run. Shorter time a peak temperature.
Do not use glucose.
I would prefer Mannitol, because I found similar experiments that use a 280mM mannitol solution (now that I know what I have to look for it is much easier 
)
That means, for your example, a 0.4M mannitol solution consists of 1l deionised water and 0.4*182g=72.8g mannitol (C6H14O6)., right?
Do you think less is ok too, like the 0.28M mannitol solution?
Either way, can I suspend both yeast and E.coli in this solution? I think so since the osmolarity is ok!
Glucose should not be used because it would assist the cells to grow and because it has no advantages, right?
Thank you so much for your help!!
)That means, for your example, a 0.4M mannitol solution consists of 1l deionised water and 0.4*182g=72.8g mannitol (C6H14O6)., right?
Do you think less is ok too, like the 0.28M mannitol solution?
Either way, can I suspend both yeast and E.coli in this solution? I think so since the osmolarity is ok!
Glucose should not be used because it would assist the cells to grow and because it has no advantages, right?
Thank you so much for your help!!
To make one liter of 0.4M mannitol solution, weigh out 0.4M of mannitol first.
0.4*182.172 = 72.9g
Add the mannitol powder into a 1L bottle (suitable for autoclaving). Then add approximately 500ml of distilled water. Swirl the mixture about until the powder is dissolved. Add a little more distilled water if the mannitol is being uncoperative. Once all the manitol is dissolved, top up with more distilled water until the 1L mark is reached. Note that less then 1L of distilled water is added, as the mannitol occupies a certain volume.
A 1M solution means a 1L of said solution contains 1mol of solute.
You many then filter sterile or autoclave (sugar type run)
I have not tried using a manitol solution that low before. I guess the best way to find out would be to conduct a small experiment and see if 0.28M is as good as 0.4M
And yes, using glucose would be the equivalent of thowing petrol on a fire. Cells are built to absorb and metabolise glucose. So it is not the best thing to use.
Ok, I'll try it that way.
Thanks for your help. I'll post how it worked when I've my results ...
cheers, J.