Science Buddies: "Ask an Expert"

I'm testing the effect of lemongrass essential oil on E. coli using the disk diffusion method. I need to make dilutions of the oil, and after doing research I found common solvents were polysorbate 20, tween 80, DMSO, etc. I want to use PS20, but they always say to add water when using it. I only want the oil and PS20. Will adding only these two make a completely soluble solution that will properly diffuse into the agar? What other solvent can i use that doesn't affect the bacteria itself? Basically, I need to find something that is miscible with the oil that will help make dilutions and have th oil diffuse into the agar to measure the effect on the bacteria. I'm also comparing these results to dilutions of sodium nitrate and potassium nitrate, but I can make those dilutions with water, which doesn't have an effect on the bacteria. I'm just worried about the EO. Please help me!

Hi GG,

The PS20--water--oil mixture should be fine. You don't need to use all oil to see an effect. A dilution should do it. Just remember to also do a control with the PS20 and water alone--no lemongrass oil. The PS20 is like a detergent that disperses the oil, which cannot otherwise dissolve in water, to form a fine emulsion. This will diffuse into the agar from the disk and have an effect on the bacteria.

I don't know if polysorbate 20 will kill bacteria or not, but i would use the lowest concentration of PS20 necessary to emulsify the oil-water mixture. You could do a pre-experiment in which you tested various concentrations of PS20 to see how toxic it is to your bacteria. As long as you use the same conc of PS20 in your control disk, the difference has to be due to the essential oil.

Does this help? If you have more questions, please ask and we will try to get you going on the right path.

Best wishes,

Sybee

Do I have to use water at all? It's just that if I want make a 5% oil dilution for example, how much water and ps 20 do I add to make it a 5% dilution? If I used only one solvent (the ps20) i could readily calculate a 5% dilution but how do I keep it 5% by adding water?

I suspect that if you use the essential oil and PS20 alone you might have problems with diffusion into the agar, not to mention that your PS20 would be highly concentrated which could have an effect on the bacteria. I agree with SciB 100%. Use a PS20-oil-water mixture and make sure in your controls you have a PS20-water control. That way you can determine how much of the antibacterial effect you see is due to the essential oil vs. the PS20.

As for making the 5% dilution, it won't be a problem at all. Based on the articles you found, you should have an idea of how much PS20 you need to get the essential oil to mix with the water. That amount will always remain constant. Then you calculate your dilution based on the total volume, not one solvent alone (water + PS20). So for example, you could make a 5% dilution of your essential oil that would be 5mL of essential oil into 100mL total (water + PS20).

I hope this helps!
JMP

Ok, I thought that's how it should work. Thank you for your help!

Is there any difference between cosmetic-grade PS20 and "lab" grade PS20? It seems like the one described as being used for cosmetics is a little more viscous, and so would it be harder to combine with the essential oil and water?
http://www.naturallythinking.com/produc ... te-20.html and http://www.makingcosmetics.com/Polysorbate-20-p148.html seem like PS20 used to make sprays, in soap etc.
This one seems more "lab" grade but I'm not sure what else they add in it, and if it will have a bactericidal effect:
http://www.piercenet.com/product/tween- ... t-solution
Can I use one of the first two links?

Hi GG,

You should buy lab-grade Tween 20, but you don’t need to spend over a $100 for it. Get it from Sigma-Aldrich Chem. Co. http://www.sigmaaldrich.com/catalog/pro ... &region=US
They sell a bottle of 100 mL of the pure, undiluted liquid for around $25.

Can you work in your school lab while you are doing this experiment? It would be so much easier than trying to do it at home.

When you get the Tween 20, you will see that it is very viscous and you can’t use it that way. Get a 100 mL graduated cylinder and CAREFULLY pour 10 mL of Tween 20 into it. If you go over the 10 mL mark, rinse out the cylinder and try again. Once you have exactly 10 mL, fill up the cylinder with sterile distilled water to the 100 mL mark. Cover the top with a piece of plastic wrap, put your hand over it to hold it tight and invert the cylinder several times, back and forth, until the Tween 20 has dissolved in the water. This will give you a 10% solution that is much less viscous and easier to use.

I remember from an earlier post that we suggested you test the toxicity of various dilutions of Tween 20 alone before you use it with the lemon-grass oil. I don’t know how much 10% Tween 20 you will need to solublize the oil, but you should aim for the LEAST amount to minimize the risk of killing E coli by the Tween 20 alone. It is a detergent, and detergents can kill bacteria. If you have time, make test dilutions of the Tween 20 at 0.01, 0.1 and 1% and also test the 10%. Use sterile distilled water to make the dilutions. You will have to do this in the lab. You can look up ‘how to make serial dilutions’ on you-tube or you can ask someone to show you how it’s done.

Did you understand how to make the spreader and spread the bacteria on the plates? Please check with us if you are not sure.

Good luck!

Sybee

Yes, I think I understand the bacterial spreading process and I have researched serial dilutions before, although I wasn't sure if I needed them for this experiment.
I will probably carry out my experiment at school only because E.Coli K-12 is a BSL1 organism and some resources say I have to carry it out in a lab. I'm not sure if my school has many supplies like a micropipette for making dilutions, though. Assuming the PS20 does kill bacteria, you just have to keep diluting the PS20 and water mixture uniformly in the same amount of bacteria in test tubes until you get a clear and a cloudy test tube, correct? In other applications, it says to make a 1:1 mixture of essential oil and ps20 but you may have to increase the amount of ps20 (depending on the density of the oil, etc.). In that case, wouldn't I need less than a 10% mixture?
Also, what exactly is the difference between http://www.sigmaaldrich.com/catalog/pro ... &region=CA and http://www.sigmaaldrich.com/catalog/pro ... &region=CA ? I'd like to use the first link ("for molecular biology") since I'm on a budget and I think 50mL should be enough, but I'm not sure what the difference is between the two?

Hi GG,

1) You should be fine with the molecular biology grade Tween 20.
2) It's possible that you'll need a higher concentration of Tween 20 to solubilize your oils, but I agree with SciB that you are better off starting out with the more dilute solution. Unless you do have the time to test concentrations of Tween 20 on your bacteria, you really do run the risk of killing your bacteria with too high of a concentration of the detergent. I strongly suspect that a 1:1 mixture of your essential oil with the detergent will end up killing your bacteria because of the high concentration of detergent, and you won't be able to see any effect of the oil.
3) I'm not entirely sure what you mean about mixing the water and PS20 mixture uniformly with the bacteria until you get a clear and cloudy get tube. If you are talking about testing for the toxicity of the PS20, I would test the toxicity in the exact same way you intend to test the toxicity of your essential oils. Plate the bacteria on a petri dish, add a disk soaked in the detergent and see if it inhibits the growth overnight. If this is not feasible (time, etc.) I guess it's possible you could test it in liquid culture, but ideally you want the setting to be as close to your real experimental setting as possible.
4) Hopefully your lab will have the pipettes you need. If not, you may have to use a dropper for plating the bacteria. In this case, 1 droplet is approximately equal to 100uL. Just be as precise as you can, trying to be very consistent from sample to sample.

Hope this helps clear things up. Keep checking back with us if you still have questions!
JMP

Thanks. I was planning on making different dilutions of the essential oil as well - 1%, 2%, 3%. If I find the lowest concentration of ps20 to solubilize the EO, do I use that same concentration for the 1%, 2%, and 3% dilutions, to keep it all constant? Or should I adjust the amount of ps20 I use based on the concentration of EO I want to make?

I would definitely keep the concentration of PS20 constant while you vary your concentration of essential oil. Otherwise you would be introducing multiple variables, which makes it much harder to ensure that you have the right controls and interpret your results.

I also have another quick question. I'm almost ready to start my experiment. If I want to make a 1% dilution of sodium benzoate in distilled water, how would I go about doing this? I couldn't just add 1g of sodium benzoate in water, could I? And if I can, I'm guessing i should probably get a very fine digital scale.... I'm making 1% dilutions of my oil and the preservative, so I need to keep that constant.

A 1% solution by definition is 1 g of anything dissolved in 100 mL total volume of liquid--water in your case. So, yes, you would dissolve your 1 g of sodium benzoate in water. Do you have a graduated cylinder to measure the volume in milliliters? One liquid ounce equals about 30 mL if you only have a kitchen measuring cup, or you could measure it using a digital scale because 1 mL of distilled water weighs exactly 1 g.

Since I want to transfer very small amounts of my essential oil (like 1 mL, 2 mL, 3mL, etc.) and Tween 20 (10 mL, 5 mL, etc.) can I use a regular syringe or should I be using a graduated pipette and a pipette filler/bulb?
http://www.hometrainingtools.com/pipett ... E-PIPE10A/
http://www.hometrainingtools.com/pipett ... E-PIPFILL/

I think a regular hypodermic syringe would be hard to dispense such small amounts of liquids, especially when they're as viscous as my oil and PS20, but the people I ask are telling me to just use the syringe....

Regular syringe should be sufficient for this amount. you just need to make sure to get the right volume of the syringe (You have to take the waste space of the syringe into consideration). Depending on the situation and the kind of container you are using, attaching needle may help dispensing the solution.

If you are concerned with accuracy, pipette would be a better choice (micropipette should be used if you want to transfer anything less than 1mL). Because your samples is viscous, it may take longer for your samples to travel in the pipette, so do it slowly when you are taking or releasing the sample, for the purpose of accuracy .

Hope it helps.
Pak

Hi GG,

Yes, definitely use a pipet, preferably a sterile 1 mL pipet but a 1 mL transfer pipet would also work. And as Pak said, be sure to let all the oil drain from the pipet.

Did you test the dilutions of PS20 for toxicity on a plate culture of E coli? You need to do that BEFORE you start making dilutions of your EO. I don’t know what concentration of PS20 will dissolve the EO, but as we said before, use the LOWEST concentration to avoid killing the bacteria.

Don’t try to pipet PS20. It is too viscous. Do as I said before and pour a little into a clean, sterile tube, then add an equal volume of sterile water, cap the tube and invert it several times to dissolve the PS20 in the water. This will give you a 50% solution of PS20 which you can pipet.

Make a table showing the volume of 50% PS20, the volume of water and the volume of EO you will mix to make the various dilutions so you don’t get mixed up. You are diluting the PS20 AND the EO, so you have to be sure you know which solution has which concentration and what the final dilutions will be.

Did you decide what dilutions of EO to test? Start with the most concentrated [maybe 10%?] and add just enough 50% PS20 to dissolve it. Then using the same concentration of PS20, make serial dilutions of the EO. Your control will be the diluted PS20 alone with no EO.

I know this can be confusing. That’s why I said to make a table before you start so you are sure which volumes of which solution to mix to make the final dilutions. If you get stuck, we’re here to help!

Good luck,

Sybee

Ok, my teacher did manage to find a serological pipette and a pump for me. Thanks!

Great! Good luck! Keep us posted on your results and please, if you have ANY questions or aren't sure if you are doing it right, ASK US! It is much better to ask a lot of questions than to make a mistake that ruins the whole project.

Sybee

"Start with the most concentrated [maybe 10%?] and add just enough 50% PS20 to dissolve it. Then using the same concentration of PS20, make serial dilutions of the EO. "

How do I know exactly how much PS20 will be exactly enough to dissolve the essential oil? Do I just have to pour some diluted PS20 in the specified amount of EO, pour in some distilled water, pour in some more diluted PS20 if the oil isn't completely dissolved, etc.? Then the amount of PS20 dilutions you would pour for each dilution of EO needed would differ.
Also, I'm not exactly sure what you mean by making serial dilutions of the EO.... So I could have, for example, a 10% dilution of essential oil (10 mL of EO with appropriate amounts of PS20 and distilled water). Then I could take a mL of this and pour in another 99mL of solvent to make a 1% diltuion? Do I need to use serial dilutions? I was thinking that I could just test 1% , 2% , and 3% dilutions of the essential oil with each's appropriate PS20 and water amounts.

A serial dilution like this? I think this is basic:
http://toolboxes.flexiblelearning.net.a ... Dilutn.htm

GG, I know this is hard to explain. It would be much easier if I could take you in the lab and SHOW you! You need to do a wider range of dilutions than just 1, 2, 3% etc, because these are too close to allow you to measure a clear difference using the disk assay. I would try 10%, 5%, 1%, and 0.1% EO. This will give you a good idea of how sensitive the bacteria are to the EO. If they are not very sensitive then you made need to try 20%. But if they are very sensitive, then you may need to do more dilute concentrations.

You need to start with the most concentrated EO solution because when you have added enough 50% PS20 to solublize the oil in that one then you know you will be able to solublize all the others.

Let's say your most concentrated EO solution will be 10%. If you took 1 mL of EO and added it to 9 mL of water you would have a 10% solution but then you could not add PS20 to it because it would increase the volume and the EO would become less than 10%. So, what you have to do is try different concentrations of PS20 with a fixed concentration of EO, 10%. I would try 10% PS20 first. So here's what you would do:

100% EO....1 mL
50% PS20...2 mL
DW..........7 mL
TOTAL.....10 mL

If the EO dissolves to make a clear solution, then try 5% PS20:

100% EO.....1 mL
50% PS20....1 mL
DW...........8 mL
TOTAL......10 mL

Now do you understand how to do it? If 5% PS20 dissolves the EO, then try 1%. If that doesn't work then you know the lowest concentration of PS20 that will solublize 10% EO is between1% and 5%.

Once you have found the best PS20 concentration, then you can use that solution to make the dilutions of EO for the bacterial test.
...................10 %............5%..............1%.............0.1%..........0%
100% EO.........1 mL............0................0...............0.............0
10% EO............0.............5 ml............1 mL.............0.............0
1% EO.............0...............0..............1 mL.............0.............0
50% PS20
DW
TOTAL...........10 mL........10 mL...........10 mL.........10 mL.......10 mL

Note that the total volume for each dilution is 10 ml. Add the same amount of 50% PS2 to each to give the optimal concentration, then add enough DW to make exactly 10 mL.

I don’t know how big or thick your paper disks are, but don’t put too much of the EO solution on each disk. You want enough to wet the whole disk but not so much that it runs off into the agar. I would do one plate for each EO solution including the one that has 0% EO. Depending on the size of your disks and your petri dish, you could put 4 per plate so you would have sufficient space between them to measure the zone of inhibition.

I hope this helps. Once you have done it in the lab it will all make better sense.

Good luck!

Sybee

OK, I looked at it some more and I think I understand now. Let's assume I wanted to make a 10%, 1%, and 0.1% dilution of essential oil (just to be really basic/simple for now).
Let's also assume that I would need the 5% PS20 mixture to completely solubilize my oil (1 mL of PS20, 8 mL of DW).
I could fill test tube A with 10mL of 100% essential oil, and test tube B with 9 mL of the buffer (going with the example, 1mL of the 50% PS20 and 8 mL of distilled water). If I pipette 1 mL of the undiluted essential oil in test tube A into test tube B and mix, I would now have a 10% essential oil mixture.
Then I would also fill test tube C with the same amount of 50% PS20 (1 mL) and the same amount of distilled water (8 mL), to produce another 9 mL of buffer just like in test tube B. I could then pipette 1 mL of the 10% essential oil mixture from test tube B into test tube C, to create a 1% essential oil mixture.
Then I would also fill a test tube D with the same amount of 50 % PS20 (1 mL) and the same amount of distilled water (8mL), to produce another 9 mL of buffer. I would then pipette 1 mL of the 1% essential oil mixture from test tube C into test tube D, to create a 0.1% essential oil mixture.

Would this be accurate? Does the 9 mL buffer in each test tubes B-D have to have the same amount of PS20 and water every time to be constant?

Please let me correct you on one thing first--the word BUFFER. This specifically refers to a solution of a salt like sodium acetate, sodium phosphate or sodium citrate that is able to maintain--buffer--the pH of a solution. In your case it would be better to say detergent or surfactant solution because PS20 [aka Tween 20] is a detergent. If all you have is water and PS20, there is no buffer.

OK, now...great! Your serial 1 to 10 dilutions are perfect. Do you KNOW that 5% PS20 will solublize 10% EO? What about 2.5% or 1% PS20? Detergents will kill E. coli so you need to use the lowest concentration possible to limit the killing. You will, of course, do a control with just PS20 on the disk and no EO, but if the detergent is very effective at killing the bacteria, then you won't see any effect of the EO.

So, your plan is to get a liquid broth culture of E coli, spread 50 or 100 microliters on each plate, add the disks with the various concentrations of EO on them then incubate the plates and measure the zone of inhibition around each disk, correct? Try to incubate them as close to 37C as you can, but be careful the plates don't get warmer than 39C because this can inhibit E coli growth. Try to find a graduate student friend who has access to a 37C incubator for your plates. It would be SO much simpler.

Good luck!

Sybee

Whoops, sorry! I meant solvent, not buffer - both the Tween 20 and the distilled water. The 5% Tween 20 was just an example - I'll definitely test to find the lowest concentration I need.
As well, since I took the route of food preservation, I was thinking I could also test sodium benzoate (it's been tested more recently now to inhibit E.Coli growth) in the same concentrations as my essential oils - 10%, 1%, 0.1%, just to make my project a little more convincing? So I would do my serial dilutions but with 9mL of distilled water each time as my solvent. I would then compare my results with the results of the essential oil zones of inhibitions. I'm just afraid that if the Tween 20 does have an effect on the bacteria, no matter how small, I can't really accurately compare the essential oil and the sodium benzoate because this might be flawed.
Thank you for all your help!

The sodium benzoate is a good idea. You need a second inhibitor for comparison. Did you do a search for the concentration of sodium benzoate that is commonly added to food as a preservative? That should be your starting point for test concentrations.

Do you have enough time to do a test of your PS20 concentrations on the E coli? That would tell you how toxic it is. Also, try googling 'e coli toxicity tween 20 ps20' and see if there is anything. You could also try 'e coli detergents'.

Good luck!

Sybee

So, I have started the first part of my experiment - finding out the lowest concentration of Tween 20 needed to completely dissolve 1mL of essential oil in 10mL of solvent.
I started with 50% Tween 20 to make my dilutions as suggested. I didn't have a 100mL graduated cylinder so I added in 10mL of Tween 20 with 20mL of water. However, it was still too viscous and I could not use my pipette to suck some up. So, I modified it to 25% Tween 20 which was a little easier and also dissolved some more in the water (10mL of Tween 20 and 30mL of water).
I created my 10% dilution of 25% Tween 20 with 10% essential oil by adding 1 ml of oil, 4 mL of 25% tween 20 (4 X 0.25 = 1, 1/10 = 10%) and 5 mL of distilled water. The resulting mixture was very white and opaque, and when I left it overnight I could see this layer between the white and a lighter layer on top (the water separated from tween 20 + oil?). So I assumed right off the bat that 5% Tween 20 wasn't going to work - I tested that too, and after one night i could see small oil droplets at the top. So today, I tried 15% Tween 20 (6mL of 25% Tween 20 [0.25 X 6 = 1.5, 1.5/10 = 15%], 1 ml oil, and 4 ml distilled water). It was still kind of cloudy, and I still couldn't really tell if there were any suspended particles - I don't think they were, but the mixture was a thick white, and many sources I read said I have to make sure it's clear to form a microemulsion. I then tried 20% Tween 20, but this time I got a different syringe for the Tween 20 and I just sucked up 2 mL of undiluted Tween 20, added 1mL of essential oil, and 7mL of distilled water. It was still pretty cloudy, but a little clearer. The same thing happened for 25% Tween 20 (2.5ml T20, 1mL oil, 6.5ml water). Then I tried 30% Tween 20 (3mL Tween 20, 1mL oil, and 6mL distilled water). This time, the mixture was very clear and you could not see any suspended particles in the solution.
The problem is, I'm not sure if 30% Tween 20 is too much. I am going to do my control testing this week at university to confirm this. Assuming it is not bactericidal to my E.Coli, I would then have to perform my serial dilutions of 10%, 1%, and 0.1%. So I take 1mL of the original 10% oil dilution and put it in 9mL of Tween 20 + water, and do the same for 0.1%. Won't that be too viscous and have too much Tween 20? Do I have to have 9mL of the same amount of Tween 20 and water, or can I just pour 9mL of distilled water? Is there any other way I can perform serial dilutions without making the resulting solutions too thick/inaccurate? Does this mean I should take a lower concentration, such as 25% or 20% Tween 20, seeing as the solution will turn clearer once I make my serial dilutions? I

https://docs.google.com/document/d/172M ... sp=sharing

^my dilutions

Hi GG,
You have made some good progress! I am a little worried that 30% T20 is going to kill the E coli or inhibit their growth. You might have to reduce your max EO concentration to 5% or maybe even 2%. You should have done the T20 toxicity test first because that is what will determine the highest concentration of EO you can use. What you would like is for the E coli to be VERY sensitive to the EO and much less sensitive to T20. If it is the other way around then you have a problem.

I did a search for "killing e coli with tween 20" and found this paper: http://www.ncbi.nlm.nih.gov/pmc/article ... 36/?page=3
They tested various detergents by adding them to liquid broth cultures rather than E coli growing on agar, but the result should be the same. They showed 50% growth inhibition at a concentration of 4% Tween20, so your 30% T20 is definitely way over the permissible concentration.

You were able to make a 10% solution of EO using 30% T20, so if the ratio stays the same, 4% T20 should be able to dissolve around 1.3% EO—maybe 1.5%. I would do the T20 toxicity test starting at 10% and working down by halves from there—5%, 2.5%, 1.25% and 0.75%.

Try to do the toxicity test right away because everything depends on that. You need to test the EO as soon as you get the results from the toxicity test, so make sure you have the agar plates and E coli culture all ready to go.

Good luck!

Sybee

OK, I was afraid 30% might be too much Tween 20.
However, if there is only 1.3% EO, that means I can't do my serial dilutions anymore, correct? (of the 10%, 1%, and 0.1% EO)

It just means that you cannot test 10% EO. Based on the T20 toxicity test, you will know the maximum amount of EO that you can dissolve and that becomes your highest dilution. Then you set up a series of dilutions of EO from there. Let's say the highest concentration of EO that you can dissolve in T20 without killing more than 50% of your E coli is 2%. Then you would dilute that by half to 1%, using the same concentration of T20, then by half again and so on. You have to use the same concentration of T20 each time so its effect on the bacteria will be the same and you can simply subtract it from each reading. Don't forget to always include a control disk with T20 only, at the same concentration.

Are you doing the test of EO concentrations today? Do it as soon as possible so you have time to repeat the experiment with the EO.

Sybee

So, I'm just going to use your example of 2% essential oil as an example to help me understand. Let's say that 2% EO is the most amount I can use. Since I concluded that 30% of my Tween 20 was what would completely dissolve 10% EO, using the same ratio for 2% EO that would be about 2% EO and 6% T20 (so, for example, I could put something like 2mL of EO and 6mL of tween 20 into 92ml of water in this example). Once that dissolves and I put in my disk dipped in this, then I would go on to 1% EO. I would STILL use 6% Tween 20, but my amount of water would change (so I could have, say, 1ml of EO, 6ml of Tween 20, and 93 ml of distilled water). Of course, this is based on the assumption that 6% Tween 20 is the maximum concentration of Tween 20 I could use without killing anything (which isn't necessarily true, I would have to find this), but would it be something like this?
I have the university booked for Wednesday afternoon so I can go find out the max. amount of Tween 20 I can use and base my ratios and experiment around that. One more quick question - if I have a very small, barely noticeable zone of inhibition around one of these disks, can I still use that concentration or do I have to find a lower concentration?
Thank you!