Science Buddies: "Ask an Expert"

Hi Trader,

The colony description sounds good; and opaque with darker spots sounds good too. I’m sure you have B. subtilis.

The “wrinkled” referred to the colony growing on the plate, not appearance under the microscope.

What do you mean by “faint?” Is the color pale, or is there no growth? If you touch a colony with a sterilized inoculating needle, and transfer this to a plate, and do a spread out the colony, sterilizing the loop 2-3 more times in between, you should obtain isolated colonies on the plate.

If you start your experiments with the homemade broth, you should continue using this medium even if the commercial nutrient broth arrives, just to keep this parameter controlled. You will be able to switch to the nutrient broth for the next set of experiments.

If you have 1 billion bacteria per ml, you will need to dilute your sample 1:10,000,000 to get 100 colonies on a plate. You can’t put 1 ml on a prepared agar plate and spread it around, otherwise it will be too wet and all the colonies will grow together; 100 uL or 0.1 ml is about the maximum volume for a spread plate. If you take 0.1 ml and dilute it in 10 ml of sterile water, this is a 1:100 dilution; if you pipet 10 microliters into 10 ml of sterile water, this is a 1:1000 dilution. You need to do a series of dilutions and plate out the dilution that you think will contain 30 to 300 colonies, plus one dilution above and below your target, just in case you miss the correct dilution. If you pipet 0.1 ml on the prepared plate, this counts as a 1:10 dilution. So, if you take your sample, and pipet 10 uL into 10 ml (1:1000), mix this dilution and then do a second 10 uL into 10 ml (another 1:1000 dilution); you be up to a 1:1,000,000 dilution. If you put 0.1 ml of this on a plate, and spread it around on the surface and incubate it, the results will be multiplied by 10,000,000 to get the original number per ml. Now, how will you do the 1:100,000,000 and the 1:1,000,000 dilutions (one below and above)? If you don’t have pipet tips for 10 uL, how will you do these dilutions? I know this is complicated now, but after you do it a time or two, it will be second nature for you.

The protocol you sent is a good standard protocol.

If there is some reason that you think your plates may have become contaminated, you could incubate them overnight. However, you don’t want to dry out your plates because this will inhibit growth. When you do your growth curves, if something has grown on one the plates, and you spread this around with your dilution, the mistake in the results will be obvious.

One option on doing plate counts is to do pour plates. This allows you to pipet a 1 ml sample into a Petri dish, pour in molten agar that has been melted by boiling and then tempered to 45 degrees Centigrade (warm but no solidified), and swirl it to mix the bacteria thoroughly. The agar is then allowed to solidify and dry, and turned upside down to incubate. Most of the colonies will be trapped inside the agar and the colonies will be easier to count. However, you should use up your prepared plates first before you try this.

Good luck on your growth curve experiments!

Donna Hardy

Unfortunately, it seems like none of the nutrient broths are available for now.

Our school doesn't have beef extract or peptone (which is somewhat disappointing...I thought those would be 'common materials'), and there appears to be no response regarding the nutrient broth order yet.

Is it possible to "substitute" for beef extract and peptone, such as having "home-made" beef-extract and peptone, and then using that to make nutrient broth?

Right now I've managed to obtain several good plates of b. subtilis and really am just waiting for the nutrient broth <_<. This is a lack of organization on my part...

Hi Trader,

Don't worry. The bacteria you are working with are not picky eaters. You can use the recipe for homemade media from the science buddies website:
http://www.umsl.edu/~microbes/pdf/homemademedia.pdf

Just use the beef buillon, sugar, and water. Leave out the gelatin, since you want a liquid culture. You bacteria will grow very well on this medium.

Donna Hardy

WOW. OK! This would definitely work.

By faint I meant that it had a hint of brown, but was almost translucent, as opposed to the recently inoculated b. subtilis which was brown and not translucent.

Just out of curiosity I incubated some of those "faint" plates overnight and they remained faint (though maybe my eyes were tricking me to make my mention that they may have been a bit browner?) -- I think that they are no longer viable.

It does strike me as weird however, because the "lawns" of the bacteria are actually not translucent, while most if not all of the streak plates are "translucent", so I'm thinking that the bacteria wouldn't be as able to survive if it were "alone"?

Another thing is that one of my lawns of bacteria seems like there is white powdery materials growing on it -- I hope I did the right thing by disposing of it? (w/ bleach overnight, of course ).

Another quick question (sorry with the list!) is about how transferring new cultures should be made -- I'm assuming that from now on, all new plates made will be streak plates so eventually we will dispose of the lawn of bacteria and that soon, there will just be streak plates? (though what worries me is that it seems like they get "faint" a lot sooner, and therefore we would need to transfer them faster? Though the hope is that after I make nutrient broth tomorrow, I would be able to spend an entire 12 hours doing plate counts so I won't have to transfer that much anymore.)

Some... bad news: I just found out that my top choice for a science internship, which was the "Summers of Discovery" Program offered by NIEHS had eligibility requirements that I didn't really fulfill... I needed to be in a U.S. institution for at least a majority of my educational background, and I've been in Shanghai almost my entire life . Aww... I have one internship left, and 3 science training programs left that all didn't shine as NIEHS did, but at least I'm eligible for them . In the case of the email reply:

"I am afraid that you would not be eligible for our program. As I indicated, it is for those who have received the bulk of their education in the US."

Would it be worthwhile to attempt to attempt to change their mind? (Though in my defense, the eligibility requirements outlined outside of the application itself stated that I be enrolled in a high school, U.S. college or university, implying that the high school need not be the U.S. -- though the U.S. part was explicitly added in the description that came along with the actual application).

Hi Trader,

Thanks for explaining your description. I think the "faint" color is a characteristic of the strain of B. subtilis, which you have. You don't need to worry about this bacterium surviving. It will remain viable for a very long time. Bacillus species are soil organisms, and are routinely exposed to wet and dry conditions, so are adapted to survival under adverse conditions.

The white powder sounds like a mold, so it's good that you discarded that plate. You don't want mold spores getting into your cultures.

You should plan to make new streak plates about once a week to make sure you have a good culture to select from. Keep at least 3-4 plates (up to 3-4 weeks old) to make sure you don't lose your culture. The Bacillus won't die out easily because is will make spores, but the E. coli will die if it dries out. Always check to make sure the colonies on your new streak plates have the appearance of the original culture, and redo the streak plate if it looks like you have picked up a contaminant.

I'm very sorry to hear about the rejection from the NIEHS program. It sounds like your application was rejected before anyone looked at it. I guess the rules were made to exclude anyone living outside of the US. I don't know how flexible the rules are, but it sounds like someone was reviewing the applications to make sure each applicant met the exact requirements and you got an automatic rejection. You probably should not spend a lot of time on the appeal, but it would not hurt to resubmit your application. If you are a US citizen, then do send a note to the person who send the rejection and explain why your application should be considered, e.g. you are extremely well qualified and you want to celebrate July 4 in the US for a change. It would definitely not hurt to explain how patriotic you are (your debate skills should help you here). If it would help any, you can say that your science buddies advisor thinks it is essential for you to participate in the program, as it will give you the experience you need to win next year's science fair.

Good luck!

Donna Hardy

YES. I can make it tomorrow.

*-- Update 2/19 --

Please reply soon... the weekend is coming up and hopefully I am able to do a test run this weekend, but there are many things I'm not sure about.

1) How many plates will I need? I was thinking about this below, and I'm thinking that if I test for the first 6 hours as a test run, typically do growth curve experiments involve 2 plates per hour or...?

2) How dilution plating really works. I'm afraid I am very unsure about many of the steps. This is my understanding of the protocol:

-->
1. If we are preparing 6 hours of testing and plan to test a total of 10 time intervals in it, then we will need 10 nutrient broth tubes (that has the oil on top removed [there was a layer of oil that hardened after being stored in the fridge -- this was from the homemade recipe]), of same volume that we need to record down.

Quick question: We would want approx. how much volume? I'm thinking that the more volume there is, the more diluted the bacteria would be in the first place. Does the culture naturally "spread itself" through the solution?

2. I best prepare 2 X the number of time intervals I'm testing of plates because two trials (or is it typically 3?) are best.

3. I incubate the 10 tubes at 37 degrees Celsius. After every time interval I do the following:

...1. Used a sterilized pipette to take 1 ml of the sample of the tube 1 (I'll use one tube for each interval) and then pipette it into sterilized water with 99 ml? And then take another 1 ml and then pipette it into a sterilized water tube w/ 99 ml, shake and then pipette 1 ml from that sample to another fresh sterilized tube w/ 99 ml for a 10^-3 dilution?

...2. I repeat the above for approx. 4 hours into it, and then increase the dilution factor to diluting 1 ml into 99 ml three times for 10^-4 (as well as having another tube that just has 10^-3) to measure the doubling time.

...3. Repeat the above for all time intervals.

...4. After we have the nutrient broth tube with the correct dilution, we pour 1 ml (only?) of the sample into two fresh petri plates and make a lawn? And then we incubate that (it doesn't matter which temperature?) for 24 h and then count the number of colonies that there are, and then times that by 100 for 10^-2 dilution, 1000 for 10^-3 dilution, etc?

But another thing that is worrying me is that

Is there any way to "reuse" the tubes above? It seems like I'm using most of the 99 ml tubes of sterilized water for only one time, putting 1 ml of bacteria then taking 1 ml out to dilute again -- is it possible that after we use it, we autoclave it and "reuse" it?

I'm basing a lot of what I know from your very helpful explanation earlier:

If you have 1 billion bacteria per ml, you will need to dilute your sample 1:10,000,000 to get 100 colonies on a plate. You can’t put 1 ml on a prepared agar plate and spread it around, otherwise it will be too wet and all the colonies will grow together; 100 uL or 0.1 ml is about the maximum volume for a spread plate. If you take 0.1 ml and dilute it in 10 ml of sterile water, this is a 1:100 dilution; if you pipet 10 microliters into 10 ml of sterile water, this is a 1:1000 dilution. You need to do a series of dilutions and plate out the dilution that you think will contain 30 to 300 colonies, plus one dilution above and below your target, just in case you miss the correct dilution. If you pipet 0.1 ml on the prepared plate, this counts as a 1:10 dilution. So, if you take your sample, and pipet 10 uL into 10 ml (1:1000), mix this dilution and then do a second 10 uL into 10 ml (another 1:1000 dilution); you be up to a 1:1,000,000 dilution. If you put 0.1 ml of this on a plate, and spread it around on the surface and incubate it, the results will be multiplied by 10,000,000 to get the original number per ml. Now, how will you do the 1:100,000,000 and the 1:1,000,000 dilutions (one below and above)? If you don’t have pipet tips for 10 uL, how will you do these dilutions? I know this is complicated now, but after you do it a time or two, it will be second nature for you.

I'm afraid my concepts once again aren't very good here. If 1 colony of bacteria means about 1 billion bacteria, and I inoculate that into a tube w/ 100 ml nutrient broth, that means that I'll have 10,000,000 bacteria per ml, and after already that I start incubating the sample?

I think I might have pipettes for 10 uLs, but if I don't, I'm thinking that 10^-6 in worse case scenario would mean that I do three dilutions of 1 ml in 100 ml? Would this be practical? Though this leads me to another question -- How can I be sure that through the diluting, the bacteria is well spread throughout the water that the 1 ml sample that I take contains the bacteria (and nothing else?)

This is the list of what I think I'll need:

- 10 nutrient broth tubes
- 4 streak plates, total of 10 individual colonies
- ?? tubes with sterilized water (If I can "reuse" them, do you know how long it might take for the UV light to sterilize each tube? Thanks)
- 10 X 2 plates of nutrient agar for creating a lawn of the diluted sample in, then incubating and counting the colonies later.

Regarding the streak plates, the morphology appears to be the same among all the plates, but in some cases there is one individual colony that is the same color, and wrinkled, but the color is a bit lighter than the rest. Would it be safe enough to take an individual colony of the isolated colony that looks the same as the rest, or would I have to make a slide out of each "half" of a colony to identify it, then inoculate the remaining half into each nutrient broth plate?

THANK YOU SO MUCH. I am very desperate -- I truly hope I can get a test run in by this weekend!

... I'm sorry if my questions are too general -- here is truly my understanding of the protocol. If there's something I'm doing "wrong", please do tell me . I wasn't able to make the weekend because I think that I'm not ready -- there are some ambiguities that I truly do not know how to solve!

1. I should make 10 nutrient broth tubes. That is done by first autoclaving the nutrient broth. Their volume is best at 99 ml? Or else I would have to dilute it a lot later.

2. I should prepare 20 tubes of sterilized water, because that would mean that I can on average, dilute the sample twice. They all contain exactly 99 mL and 9 uL, making it 999 uL?

My biggest question remains:

If I incubate the bacteria for 1 hour and then test the bacteria population, I think that it would still be in the lag phase, so wouldn't there still be one colony in the nutrient broth tube? -- Would the "1 billion" bacteria only result at the stationary phase? Or have I completely confused the concepts and should know that I only start counting the population after a 24 h incubation?

3. After the appropriate dilutions, I make 2 lawns of bacteria from each nutrient broth, and then incubate that for 24 h, and then if the calculations are correct, there are "dots" on the plates hopefully between the numbers of 30 and 300.

I apologize if I did something in the post above that made you go online but not reply to my thread. If that's not the case, I apologize for misunderstanding that. -- I truly want this to work out.

Is there any way to "reuse" sterilized pipettes?

Because I'm thinking that I pipette 1 uL into 9.9 mL tubes 1 times + 1 mL into 9 mL sterile water, then take 1 uL of that to 1 mL of that. But that'll take a lot of pipettes. Currently I have 10 mL glass pipettes that measure to the uL, but it would be a waste to throw the entire thing away after one use. So

Is there any way to reuse pipettes?

As for the original tubes, please help me understand: It IS ok if we have 20 20 mL nutrient broth tubes (in normal test tubes) for 20 interval tests, and we DO begin our first dilution of 1 h after incubation, after exactly 1 h after incubation after inoculating right?

What I'm not sure about is whether the 1 h will produce enough bacteria to be counted.

Thank you very much!

Hi Trader,

Unfortunately, I don't know of any way to reuse pipettes. There would be too much carryover going from a high concentration of bacteria to a lower concentration. If you ever have the occasion to pipette multiple samples, and can start with the low concentration and then go to a higher concentration, then it's OK to reuse the pipette for multiple samples. Otherwise, it's not a good idea. What type of pipettes are you using? Do you have a handheld pipettor with disposable pipette tips?

I'm not quite sure what you mean by "20 20 mL nutrient broth tubes (in normal test tubes) for 20 interval tests." You should have one nutrient broth culture growing that you take samples from periodically to do the plate count. These bacteria can double in number every 20-30 minutes, so a one hour interval between plating is not too short of a time interval between samples.

Donna Hardy

OH.

What I thought was that there were 20 tubes, each w/ 20 ml of nutrient broth in it. And then I inoculate each w/ b. subtilis and then every hour, take one at a time and do dilutions so that I could control the volume. Does this mean that I have one 50 mL or so nutrient broth, inoculate the bacteria and after 1 hour take 1 uL?

Does this mean that I only have one 20 ml tube and I take 1 uL sample of it from each? It would be OK then that the volume periodically decreases, increasing the concentration of the bacteria population in addition to its growth?

As for the pipettes, I have ones that are made out of glass and are quite large actually -- they measure 10 mL (though have uL increments). Since I'll be pipetting from a higher concentration to a lower concentration, it'll seem like I'll have to either find an alternate source of pipettes (esp. the plastic "one use" small ones) or find someway to reuse them.

Thanks!

Hi Trader,

I assume you are using 20 tubes because you don't have a larger container. Since you will only be taking 1 uL of sample, you could do the growth curve study using one 20 ml tube. Using 20 separate tubes will really be generating 20 different growth curves. Normally, you would use just one tube or flask, and periodically take samples from the same container. Even if you take 0.1 ml ( 100 uL) for a sample and do 10-12 samples, this won't significantly affect the volume of the original tube.

I have been thinking about your pipette problem. You can save your pipette tips; sterilize them in a 1:200 dilution of bleach for a few minutes, in 70% ethanol for 1 hour, or in 1 M NaOH for an hour. This will kill the bacteria on the pipette tips. You will then need to sterilize the tips for reuse. If the pipettes have been irradiated to sterilize them originally, they will be brittle and you can't autoclave them, but you could use 70% ethanol and rinse them well in sterile water. You can't leave any residual ethanol, bleach or NaOH on the tips because these will kill your B. subtilis and E. coli. Be careful with the bleach; sodium hypochlorite is a strong oxidizing agent, and will quickly degrade the plastic pipette tips. If you use bleach, just soak the tips for a few minutes, and immediately rinse in water. Does this give you any ideas for use with your available resources?

Donna Hardy

Interesting -- I have only prepared 9.9 mL tubes (for 1 uL dilutions so that it can be a 10^-2 dilution) as well as 9 mL tubes (for 1/10 dilutions) of sterilized water.

I do have access to containers that are large enough but I am not sure what type of a container we are looking for. I'm thinking that we probably wouldn't be wanting to use beakers (unless it has been done before and/or is what we are supposed to use? Right now my nutrient broth is in an Erlenmeyer flask as I have just autoclaved it -- I'll do a test run by taking an identified individual colony, dipping it in the nutrient broth, and then somehow shaking it (I'll check on the shaker) --, then in around maybe 12 hours I'll expect the population to be about 1 billion -- perhaps I'll do a dilution then to get a feel of the technique.

>> If I don't have a shaker, is it OK if I leave a magnetic rod inside the Erlenmeyer flask and then place it on a stirrer plate and leave that on? Though then I wouldn't be able to incubate it...

Thank you for the reusing technique.

There's another problem I'm not too sure about -- right before I pipette 1 uL of a solution with the bacteria into the plate, should I add 9.9 mL of sterile water so that the bacteria is spread evenly?

Hi Trader,

An Erlenmeyer flask with a sterile cover is a good container for the growth curve. If you used the 20 ml tubes, it would be difficult to aerate the culture. I don't know how you would keep a sterile cover on a beaker. Leaving the magnetic rod in the flask and placing it on the stir plate would be a good way to aerate the culture. However, if you can't keep the culture mixing during the growth curve, you can just keep the culture static. The bacteria will grow slower, however. I've never done growth curves on static cultures, so I don't know what the doubling time would be without oxygen, but I'm sure it would be slower than a 20 minute doubling time after 3-4 generations. Incubating at ambient temperature at about 18 degrees C instead of 37 would also slow the doubling time to maybe 60-70 minutes. Mixing the culture during the growth curve is one of your controlled variables, so you should incubate under the same conditions (mixing, temperature, composition of medium, etc) and continue doing whatever you do on your first experiment. Is there room in the incubator for the stir plate if you tape the door of the incubator shut? You will have to consider your resources and make an executive decision on what to do about growing cultures static or with mixing.

It usually works well to use about 0.1 ml on the surface of the agar plate. This gives enough liquid to spread the sample over the surface of the agar, and is a small enough volume that the surface of the plate will dry out before you invert it for incubation. Using 9.9 ml would leave the surface of the agar wet and would allow the bacteria to grow altogether.


Donna Hardy

I'm pretty sure I don't have a sterile cover ... I can double check on that.

Will plugging the Erlenmeyer flask with cotton and covering it with aluminum foil work to keep the solution sterile? Though I'm thinking that it wouldn't because I already autoclaved it and the nutrient broth appears to be a bit murky. I'll double check on that by making it one of my first tries in understanding just about how much bacteria would be in there (hopefully it represents maximum bacteria). At least I'll have an idea.

As for inoculating loops, I just have a small question. Mine are plastic so I'll have to sterilize them every time I use them. Would soaking them in ethanol overnight sterilize it? What I'm a bit not sure about is what to do after it stayed overnight. Sorry, my sterile technique concepts are really bad. Do I wash it with tap water (though that would add bacteria into it again) and then dry it with a towel (possible contamination everywhere!).

As for taking samples out of the Erlenmeyer flask, if I have a sterile cover, how can I ensure that the nutrient broth isn't contaminated with bacteria other than b. subtilis after I remove the cover to take a sample to dilute the plates with?

Thank you! I've prepared 9.9 mL tubes as well as 9 mL tubes. I love this.

Just out of curiosity I think I'm going to find out what bacteria exists in the air. Perhaps open a petri plate for about 5 minutes, and then see what settles in. Perhaps I can learn to identify these bacteria to know whether or not my plates are actually contaminated.

As for identifying the b. subtilis, I made a slide out of one of the individual colonies in the streak plates, but I'm afraid all I managed to see under the microscope was several long S shaped "string-like" things. There were also several round, very dark dots at the lowest magnification. I'm sure this is not bacteria as even under x1000 magnification b. subtilis is very very small. I redid the slide with another individual colony, and so I also wanted to clarify, while previously I made a slide out of a lawn of bacteria, when making a slide out of one half of an individual colony from a streak plate, how is it different from a lawn of bacteria? Would it be just like the lawn of bacteria (with rods everywhere), only in a smaller proportion?

One last question (thank you very much for bearing with me) -- When microbiologists do simple experiments like these, they do confirm each new streak plate by making a slide every time right? Even though the morphology between the old plate and the new streak plate looks almost identical?

Thank you so much!

Hi Trader,

You have some good questions. I can tell that you are good about thinking about the details of your experiment. You will appreciate it sometime in the future when you have access to a lab with all of the equipment you need.

If you use a cotton plug covered with aluminum foil on the Erlenmeyer flask and autoclave it with the nutrient broth, the cotton and aluminum will be sterile at that point. If you carefully remove the cotton/aluminum foil, and don't expose it to air or set it on a surface (kind of hold it by the top without touching the surface that will go back into the flask), it will remain sterile enough to get through an experiment. You might want to autoclave extra cotton plugs and wrap them in aluminum foil to have on hand in case the original gets dropped on a non-sterile surface.

Soaking your plastic inoculating loops in 70% ethanol for one hour or longer will sterilize them, and it's chemically compatible to do this overnight. You would need to dry the loop and make sure the ethanol is gone before you use the loop, however. Using a dry towel or paper towel that had been wrapped in aluminum foil and autoclaved would be a good way to avoid contaminating the loops. Tap water or a non-autoclaved towel would contaminate the loops.

During the growth curve, the E. coli and B. subtilis should be the most abundant organisms present, and a stray contaminant will not affect your results significantly. If contamination occurs early, however, you will see a difference in the colony morphology and other characteristics of the culture. Don't put anything that is not sterilized inside the flask except the starting cultures you want to study.

If you leave a Petri dish exposed for 5 minutes to open air, you are bound to pick up some contamination. Mold spores are the most common air contaminant, and Bacillus species would be common also. If you want to have a little fun, sprinkle a small amount of dirt from the garden into a Petri dish. You will grow some UV-resistant pigmented bacteria and probably get an interesting assortment of different colors.

I wonder if you are putting too much sample on the microscope slide. Try transferring just a small amount of a colony into a drop of water. It should be just slightly cloudy before you air dry it. If you can see a heavy visible white film on the slide before you heat fix the slide, then try adding more water to make the bacterial film thinner. This should give you a slide with a single layer of long thin rods under highest magnification. Your slides should look like the pictures you have seen.

It is best practice to do something to confirm your culture with every experiment. You are becoming familiar with the appearance of your cultures, so doing a streak plate is fine and verifying that the appearance of the colonies is as expected is good. It would be OK to skip this step if you are doing several experiments in a row.


Donna Hardy

Thanks for the reply! I tried a test run today and I've made 10^-2 and 10^-4 just to check what I should be expecting.

I think I'm going to avoid diluting 0.1 mLs, because the pipette only has increments of 0.1 mL. I'm going to stick to using a total of 6 tubes for 1 ml in 9 ml dilutions so that it'll be more accurate.

I've dropped approx. 0.1 mL in a petri plate and the water only managed to occupy about 1/3 of the plate -- is this OK? I'll have to see soon .

As for the inoculating loops, I suggested the idea of autoclaving towels but my teacher said it would work if I leave the inoculating loops in ethanol and then let it air dry. I'm a bit skeptical about this (b/c wouldn't exposure to air immediately contaminate it?). Would this work?

I know I'll have to reuse pipettes soon. In the two dilutions I made today as a test run (I've never done it before), I wanted to sterilize the pipettes through leaving the pipette tips in ethanol. But then, the surface inside would still be contaminated with the diluted nutrient broth. I think someday I'll have to close them up in ziplock bags and autoclave them altogether?

I also wanted to make sure -- I've previously wrapped the cotton plugs of the 9 mL sterilized water tubes with aluminum foil. After autoclaving, I do remove the aluminum foil right? I'm not sure about this because if I don't remove the aluminum foil, I almost can't "replug" the cotton w/ aluminum foil plug back in because previously it was very well wrapped. But I also unwrapped one cotton plug with aluminum foil today and felt that the cotton was a bit wet. I'll double check -- maybe it was me thinking that it would be wet?

I also wanted to make sure on another thing -- in terms of the entire process, if I'm going to make 10^-2, 10^-3, 10^-4, 10^-5, 10^-6, only 5 tubes will be needed right?

That means that I'll need to pipette the entire solution of the 10^-2 solution and pipette 1 ml of it in another 10 ml solution, and then take a new pipette and take 1 ml of that solution into the next 9 ml tube of sterile water. This is very cool .

Hi Trader,

Let me know what happens in your test run. This is exciting that you are about to get some results.

When you are more experienced in pipetting, you will be able to pipette 0.1 ml accurately and reproducibly, but for now you are probably correct to do the 1:10 dilution with 1 ml of sample. I like your cautious and careful approach to this. It sounds like you have traditional pipettes, and not the small micropipettes.

I don't know why 0.1 ml only covered 1/3 of the plate. Did you try spreading the 0.1 ml sample over the entire surface with a spreader of some sort? You need to spread out the sample over the whole surface so the individual bacteria will have room to grow into colonies that you can count. Are your agar plates dry? If the sample won't spread over the whole surface, add another 0.1 ml or so of sterile water and try spreading it again.

Your teacher is right. Air drying for a short time is acceptable. Just make sure the loops are completely dry before you use them. If one stray contaminant land on the loop while it is drying, it is going to get lost in the billions of bacteria you will be transferring.

Sorry. I forgot to mention that you have to wash the pipettes in soapy water to remove the nutrient broth and other residue. After you use the pipette, soak if for a few minutes in disinfectant to kill everything. Then, the best way to wash traditional pipettes is to have a long thin reservoir of hot soapy water than you can completely submerge the pipettes in. Devise some way to hold a bunch of the pipettes vertically, then dip them in the soapy water and then raise them up and let the soapy water drain from the inside of the pipette. Repeat this a few times until the pipettes look clean. Then rinse in 3 changes of tap water, and finish with a final rinse in distilled water. Then soak them in the 70% ethanol again for an hour or autoclave, if they are autoclavable to sterilize. You will have to adapt this procedure to your available resources, of course. Everyone in a lab has to spend time washing glassware for their experiments; it's really important to do experiments with pipettes and containers that won't add anything to your samples.

You should leave the aluminum foil on the tubes and flasks until you are ready to use it, but yes you have to remove it in order to remove the cotton plug. If the cotton was wet, your container may have been too full for autoclaving, or you may have released the pressure on the autoclave too quickly. The autoclave should depressurize under very slow exhaust to avoid boiling the nutrient broth/water. If the pressure is released quickly, you could lose all of the liquid from the containers.

For your dilutions: If you take the original sample and pipette 0.1 ml into 9 ml; this is a 1:10 dilution. If you take 0.1 ml and pipette it onto the plate, you label the plate 10^-2. If you then mix the 1:10 dilution and pipette 0.1 ml into the next tube, this tube will be the 10^-2 of the original, and a 0.1 ml sample transferred to the Petri dish will be the 10^-3 dilution. So, yes you are correct. You will only need 5 tubes for this series of dilutions.

You are doing great!

Donna Hardy

After 24 h incubation at 35 C, it seems like there was no growth on the plates. In the 10^2 (I made a 10^-2 and 10^-4 plate), I thought I saw individual "dots" on the cover and not on the actual agar; maybe that was me seeing things?

I'll incubate it for another 24 h and see what happens. This is based on the nutrient broth that did look murky (though it was autoclaved with the cotton plug -- I thought it was contaminated b/c it was a bit murky, but if it's not my plates, then the dilution plating results appear to say that it isn't contaminated?)

I also did another dilution on a sample of known contamination so I've diluted 10^-2 and 10^-3.

I think the soap technique may be a bit too tedious -- I think I might soak the pipette tips in ethanol, and then let them air dry, put them in a ziplock bag, then autoclave them all together sometime? The soap idea seems like it'll take up a lot of sink space, though I'm pretty sure it'll be OK if I go through the process all at once some time.

I'm thinking that the petri plates may be too dry. I did a sample experiment where I left the cover open for about 15 seconds, and then closed it, and incubated the petri plate at 35 C. I couldn't find anything on the agar after 24 h...

I made sure that I pipetted about 0.2 mL of the diluted solution and it still couldn't cover the entire plate.

Those plates are also over a month old, and I'm pretty sure I have not followed the proper way to store them. Just to check (I'll make a new batch and start with these now), the plates should be stored in ziplock bags, in the fridge right, for up to a month?

Also regarding the structure, I find that it would be very convenient that after I've pipetted some of a 1/10 dilution for example, to pipette 0.1 mL into a plate (a fresh batch that'll hopefully spread), and then while I wait approx 15 seconds to prepare for a second dilution, leave the pipette (with some of the 1/10 dilution) out in the open air. After about 15 s, I'm ready to pipette 1 mL into another 9 mL tube. I'll do my best to be fast in those minutes, but is "OK" right?

Hi Trader,

I think you are right. Dry agar plates will not support the growth of bacteria. I doubt if the plates will grow with a second day of incubation. 0.2 ml of diluted sample should definitely be able to cover the surface of a plate. You should definitely prepare new plates, and include a positive growth control with a heavy inoculation of your cultures to make sure they will grow. What was the starting culture that you made the 10^-2 and 10^-3 dilutions for? Was this from an agar plate, or from an overnight broth culture?

Did the murky nutrient broth smell bad before you added the bacteria to it?

Yes, I think your protocol of doing the dilution and leaving the pipette exposed for a few seconds will not cause any problem with contamination. You don't have to rush, just keep working and don't stop to take a break until all the dilutions of the sample are plated. You should mix the dilution before your pipette the 0.1 ml volume onto the plate. After you have pipetted all of the dilutions for a sample you should use a sterile spreader (glass or wood) and push or spread the 0.1 ml sample over the entire surface of the plate so it is evenly distributed. You can use one spreader if you start with the highest dilution sample and work back towards the highest dilution.

Do you have a thermometer in the 37 degrees Centigrade incubator? Temperature could be a cause of no growth also.

Donna Hardy

W00T!

It turns out that even after 48 h, there are no colonies -- I think that's because the original nutrient broth (the one where I autoclaved, but when I took out, thought was very murky and almost had a "dust like" string in the broth before I swirled and made it murky) is probably NOT contaminated w/ b. subtilis.

I'm thinking this because I've done another incubation of 10^-1, 10^-2 and 10^-3 dilution plating with nutrient broth known to be inoculated w/ b. subtilis, and after 24 h incubation of around 35 C, there are dots! I've never been so excited haha

I can see where the liquid has spread (and it has definitely not spread throughout the entire plate), and I can see that there are a lot of dots, too many to count in 10^-1. In 10^-2 the dots are much more spread out. I'll count them soon and see what the difference is. Strangely enough though, I was looking most forward to my 10^-3 yet I found nothing. Perhaps I mixed up the tubes previously? Hmm...

Either way, at least I know what to do now. My teacher suggested using the dry plates up anyways (I have 22) as part of a "real test run" -- is it OK if I spread 3 mL (I estimate will be able to cover the entire 'dry' plate) in each plate in the "real test run"? After this test run, I'll be able to get to the real thing.

I was really expecting there to be no colonies (b/c I thought there would be some little mistake I made), but there are dots!! Evenly spaced dots! What's best is that the lack of colonies in the now "confirmed" no-contamination, "properly autoclaved" nutrient broth means that I probably had an OK grasp of the sterile technique involved!

This is great.

Hi Trader,

This is great. It's exciting to get your first results, and dots are good because it means your colonies are growing! Good for you. I'm so happy you finally have something, even if it's too many to count (TNTC). You'll be able to tell when you get the right dilution because there will be between 30 and 300 individual colonies. Yes, if there is TNTC on the 10^-2 and nothing on the 10^-3, it probably means you made a small mistake. Did you mix each tube thoroughly before you pipetted? Don't worry, the mistakes will go away as you get more practice. Your sterile technique is obviously excellent.

Yes, please follow your teacher's advice as he is there and can see what's happening. The bacteria don't grow on the dried plates because there's not enough moisture. You could try adding a larger volume of water to the dry plates to rehydrate the plates. The surface would have to be dry before incubation however. Or, after you prepare the dilutions on the plates, you could add an overlay of 5 ml of freshly prepared agar that has been tempered (cooled) to about 56 degrees Centigrade. Pour a little molten agar on the plate, swirl it around immediately to cover the surface, and let it solidfy and dry before inverting for incubation. If the moisture content is high enough, the bacteria will grow.

Donna Hardy

Sorry as I approach the deadline I'm thinking some things over.

Some things about previous posts:

After you have pipetted all of the dilutions for a sample you should use a sterile spreader (glass or wood) and push or spread the 0.1 ml sample over the entire surface of the plate so it is evenly distributed. You can use one spreader if you start with the highest dilution sample and work back towards the highest dilution.

This may be the reason why the 10^-3 didn't show anything -- all I did was swirl the flask briefly and I didn't use a sterile spreader -- when I put in 0.2 ml, I only covered the petri dish up and moved it quickly in a figure 8 pattern to attempt to get the liquid over. According to my dilution plating protocol, it says to pipette 1 mL from a 10 mL solution into a plate -- does this mean that I am pipetting too little to start with? (Though I'm also sure that my plates are too dry also).

Also with regards to sterilizing the pipettes for reuse again -- with the process of using the soapy water and the 3 changes of tap water, to finalizing a rinse with distilled water and ethanol, would having a cotton plug in the end of the pipette change any methods? Because I realized that when I soaked the pipettes in soapy water, remains of the soapy water would be in the cotton, and would be part of the distilled water run and the ethanol...

Would it be OK if I wash the pipettes through 3 runs of water, put them in a ziplock bag, and then autoclave them? I'm thinking the ziplock bag because they'll remain sterilized until I open the bag for reuse again.

Hi Trader,

It's good you are thinking about the deadline. Try not to do experiments too close to deadline so you have time to do a good write up. You can start preparing sections of your board now and plan you layout so you will know how much room you have for results and conclusions. I think you have the question, hypothesis, materials, methods, and bibliography written, but you might want to look over content and make final revisions now. Your teacher probably gave you an outline of everything that needed to be included, so look that over and make sure you haven't forgot to include any details.

I don't think that your spreading technique caused the lack of growth on the 10^-3 plate. The bacteria would have grown if they were there and the plate wasn't too dry. Your protocol says to pipette one ml of sample into 9 ml of water to make a 1:10 dilution. If you pipette 1 ml of the 1:10 dilution into the Petri dish, you would multiply the number of colonies that grow by 10 to get the original number of microorganisms in the sample. If you pipette 0.1 ml of the 1:10 dilution into the Petri dish, you would multiply the number of colonies by 100 (1:10 x 1:10) to get the original number. If you pipette 0.2 ml of the 10^-3 dilution into the Petri dish you would multiply the number of colonies by 5000 (1:10 x 1:10 x 1:10 x 1:5). Think about this and make sure you are making the dilutions you intend to make. If pipetting 1 ml gives you the dilution you intend to plate, and if this doesn't leave too much liquid on the surface of the plate, then that might be the best way for you to do this. I would avoid using anything other than 1 ml or 0.1 ml, as this would give you an odd dilution and make calculations more difficult. If 0.1 ml gives you the dilution factor you want, and it the liquid volume is insufficient, then add sterile water to the plate to increase the volume.

If the pipettes look clean enough, then just rinsing with water should be good enough. If you do submerge them, you will have to remove the cotton. If you autoclave the pipettes and they stay clear, then they are clean enough. The zip lock should keep them sterile until you need to use them.

Donna Hardy

Thanks for the protocol on sterilizing and reusing the pipettes!

I was wondering if we could re-use petri dishes? I know I'll be using a lot and it would be cool to have a way of reusing them. But then I'm thinking that there would already be growth on the actual plate, and the entire agar would have to be removed. That is probably not the best idea.

I think I know why the supposed "10^-3" didn't have anything appear up. I was diluting 1 ml in 9 ml, and then taking 0.1 ml from it, thinking that it was a 1/10 dilution. But actually that is already a 10^-3 dilution, so what I thought was a 10^-3 dilution was probably a 10^-7 dilution or something? Perhaps I also made a little error.

I just wanted to make sure: I think I can only swirl the test tube briefly to attempt to mix the bacteria through the dilution -- would this be OK enough, esp for later dilutions where the bacteria would be a very small part of the dilution?

Thanks

Hi Trader,

I've never tried reusing Petri dishes. You would have to dump out the agar into a bucket of bleach, soak the Petri dishes in a disinfectant that wouldn't damage the plastic, and then clean them and sterilize and rinse them in sterile water. It would be a lot of work, but since you are working with non-pathogenic organisms, it would probably be OK. If I were you, I think I would try to get as much data as possible with the sterile Petri dishes you have available, and stop the project and write up the results. You are getting very valuable experience just doing the experiments, even if the results are not turning out perfectly.

Good for you! You figured out what happened. If you transferred 0.1 ml into 9 ml, that is close to a 1:100 dilution, so yes, your sample was much more dilute that you thought.

You need to try to mix the sample with the water as thoroughly as possible. Otherwise, your next dilution will be inaccurate. Try putting 0.1 ml of food coloring in a 9 ml tube of water and see what you need to do to get good mixing. It's OK if it takes some time.

You are welcome!

Donna Hardy

I'm testing to see approx what number of "max viable organisms" I'm looking at just to make sure that I'm doing the dilutions right.

As for the previous nutrient broth which I thought was contaminated, and thought that it wasn't contaminated after all, I'm thinking that perhaps it is contaminated after all. I took a close look at the plates today and there is a circle of "dots" around the edge of the petri dish, and then near the center there isn't any. Near the center of the cover however, there are little dots, actually on the agar-less cover. Perhaps this is water dripping down from the agar? But I didn't put that much water...

Either way, I'm almost pretty sure the nutrient broth (that I said looked murky when I took it out of the autoclave) is contaminated, but I'll do a double double check just to make sure . But then I did use a cotton plug and aluminum foil, so I don't know why it doesn't work ...

I also wanted to clarify -- we would want to keep volume constant as well right? That means if our first trial tests bacteria in 1 h, in 500 mL nutrient broth, then we make sure it's 500 mL the whole way? I'm thinking because a difference in volumes would mean difference in concentration of bacteria.

Oh no...then does that also mean that we would want to make sure we have roughly the same amount of bacteria inoculated into each 500 mL nutrient broth to make sure that we maintain that concentration? I'm thinking this because we'll be taking a sample from that concentration, and if it is altered, then we would have different population reports...

I also wanted to double check on the accuracy of "similar morphologies". So far all my plates have "similar morphologies", but when I checked again today, the first time at 40X I saw rod-shaped organisms and at 100X (w/ oil emersion) I couldn't properly focus it so I tried again.

When I used the lens paper to rub the oil off, a small patch of gray remained (as opposed to a large patch of gray in the microscope trial #1) so this time I focused only on the gray patch.

I'm pretty sure I got the right sample this time because now I managed to easily increase in magnification, including getting the 100x mag. BUT... whatever the slide consisted of wasn't rod-shaped at all. Whatever it was a sample of was not of constant size, and almost seemed to have little "round dots" (they were relatively large actually. The largest one took up approx. 1/4 of what I could see, while the smallest one probably 1/100) that overlap, and have varying circles, ovals (though generally round shaped.

I'm sure this wasn't b. subtilis... But then I'm also pretty sure no bacteria can be that big. I'm still pretty sure what I have was b. subtilis because of the similar morphology, but I just wanted to know -- usually how "accurate" will a "similar morphologies" observation hold for making sure that we have the right culture?

I also recall in microscope trial #2 that in the 40X magnification, prior to getting the finest focus for that level, there were little rod-shaped things that made up each of the "round dots" described above. I thought that it looked a bit out of focus, but now that I think back, perhaps I was in the wrong focus the entire time? Because the round dots I saw in 100x had a very clear body, and had only a black outside lining.

As for re-using the plates, I think I might drown the plates in disinfectants overnight, and then clear the agar, dry in ethanol. I'm not sure about this step though: would it be OK to let the ethanol airdry after a day's worth of soaking, and then close the petri plate, then pour agar into it again?

If this works, then I can pretty much go all the way through! .

As for pipetting, I've tried to pipette 0.1 ml but the smallest amount I've managed to pipette accurately is still 0.2 ml. I have one of those pipette pumps that has only a round button for me to push to pipette the liquid out, and even if it's a brief push, 0.2 mL is already out. I think I'm going to stay with making 1/10 dilutions.

And laaast thing: when I make sure that there is no ethanol on inoculating loops or the plates (if the above method works), that means that it is completely dried right? Or does that mean that I'll have to wash with sterilized water? Though what confuses me is, after we wash the surface w/ sterilized water, won't it still be exposed to air and then will still be contaminated?

I know that my project isn't original at all, but it's still something I can't really find the answer to on the Internet (what is meant when people say optimum temperature?) But I've already learned so much. It's great to go through everything now with more efficiency than before .

Thanks once again

Hi Trader,

You have lots of good questions. A circle of dots around the edge of the Petri dish sounds like contamination from somewhere. A free drop of water in a Petri dish will pick up bacteria that you have inoculated and spread them all over. When preparing a plate for incubation, it’s important that the entire surface liquid is evaporated, and the plate inverted for incubation. I thought your plates were dried out and the sample did not cover the agar surface. Where did the extra water drop come from?

Did the nutrient broth turn murky a few days after you autoclaved it? If the cotton plug, and aluminum foil cover was still intact, this probably means that the autoclaving didn’t sterilize the broth. How long did you autoclave the broth? Do you have a record of the temperature during the autoclave cycle? That would give you some clues about what may have happened.

It’s not critical to keep the original sample volume constant. The numbers are bacteria are counted as organisms per ml, so it doesn’t matter if you remove some volume for samples and the whole culture should be homogeneous, so a 0.1 ml or 1 ml sample will represent the number of bacteria in the whole culture. You’ll get the same results with 400 or 500 ml of nutrient broth.

After you inoculate the sample, the number of bacteria will be increasing over time, so you won’t be maintaining the same concentration. The number of bacteria in the sample will change, so you will get a different number every time you take a sample. It’s like a moving target. Let me know if I have not understood your concern on this question.

It sounds like you are not able to focus properly when you switch to high power. Try focusing on the 40 x magnification, and then switch to the 100x power without moving the slide and then you should need just a minor focus adjustment and maybe a higher light intensity to focus on the cells at high power. Your description of the high power field sounds like artifacts and air bubbles. Try again and be patient with the fine focus adjustment knob. You will know when you have the right focus. Hopefully, there is not a problem with the high powered lens on your microscope.

Yes, if you let the ethanol air dry, the Petri dishes should be good to use. However, do minimize the time of exposure to open air and dust conditions to minimize contamination in the ethanol-soaked plates. Residual ethanol will kill the bacteria, so you have to make sure it is evaporated.

Next time you pipette, after you draw in the sample with the pipette pump, take the pump off the top of the pipette, and quickly put your index finger on the top of the pipette. I think you will be able to let 0.1 ml out of the pipette accurately with hand control. You have to use the pipette pump to draw in the sample, but you can let it out by hand, and this will give you better accuracy. Practice a few times, and see if it doesn’t help. Pipetting technique always improves with practice, but it sounds like your pipette pump is not working well. Are you using the round rubber type?

Optimum temperature refers to the temperature that allows the highest rate of growth.

Keep going. You are making progress, and your project is much more original than you realize. Your preserverance in overcoming all of the obstacles all of the obstacles you have encountered on this project is amazing.

Donna Hardy

I think I'll be ready to do a test run of 6 hours soon (12 hours will be pointless, as b. subtilis enters the stationary phase at approx 5 - 5.5 h apparently, making that the highest amount of viable organisms).

Tomorrow I'll try to see what happens in the first 4 hours. It'll be probably the 3-5 h that will be the most critical because those will be the doubling times... I'll have to see when the log phase really kicks off .

I checked my autoclaved tubes of 9 mL sterile water today, and it turns out that they aren't close to 9 mL of water anymore o=. Does this normally happen? (I checked the average volume of 4 tubes and found it to be around 7.9 mL... that means that I'll have to dilute 0.79 mL into approx 7.9 mL of distilled water to still get the same dilution ...)

But just to increase the accuracy, is there any way to "prevent" losing water during autoclaving? I'm thinking that some of the water is in the cotton, but that's expected because it's a very high temperature in there.

Another possible reason is sometimes, I prepare the 9 mL of distilled water in the tube, plug it w/ cotton, and then autoclave it the next day (b/c there's always something I'm autoclaving and sterilizing water isn't priority yet =P) -- I think it would be wrong to assume that plugging the tube w/ cotton would "slow" down cotton right?

I also have a question regarding it not being critical for volume to remain constant. What I'm thinking is, if there's a 200 mL and a 500 mL nutrient broth inoculated with the same amount of bacteria, then it's population would be the same. Do you mean that it's not critical b/c we can take 1 mL from each sample, and even though 1 mL from the 200 mL nutrient broth will have more bacteria per mL, we can just times it by 200 and expect to get the same number as population of 1 mL in 500 mL X 500?

I just realized this a short while ago ... that's after I attempted to test for the "maximum population" with a 20 mL sample of bacteria. I thought that making 10^-6 dilutions would work, but even with that it's TMTC. But the cool thing is, I can definitely see the bacteria all spread out! If I had the time...and the energy, I could actually count the colonies! .

Since I want to use as little plates as possible, I'll try to get a better idea of the exact population of bacteria after so and so time so I'll do the necessary amount of platings .

OOOH. And I can pipette 1 uL NOW!!!! I'm so happy!!! . Now I'm actually using just 4 tubes for 10^-1 to 10^-6. Just to make sure: it's OK if I use 1 tube from the original broth w/ bacteria, to get 1 mL into 9 mL for a 10^-1 solution, same pipette for 0.1 mL into a 9.9 mL (which based on what I've discovered with my 9 mL tubes today, I know that it's much less than 9.9 mL ), same pipette for 10^-3, 10^-4 dilutions, same pipette for 10^-5, 10^-6 dilutions, and then use an entire pipette to pipette 1 mL of each of the sample, stating from 10^-6 all the way to 10^-1?

Just to double check - When I take 0.1 mL of the original broth, dilute it into "supposedly" 9.9 mL of distilled water, shake it (would this be sufficient for making sure that each mL has same number of bac?), then take 1 mL from the new 10 mL solution ... that is a 1:100 dilution right?

w00t . I'm a bit worried about the possible error with the water lost during autoclaving though . And I do hope that this time I can properly autoclave the nutrient broth .

As for the extra drop of water, I am almost sure that there was no water that touched the petri plates after dropping the 1 mL (which was immediately soaked up) so I'm not sure what happened there either.

I think for the nutrient broth that I now think it's contaminated, I remember that I started to autoclave it, and then there was an issue with the power supply that made me have to wait another day. While the broth is in there, whatever bacteria that is in there would have grown. I sterilized it again, but I'm thinking that whatever bacteria was in there is what is murky? Though, then when I dilute the plates nothing should appear, and yet there are colonies... I autoclaved it for 20 minutes at 121 C... I think that's the standard autoclaving time and temperature?

One quick question with ethanol -- I'm a bit anxious about letting the plates out in the open air for too long either. Is it OK if I briefly shake any ethanol off, and then put the cover on the petri plate so that whatever remains inside will dry through the night? It seems like this technique of letting the ethanol "air dry" inside (even when the cover is closed) works.

THANK YOU very much for bearing with me!

Hi Trader,

If your tubes are about half full with a 9 ml volume, you should not lose any volume during autoclaving. Autoclaving is done under pressure, and if the pressure is reduced too quickly, the tubes will boil over and get the cotton wet. Next time you autoclave, be sure and set the autoclave for slow exhaust and when the cycle is finished, don't open the door right away. Then, open the door very slowly to prevent boil over. It usually takes about 45 minutes for slow exhaust so check and make sure the slow exhaust cycle is working properly. For dry items, the autoclave can be run with fast exhaust, but slow exhaust must be used for any liquids.

If you wanted to have identical growth curves with 200 and 500 ml flasks with nutrient broth, you would inoculate the 200 ml flask with X organisms and the 500 ml flask with 2.5 times X bacteria. Both flasks would then start with the same number of bacteria per ml, and if incubated at the same temperature and aeration, will have the same growth curve. If you put X bacteria into both flasks, then the 200 ml flask would start at X divided by 200 bacteria per ml, and the 500 ml flask would start at x divided by 500 bacteria per ml. If incubated at the same temperature and aeration, the bacteria in the two flasks would double at the same rate, but the 500 ml flask would reach the maximum number a little more than one generation time (20 minutes) later than the 200 ml flask. All flasks would reach the same maximum bacteria per ml at some point.

Try doing at least 3 dilutions per sample; you might miss, but at least you'll have a < or > number.

Everything about lab work gets easier with practice. Congratulations on perfecting your pipetting technique. However, I think you mean that you can pipette 100 uL, or 0.1 ml, not 1 uL.

If you can pipette 800 uL or 0.8 ml, you could use the 7.9 ml dilution tubes and call that a 1:10 dilution. If you could pipette 80 uL or 0.08 ml, you could make a 1:100 dilution. The arithmetic will be too complicated if you do an odd dilution.

If you autoclaved the murky nutrient broth after something grew in it, then it will be sterile, but it will contain waste products for the dead contaminants and won't support growth as well as non-murky medium. Next time you have a power supply problem, put the medium in the refrigerator to retard growth. 121 degrees Centigrade is a standard autoclaving time, so that should have worked.

Your idea with the Petri dishes is probably a good one. You could also shake off the ethanol, and then set the lid of the Petri dish on top of the bottom, just slightly offset so a little air could get in and dry the residual ethanol. This would probably take just 20-30 minutes, and the plates would have minimum exposure to the air.

Good luck tomorrow!


Donna Hardy

I'll have to wait until tomorrow. The other science fair member(s) and I have been working after school late in the lab lately, and we've just been told by a science teacher that we are not allowed to do that...

That means that I only really have 2 hours to work afterschool! But I know that I should be doing these experiments under supervision...

I'll have to wait until tomorrow. What's worse is that the teacher who normally supervises the science fair is away for the weekend! It may come entirely down to what happens next week...

I do have a few questions:

To avoid losing water while sterilizing the tubes, what is meant by "depressurizing too fast"? I've checked out images of autoclaves and what my teacher calls a pressure cooker looks nothing like it . Mines is a big steel (I think it's steel?) cylinder that has a metal cover at the top. There are bolts to fasten the cover to the rest of the pressure cooker, and knobs to make sure that the cover stays on tight. Then there is a temperature control knob at the bottom, a pressure guage (which my teacher tells me to look carefully to make sure it doesn't go over "20"... i think it's pounds per square inch?), and a metal thing that I tip over when steam comes out,... Is there something above that is called a slow cycle ... controller that I've missed? Or else when I use the autoclave (I'm now thinking that it's actually a pressure cooker?), I make sure the cover is tight, turn the temperature to the highest, wait until steam comes out of the metal thing that sticks out, and tip it over so that pressure starts building. When the pressure guage points to 20, I control the temperature to make sure it stays at 20 for 20 minutes, and then turn the temperature down.

When I open the autoclave (after 24 h wait), I tip the thing back up and a long rush of steam (cold) comes out... would this be depressurizing too quickly? In that case, how might I be able to open it without depressurizing too quickly?

Hopefully I'll be able to convince a teacher to stay with me from 3 - 9. Maybe get 6 hours in and hope that the stationary phase is entered in the right time. 37 C .

I have many sources that say that the optimum temperature for b. subtilis is 30 - 37 C, and I think this probably varies because of the strain or the environment which it is in (such as a different type of agar or pH?). In that case, if I happen to find that the optimum temperature for lowest doubling time (which, please correct me if I'm wrong, is meant by "optimum temperature) and find that it is different from the optimum temperature that yields the highest amount of viable organisms, wouldn't that be inconclusive anyways because the b. subtilis optimum temperature is varied by so much?

Would that mean that all I can really conclude, if I find the above, is that given this strain, and nutrient agar, the temperatures for this and that are different?

-- OK it looks like I'll probably only have 3-6 available. When doing growth curves, is it "workable" to have "inconstant" time intervals? I'll try to run back and forth from the science halls between classes so I can do a test every 80 minutes or something...But the timing between each will be awkward.

Also, as there are sometimes only 5 minutes between each class -- will it alter results significantly if I dilute 1 mL into supposedly 9 mL of distilled water, and leave it in normal room temperature for up to 5 hrs before I actually have the time to dilute it more and make them into plates? -- will the bacteria in there still represent the bacteria in 1 mL, diluted 1:10, at that time period?

Thanks