MicrobeHunter.com Microscopy Forum

Tried glycerin to slow down protozoa. Environmental disaster.
What wasn't killed outright, exploding or horribly deformed, looked like this one. (A few were slightly more mobile)
The survivors were flattened to the point that all internal functions are shut down. All trichocysts seem to have fired, or the alveoli are swollen.



Radazz

When adding any of the viscous liquids to slow protozoa motion, better add very small quantities. Even a few percent are, in my experience, too much. Also, I think that methyl cellulose and similar polymers are better than glycerol, since their osmotic effect (for the same volume or weight) is much lower. There is a recipe by charlie g, cited by 75RR in a previous post from December 23, 2018. Search "cellulose" in the forum.

I just had a chance to try out copper sulfate and copper acetate for, as the instructions say, "immobilizing fast-moving ciliates."

Results:
Copper sulfate (one drop per drop of pond water) - indeed all ciliates on the slide were completely immobilized.

Copper acetate (apx 1/4 drop per drop of pond water) - same result. Ciliates immobilized.

The water drop feels pretty dead with all the pond favorites immobilized, but I suppose if there is a need to have ciliates completely immobilized to study an internal structure or take a photo, this is a viable method.

On an unrelated note, I may have gotten a photo radar ticket today. Darn it. if so, there goes microscope/hobby money down the drain for penance of my crime. But there's hope 'til it arrives in the mail.

Sauerkraut wrote:I just had a chance to try out copper sulfate and copper acetate for, as the instructions say, "immobilizing fast-moving ciliates."

Results:
Copper sulfate (one drop per drop of pond water) - indeed all ciliates on the slide were completely immobilized.

Copper acetate (apx 1/4 drop per drop of pond water) - same result. Ciliates immobilized.

I should have mentioned that the solutions are from the Innovating Science stain kit purchased from AmScope. Copper(II) Sulfate 1% aqueous solution. Copper (II) Acetate 3% aqueous solution.

Please try 1.5% methyl cellulose..50/50 mix of your specimen fluid and this viscosity tool. Please purchase the larger size rectangular cover slips so you never approach the wetmount cover slip borders with 'tight working distance' high mag objectives.

You can enjoy ciliates, large bacteria slowed down yet still patterning their usual body motions and ciliature wave propagations...a Parameciums 'fields of swaying ciliature' are beguiling..especially when the protist elects to change direction of body motion.

Enjoy live protists and use your oil-immersion objectives, with this 1.5% methyl cellulose viscosity tool. Charlie guevara

charlie g wrote:Please try 1.5% methyl cellulose..50/50 mix of your specimen fluid and this viscosity tool. Please purchase the larger size rectangular cover slips so you never approach the wetmount cover slip borders with 'tight working distance' high mag objectives. --Charlie guevara

Thank you. Good to know. I only tried the other reagents because they unexpectedly came with a staining kit. Trial and error. Mostly error.

Hi together,
we had our group meeting today and met in a restaurant afterwards with a couple of members. One topic was slowing down fas placton animals. One member reported a couple of methods he has applied with success:

-Osminumtetroxyd-fumes (Absolutely not recommended, highly toxic, he found a bottle in a hospital cupboard in Thailand and made a few tests)
-Cocaine-solution (availability....)
-Nicotine-solution for e-cigarrettes
-There was one more, I think it was PVAC = white wood glue, not sure, have to ask

Bob

Hi Charlie g, Sauerkraut and Bob,

Osmium Tetroxide is a lethal poison, and in addition, can cause blindness.
here is a rational for charllie g's protocol: the viscosity of a 2% solution of methyl cellulose in water is ~400 cp. That of water is just ~1 cp. Dilution of the 2% MC solution, first by 2 (to make 50%) then by 67 (to make 1.5%) yields a viscosity of ~3. So it will slow down the cilliate motion by a factor of ~3 - very reasonable in my opinion. 2% MC is being sold as food additive and for other commercial purposes. It is very safe to use.

Hobbyst46 wrote:Hi Charlie g, Sauerkraut and Bob,

Osmium Tetroxide is a lethal poison, and in addition, can cause blindness.
here is a rational for charllie g's protocol: the viscosity of a 2% solution of methyl cellulose in water is ~400 cp. That of water is just ~1 cp. Dilution of the 2% MC solution, first by 2 (to make 50%) then by 67 (to make 1.5%) yields a viscosity of ~3. So it will slow down the cilliate motion by a factor of ~3 - very reasonable in my opinion. 2% MC is being sold as food additive and for other commercial purposes. It is very safe to use.

Here is a link to show that it does exist and is not overly expensive.

https://www.homesciencetools.com/produc ... ose-30-ml/

As far as I know methyl cellulose is what is used to put wallpapers up the wall, you get it in DIY or decorating markets.

Bob

MicroBob wrote:As far as I know methyl cellulose is what is used to put wallpapers up the wall, you get it in DIY or decorating markets.

Bob

So it is. It is a matter of quantity and convenience.

Note: buying a bottle of methyl cellulose solution is a lot more convenient than buying the powder and dissolving in water. It is soluble in cold water (NOT hot water), but only after prolonged stirring.

Note that, as used in wallpaper paste, it typically has a fungicide added.
Tom

tgss wrote:Note that, as used in wallpaper paste, it typically has a fungicide added.
Tom

That will really slow them down!

Hobbyst46 wrote:Note: buying a bottle of methyl cellulose solution is a lot more convenient than buying the powder and dissolving in water. It is soluble in cold water (NOT hot water), but only after prolonged stirring.

I prepared 1.5% solution in water and and after a few days of stirring it seems to be all in solution but I see a lot of tiny dots/fibers floating around. Any idea how to get rid of them?

Wes wrote:

Hobbyst46 wrote:Note: buying a bottle of methyl cellulose solution is a lot more convenient than buying the powder and dissolving in water. It is soluble in cold water (NOT hot water), but only after prolonged stirring.

I prepared 1.5% solution in water and and after a few days of stirring it seems to be all in solution but I see a lot of tiny dots/fibers floating around. Any idea how to get rid of them?

I am afraid filtration is the only way. The ideal filter would be a disposable syringe filter, that looks like a top, and has Luer tips. It is a standard item in laboratories. One fills the syringe with the raw polymer solution, attaches the filter onto the syringe tip and pushes the syringe plunger slowly.
An the absence of such syringe filter, one may try gravity filtration through a funnel of coarse filter paper, or cheese cloth, coffee filter; a piece of ladies nylon stocking (high Denier) might help as well, perhaps.

Hobbyst46 wrote:

Wes wrote:

Hobbyst46 wrote:Note: buying a bottle of methyl cellulose solution is a lot more convenient than buying the powder and dissolving in water. It is soluble in cold water (NOT hot water), but only after prolonged stirring.

I prepared 1.5% solution in water and and after a few days of stirring it seems to be all in solution but I see a lot of tiny dots/fibers floating around. Any idea how to get rid of them?

I am afraid filtration is the only way. The ideal filter would be a disposable syringe filter, that looks like a top, and has Luer tips. It is a standard item in laboratories. One fills the syringe with the raw polymer solution, attaches the filter onto the syringe tip and pushes the syringe plunger slowly.
An the absence of such syringe filter, one may try gravity filtration through a funnel of coarse filter paper, or cheese cloth, coffee filter; a piece of ladies nylon stocking (high Denier) might help as well, perhaps.

I tried that. It was impossible, not even a single drop came out and I pushed to the limit of the syringe/filter. I ended up spinning it very fast which made a ton of difference, I could see all the crap pelleted at the bottom and a very nice clear solution on top which I collected and placed in a separate vial. It seems clean looking at it through the scope.

In terms of sample preparation is it a 50/50 mix of sample water with protozoans and methyl cellulose solution?

Wes wrote:

Hobbyst46 wrote:

Wes wrote: I prepared 1.5% solution in water and and after a few days of stirring it seems to be all in solution but I see a lot of tiny dots/fibers floating around. Any idea how to get rid of them?

...

I tried that. It was impossible, not even a single drop came out and I pushed to the limit of the syringe/filter. I ended up spinning it very fast which made a ton of difference, I could see all the crap pelleted at the bottom and a very nice clear solution on top which I collected and placed in a separate vial. It seems clean looking at it through the scope.

In terms of sample preparation is it a 50/50 mix of sample water with protozoans and methyl cellulose solution?

Congratulation on the successful centrifugation. Yes, this dilution is charlie g's protocol AFAIK.

Wes wrote: I ended up spinning it very fast which made a ton of difference, I could see all the crap pelleted at the bottom and a very nice clear solution on top which I collected and placed in a separate vial. It seems clean looking at it through the scope.

Hi Wes,
so you first centrifugated first and then filtered with the syringe filter Doron has suggested?

Hi Doron,
can you measure the mesh size of these filter tips for me and maybe post a link to them?
I have never had much contact to laboratory equipment and don't know what nice things are used there.


I always like recipes for microscopy material that is widely available. There remain so many special ingredients anyway and I have the impression that this difficult availability is often what keeps people from exploring new microscopy topics.


Bob

MicroBob wrote:

Wes wrote: I ended up spinning it very fast which made a ton of difference, I could see all the crap pelleted at the bottom and a very nice clear solution on top which I collected and placed in a separate vial. It seems clean looking at it through the scope.

Hi Wes,
so you first centrifugated first and then filtered with the syringe filter Doron has suggested?

I only centrifuged. I didn't see the need for filtering after the centrifugation step because there was nothing visible in the solution under high magnification i.e. clear solution was obtained. I doubt filtering would've worked after the centrifugation step but then again I used a 0,2 µm filter so it might work with a less stringent filter.

MicroBob wrote:

Wes wrote: I ended up spinning it very fast which made a ton of difference, I could see all the crap pelleted at the bottom and a very nice clear solution on top which I collected and placed in a separate vial. It seems clean looking at it through the scope.

Hi Wes,
so you first centrifugated first and then filtered with the syringe filter Doron has suggested?

Hi Doron,
can you measure the mesh size of these filter tips for me and maybe post a link to them?
I have never had much contact to laboratory equipment and don't know what nice things are used there.
I always like recipes for microscopy material that is widely available. There remain so many special ingredients anyway and I have the impression that this difficult availability is often what keeps people from exploring new microscopy topics.

Hi Bob,
Here is a photo of the filter. They are sold in quantities, I only have a few. It is a polymeric membrane - nylon, polycarbonate or other (possibly Teflon as well, although I have never seen one), diameter of about 25mm. within a plastic fixture, with Luer (syringe) connectors on both sides. There are other sizes and variants. The polymer is a very uniform mesh, and hole diameter is most often 0.2 or 0.45 micrometers (there are other hole sizes). The mesh size is hence printed on the filter and package, as hole diameter. One popular application is sterilization, since they can filter off microbes from a liquid. Sold by all chemical/biochemical/biological lab suppliers.
I had thought that the issue posted by Wes could be solved with such membrane filter, but apparently the viscosity of his cellulosic solution was too high. So, a very high pressure on the syringe plunger. According to my experience, very high pressure might break through the filter and destroy it, so the filtrate is the same as the unfiltered solution. But it depends on the liquid in question. With cellulosic materials, the viscosity rises steeply with concentration.
A centrifuge in such a case is a very good idea.

I too prefer home-made and easily available over lab-supply chemicals and accessories, but sometimes there is no choice...

I've got a hand crank centrifuge for two quite big glasses, ca. 15x100mm. This will probably do. Otherwise we have an electric one in our group lab available.
Centrifuging is a nice process step, not messy, no fumes and no waste.

Thank you for showing these filters, Doron! I have heard of membrane filters but have not seen on in person. For an outdoor holiday in wilderness I once looked for water purification methods and there a pumps which probably employ such a filter, only bigger.
My idea was whether my 25µ stainless mesh might do but 0,xµ is a completely different thing. There are probably quite a few experiments that can be done with such a filter, e.g. removing pigments from fluids. Such a membrane could also separate inhabitants in a plancton aquarium while allowing exchange of oxygen and minerals.
With a thin syringe one can produce quite a pressure, I can imagine that even the filter housing could be crackes apart if one would really try.
As primary school boys we did do target shooting with syringe needles, powered by the compressed air from thin syringes... No recommended safety toys!

Bob

Here’s a YouTube link that compares methylcellulose to physillium seed husk.

https://m.youtube.com/watch?v=vowfgDu8Fz4

Think I’ll give the seed husks a try and see how that goes.

Heather

Sauerkraut wrote: ↑

Thu Jun 06, 2019 11:04 pm

Here’s a YouTube link that compares methylcellulose to physillium seed husk.

https://m.youtube.com/watch?v=vowfgDu8Fz4

Think I’ll give the seed husks a try and see how that goes.

Heather

Heather-- I just came across this thread. Just in case anyone might still be reading it..... Did you ever try out the physillium seed husks ?
I like this idea much better than MC. Good old Metamucil
(I know Metamucil has additives )

https://www.youtube.com/watch?v=vowfgDu8Fz4

In the video the Psyllium jelly seemed to work well, with ciliates active yet slower than with Methylcellulose. There was however some distortion as they traversed what seemed to be thicker areas of jelly.

mintakax wrote: ↑

Fri Oct 04, 2019 5:08 am

Sauerkraut wrote: ↑

Thu Jun 06, 2019 11:04 pm

Here’s a YouTube link that compares methylcellulose to physillium seed husk.

https://m.youtube.com/watch?v=vowfgDu8Fz4

Think I’ll give the seed husks a try and see how that goes.

Heather

Heather-- I just came across this thread. Just in case anyone might still be reading it..... Did you ever try out the physillium seed husks ?
I like this idea much better than MC. Good old Metamucil
(I know Metamucil has additives )

Dan,

I tried the physillium seed husks and did not have good success. My assumption is that the person who posted the video has a specific preparation method. It's been a while but my recall is that the husk solution was too thick and and created too much image distortion (cloudy). But it's possible that with the correct experimentation, this is a viable method. If you try this method and have success, please post methodologies.

I may also try to revisit the husks and try to figure out how to make it work.

Hi together,
I asked our group member what to use in this case:

- High molecular weight
- Quince (the fruit) slime
- Polyehtyleneglcol
- perhaps PVAC glue, he hasn't tried

Not suitable:
- substances of low molecular weight like glycerol as they have an osmotic impact on the organism

I hope this helps.

Bob

Not suitable:
- substances of low molecular weight like glycerol as they have an osmotic impact on the organism

Good to know

Sauerkraut wrote: ↑

Fri Oct 04, 2019 8:13 pm

mintakax wrote: ↑

Fri Oct 04, 2019 5:08 am

Sauerkraut wrote: ↑

Thu Jun 06, 2019 11:04 pm

Here’s a YouTube link that compares methylcellulose to physillium seed husk.

https://m.youtube.com/watch?v=vowfgDu8Fz4

Think I’ll give the seed husks a try and see how that goes.

Heather

Heather-- I just came across this thread. Just in case anyone might still be reading it..... Did you ever try out the physillium seed husks ?
I like this idea much better than MC. Good old Metamucil
(I know Metamucil has additives )

Dan,

I tried the physillium seed husks and did not have good success. My assumption is that the person who posted the video has a specific preparation method. It's been a while but my recall is that the husk solution was too thick and and created too much image distortion (cloudy). But it's possible that with the correct experimentation, this is a viable method. If you try this method and have success, please post methodologies.

I may also try to revisit the husks and try to figure out how to make it work.

Heather,
I have been experimenting with PSH for the last several days and I am struggling to get consistent results. One time I think its slowing down the organisms, the next I'm not sure. I am using the "jelly" at 100% concentration, but the actual thickness of the jelly is hard to quantify. I am going to continue with the experiments and I'll eventually post some video of results.

Today I tried MC for the first time with disastrous results. My technique for both the PSH and MC is the same: I have several drops of a sample in a small petri dish and using a stereo microscope, I slurp up an organism or two into a tiny capillary pipette (0.3mm ID). I then put a drop of the whatever substance I'm using on a slide and then transfer the contents of the pipette (a very tiny drop) onto the drop on the slide. I check the slide under the stereo scope to make sure the organisms are there, then gently drop a cover slip on and recheck using the stereo scope.
For the MC I mixed 50/50, 1.5% MC and spring water. Under the stereo scope I could see the organisms swimming around in the tiny drop of sample water surrounded by the MC mixture. When I put a cover on and the suspensions mixed, there was no movement of the parameciums and high magnification confirmed they were dead. This happened twice.
Could it have been the spring water or the MC or my technique ?

mintakax wrote: ↑

Tue Oct 08, 2019 5:22 am

For the MC I mixed 50/50, 1.5% MC and spring water. Under the stereo scope I could see the organisms swimming around in the tiny drop of sample water surrounded by the MC mixture. When I put a cover on and the suspensions mixed, there was no movement of the parameciums and high magnification confirmed they were dead. This happened twice.
Could it have been the spring water or the MC or my technique ?

What is the source of your methyl cellulose? See quote below ...

tgss wrote: ↑

Sun May 26, 2019 12:03 pm

Note that, as used in wallpaper paste, it typically has a fungicide added.
Tom

See link: viewtopic.php?f=13&t=6752&p=60342&hilit#p60342