I am looking for a fluid solution to close the lateral borders of a coverslip against the diffusion of air under the slip. I used nail polish before. This works fine, but I would like to use something less toxic. It is important, that I must not apply even minimal pressure to the coverslip. Best would be some chemically clearly defined fluid that will not interact with my specimen (living platelets, very sensitive).
I know some people use VALAP, a mixture of vaseline, lanolin and paraffin wax, to seal coverslips. It's non-toxic, but it's not fluid though. It's basically wax so you have to heat it up and slowly dab it on around the coverslip. It's definitely inert and won't interact with your specimen.
Here's the recipe that our Imaging Center recommends for use.
https://nic.med.harvard.edu/VALAP
Hello, I had this kind of problem before as well when I was working on protoplasts which are also very delicate in handling. What I first did was putting tape on the glass slide and cutting a frame out of it. This creates some kind of "pool" for your specimen. In addition, since you only have your specimen + medium inside of it, it prevents the liquid from evaporating. (The cover slip is laying on the tape-frame)
What you can do in addition is to seal the edges around the coverslip with spray-adhesive! It is defenitely less toxic than nail polish. Spray-adhesive turned out to be very good for our setup and maybe this is also non-invasive enough for your samples (and the ingredients are given on the bottle). You may want to buy the medical grade spray adhesive.
Best,
Falco
We use Permount, Crystal mount from Sigma also might work for your samples.
In relation to Falko Krügers response I can highly recommend "Secure Seal Imaging Spacers" (http://www.gracebio.com/life-science-products/imaging-microscopy/secureseal-imaging-spacers.html). These pre-cut double-sided tapes are, at least in my hands, inert in live cell experiments with mammalian cells, as well as plant cell and yeast protoplasts. They form a sealed cavity between the slide and coverslip with a depth of 120 µm. To cut costs, standard double-sided tape, cut with a razor blade, may work for your application as well. However, I have observed toxic effects on cells in live cell experiments, which I attribute to leakage of some organic solvents from those "standard" tapes. With "Secure Seal Imaging Spacers" I do not observe any toxic effects.
What I have done in the past is to thermal plasma treat glass slides, cut a small doughnut ring of silicone (
Nail polish contains butyl or ethyl acetate and will kill live cells in seconds. I just mention this, as Max-Joseph Kraus asked for a non-toxic way to seal coverslips. The solvents in nail polish sometimes even interfere with membranes of fixed cells. As always, however, all this depends on the specific application.
You may try Bee wax which is pure ,non toxic ,natural substance without any aroma and having a low melting point of 40-50 degree Ceslsius.It can be applied in place of a nail polish. Dr Ramesh Bhonde
For this purpose we use dental cement (http://www.picodent.de/artikel/831-picodent_twinsilsupsup_22.html) - it is (expectedly) inert and you can easily remove your seal to recover valuable living samples after imaging.
Josef
Josef Gotzmann - that looks like an interesting solution. Is it a two part product that you mix before application?
@ Matthew: yes, you mix a blue and yellow "solution" (quite viscous) and using a toothpick or a cotton-swab you apply it. you can buy solution with different hardening times - 1min, 5min, 20min - whatever you need. And you can easily remove it without any remnants.
Josef
@all: Thanks a lot for your great answers. That really helps me a lot!
I will try first VALAP and bee wax. Dental cement sounds also very interesting to me. The double-side-tape solution is not suitable for our application, as we need the smallest possible film of fluid (Platelet rich plasma) between the slide and the coverglass.
By the way: Nail polish worked really excellent. It is very easy and fast to apply. One could see the cytotoxic effects at the outermost part of the specimen, but I could not see any cytotoxic effects towards the center of the slide. Anyway, I don't want to use it anymore, as nobody can know, what effects it really does also at more distant parts.
I had some problems to get the right ingredients for VALAP. At my local pharmacist there was only fluid paraffin available. With this, the Valap gets to runny. But I could persuade the guys, that there must be something like paraffine flakes (as stated on the linked site), and they finally found it. I will recieve the first sample tomorrow.
I also think, Bee wax could be a very usable solution for us.
Again, thank you, for all of you're results. Best regards from Munich. Max
2-3% low melting agarose. Solidifies in seconds on the cold slide. Inert. Transparent
DPX (Dibutyl Pthalate Xylene) can also be applied with little care! The margins of cover slip should be dry if DPX is used for sealing.
If you cannot apply pressure, I'm wondering if you've used spacers to protect your sample? I've also used low melting agarose gel, double sided tape and also Parafilm (it melts at 60C and it seals so well that we use it for flow cells).
Hi,
Glycerol jelly may be a good solution. You can add thymol or equivalent to inhibit microbial growth. You can easily find formulations, or I can send you some. It only needs to be warmed to about 60oC. Once cooled, it sets solid and can be ringed with a permanent sealant. No pressure is required - it was used by Victorian mounters for pressure-free mounts of insects! If your require minimal toxicity, you can omit the thymol. It is used as a fluid mountant and will seal the whole slide. Ringing is optional for longer term storage.
You could also use glycerol jelly as a non-toxic ringing sealant for an aqueous mount.
tony ([email protected])
Hi,
How about using the Gene Frame from ThermoScientific.(http://www.thermoscientificbio.com/plastic-consumables/gene-frame/)
It is like a spacer but it is adhesive on both sides, so you can adhere the coverslip and the slide, and then it is sealed. You can fill whatever medium inside with the precise amount corresponding to the size of it. And, you do not need to press on top of it. They provide different sizes so you can have several selections.
How about some super glue? They use it for some types of surgery and so it is likely not very toxic? You would need to let it cure a bit so that it would'n t flow across the cover slip due to capillary action.
The agarose and glycerol gelly sound like good choices, but I don't have any experience with these. Good luck
We tried a silicon vacuum grease quite satisfactorily and ever at hand in the lab
we use vacuum grease too- High vacuum grease (Dow Corning Cat. No. 1597418)
Hi
In the past I've used the putty to colophony (of Noyez):
20% lanolin heats in porcelain dish (on the sand bath), for about 30 minutes.
Add 80% colophony and them 10% of paraffin or beeswax, stir with a glass rod to obtain a homogeneous fluid mass.
Pour the hot fluid mixture intopaper or metal molds.
The blocks thus obtained are preserved indefinitely.
To close the cover slip, heat the pad and wash the resin along the perimeter of the coverslip.
Best Regards
Fabio Nobili
There is a bunch of them on market; you should figure out which fit better:
http://www.dako.com/us/download.pdf?objectid=105416003
http://www.microbehunter.com/2010/01/23/an-overview-of-mounting-media-for-microscopy/
http://ibidi.com/xtproducts/en/Cells-Reagents/ibidi-Mounting-Medium
http://www.lmscope.com/produkt22/Einschlussmittel_en.shtml
http://www.thermoscientific.com/ecomm/servlet/productscatalog_11152_L11107_82082_-1_4
Please see overview at:
http://nic.ucsf.edu/dokuwiki/lib/exe/fetch.php?media=mounting_media_tests.pdf
Regards
I have used entellan. It helps with the lifting of the cover glass. I dont know if it's compatible with your samples because it's mainly for long term preventing of dryness to mounts especially in histology
@all: Thank you again for the numerous answers. I'am quite impressed by the diversity of possibilities and the creativeness of the different researchers. For me the question results more and more in the question of ease of handling and low cost. I made a small table to summarize the different answers. Probably this can help others with their decisions. To date, there a 21 different solutions! As my question for optimal sealing of coverslips is part of a larger research project I will spend some more time to test the different possibilities for our application. Thank you again. All answers were very helpful for me. Regards. Max
Dear all,
not knowing if the Microscopy-UK website could be of help anyway, for your convenience (microscopical amateur as well as professionist) I would like to add the following:
http://www.microscopy-uk.net/search/search.php?zoom_sort=0&zoom_query=mounting+media&zoom_per_page=10&zoom_and=1, where the search phrase "mounting media" resulted in:
74 results found containing all search terms.// 8 pages of results.
@ Max-Joseph: thanks for providing the summary-pdf (hoping you will share another one with the result of your trials...) Best regards and wishes, Wolfgang
Hi
I read the all recomended methodology for this purpose.However, it seems that you have an altenative way for elimination of needing for coverslip! Ok, you can use the catadioptric and long distance objective for direct observation without mounting the coverslip. All the best, Reza
Dear Mohammad Dadpour,
thank your for your comment. I've never used a catadioptric objective before. Do you know the maximal magnification of these objectives? I usual work with a 100-fold oil-immersion objective. Regards. mjk
Re: cadadioptric systems
Dear Mohammad, dear Max,
Interesting approach, but: I doubt that catadioptric lens system(s) (as you proposed) [cf. e.g. http://en.wikipedia.org/wiki/Catadioptric_system or
http://www.dpreview.com/forums/post/32050684] can be used easily for the task (using ) Max really needs.
@Mohammad: could you specify more in detail (or give a reference) how the / your proposed system works (in terms of “microscopical routine work”)?
I could not find (with a short google-search) any valid data on < true resolution > (which should be a better term of what Max needs contrary to “magnification”!) of catadioptric (lens) systems (cdls) and not knowing better from two sources (see below), it seems that “resolution” of such cdls is not simply comparable to “resolution” as defined for more than a centennium (~1873/1876 Abbe and Helmholtz).
The only source I found for “catadioptric systems AND microscopy” was @ http://encyclopedia2.thefreedictionary.com/Microscope :
. @ http://www.answers.com/topic/optical-microscope one can find the information:
A perhaps practical example for using catadioptric system (catadioptric zoom lens ) is included with/in:
< Dual modality endomicroscope with optical zoom capability> by DG Ouzounov - 2013 @ http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3771821/ .
It could be well that I misunderstand the whole matter [despite the roots of the technique were founded in the 1820 ies by Fresnel, a was established in 1859 by Foucault, confessing that even as a long practizing (as well as teaching) microcopist I never was tought or had heared about before, perhaps also because the technique was / is used mostly for (astronomic) telescopes too].
@Max: you need for sure not only but specially “resolution” (I guess your x100 Oil immersion objective has a Numerical aperture NA equal to or even greater than 1.3 (right?) for which (knowing the “secondary magnification of your ocular, wavelength of used light, for imaging eventually the “camera factor” too) you can calculate your actual “point-to-point resolution” with the formula of Abbe: or considering other methods like far-field light microscopy / fluorescence techniques / STED microscopy ( for principle cf. e.g. http://www3.mpibpc.mpg.de/groups/hell/publications/pdf/Topics_5_.pdf ) which theoretically achieves 20-50 nm lateral resolution, or other super-resolution techniuqes (cf.e. g. http://en.wikipedia.org/wiki/Super_resolution_microscopy ).
Last comment:
CAVE CONFUSION of
Petroleum jelly, also known as Vaseline. Vey low cost, non toxic, easy to remove the slip. The only hassle is filling a syringe without trapping bubbles, in order to apply a thin line around the sample.
A vaseline work up that works well for me (a time ago):
- make a thin spread of vaseline on parafilm
- pick up some vaseline on three edges at the underside of the coverslip
(hold with a tweezer on the fourth clean edge)
- make sure to close off at the corners too
(at this stage you can pick up parafilm spacers at the corners if needed, they will stick)
- put on the microslide, sealing off the 3/4 sides of the cell culture, push
- clean off liquid leaving the slide-coverslip area with ckleenex or filterpaper
- close the fourth edge (which always is a bit messy, use syringe like Sharona says or spatula)
- keep fourth side far away from objectives if you use an upright scope
hope this helps ;)
Hi Max,
I was wondering if you could find a good solution for sealing your slides. We use the same method (nail polish), but I'm also looking for a better way. We found that letting the mounted slide air dry (with a fan) for an hour or two have better results.
I also found this: https://www.youtube.com/watch?v=4Nurc1hSN4o
The problem with the Vaseline base sealing solution is that we can't store the slides in the freezer.
I'd appreciate if you would share your findings.
Thanks,
Nasim
https://www.youtube.com/watch?v=4Nurc1hSN4o
Dear Nasim. Thank your for adding your answer to my question and for sharing the link. I could not find any better solution than nail polish for the sealing of my slides. I tried all of the proposed methods (thank you everybody!), but still nail polish seems to be the best solution for my needs.
As to my experience the quality of the slides mainly depends on the amount of sample solution. For example with 22x22 mm cover slips I use exactly 10 µl of aqueous sample. This results in a complete dispersion of the solution beyond the cover slip, while there is no excess fluid on the borders of the cover slip. As soon as there is to much sample fluid (as for example 15 µl), the cover slip starts to "swim" on the fluid and it therefore moves away when I try to seal it with the nail polish brush.
As to reduce the influence of the toxic nail polish onto the specimen, I started to only examine the center part of the slide that is sufficiently far away from the sealed borders. Furthermore I use bright red nail polish and therefore I can always exactly see the parts of the specimen that came into contact with the nail polish. Especially for larger (to large) amounts of aqueous specimen, I have seen the red nail polish diffusing between cover slip and slide and therefore contaminating the specimen.
Best regards. Max
How about biological grade silicone, which is used in aquariums and solvent free, thus non-toxic? Gold label is a good one.
You can try sealing the coverslip with DPX solution using a fine brush. The slide as well as coverslip margins should be free from water before applying DPX.
@Ana De Moura Queiros: thank you for your interesting note on biological grade silicone. I tried to order "Gold label" Silicone, but they do not ship to Germany (a company from scotland ships it's "Gold Label" products to Germany, but this is a different story and I prefer single malt anyway ... -:).
Therefore I ordered Juwel Aquarium Silexo, a black coloured biological grade silicone, that is made "for sealing and bonding glass". It comes in a tube and has a intense smell somehow similar to vinegar. It has a consistency similar to toothpaste. I tried to seal my coverslips by applying the silicone using a small brush - that did not work, due to the stiff consistency of this silicone. I tried to fill the silicone in a syringe to get a smaller tip. But this way, the silicone was very difficult to apply to the right spots of the coverslip.
On the other hand, after hardening, the silicone would do exactly what I need. On the package insert it says: "Silexo is a high-performance silicone for use in aquariums. After hardening, it is completely odour-less and does not release any toxic substances into the water" Are you probably aware of a biological grade silicone, that has a more fluent consistency and is therefore easier to apply? Thank you for your answer. Best regards from Munich. Max
@Karumanchi Sambasiva Rao: Thank you for adding your answer to my question. Indeed, you have had recommend Dibutylphthalate Polystyrene Xylene back in 2013! I have had no possibility to really test this. But according to the safety data sheet available from Sigma-Aldrich I would expect DPX to have somehow similar characteristics as nail-polish. Is this correct? Are there data on the toxicity of DPX on living cells? I just orderd a small bottle of DPX to test.
@ Pablo Bolaños-Villegas: thank you for your answer. The problem for me here is that I need a very small amount of specimen (10 µl) dispersed completely between the slide and the cover slip. If I understand right, you put the ring binders (what is this exactly?) between the slide and the coverslip? This will increase the distance between slide and cover slip to much for my application. Or do I get this wrong?
@ Ramesh Bhonde: I also thank you for your answer. I found molten bee wax a very attractive solution. The only point with this is, that the bee wax has to be heated to be properly applied to the coverslip. And with this very short distances on the microscope slide I am not really sure on the influence, that heat conduction has on my specimen.
I don't know if you are aware of this product
https://biotium.com/product/covergrip-coverslip-sealant/
Dear Konstantions,
fantastic! No, I was not aware of this product. This seems to be exactly what I was looking for. I just ordered a small bottle to test. THANK YOU!
You're very welcome. I'm facing the same problem as I'm currently starting a new research project in probiotics and I need to run long-term time-lapse microscopy...
Here another product that I found also useful, although, I have not tested yet
http://www.strettonscientific.co.uk/cytobond-removable-coverslip-sealant.html
I wonder are you aware of the product iSpacer from SunjinLab (https://www.sunjinlab.com/product-category/ispacer/)? I love it and daily use it in our lab. It's cheap, ready-to-use, and very easy to handle. It can form a secure chamber to hold the aqueous samples and even can be stored in 4 degree without leakage.
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