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Acid Wash, Autoclave, Flame or Coat? Slide Basics Explained

Posted in: Microscopy and Imaging
Acid Wash, Autoclave, Flame or Coat? Slide Basics Explained

There are about as many protocols to prepare coverslips as there are ways to make tuna casserole. You can spend from 5 seconds to 2 days, depending on what your lab prefers. But in the end, what’s really needed? I’ve tried many protocols over the years and I’ve questioned some steps. Here I share my opinions on some coverslip preparation techniques – from the seemingly absurd to the down-right reasonable to help you to separate the helpful from unjustified traditions in your experiments.


Acid Washing

Acid washing is often recommended by the manufacturers of coverslips, such as Corning, because it is not only reported to enhance the cleanliness of the coverslip but the acid also etches the glass making it a better substrate for cellular attachment. And while I have done it in the past, many people skip acid washing their coverslips. For the most part acid washing is a step that can be reserved if quicker methods of cleaning and sterilizing aren’t working for you, or if you are having attachment problems.


I was surprised to find this step in protocols, but with the advent of sonicating toothbrushes I see how people use the logic that sonication scrubs. But I find it hard to believe that sonicating is going to remove anything significant from an otherwise clean and polished coverslip (comment below if you have evidence otherwise). So I’d skip sonicating unless you are desperate to get rid of some sort of observed particulates not removable by other means.

Phosphate Buffered Saline versus water.

Some people recommend rinsing your coverslips in PBS instead of water. And while the coverslip does appear to coat better when there is salt in the water, I do not use PBS. Instead I use pure water because I do not want the possibility of salts drying onto the coverslips, since salt could carry over and cause problems in future steps.



Many labs consider autoclaving a necessary step for complete sterilization. In this method, typically after various washes, coverslips are placed in a glass petri dish and sent through the dry cycle of the autoclave. The upside of autoclaving is that you are sure your coverslips are tissue culture sterile, which is necessary if you plan on growing cells on these. The downside is that autoclaving takes time. Also, don’t autoclave fancy commercially-purchased precoated coverslips.


70% ethanol is frequently used for sterilizing coverslips. However, for sterilization ethanol is inferior to autoclaving, therefore it is often followed up by exposure to UV light or flaming. Doing both may still be faster than autoclaving and the subsequent cooling. Remember though, if you choose to use this method, some microorganisms can persist in 70% ethanol, so do not reuse your alcohol.


While ethanol sterilization is typically performed at a concentration of 70%, flaming is often done with a higher concentration of ethanol, such as 90% (although 70% will still catch on fire). Flaming is nice because it is super-fast and better at eliminating spores and other hard-to-disinfect microorganisms than ethanol alone. However, because one of the most dangerous hazards in the laboratory is the Bunsen burner in the tissue culture hood, some universities have outright banned the practice. Not only is it dangerous due to the open flam aspect but also because some people (*cough cough* the summer undergrad) accidentally turn the gas UP till the flame goes out, thinking they have turned it off when in reality they are creating A BIG BOMB. Don’t let this happen to you, and as fun as it may be, I recommend that you just skip the flame when in the tissue culture hood.

UV Light

This is my favorite, it’s quick, it’s easy and can kill just about any microorganism in its path: Yes, UV light is the way to go for fast and efficient sterilization. After cleaning your coverslips and giving them a preliminary sterilization in 70% ethanol, pop them under the germicidal UV light of your tissue culture hood for 15 minutes and voilà! you have very sterile coverslips. Incubations under the UV light vary from 15 minutes, up to 1 or 4 hours, or even overnight. The only negative to this method is that your lab mates are going flock to that blue light like a moth and will be annoyed that the hood isn’t being used for tissue culture work. They may even move your coverslips to the side and begin working, compromising all your sterility and cleanliness. So take care to coordinate and explain what you are doing to your lab mates ahead of time.



This is a tried and true method. Coating with poly-L-lysine provides a positive charge for cell attachment and is used for a wide variety of cell types. If you are working with a cell type that does not attach to glass easily, like HEK293 cells, coating your coverslips with poly-L-lysine can make the difference between having one cell left after treatment, or a million. Some prefer to use poly-D-lysine since it is resistant to breakdown by cell-released proteases. Regardless, it IS important to spend the time rinsing off excess poly-lysine to prevent its accumulation into the media which induces cellular toxicity.


As a major constituent of extracellular matrixes, collagen I, II and IV can be used to coat coverslips for a wide variety of cell types including primary cells, myocytes, chondrocytes and endothelial cells. Do be sure that you are using the recommended collagen as there are many different types, and take care to sterilize your coverslips properly.


Fibronectin is another major component of the extracellular matrix. Fibronectin can serve as a substrate for a wide variety of cell types including neurons.


As an important component of basement membranes, laminins are also a good substrate for many different cell types. They are especially well known as a substrate for neurons and in assays for chemotaxis.


Nail Polish

Why do people bust out the nail polish and paint it around the edges of their coverslips? Some say 1) to prevent the sample from drying out 2) to prevent oxidation 3) to prevent the mounting medium from leeching onto the objective or 4) to keep the coverslip in place. It’s a mixture of all those reasons, but the reason varies depending on the type of mounting medium and the time after mounting. If you are using a mounting medium that does not harden, it might not make any difference when you seal it. But if you are using one that hardens, such as Prolong gold, and you seal it before it hardens, it may never cure properly. So make yourself familiar with the recommendations for your mounting medium, and also to check if there are any reports of the nail polish quenching your fluorophore of choice


This homemade solution isn’t toxic like nail polish and is recommended in many protocols especially those that involve live imaging. The anacronym is derived from its ingredients which are Vaseline, Lanolin and Paraffin wax. I’m not sure why anyone wouldn’t use VALAP over nail polish, unless it’s a time issue (or you just secretly like to get high off the smell of nail polish).

Other Epoxy

In theory you could use all sorts of glue to hold down your coverslip. But when using an unknown/untested epoxy you run the risk of it quenching your fluorophores or interfering with the anti-quench agents in your mounting medium. Not a risk worth taking.


That’s right, good old paraffin wax. Why not? It may harden quickly but at least it’s non-toxic and a simpler than VALAP (just melt). Now if only you had some melted on-hand, like all the time.

Do you have your own insights into coverslip preparation? If so, please share below. But let’s focus on facts and experience – not unjustified traditions.

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  1. Zach Hough on December 7, 2021 at 11:56 pm

    While I’m open to new suggestions (hence reading this article) my lab’s approach is to autoclave a large number of 1.5mm thick coverslips in a glass beaker covered in aluminum foil and then store them under our culture hood for later use. They seem to stay sterile so long as they are not opened to air outside of the hood (probably also helped by repeated UV exposure). This way they are ready to go whenever we need them. Next we add 0.5mL of 0.01% Poly-L-Lysine to each well of a 12-well plate, then transfer the coverslips to the wells and float them on top of the Poly-L-Lysine. If coating both sides is desired, they can be flipped over or submerged (though you may need >0.5ml to fully submerge). They incubate in this solution for 15min per side, after which the poly-l-lysine is removed with a pipette and saved for reuse later on (we typically get 2-3 uses out of it before discarding). Any excess is aspirated and then they are allowed to air dry until there is no liquid left (anywhere from 15min to 1hr). I will often pre-coat coverslips and store the covered plate (once dry) in the fridge for use at a later date (usually within the week). Then I simply plate my cells into the wells as I normally would, making sure the coverslips remain fully submerged (no bubbles) before placing in the incubator. It often takes them an extra day to fully adhere and they divide more slowly during this time. If treatments are applied or media changed before they are fully adhered, many will likely wash off. Once fully adhered, I apply my treatments and allow them to incubate. Sometimes bubbles form under the coverslips, so I will push them down gently with a pipette tip to fully submerge. Next, I will either fix the cells and follow staining protocols or do the following for live cell imaging. I will first apply melted valap with a brush to form a small spacer/reservoir on a glass slide. Then I will fill the reservoir with PBS +Calcium and magnesium containing 10% glycerol. I place the coverslip, cell side down onto the valap reservoir, leaving a small gap on one side. Add more PBS/glycerol with a pipette to push out bubbles and fill completely, then slide the coverslip over gently. Finally, I seal the coverslip onto the slide with more valap around the edges and begin imaging. This should allow the cells to behave normally and remain attached for at least an hour or two, perhaps longer if they are able to be maintained at 37°C while imaging. Note that media without phenol red can be used in plate of complete PBS provided it is compatible with your live-cell stains and may increase the cells’ longevity. Phenol red will autofluoresce and result in high background, especially at 488nm. I hope this helps anyone looking for a more complete protocol. The general advice provided in this article is very helpful and pretty consistent with my labs practices and experiences.

    • Thomas Warwick on October 20, 2023 at 3:37 pm

      Very comprehensive, thank you, Zach.

  2. zac on October 4, 2018 at 4:55 am

    Exposure to UV light does not *reliably* kill ALL bacterial or fungal spores. 70% ethanol also does not reliably kill ALL spores. Both do kill vegetative forms of both bacteria and fungal cells (IF the concentration of the ethanol is above ~70%). But neither UV light nor ethanol solutions sterilize since sterilization is the RELIABLE and COMPLETE removal of all living cells. It is a common misconception in the tissue culture field that 70% ethanol and UV light sterilize. They do however vastly decrease the microbial bioload in tissue culture environments.

  3. Tariq on September 7, 2018 at 12:48 am

    Thanks for the tips. I like the UV sterilization. Because it’s fast 🙂
    I wonder if I got it right.
    Clean (not sure about this; clean with water ? Acid?), then treat with 70% ethanol and then expose to UV?
    I would appreciate being corrected if I misunderstood something.
    My way is to soak the coverslips in 20-30% HCL 30min to corrode the surface and enhance attachment, wash with MQ, transfer to wells if to be used immediately (or store in 70% ethanol for later use), sterilize with 70 ethanol in the wells, remove ethanol and UV sterilize about 15min, coat and culture cells.
    What do you think.

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