Many cell lines commonly used in research are adherent, meaning they attach and grow on a surface rather than just hanging out in suspension. If you wish to perform imaging of adherent cells, such as to undertake cancer cell microscopy, you need to fix them to your microscope slides. Luckily for you- this is fairly easy to do. Thanks to the fact that adherent cells are, well, adherent, and like to grow on plastic or glass. The same stuff your microscope slides and covers are made off!
Below is a step-by-step guide to growing your adherent cells on your microscope cover slips and ‘fixing’ them for excellent imaging results.
Things You’ll Need:
|Adherent cell line||Pipettes & tips|
|Methanol or paraformaldehyde||Cell incubator|
|Sodium hydroxide||Large beakers|
|Poly-lysine or gelatin||Cover slips (see this article about cover slip sizes)|
Step #1: Use sterile techniques.
It should go without saying, but I have to say it anyway as it is so important. Whenever you are doing cell culture you must use sterile techniques. This means that everything that goes into the cell-culture hood must be sterilized, and once in the hood you must use good technique to prevent cross contamination. In short, spray everything with ethanol, only open flasks in the hood, wear gloves and be careful not to let your pipette tips touch anything.
Step#2: (Optional) Clean your cover slips.
Your cover slips should be fairly clean out of the box, but they may have some dust on them. This dust can interfere with your ability to evenly coat your slides and thus impair your cells from evenly sticking to your cover slips. Therefore, you may choose to wash your uncoated cover slips. (Note: Do not wash pre-coated cover slips).
Washing Protocol: To wash your cover slips use a cleaning solution of 57% ethanol, 10% sodium hydroxide. Be careful and remember your PPE when handling. Once made, place your cover slips in a beaker and place this beaker in another container. This secondary container is to prevent the basic solution from accidently splashing onto your bench top or onto other equipment. Pour your cleaning solution over your slides and shake (gently!) for 2 hours at room temperature. Then rinse your slides well with distilled water. When done, keep your cover slips clean by keeping them covered as much as possible and only handling them on the edges with gloved hands, or with forceps.
Step#3: Sterilize your cover slips.
Regardless of whether you choose to wash your cover slips or not, they need to be sterilized. To sterilize your cover slips before use, place them exposed in your cell-culture hood with the UV light on for 20-30 minutes.
Step #4: Coat your cover slips.
Following sterilization, you need to coat the cover slips with either poly-lysine or gelatin. These coatings are necessary to help your cells stick to the glass surface (see also Catriona’s article on slide coatings). To coat, place your sterilized cover slips into the wells of a sterile 24-well cell culture plate — one cover slip per well. Then follow the directions below to coat with either poly-lysine or gelatin.
Poly-lysine coating protocol: Nearly all types of adherent cells will adhere to a poly-lysine coating, making it the most popular coating choice. To coat your slides with poly-lysine, add enough 1:10 poly-lysine solution* to cover the tops of each of your cover slips. Incubate your slides with this poly-lysine solution for 2 to 24 hours. Then aspirate off any remaining solution, rinse a few times with sterile PBS** and let your slides air dry in the hood for 15 minutes. It is important to know that the poly-lysine is bound to your cover slips only by electrostatic charge- it can be easily rubbed off. Therefore it is important to handle your slides carefully once they are coated. Coated slides are best used <4 months after coating.
*You can make this 1:10 poly-lysine solution (0.1-1 mg/ml of poly-lysine in 0.15 M borate buffer, pH 8.3, filter sterilized), buy it premade, or alternatively you can purchase cover slips that are pre-coated with poly-lysine. Depends on what you have more of: Time or money?
**PBS (phosphate buffered saline) is a common buffer used in cell culture and histology. The osmolarity and ion concentration of PBS is the same as human tissues, so it will not cause your cells to shrivel or explode. PBS is 137 mmol NaCl, 2.7 mmol KCl, 10 mmol Na2HPO4, and 2 mmol KH2PO4 at pH7.4. Sterile filter your PBS.
Gelatin-coating protocol: Cover your cover slips with a 0.1% gelatin in deionized H2O solution. Let stand for 10 minutes. Then aspirate off and let your slides air dry in the hood for 15 minutes. Once dry, the coated cover slips can be stored for future use.
Other coating methods: Almost any sort of extracellular matrix protein can be used to coat your cover slips, including collagen, fibronectin and laminin. But few offer the universality of the poly-lysine or gelatin coatings. For example, fibronectin only works with endothelial cells, fibroblasts, neurons and CHO cells, but not leukocytes or myoblasts. In contrast, the gelatin or poly-lysine methods are compatible with most adherent cell types, leaving you one less thing to worry about.
Step #5: Plate your cells.
Place your sterile and coated cover slips into a new sterile 24-well culture plate. Split your adherent cell line as you normally would with growth media. Plate your cells at your normal confluency (~10%) onto the surface of your cover slips. It will take ~400 to 500 µl of media to cover your cover slip. When done, replace the lid of your 24-well plate, and place your cover slips and cells back into your 37°C humidified cell incubator. Check your cover slips daily until your cells are ~70% confluence, usually in 2 to 3 days.
Step #6: Aspirate your media.
When your cells are 70% confluent (or more or less depending on what you want to image). Use the culture hood vacuum to remove the old growth media, leaving your adherent cells stuck to your cover slips. Immediately after aspirating proceed to the next step. Do not allow your cells to dry out.
Step #7: Fix your cells.
Next you need to fix your cells. The goal of fixation is to halt your cells decomposition and freeze cellular proteins and subcellular structures in place. There are two common classes of fixation: 1) Organic solvent methods and 2) The cross-linking method. The goal of both methods is to denature your proteins. Sadly, there is no way to anticipate the best fixation method for your staining or immunohistochemistry needs. Instead you will likely need to test a variety of fixation conditions for your particular situation. See this article for more on troubleshooting fixation.
Organic solvent methods: In this method, organic solvents such as alcohol or acetone are used. These organic solvents work to preserve your samples by removing lipids, dehydrating your tissue, and denaturing and precipitating the proteins in your cells.
To fix with organic solvents, use ice-cold methanol, ethanol or a 1:1 mix of ethanol and methanol to cover the cells on your cover slips. Once covered, incubate your cells in the freezer (-20°C) for 5 to 7 minutes. Do not worry about keeping your cells sterile at this point – you are killing them!
Cross-linking method: In this method, paraformaldehyde is used to form covalent chemical bonds (or cross-links) between the proteins in your tissue and their surroundings. This method usually provides the best preservation, especially of soluble proteins, but it can also ‘mask’ your antigens. If your antigens are ‘masked’ this means that the cross linking is preventing your antibody from recognizing your antigens. There are numerous methods to ‘unmask your antigens’ after cross-linking, using various combinations of proteinases, heat and chelators. For a full piece on unmasking your antigens see Catriona’s article.
To fix by cross-linking, cover your cells with 2 to 4% paraformaldehyde solution (diluted in PBS**). Incubate your cells in this solution for 10 to 20 minutes at room temperature. Note some cells can be damaged by the abrupt change between the culture media’s osmolarity and the fixation solution’s osmolarity. If this is your case, you may want to spike your growth media with ~500µl of 4% paraformaldehyde, wait 2 minutes, aspirate, then cover your cells with pure 2 to 4% paraformaldehyde for 10 to 20 minutes.
Step #8: Rinse.
Gently rinse your fixed cells with PBS** to remove any fixation agent. Be careful (pipette gently!) when applying your PBS so you do not disrupt your fixed cells. When done, your cover slips with their fixed cells can be stored under PBS in the refrigerator for up to three months.
Step #9: Proceed with staining and imaging.
Your adherent cells are now fixed to your cover slip and ready for staining, mounting and imaging when you are.
Good luck and happy imaging!
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