Notes on Imaging Chambers and CoverSlips
When you are preparing samples to bring to the NIC, please ensure that your cover glasses and or glass-bottom chambers are #1.5 glass. This glass is 170µm thick (high quality glass will have a tolerance of ±5µm) and most microscope objectives are designed for this glass thickness. In fact the coverslip is usually the first lens considered when the lightpath is designed. The thickness of the coverglass is extremely important for using high NA oil-immersion objectives. Multiple companies supply high-tolerance #1.5 coverslips and imaging chambers. #0 and #1 coverslips should only be used when extra working distance is absolutely essential (they are 100µm and 150µm thick respectively), and when they are used, a member of the staff should be notified so that an objective can be placed onto the system to compensate for the non-standard coverslip thickness. When plating cells onto coverslips or into live cell chambers, it may be necessary to modify the glass to enhance cell attachment and/or morphology. Poly-lysine, fibronectin, and collagen are all suitable for this purpose.
Live cell imaging chambers that are compatible with our incubation systems are 35mm dishes (available from multiple suppliers), and Lab-Tek II Chambered Coverglasses. ibidi chambers, with a footprint similar to a standard micrsocopy slide are also compatible with our stage-top incubators. However, if you wish to perform DIC imaging, please consider the top/lid of your imaging chamber. DIC requires polarized light. The birefringence of most plastics prevents the maintenace of polarization. So, please ensure that any imaging chamber that you have comes with a DIC compatible lid or we will have to remove it for performing DIC.
When mounting coverslips on slides, you will get better results by putting the coverslip directly in the middle of the slide. Because we are using inverted microscopes, the mounting of the slide can be affected by having a coverslip that hits the slide holder. When the slide is sitting on the holder on one end and a coverslip on the other, the sample will be held at an angle, which will show up in tiling experiments. This is especially important when using the MadCityLabs piezo Z and 3 inch slide holder on the A1 confocal. Putting coverslips in the middle will also prevent you from smearing oil onto the slide holder.
Immunofluorescence is a workhorse technique for many users. However, there are multiple considerations that need to be kept in mind.
Antibody specificity - Please ensure that you’ve done adequate controls to confirm that your primary and secondary antibodies are specific for your antigen. This can be done by doing a) a knockout or knockdown control to ensure that your primary is actually recognizing your protein of interest - b) a no-primary control for your secondary to see how it stains your cells - c) a co-stain where you omit each primary antibody while including all secondaries to ensure that your secondaries are not reacting with the incorrect primary antibody. Purchasing highly cross-subtracted antibodies such as the donkey secondary antibodies made by Jackson immunoresearch is usually essential when trying to use primary antibodies from closely related species (i.e. rat and mouse).
Antibody dilutions - When you are setting up staining protocols, it is useful to determine the lowest dilution of primary and secondary antibodies that produce a real signal. Using too much primary and secondary antibodies can lead to aberrant staining.
Fixation - There are many different fixatives that can be used for immunofluorescence. Paraformaldehyde (PFA), methanol, glutaraldehyde, acetone, and glyoxal have all been used successfully. It is frequently beneficial to test several different fixatives to find out which one best presevers your structure of interest as well as the antigenicity of the protein that you are looking for. It is also possible to combine fixatives to enhance their efficacy. For instance, a mixture of 90% methanol with 10% (by volume; 3.2% concentration if working from a 32% stock) PFA works very well for fixing the cytoskeleton. Note that PFA and glutaraldehyde work the best at a basic pH so using a basic buffer like PHEM at pH8.0 or adding some sodium bicarbonate to MeOH + PFA will enhance the fixation. After fixation it is usually best to get rid of unreacted aldehydes by washing with a buffer with a primary amine like Tris or by incubating with (freshly-prepared) sodium borohydride. Fixative quality also impacts how well it works. Freshly prepared PFA or PFA or glutaraldehyde diluted from a freshly opened concentrated stock will produce better results.
Mounting - Although mounting media is usually the last thing that scientists consider when performing IF, the selection of the proper mounting medium is essential for getting the most out of the samples that you work so hard to optimize. Ideally, you want a medium that will preserve staining and combat photobleaching during imaging.
You should consider the refractive index (RI) of the medium. If the RI is too high (such that it matches or is close to the glass of the coverslip; RI = 1.5), the Perfect Focus System will not work properly since the PFS is looking for the change in RI going from the coverslip to the medium.
The ability of the medium to maintain staining is important and should be tested. We have observed that some types of staining are very labile in one type of medium but permanent in another.
Some media will harden over time, but others won’t. If you select a hardening medium, you should evaluate whether this causes the sample to flatten and/or contract.
If you seal your coverslips with nail polish, please ensure that the nail polish is fully dried before you start imaging. CoverGrip™ by Biotium is superior in many ways to nail polish. Nail polish can contain chemicals that can leach into the mounting medium and either fluoresce or diminish your sample fluorescence.
Using mounting media with DAPI is inferior to incorporating DAPI into your secondary antibody staining and creates more background.
After your coverslip is mounted and stable, you should wash its surface with distilled water to avoid contaminating the immersion oil with salts.
Selecting Fluorescent Proteins
UC San Diego has a history of developing fluorescent proteins, recognized by the awarding of the Nobel Prize to the late Roger Tsien in 2008. When designing experiments with fluorescent proteins, it is important to consider what you wish to see. GFP and mCherry are the most common red and green proteins that are being used in labs, but there are many better options available (see https://www.fpbase.org/spectra/). For the lasers and LEDs available in the NIC, mTagBFP2 (blue), mTurquoise2 (cyan), mEmerald/sGFP2/mNeonGreen (green), mVenus (yellow), mScarlet-I/mCherry (red) and miRFP670 (far-red: requires biliverdin as a cofactor) are all recommended (NB mTurquoise2 and mVenus can only be used with Cyril and Krieger).
SNAP, CLIP, and Halo-tags can also be used with a variety of very bright cell permeant and impermeant organic dyes. Additionally, it is possible to use photoactivatable and photoconvertible proteins such as PA-GFP or mEos3 with our DMD system on the spinning disk/TIRF system.
Finally, when tagging your favorite protein, the length of the linker should be considered. It may be useful to test several linker lengths and positioning the tag at the N- and C-terminus to create a protein that behaves like the endogenous. When designing CRISPR strategies to add an FP to an endogenous locus, it can be very useful to add multiple copies of the fluorescent reporter to more easily see small numbers of the endogenous protein.