To image a thick sample, it is crucial to match the refraction index of the sample with that of the immersion medium between the sample and the objective. Typically, life samples are in an aqueous solution like culture medium which has a refraction index of 1.33. Unfortunately organoids often have a higher refraction index closer to 1.44 therefore as one images deeper into the organoids, light scatters due to the refraction index mismatch and the images become blurry.
This paper presents a product that has a high RI and is compatible with cell culture. Good to keep in mind for those who image organoids over time.
On Wednesday (13th) at 9:30 in Lipid seminar room in Neo (KI Flemingsberg), please come and enjoy a short seminar presenting a new way to label organelles in live cells.
LabLife will present their product called Viromer Cytostain.
We will stream the seminar live so you can follow it even from your desk! 🙂
Do not miss the seminar tomorrow morning at Scilife: the inventor of the CUBIC technique, Etsuo Susaki, will present his technique which can clear fatty and dense tissues.
Click here to know where and when.
And here is the link to the updated Cubic protocol.
Together with the coverslip and the immersion medium (oil, water, glycerol or air), the sample mounting medium is part of the design of a microscopy objective. Matching the refraction index of the sample to the one recommended by the manufacturer of the objective will make the sample transparent for the objective, drastically improving fluorescence microscopy in samples thicker than a couple of um (i.e. anything except fluorescent beads!).
Not matching the refraction indices is equivalent to watching something through a wet window… Far from optimal! :-/
The refraction index recommended by the manufacturer is the same as the RI of the immersion medium: 1.52 for an oil immersion objective, 1.47 for a glycerol objective, 1.33 for a water objective, 1 for an air objective.
This article compares 7 mounting media and their effect on the refraction index of brain samples. CFM3 seems to be a cool mounting medium. The company that produces it has partially paid for the study but it sounds worth a try anyway!
In the same vein, this article presents a non-toxic way to change the refraction index of cell culture medium (not the sample) to improve imaging of live samples. Sounds pretty promising to grow live organoids which quickly become opaque. This will also be very useful when clearing samples as the sample chamber on a light sheet microscope is big so this is a cheap way to fill the chamber for imaging. 🙂
If you try any of these 2 chemicals, please leave a comment to let us know how it went! 🙂
Poor PFA fixation often causes trouble in antibody staining. Folded cells, poorly preserved cytoskeleton… These artifacts appear when the stock of PFA gets older and degrades. Buying ready made PFA solutions, most of which contain 10-15% of methanol, can also lead to low labelling with some antibodies.
Glyoxal seems to be a good alternative. It had the added advantage that it is less toxic.
Check this article to know more. 🙂
The Live Cell Imaging facility is back with its intensive microscopy course! In 2018 we moved to a new building so there was no course but we will strike again in Jan-Feb 2019!!You will definitely learn tons at our course. Have a look at the program and judge for yourself.
As usual we run 2 courses in parallel:
#2870, 6 points, is the full course with all lectures, workshops, demos… This will run 22/jan-08 feb, 3 days/week 9:00-17:15.
#2871, 4.5 points, is the same minus some workshops and demos. This course will run at the same dates but 10:00-15:00.
The rest of the time is used for home assignments.
All lectures are open to the public without any registration so tell your colleagues!
Course applications are open from today and until the 15th of November. The full course (2870) has only 16 spots available so just go for it NOW! 🙂
As you all (nearlyish) know, one should never place a sample on a thick glass slide and add a coverslip. Instead, the sample should be placed on the coverslip then covered with a thick glass slide. And the coverslip should be 170 um thick (also labelled thickness #1.5).
Why is that? Because the coverslip is part of the design of the objective and all objectives from all manufacturers are designed to image through 170 um glass and assuming that the sample is directly in contact with the coverslip.
What about superfrost slides that one uses to make sure tissue sections don’t float away during antigen retrieval? No worry! You can make your own superfrost coverslips. It is cheap and you can prepare tons at the same time. Here is the protocol (and pasted below).
Not convinced? You will only see the difference when you compare side by side! The images of your tissue will be much sharper if the sample is on the coverslip because when you put your sample on the slide, either the thick glass or the mounting medium end up between the objective and the sample. The objective is not designed for this. ?
Here is the protocol:
- Gelatin-coating solution: 1 L deionized H2O, 5 g gelatin, 0.5 g chromium potassium sulfate dodecahydrate CrK(SO4)2 · 12H2O
- Filter units
- Histological slides
- Hot plate with magnetic stirrer
- Slide racks
- Staining dish
- Prepare the gelatin-coating solution by dissolving 5 g of gelatin in 1 L of heated, deionized H2O (temperature should not exceed 45 °C).
- After the gelatin has dissolved, add 0.5 g of chromium potassium sulfate dodecahydrate. Chromium potassium sulfate dodecahydrate will positively charge the slides allowing them to attract negatively charged tissue sections.
- Filter this solution and store at 2-8 °C until use. It is recommended that this solution be filtered again immediately before use (adjust to room temperature before filtration).
- Place the histological slides into metal racks.
Note: The slides should be cleaned by washing them in soapy water and rinsing them thoroughly, first in tap water and finally in deionized water.
- Dip the racks containing the slides 3 to 5 times (~5 seconds each) into the gelatin-coating solution.
- Remove the racks containing the slides and let them drain. Blot excess solution from the racks onto filter paper (gently tap the racks against the filter paper for better drainage).
- Place the racks containing the slides on the lab bench and cover them with paper towels to protect them from dust.
- Dry at room temperature for 48 hours.
- Dried slides can be put back into the boxes that they arrived in and stored at room temperature until use. Slides intended for cryostat sections can be stored at -20 °C.
Have you ever heard about Superfolded GFP? It is 50% brighter than GFP! And mScarlet is almost 6 times brighter than mRFP! How do I know? I look at this fantastic database called FPBase.
You can see which fluorescent protein is monomeric, sort them by excitation and emission or find which bleaches least or maturates fastest! Great tool! 🙂
Fluorophores are constantly being developed. If you make a new plasmid, make sure you check that the one your lab has been using for trillions of years is the very best one!
If you want to image a large piece of tissue, it is sometimes difficult to get it to stay still in a dish while you are imaging it, especially if you have medium on top.
One nice way is to make a silicon well around it, fill the well with medium then add a coverslip on top. This allow you to keep your sample for a long time.
Twinsil by Picodent works nicely. 🙂
Yet another chance to try RNA labelling: The FENO facility, here in Flemingsberg, has purchased a machine to multiplex RNA scope. They will present it on the 16th of October. 🙂
Here is the announcement.