Here protocols are listed which are useful for work in our lab

Sealing Samples

Many samples are prepared directly on the coverslip and then embedded and sealed. Sealing is important for protecting the sample from the microscope immersion oil but especially also for protecting the objective from coming via the immersion oil in contact with the fluorophores of the sample, which would yield to a permanent fluorescent background of the objective.

Three main approaches are used to seal the sample:

  • Nail polish

Place the coverslip with the sample pointing towards the slide onto the slide. Dip the brush of the nail polish into the bottle and then dry it a little bit by squeezing gently against the inner side of the bottle neck. Place four small dots of nail polish first on the corners of the coverslip to fix it a little bit. Be careful to not pull the brush over the coveslip as it may leave a thin pulled thread of nail polish behind across it. The nail polish should not be too old, as it will be to solid then and hard to handle.

Once the corners are a little bit fixed by the four dots, seal the edges relatively quickly. It is important that the nail polish always completely covers the edges. This means a little of it has to by visible over the coverslip at each of its sides. Not just at the side seemingly touching it! You will notice that the capillary forces (at dry samples) will quickly start to pull the nail polish under it. This will destroy part of your sample, but if you don’t wait too long, and all capillary effects go from all sides, pressure builds up inside and there will be some part in the middle left, with still air under the coverslip.

Then leave the sealed slide to dry for at least 20 min. With your fingernail (flat topide) you can carefully test, whether the nail-polish is still sticky. For uncritical samples such as beads, you can also place the sample for drying onto a hot plate. The nail polish has to be dry before you put it under the microscope, as the objective can easily tough the nail polish which will then lead to a very serious deterioration of the objective. Nail polish is very hard to clean. On the sample, you can carefully use acetone but do NOT use it on objectives, as it dissolves the glue of the lenses. Objective lenses should be cleaned with Mr. Muscle, Fish-Water, lense cleaning fluid or Isoprobanol. Only of there are no more other options (because the residue still did not vanish from the front surface) and the issue has been discussed with your supervisor, one can very very carefully and shortly wipe the objective front with a tiny bit of acetone.

If you pre-embedded the sample (e.g. with a glycerol mounting) it may be important to think about the acidity of the nail-polish. The nail polish can make your embedded sample acidic! Rumor (E.E., KCL) has it that Swizz nail polish is less acidic than the Austrian version.

Nail polish is (due to the acidity) NOT recommended for live cells.

  • Dental glue

Instead of sealing with nail-polish, a silicon-based two-component material used in dentistry (for impressensions) can be used. Mix it in equal parts on a little dish and quickly apply it using a wooden stick around your sample similar to the steps for nail-polish above. It sets quite quickly (1 min liquid enough to use, ~5 min until dry) so you need to hurry or make two batches for using it. The advantage of this is that it is gentle to live cells and quite easy to pull off again for remounting your samples.

Dental glue is recommended for living cells and for dSTORM samples (as they can easily be reimbedded with fresh medium a few days later).

  • Velap

Velap is equal volumes of petroleum jelly (also called vaseline), lanolin and paraffin. The material has to be first heated on a hot plate so it is liquid and then applied with a wood stick to seal the sample, where it becomes solid by cooling when it contacts the sample. It is handy to work with and gentle to the sample.

Recommended for live cells.

Fluorescent Plane Samples

Fluorescent planes are the standard way of judging the confocality of a fluorescent microscope system. Such samples can be generated by spin-coating a labelled polymer onto a #1.5 coverslip (this is how the confocal check calibration samples we have from the Brakenhoff lab were generated) or by pulling the coverslip out of a Langmuir-blodget layer.

However, the cheapest and easiest ways is to simply dry some of the ink of a fluorescent text marker pen onto a coverslip:

  1. Have two #1.5  coverslips handy
  2. touch one coverslip a few times (e.g. 3) gently from one side with a green fluorescent marker pen, each tough leaving a little bit of marker ink behind.
  3. quickly put the second coverslip against the first one. The residues will become wider by cappilary forces.
  4. move the coverslips gently in circles. Be sure that the ink residues never reach the side of a coveslip.
  5. Grip both coverslips and rapidly pull them apart in a sliding movement sideways. REMEMBER WICH  SIDE IS THE TOP SIDE!
  6. Put the coverslips with the coated side pointing upward onto the table for drying
  7. Let them dry (~10 min) completely
  8. Mount them on a slide (with rounded edges) and seal with nail polish (see below).

Various marker pens have quite different compositions. Most consist of nm-sized beads (200-500nm estimated diameter). The orange color is sometimes even a mixture of green and red fluorescent beads. If you look around, you find areas where you can see the beads individually but also areas consisting of hexagonally self-assembled mono-layers. Such layers or depending on the application also the thicker multi-layer areas can serve as fluorescent plane samples

Beads

Beads can be obtained from various commercial manufacturers (e.g. multispec beads, Kisker, Invitrogen, …). For superresolution microscopy they should have a diameter of less than 100nm. However, if 200 nm or more are OK for the topic of interest, they are also contained in many of the fluorescent text marker pens.

Dilute the beads in water by about 1:10000 from the stock solution (usually done in two steps 1:100). Use a small pipette and please about 5×5=25 tiny drops (as small as possible, 0.5 µl each) side by side on a coverslip. If they melt together, you can try pulling them apart again with the tip of the pipette. Let them dry (~10 min) with the coated side pointing upwards. The dried drops will display a nice gradient of concetration (like coffee-stains) each with a ring of densely coated beads and only very frew in the middle or outside. This way it is good to find the right concentration for your experiment e.g. for measuring point spread functions.

Embedd the sample (e.g. using the antifade protocol) with antifade and seal (see above).

Fixing Pollen Grains

(Protocol communicated by Dr. Brad Amos, Cambridge)

This has to be done in a fume cupboard with eye protection because of the use of strong acids! Handling the strong acids and acetic anhydride (as in this protocol) is very dangerous! They should be used in tiny quantities and diluted into at least 20x the volume of water for washing. An Eppendorf centrifuge is fine for spinning a small quantity of the pollen grains down but a bigger bench centrifuge with 50ml tubes is good for the washing stages. Eye protection with goggles is essential. If you are not used to handling chemicals in a fume hood, you should not work alone and the partner should be ready to hold an acid-splashed eye open immediately under a cold water tap for 10 or more minutes until medical help is available. This is even more necessary for protocols with concentrated alkali and heating, which some people find necessary.

1. Remove the anthers from the plants and store them in glacial acetic acid. They can be stored indefinitely. With some plants such as Passiflora, Cobaea and Lilium the anthers are large and can be handled singly with forceps. With Taraxacum ( Dandelion)  and other compositae it is best to cut into the central part of the flower ( actually hundreds of small flowers, crush in acetic acid and then filter out large particles to obtain a thick suspension of the small but very attractive pollen grains. I have attached an SEM photo from the web.

2. Spin down the pollen grains in a centrifuge tube, pipette off the acetic acid and add a few drops of concentrated hydrochloric acid and leave for at least 5 min before adding distilled water and spinning down again and washing with more distilled water.

3. Repeat with concentrated nitric and then sulphuric acids.

4. Finally, wash in distilled water and dehydrate through a series of ethanol/ water mixtures, e.g. 70% ethanol,,90%. 95%. two changes of absolute ethanol.

5. Pass through two changes of xylene.

6. Mount in Histomount (Fisher) or a similar hydrophobic resin permanent mountant.  Use a highly concentrated suspension of pollen grains that will be quite numerous even when spread into a thin film under the coverslip. Dandelion pollen is easy to get in large quantities.

The procedure in the link below is one which is used widely and involves a highly reactive compound, acetic anhydride, which tends to boil and spurt if added to water but is OK with acetic acid. It is definitely a fume-cupboard and eye protection job.  I think it is probably worth using the acetic anhydride, in spite of its nasty reactivity, because it tends to enhance the autofluorescence of the pollen , which is a red emission excited by green illumination. The autofluorescence is thought to be due to some kind of carotenoid pigment which is stable enough to survive the strong acids and is even present in fossil material.

 

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