These methods pertain to cleaning diatoms from unialgal cultures for qualitative purposes. For cleaning diatoms from sediment samples, including quantitative estimates, useful resources are the UCL Geography Dept, Diatoms Ireland and Diatoms of North America.

The main difference between sampling from cultures and from field samples is that in most cases cultures are unialgal and contamination is self evident. However because diatoms are ubiquitous and physically resilient cross transfer from preparation tubes or even tap water can seriously compromise species presence and abundance data in field samples. Therefore all steps should be taken to avoid contamination. In terms of good practice these steps should also be applied to preparing diatoms from cultures.

 

Diatom mounting media

 

Earlier editions of this website included links to the merits of different mounting media and their interplay with optical resolution and contrast compiled by Gene Stoermer and F.A.S. Sterrenberg and presented in the Diatom-L list (Indiana University). These links are now dead so users are directed to the Diatom-L list (password required).

Several different mountants with different refractive indices (R.I.) are available to mount diatoms for light microscopy but those with a high R.I. are generally best for mounting diatoms. For most diatoms high R.I. mountants provide greater contrast in the microscopic image and a better depth of field for a given Numerical Aperture (N.A.). A general refractive index for diatoms is about 1.43 so they would be invisible in a media of this index. However, one of Sterrenbergs most critical comments is that “contrast depends on the difference in RI not its absolute value,” (also note that the mountant RI affects contrast, not resolution). Therefore some heavily silicified diatoms with a high R.I. may actually appear better in low R.I. mounts (eg R.I. of air = 1.0).

In ANACC we use Naphrax which has a R.I. of 1.65 in solution but increases to 1.73 when the solvent toluene is heated out of the resin during slide preparation (note – due to air transport prohibition on products containing toluene, Naphrax may not be readily available in all countries. Pleurax is a substitute).

Safety

Several different chemicals are available to remove the organic matter from diatoms and leave the silicon frustule intact. We use Hydrogen peroxide (H₂O₂) which is a strong oxidizing agent. Rubber gloves and eye protection should be used when handling it and spills should be diluted with water before using paper towel to absorb the spill. Toluene is a hazardous agent therefore Naphrax should always be handled inside a fume cupboard particularly as the procedure involves bubbling off the toluene on a hotplate.

Digestion procedure for both Light microscopy and TEM

Glass centrifuge tubes are commonly used but suitable plastic centrifuge tubes (eg Falcon™) that can withstand boiling hydrogen peroxide may also be used. Diatoms are relatively robust compared to other microalgae so the centrifuging method is suitable, especially for multiple samples. An alternative filtration method could be used if excessive damage or significant cell loss at the rinsing steps is encountered.

1. Centrifuge 10mL – 15mL of culture for 4 mins x 3000 – 4000 rpm. Remove media. If necessary, repeat to obtain pellet of cells.

 

2. Resuspend pellet in distilled water. Centrifuge at 3000 – 4000 rpm x 4 min. Remove water and repeat twice.

 

3. Resuspend pellet in 1 – 2mL Hydrogen Peroxide (30% w/v). Mix well.

 

4. Place tubes in a rack in a water bath (or in a large volume beaker with distilled water on a hotplate) at room temperature and increase to 80-90 ⁰C for 1 – 1 ¹/₂ Allow to cool.

 

5. Fill tubes to ~10mL with distilled water (… this seems to help the cells to sink during centrifuging). Centrifuge at 3000 – 4000 rpm x 4 min. Remove acid. (The cells are “colourless” and don’t pellet as well so more care is needed to prevent removing too many cells). At this stage a small drop of sample can be taken and observed under the microscope to determine how efficient the oxidative step was and give an indication of the diatom density.

 

6. Rinse pellet twice with distilled water, centrifuging each time.

 

7. Remove pellet and add to about 1mL of distilled water in a clean 1.5mL Eppendorf tube and label. Note only experience will determine whether samples may need further concentration or dilution to obtain a good density of cells on either the microscope slide or formvar coated grids. Generally samples should be neither milky nor totally clear but when held up to the light fine particles in suspension should be seen.

For Light Microscopy

Work in an environment free from dust and air currents and ensure that slides and coverslips are clean before mounting. For each sample make up to 3 slides which allows for poor mounting, archiving and breakages.

 

equipment : preplugged pasteur pipettes, glass slides, 13 -19 mm diameter round coverslips, forceps, settling tray or container, Naphrax, hotplate.

 

1. Place up to 0.5 mL of mixed suspension onto each coverslip, cover and leave undisturbed for up to two days as this is the gentlest and most efficient means of obtaining even cover of diatoms. Alternatively, coverslip samples may be settled for 15-30 minutes and then dried more rapidly using very low heat on a hotplate. This gives lower quality but still acceptable slides for qualitative analysis). Note this is an important step and counter intuitive to normal “wet” slide preparation, however drying on the slide leads to poor results as “the spherical correction of a high NA objective does not know its head from its tail” – Sterrenberg.
2. Heat a hotplate in a fume hood to 130 ⁰
3. Place 1 drop of Naphrax on a glass slide and using forceps invert the coverslip with the dried diatoms over the drop.
4. Heat the slide on the hotplate for 15 minutes to drive off the toluene in the Naphrax.
5. Allow the slide to cool and use a fingertip or forceps to ensure the coverslip can not move, otherwise the slide will need to be heated for longer.

 

 

For TEM

1. Centrifuge the Eppendorf tube and remove most of the water.
2. Put a small drop of water/cells onto a microscope slide and check under the light microscope that there are actually cells in the tube (… and that you haven’t sucked all of them out during rinses).
3. Prior to putting samples on grids, re-centrifuge the tubes to collect/pellet the cells in the bottom of the Eppendorf tubes.

 

 

Loading grids

1. Make sure the formvar film is dry before loading samples on. It is also wise to check the quality of your grid/film under the light microscope.You can buy Formvar coated grids or prepare your own as detailed below (former more expensive)

 

2. Add one drop of cells onto the grid and leave for a few minutes to allow the cells to settle.

 

1. Gently draw off the water. This is best done using “points” of filter paper. Make these by cutting the filter paper into triangles. e.g………….

1. Check the grid again under the light microscope. This is crucial as it is a waste of money (not to mention frustrating) to put dodgy grids or empty grids under the TEM.

 

Preparing Formvar coated grids

Mix formvar powder in 1-2 dichloroethylene and make up to 0.3 to 0.8% (typically 0.6%).

As with all TEM prep, work as dust free as possible

Over a beaker, pour Formvar down the face of a glass microscope slide. The Formvar sets very quickly. Use a scalpel to trim around the edges of the Formvar. Then place the slide at an angle into a basin of MilliQ to float the Formvar off the slide. Using forceps place grids onto the film and then remove the film from the water by overlaying it with a section of Parafilm backing paper (not the parafilm itself). The grid laden paper can then be removed to a clean storage place (eg peri dish).

While this section concentrates on diatom frustule cleaning for TEM mounting the cleaned material is also suitable for SEM examination.

 

References

Flemming, W. D. 1954. Naphrax: a synthetic mounting medium of high refractive index. New and improved methods of preparation. J. Roy.Micros. Soc. 74:42.

Pickett-Heaps, J. D. (1998). A rapid, highly efficient method for collecting, fixing and embedding planktonic and other small cells for electron microscopy. J. Phycol. 34, 1088-1089.