Евгений, I keep all my samples in  glycerol. No problem up to now.  But my samples are fossil

The long term storage requirements of modern pollen reference collections seem to be poorly understood (at least in Australia). My limited experience with Australian reference collections established in the early 1970s indicates that many of the old slides have deteriorated terribly – though I believe none of this early material was suspended in glycerol, but in silicone oil.  I am not sure there is a consensus on the mechanism through which the pollen deteriorated, but I recently came across a pollen reference collection mostly made in the 1950s, which had for most of the intervening period been kept in a cupboard, completely unused. In general the pollen is still in exquisite condition. Some colleagues suggested that the difference can be explained by the extent to which the slides have been exposed to light – though whether the incident light of an ordinary laboratory is sufficient, albeit over decades of routine consultation of the reference collection; or whether it is the the intense light of the microscope that photo-degrades the pollen, I am not sure. I would be interested to hear from palynologists working with much older reference collections: do any of you have slides made prior to the 1960s, still in good condition?

In general I try to reduce the quantity of flora material, ideally to anthers only, in order to minimize the need for a KOH step. Then simply acetolyse, dehydrate in ethanol, suspend in glycerol.

Andrei and Kale, thank you very much.

Kale, our oldest reference collection was made in early 1980s, sealed with Canada balsam, and half of its samples have faded so far.

My experience with old reference slides is that the pollen is better kept in silicon oil than in glycerol. I have been told that the worst probems with glycerol happen when the glycerol is not pure enough and still contains some water.

Hi there, I don't think it is glycerol what destroys pollen grains but micro-organisms that develop there afterwards...I have looked at fossil pollen samples from the 80's that contained glycerol and some of them were full of "bugs" that you could see through the microscope.

Hi,

I have analyzed slides stored in my reference collection. Slides made in glycerol/glycerine jelly swell by as much as 50% after a few decades. Those effects are not immediately apparent, but after 10 years become noticeable. With the swelling there is also a loss of detail in surface pattern.

Silicone oil is more durable, but air-bells can form and there are also problems with bacterial decay. A further problem is that if you make your regular counts in glycerol, the silicone oil mounted material will apper smaller and may have a rather deflated appearance.

We now use a 24 hr soak in glycerol, followed by mounting in Syn-Matrix. This gives excellent results that should be inert over the long-term. The only downside is that it is a fixed mount and grains cannot be rolled. Having sufficient grains on the sample generally overcomes this problem, and a few grains can be manually oriented if need be prior to adding a coverslip.

Lydie, Gonzalo and Mark, thank you for your valuable answers.

Best wishes.

We have an extensive modern pollen reference slide collection in our NZ lab - many of the slides were made as far back as the 1950s and through the decades to the present. Some of the earliest slides made are still perfectly useful. The pollen is mounted in glycerine jelly (42g gelatin,  6g phenol (CARE: phenol is very toxic), 114ml dH2O, 157ml glycerin (glycerol) - the phenol prevents any fungal or bacterial growth on the slides).

It seems to me that the trick for longevity is keeping these mounts completely airtight, so the crucial factor is what you use to seal the coverslips. Some of the very oldest slides we have had their coverslips sealed with a black lacquer paint and the pollen on these are still perfectly preserved and not faded. We have fossil slides made up this way from the same era, and amazingly they are still perfect too. Later in the 1970s some coverslips were sealed with a nail varnish and these are terrible - the varnish develops micro-cracks and the slides dry out or get air bubbles and the pollen in these has deteriorated badly. Lately, we have been using wax to seal the coverslips and these appear to work very well at sealing the coverslips too. We have 20 year old ones made like this and they are still perfect, although time will tell how well they will preserve into the future.... hope this is of some help.

Janet, thank you very much for your help.

I agree with you that it is the access of air that may cause the deterioration of grains.

Cushing wrote extensively about this. Aerobic decay causes bacterial growth and the atack of bacteria starts as pin-pricks and, with time, can cause grains to appear to be punctate. Glycerol and glycerine contain residual water that is absorbed by pollen and causes the observed swelling.

We store our modern pollen samples in Silicone Oil (Dow Corning, 220 fluid, 1,000 cs. viscosity) in glass vials with screw top lids. We then mount our samples on slides and seal the slide cover with nail polish, and store them in cabinets away from light. We prefer to have the ability to move the pollen for imaging and identification purposes, so we do not typically use permanent mounting techniques. Samples collected in the early 1980's still look great, where some are more degraded. However, they were not in good condition at the time of mounting, so it appears that it is not the results of the mounting agent. However, we do keep extra material in vials (in Silicone) so we can make new slides for our reference collection if needed.

Thank you very much, Alwynne.

That's a very practical article.

Dear Rohozin

When I started to work with pollen I used KOH to stabilize the pollen grains. But the color go out. Today I use only acetolise method to process pollen from flowers, honey and more. It is the best way to conserve  it. the slides I use glycerin mixed with phenol to protect against fungal attack.

Hi there, I just wanted to add an update to this post, for future readers researching pollen analysis methods. We (Quaternary Environments, Royal Alberta Museum (RAM)) no longer use nail polish as a sealant for our reference and working slides, as it is destructive to the pollen long-term based on literature (Cushing 2011) and discussions with Conservation specialists here at the RAM. We now use Acryloid (Paraloid) B-72 (conservation grade polymer/adhesive) or acrylic paint (art suppliers). The paint is the easiest to use, and provides a good seal. B-72 is easier to use if mixed with ethanol vs. acetone, which makes it less stringy when applying. It also provides a very good seal. Having a good seal, with a conservation-grade sealant, is likely one of the most important factors in preserving material long-term, both preventing biological and chemical degradation to the pollen. As discussed above and based on some users feedback, Syn-Matrix sounds like it may be a good potential alternative to other slide mounting mediums; however, fixed mounts like Syn-Matrix and others would only apply to potential use for preparing reference slides (with sufficient grains in both polar and equatorial positions). If a project requires counting and identifying pollen from slides, you need to be able to move the palynological material. Also, as I do not have experience using Syn-Matrix, I wonder how it impacts the optical properties when imaging pollen from these fixed mounts? Once pollen is fixed, if you are gathering measurements of pollen from reference material, it would be important to have extra pollen residue to put in oil, so you can obtain both polar and equatorial measurements from the same grain plus multiple grains. Some things to ponder while deciding on mounting mediums and sealants for you collections and projects. Moore/Webb/Collinson, Pollen Analysis (Blackwell Sci. Publications) is a good resource discussing general pollen analysis methods and techniques.

https://www.uco.es/rea/infor_rea/manual_eng.pdf

Diana Tirlea when you seal your samples, you use silicone oil? And you don't stain them correct? Im used to using fuchsin gel as it helps us extract pollen from pollinators --so it's preferred. But Im wondering if I need to add phenol if I seal the samples and refrigerate the gel.

Hi, yes, we do stain the pollen at the end of acetolysis using Safranin (red/pinkish colour). Silicone is added to the stained pollen, which is then subsampled to make a slide and sealed with B-72 or acrylic paint. Based on Parrish (2004, see below), you do not have to use phenol in the gel mixture if you refrigerate your gel mixture. However, as Jose Oteros manual and Sílvio José Reis da Silva comments above indicates, for long-term storage of your samples / slides using fushin gel, phenol is used when making the mixture to help preserve the samples. So it would be good to know if you plan to keep your samples long-term or just need the gel to sample pollen from insects, which you then count only (sample is exhausted). If you keep your samples in the fridge (without adding phenol) at 3 deg C, it will reduce microbial growth - but you should monitor your samples. I store wet soil and macrofossil samples (in water only) in the fridge for a few weeks during processing, and they are just fine. However, in my experience, even when samples are well-sealed and stored in the fridge they can become susceptible to microbial growth if stored too long, including just a month for some samples. I would suggest you reach out to people using phenol to get further feedback on how long gel preserves in the fridge without pheonl added and determine if you need to keep these long-term. Berry J. Brosi, Ph.D. (Department of Environmental Sciences Emory University brosilab.wordpress.com) uses gel in the field for removing pollen from insects and may be a good person to reach out to.

Some other resources which may help in regards to preparing fresh pollen using gel:

1) Gretchen D. Jones POLLEN ANALYSES FOR POLLINATION RESEARCH, UNACETOLYZED POLLEN Journal of Pollination Ecology, 9(13), 2012, pp 96-107

2) Keith S Delaplane, Arnon Dag, Robert G Danka, Breno M Freitas, Lucas A Garibaldi, R Mark Goodwin & Jose I Hormaza (2013) Standard methods for pollination research with Apis￿mellifera, Journal of Apicultural Research, 52:4, 1-28, DOI: 10.3896/IBRA.1.52.4.12

3) Judy Parrish 2004 Teaching Issues and Experiments in Ecology - TIEE Volume 2 (tiee.ecoed.net) EXPERIMENTS Pollination Ecology: Field Studies of Insect Visitation and Pollen Transfer Rates

Thanks for the references and the explanation. I appreciate that. I am hoping to avoid using phenol. I guess I was wondering do you still need phenol with the gel if you seal the slides? I did my part of masters with gel (but didn't make it) and Ive learned since that we did not use phenol. Recently I went back to my old lab and found some petri dishes of the gel...there were no obvious signs of mould but I only viewed it with the naked eye. I assume its no good. For my purposes now, I would like to create a reference collection. One that could theoretically last for a long time...but I also have limited resources right now with COVID and our lab just starting up. Ive read much of the Jones paper but Ill try to get the others.