PTA colour change, agarose and destaining

On 18/03/2019 21:38, Gonzalez, Brett wrote:

Hi Sarah…

My name is Brett Gonzalez and I am a postdoc at the Smithsonian National Museum of Natural History, previously a Ph.D. student with Katrine Worsaae. I am not sure if you remember, but last year I emailed you regarding some general advices towards integrating CT work into my research. Here at the museum we have a newly installed GE nanoCT and since the technicians are still technically new, I am hoping you could potentially assist once again with my questions.

I work on scale worms and since they soft bodied and fragile, I wanted to integrate alternative methods for scanning aside from just placing drying them or leaving in ethanol or other liquid. Several papers, including some where you have worked on, have used low-melting agarose to imbed the animals prior to scanning. The agarose percentages I have seen range from 0.5%-1.5% with very few other specifics. I have now tried twice, the most recent being with a 0.5% agarose embedded animal, and the entire pre-scan viewing is opaque or nearly. The animal cannot be seen. Can you think of anything in the embedding process that I am doing wrong that would prevent the X-rays from penetrating the agarose and the specimen? The agarose is prepared in 1% TAE buffer mixed with di-water. 

My only thoughts are that when putting the specimen in the agarose, the warm temperatures are causing the PTA to come out of the animal and disperse among the agarose. Could this be the case or is the agarose maybe wrong brand or age or something else? I would really like to use agarose so that specimens without chaetae don’t move during the long scans.

The only other question I have is that I had a specimen turn from ivory color (in ethanol) to blue/brown after a scan, but only in the portion being scanned. The specimen eventually turned back to the original ivory color upon upon placing in new ethanol. Have you seen this before and is this somewhat normal in liquid mounted specimens or is it an energy issue when running the scan? I have not been able to see any literature or mention of this either.

Sorry for such random questions but would really like to keep going with this technique in order to investigate muscular innervations in swimming scale worms and other annelids. Any help on the issue is greatly appreciated.

Thank you for your time.

Cheers,

Brett

--
Brett C. Gonzalez, PhD.
Postdoctoral Fellow

Smithsonian Institution
National Museum of Natural History

Hi Sarah & Brett,

Random questions are sometimes the best ones. Starting with the colour change: I often see PTA-stained regions change to blue-green-brown under X-rays, then revert after some time to the original whitish. I have assumed that this is caused by an oxidative change in the tungsten. I have not seen any effect on the scan quality. 

PTA can leach out into the agarose (where it can also turn green), but usually does not if the sample has been rinsed after staining. PTA binds strongly to proteins under acidic conditions, and this seems to be permanent if the pH stay low. PTA staining can be removed with a slightly alkaline buffer, or even with PBS, as the PTA polyacid molecule dissociates into smaller tungstate species at higher pH. 

Which brings us to the agarose: I always use agarose in water (usually 0.5%-1.0%), unless I want to keep the tissues in a buffer for some reason. I have seen PTA staining fade in agarose in PBS; iodine staining is OK. I use low-melting temperature agarose, so that it can cool to below 37°C before immersing the specimen (it gels around 33-35°). 

So I imagine the problem is the TAE, which has a pH of 8 or so as I recall, plus a chelating agent (EDTA). My guess is that your agarose effectively dissociated and dispersed the PTA more or less uniformly. Aqueous agarose might solve the problem. 

Another fun trick for embedding fragile samples is to use CyGel, a thermoreversible gel which solidifies at room temperature and melts in the fridge. This avoids the problem of removing agarose from delicate specimens: just wash in cold buffer or ethanol. However, it's really expensive. (http://www.biostatus.com/CyGel/)  

Another way to remove agarose is to drop 6M potassium iodide over the specimen while brushing off the agarose as it dissolves (this helped with a centipede holotype - lots of breakable legs... Akkari et al. 2018) 

Hope this helps. With your permission, I will also post you message and this reply to my blog (http://microtomography.blogspot.com/). 

Best,

Brian 

Akkari N, Ganske A-S, Komerički A, Metscher B. (2018). New avatars for Myriapods: Complete 3D morphology of type specimens transcends conventional species description (Myriapoda, Chilopoda). PLoS ONE 13(7): e0200158. 

https://doi.org/10.1371/journal.pone.0200158

Destaining: PTA

Actually, PTA staining can (mostly) be removed after scanning. I had a student (Hannah Schmidbaur) do some studies on this, and I have done some more experiments. The short answer is to wash out the PTA with a slightly alkaline buffer, e.g. PBS with 0.01M NaOH (figure below; third row). The destaining takes about as long as the staining did (I think), and you should make sure there is enough destaining buffer (at least 10X the volume of the tissue). And of course the only way you can see if the PTA is gone is using X-ray imaging.

Hannah's presentation from the Bruker MicroCT user meeting 2015:
https://www.bruker.com/fileadmin/user_upload/8-PDF-Docs/PreclinicalImaging/microCT/2015/uCT2015-21.pdf

The 2018 Euro Evo Devo meeting!

http://www.evodevo2018.eu/ehome/index.php?eventid=232389

Zeiss/Xradia users meeting!

It's going on now!
Check out the tweets at
https://twitter.com/zeiss_micro

My ToScA talk

Here is a pdf of the slides from my ToScA presentation.

https://www.dropbox.com/s/nxql9qu5tvcky9c/Metscher_ToScA-2017c.pdf?dl=0

and the video collage is now on YouTube at

https://youtu.be/85wocaNifP4

Plant CT

Several people asked about published work on contrast-enhanced microCT for plant specimens. Here is an article from a group in Vienna, and a couple of more recent ones, as well as a couple of pictures I made using vascular contrast agents on wild and domestic cereal plants (details on request). 

Staedler YM, Masson D, Schonenberger J. (2013).
Plant Tissues in 3D via X-Ray Tomography: Simple Contrasting Methods Allow High Resolution Imaging. PLoS ONE 8(9): e75295.
http://www.ncbi.nlm.nih.gov/pubmed/24086499

Saoirse R. Tracy, José Fernández Gómez, Craig J. Sturrock, Zoe A. Wilson and Alison C. Ferguson. 2017. Non-destructive determination of floral staging in cereals using X-ray micro computed tomography (µCT). Plant Methods 13:9
https://doi.org/10.1186/s13007-017-0162-x

David Rousseau†, Thomas Widiez†, Sylvaine Di Tommaso, Hugo Rositi, Jerome Adrien, Eric Maire, Max Langer, Cécile Olivier, Françoise Peyrin and Peter Rogowsky. 2015.
Fast virtual histology using X-ray in-line phase tomography: application to the 3D anatomy of maize developing seeds. Plant Methods 11:55
https://doi.org/10.1186/s13007-015-0098-y

ToScA Workshop (Life Sciences): Microtomography for life sciences research

This workshop is offered twice on Wed. 6 Sept. and will introduce some methods for enhancing x-ray contrast in non-mineralised tissues and techniques for mounting biological samples for microCT imaging, especially embryos and other soft tissues, insects and other invertebrate specimens, and any samples of particular interest to the participants.

We will begin with some principles of x-ray imaging, discuss various types of samples and applications, and then work with your own interesting specimens.

You are invited and encouraged to bring your own samples to the workshop! Fixation and staining can take days, so you might want to prepare your samples ahead of time.

Please also bring pertinent questions, including issues concerning image analysis, publishing, and archiving!

Details of stains etc. are given in the accompanying blog entry. And you may of course email me with questions: brian.metscher@unvie.ac.at

1) General advice about sample preparation for microCT imaging  

Fixation:

The best fixation for microCT is the one that preserves the features you need to see. I have had good results with most of the common fixation procedures, but the properties and effects of the fixative must be taken into account for contrast staining. Usually most relevant are shrinkage, decalcification, removal of lipids or carbohydrates, and protein condensation or precipitation.

Preservation:

Samples can usually be stained effectively after storage in 70% ethanol or in formalin. If you want to use an aqueous stain, transfer the sample back to an aqueous solution; likewise for alcoholic stains. Dry samples are easy (they're dry).

2) Some tips for preparing different sample types

Insects & other arthropods:

I have had good results from alcohol-fixed crawlies by re-fixing them in alcoholic Bouin's (1:1 Bouin's:ethanol/IMS) for a few hours or longer and then dehydrating to ethanol (absolute but not anhydrous, i.e. 96-100%), followed by staining in I2E (below).

Others have made excellent images of critical-point dried insects (better than HMDS; Sombke et al. 2015).

Dry insects can be scanned easily, but the internal anatomy is dodgy. Chitinous structures usually come out beautiful. Pins can be a challenge, but scans can be done with pinned insects.    

Embryos and other squishy samples:  

My favourite fixative for soft stuff is 4F1G (4% formaldehyde and 1% glutaraldehyde in phosphate buffer, or other appropriate buffer, like PBS, or whatever your samples are happy in). The actual concentrations of the two fixatives are not crucial; I usually just add glutaraldehyde to 10% NBF and I'm done. The glut must be high-grade and fresh - otherwise it polymerises and becomes less effective.

PTA and iodine both give good results. Shrinkage can be a problem...   

Little fish and anything that sort of resembles a little fish:  

Mostly the same as embryos, but pay attention to whether you want to see e.g. brain, visceral organs, muscles, bones, teeth, etc. Staining with PMA can allow nice distinction of mineralised tissues and soft tissues; PTA in methanol has given good images of hearing structures (Schulz-Mirbach et al. 2013a, b).

Samples in methanol:

Samples preserved for nucleic acids work are often stored in methanol, typically after aldehyde fixation. These can be stained easily and effectively with  PTA in absolution methanol (van Soldt et al. 2015).

3) Mounting samples for microCT

The sample must be immobilised and held on a vertical rotation axis for the duration of the scan. I often use 0.5-1.0% agarose to embed (not infiltrate) samples in narrow plastic tubes or micropipette tips. Other schemes can work also, and may work better for some kinds of objects: bits of sponge and soda straws have helped on occasion.  Other friends of sample mounting include Legos, Parafilm, UHU Patafix (Blu Tack), and a hot-glue gun.