[Nhcoll-l] [EXTERN] Preserving wet specimens in the field?

Thu Mar 27 04:55:50 EDT 2025

Hi Tonya,

Just to add my pennyworth!

Wrapping in cloth soaked in formalin or preservative works well and I used to do this for sending out loans when I worked at the NHM years ago.  With injecting I always twist the needle on and off which can prevent unwelcome squirts if the needle suddenly detaches if some pressure is required to press the plunger. Also try and use some wire in the needle whilst ’sticking it' into the tissue to prevent tissue blockage, then remove the wire prior to twisting the needle onto the syringe.  Dirk’s info on muscle behaviour and David’s of multi-injecting are both good and (I may have missed a point here) always inject the gastric cavity as a priority to prevent autolysis.

With all good wishes, Simon 

Simon Moore MIScT, RSci, FLS, ACR
Conservator of Natural Sciences and Cutlery Historian.

www.natural-history-conservation.com

> On 27 Mar 2025, at 07:22, Stemmer, David (SAM) <David.Stemmer at samuseum.sa.gov.au> wrote:
> 
> OFFICIAL
> 
> Hi Tonya,
> 
> The SA Museum and our Department of Environment developed a Vertebrate Survey Manual a few decades ago when we were conducting regular joint surveys. Some bits may be a bit out of date, but it covers all aspects of going out into the field to catch and collect vertebrate specimens, including preservation. I hope you (and maybe others) find it useful:
> 
> Guidelines for Vertebrate Surveys in SA
> 
> I usually don't inject larger mammal specimens into the muscle, I inject the formalin under the skin. It initially creates a bit of a lump under the skin, but it eventually gets absorbed by the muscle and you won't notice after the fixation period. You still have to be careful with formalin squirting back at you when retracting the needle, so I usually do more small injections rather than a few large ones as it tends to then just trickle out after retracting the needle.
> 
> Cheers,
> David 
> 
> David Stemmer
> Collection Manager, Mammals
> South Australian Museum
> North Terrace, Adelaide SA 5000
> Tel +61 (0)8 8207 7531, Mobile +61 (0)421 754 848
> David.Stemmer at samuseum.sa.gov.au 
> www.samuseum.sa.gov.au <Outlook-wcb0zzbg.png>This email and any attachments may contain confidential information. If you are not the intended recipient any use, disclosure or reproduction of the contents is unauthorised. If you have received this email in error please notify the sender by return email. This email and any attachments should be scanned to detect any viruses and no liability for loss or damage resulting from the use of any attached file is accepted.
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> 
> From: Nhcoll-l <nhcoll-l-bounces at mailman.yale.edu> on behalf of Dirk Neumann <d.neumann at leibniz-lib.de>
> Sent: Thursday, March 27, 2025 5:11 PM
> To: Haff, Tonya (NCMI, Black Mountain) <Tonya.Haff at csiro.au>; nhcoll-l at mailman.yale.edu <nhcoll-l at mailman.yale.edu>
> Subject: Re: [Nhcoll-l] [EXTERN] Preserving wet specimens in the field?
>  Hi Tonya,
> 
> the cautionary remark for injecting muscle tissue was because of the pressure you build up within the tissue. When removing the syringe, the formalin tends to spray out in a high fountain again (depending on the pressure that build up inside the tissue), and because usually you are closer with your head at the specimen when injecting, there is a risk that the formalin sprays into the face or eye. Therefore, it is advisable to keep the syringe more horizontally and not vertically when injecting, and to cover the hole with a finger/thumb, when removing it.  Also, depending on the 'rigidity' of the muscular tissue matrix and the pressure you apply, the injected fluid can disrupt the muscular matrix (tearing of the tissue) or formalin pockets between the skin and the muscle could build up.
> 
> For fixing the brain (and knowing the skull morphology of the specific species in advance), an easy way to reach the brain would be to inject the formalin through the orbit of the eye (not through the eye, but leading the syringe besides the eye through the foramen). But applicability of this option surely differs from species to species.
> 
> As for the wrapping, it depends how you wrap; the specimens should stay moistened but don't need to be soaked (which you usually would not do anyway, because you would loose too much formalin for subsequent fixations - especially, if you are limited with the volume of formalin that you can carry into the field to start with).
> 
> Depending on the temperature, you loose a good proportion of your formaldehyde gas each time you open your container with the wrappings. If more specimens are included in a wrapping (which usually is the case), you would prefer to but the small delicate ones into the middle and shield them with larger ones (so they would be still in the formalin vapour, but not in direct contact with the formalin cloth. 
> 
> The warmer it is, the fast is the damage. Usually, the damage I observed was when working on the Nile at extreme temperatures (i.e. > 40°C). Keeping the wrapped as cool as possible also helps (e.g. in the shade, in well ventilated rooms, etc.).
> 
> Hope there are some useful ideas that help
> 
> Cheers,
> Dirk 
> 
> 
> Am 27.03.2025 um 00:00 schrieb Haff, Tonya (NCMI, Black Mountain):
> Thanks Dirk,
>  
> That is great and useful information, and the book chapters look useful across a range of topics, thank you. Your chapter, albeit on fish, is especially relevant and detailed, and I think will be of great help.
> Reading your email and your chapter, I am relieved as it is way easier to carry formalin to fix in the field and step up back at home than to carry ethanol, and to worry about the timing of stepping up in the field. But to clarify, do you think I don’t need to worry about specimens getting overly fixed/darkened/acidified, if they are simply wrapped in formalin-soaked cheese cloth after fixation in the field, without first being washed?
> While I have your (and others) attention, may I ask a few fixation related questions?  I would love to hear what people have to say about the fixation of vertebrate brains – I am always concerned that the brain is going to deteriorate because of poor/slow formalin penetration past the skull, and it is difficult to inject into (and hard to tell if damage is being done). Separately, I notice that your chapter cautions against injecting formalin into muscle, but I have several manuals that do suggest injecting directly into muscle. I wonder if this is different for fish, or if there is a size at which the need to inject into muscle to avoid decay outweighs the cost?
>  
> Thanks again,
>  
> Tonya
>   ------------------------------------------------------------------------
> Dr Tonya Haff
> Senior Collection Manager
> Australian National Wildlife Collection
> National Research Collections Australia, CSIRO
> Canberra, ACT 2602 Australia
> +61(0)419569109
>  
>  
>  
>  
> From: Nhcoll-l <nhcoll-l-bounces at mailman.yale.edu> On Behalf Of Dirk Neumann
> Sent: Wednesday, 26 March 2025 5:31 PM
> To: nhcoll-l at mailman.yale.edu
> Subject: Re: [Nhcoll-l] [EXTERN] Preserving wet specimens in the field?
>  
> Dear Tonya and all,
>  
> there is a 2 volume book publication on field recording techniques and protocols that is available as free PDF as well; albeit you need to scroll down on the webpage to access the individual chapters directly; maybe this is a useful resource in general: http://www.taxonomy.be/gti_abctaxa/volumes/volume-8-manual-atbi/
>  
> Processing of formalin preserved fish is described in there, including the return and relevant details to be considered if you are transporting your material back in the checked luggage (IATA compliance).
>  
> In general - and depending on the size of the specimens and the temperature (e.g. tropics), you need to be careful to not leave the specimens for too long in the formalin. Especially delicate structures in small fish (e.g. bony pores and canals of the lateral line on the head) can corrode fast in the acidic formalin, leading, e.g. to an open canal system in lamp eye fishes - which are usually closed. Terrestrial vertebrates probably would be injected with formalin.
>  
> Usually (for specimens up to 15-20 cm - again, depending how bulky they are, and how many specimens are in your fixation box), up to three days should be enough. after this, specimens are removed and wrapped in formalin soaked cheese cloth and kept wrapped in tightly sealing PE drums until back home, where you would stage them.
>  
> For carrying formalin, we usually carry more concentrated formalin in the allowed concentration (IATA) into the field and dilute there with clear/clean river water (osmolarity). Depending on where you are, keeping the cooling chain might be an issue, and carrying dry ice or even LN is also not easy and restricted (dangerous goods).
>  
> Hope this helps ...
> With all best wishes
> Dirk
>  
>  
> Am 26.03.2025 um 03:02 schrieb Haff, Tonya (NCMI, Black Mountain):
> Hello all,
>  
> I am wondering if any of you collect vertebrate specimens in remote field locations which are destined for preservation in ethanol. If so, I would love to hear about your workflows in the field.  For example, do you fix your specimens in the field, and if so, do you carry around ethanol to step them out of formalin while you are away, or do you wait until you are back in your institution? And if so, what do you do with fresh specimens in the interim?
>  
> We are interested in collecting more specimens (terrestrial vertebrates) to be preserved in spirit. We keep running up against logistical issues related to trying to avoid freezing specimens and then prepping them when back in the lab. Typically we run trips that can take us into the field for weeks at a time. If we want to fix and preserve more than very tiny things while we are on a field trip, I think it means we must carry around with us quantities of formalin and ethanol that are both potentially hazardous and that take up a lot of space. Alternatively, freezing specimens until we are home seems to me by far the easiest and most efficient thing to do from a logistics point of view, but I know that it’s preferable to preserve specimens immediately after death, and not have an intermediate freezing period.
>  
> I would love to hear any thoughts, suggestions, experiences, recommendations or references anyone may have about this topic.
>  
> Thanks!
>  
> Tonya
>  
>  
>  
> ------------------------------------------------------------------------
> Dr Tonya Haff
> Senior Collection Manager
> Australian National Wildlife Collection
> National Research Collections Australia, CSIRO
> Canberra, ACT 2602 Australia
> +61(0)419569109
>  
> 
> 
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> Stiftung des öffentlichen Rechts;
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> Sitz der Stiftung: Adenauerallee 160 in Bonn
> Vorsitzender des Stiftungsrates: Dr. Michael Wappelhorst
> 
> --
> ****
>  Dirk Neumann
> Collection Manager, Hamburg
>  Postal address:
> Museum of Nature Hamburg
> Leibniz Institute for the Analysis
> of Biodiversity Change
> Dirk Neumann
> Martin-Luther-King-Platz 3
> 20146 Hamburg
> +49 40 238 317 – 628
> d.neumann at leibniz-lib.de
> www.leibniz-lib.de
>  --
> Stiftung Leibniz-Institut zur Analyse des Biodiversitätswandels
> Postanschrift: Adenauerallee 127, 53113 Bonn, Germany
> 
> Stiftung des öffentlichen Rechts;
> Generaldirektion: Prof. Dr. Bernhard Misof (Generaldirektor), Adrian Grüter (Kaufm. Geschäftsführer)
> Sitz der Stiftung: Adenauerallee 160 in Bonn
> Vorsitzender des Stiftungsrates: Dr. Michael Wappelhorst
>  
> --
> Stiftung Leibniz-Institut zur Analyse des Biodiversitätswandels
> Postanschrift: Adenauerallee 127, 53113 Bonn, Germany
> 
> Stiftung des öffentlichen Rechts;
> Generaldirektion: Prof. Dr. Bernhard Misof (Generaldirektor), Adrian Grüter (Kaufm. Geschäftsführer)
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> natural history collections to ensure their continuing value to
> society. See http://www.spnhc.org for membership information.
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