Blood sampling of hamsters

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The text on this page is taken from an informal compilation of opinions of contributors to the online VOLE List. As such, they are not peer reviewed and may contain differences of opinion. Those wishing to contact the list may contact Adrian Smith.

We recently attempted to refine our hamster blood sampling process. Traditionally we've been using saphenous sampling and decided to try Rodrigues, M. V., et al. (2017) gingival sampling technique described in PLoS ONE (there were also scheduled pharyngeal swabs and nasal washes).

Initially we followed the anaesthetic technique described in the paper (200ml/kg ketamine and 10mg/kg xylazine ip, also described by Flecknell, 2015 and Wolfensohn and Lloyd, 2013) but found the anaesthesia to be hugely variable, with some of them becoming extremely deep with prolonged recovery times over which we struggled to keep them warm and others taking ages to go to sleep, consequently we lost two of the first 16 we anaesthetised.

We attempted acepromazine at 2.5mg/kg ip (Flecknell, 2015) but found it had barely any sedation at all, so we moved to 50mg ketamine + 0.125mg/kg medetomidine ip (a half dose of the one described by Flecknell, 2015, and Wolfensohn and Lloyd, 2013) and found it much more reliable with them much lighter and easier to keep warm on recovery and lost none of the remaining animals.

We certainly improved our recovery period and keeping them warm quickly as we progressed, but the ip route seems intrinsically variable - is there an im approach?

We have used 80mg/kg Ketamine + 10mg/kg Xylazine successfully in hamsters for the last couple of months, both in females and males. Animals lose the righting reflex already, no need for more ketamine. We have also collected oral/nose swabs and blood samples from the saphenous vein. No problems so far with females (naïve and infected). However, in the last couple of weeks we have seen infected males on the way of recovery with varying sensitivity to anaesthesia. And two days ago, males were fine but a naïve male (pre-bleed, similar weights as females) was more deeply affected but it recovered fine (perhaps this time an individual sensitivity?).

I appreciate you need injectables because of your bleeding site. Of course, I'd prefer the use of gas anaesthesia (but we are collecting oral/nose swabs, risk of cross contamination and technical difficulties in containment) or just restraint (but they have strong short legs and Health & Safety infectious risks). I'm revisiting all these, but I suspect infected animals may struggle with either of those options too.