Glowing Green Lizard Heads: Tips and Tricks of Skeletal Preparation

It’s October and that means Halloween is approaching. What says Halloween better than skeletons? They are everywhere this time of year! Therefore, I think that it is the right time to post a few ideas about how to prepare, label, and visualize skeletal material for studies of anole biology. (Make a few yourself and decorate your next Halloween party with them!) If you would like more detailed protocols please email me directly. If you have additional tips and tricks please add them to the comments section below.

Museum collections are filled with dried skeletal material

Dry Skeletons
Dried skeletal preparations are common in most museum collections. Sometimes this marks the fate of a damaged specimen or an animal that perished unexpectedly, but often these have been purposefully built to represent the taxonomic or morphological diversity of a group. Regardless of their origins and use, dried skeletal material makes up a significant portion of our museum collections and great effort should be taken to continue building them with well-prepared material.

Skeletal preparations are often laborious because the skin, organs, and muscles need to be thoroughly removed and the skeleton precisely disarticulated. This can be especially problematic for small and fragile specimens and exceptionally large specimens with large muscles. Rather than contend with these problems and the risk of breaking small bones, I let enzymes or chemicals do the difficult work for me.  A weak solution of potassium hydroxide (0.5 – 3% KOH) tends to work well for both preserved and fresh material. Just skin and eviscerate the specimen and submerge it in ample KOH until the muscle appears clear. At this time the skeleton can be easily disarticulated and laid out to dry under a stream of moving air. Typically a preserved specimen the size of an adult male A. carolinensis can be prepared in about a week with only about 10 – 15 minutes of actual hands-on time (inset image, bottom right). Fresh material can be prepared in as little as 2-3 days. One word of warning, KOH will eventually dissolve all of the soft tissues, leaving only a pile of disarticulated bones for you to examine. Be sure to carefully monitor the progress of your KOH treatments!

Cleared and Stained Skeletal Preps

Cleared and stained A. carolinensis: a mid-stage embryo, juvenile, and adult male

The vertebrate skeleton is made from a combination of bone and cartilage and while adults are made of well-ossified bone, juveniles and embryos are primarily cartilage. Cartilage is  destroyed during desiccation, making dried material of small individuals virtually impossible. Therefore, the best way to visualize skeletal morphology of young specimens is through a process called “clearing and staining” which avoids desiccation by storing the prepared specimen in glycerol.

There are a multitude of clearing and staining protocols available but many are geared towards the nuances of particular groups. Mice, for example, are very fatty and this fat needs to be removed with acetone before clearing and staining can be completed. For most post-hatching anoles I use a protocol similar to Dingerkus and Uhler (1977) which uses alcian blue 8GX to stain cartilage and alizarin red S for stain bone (also see Hanken and Wassersug 1981 and Taylor and Van Dyke 1985 for slightly simplified protocols). For large specimens clearing can be done with KOH, as used above, or something more aggressive such as the digestive enzyme trypsin.

One caveat of these protocols is that alcian blue needs to be dissolved in an acidic solution and acidic solutions rapidly leach calcium from bone leaving nothing for the alizarin stain to bind to. For large specimens this is not a problem, but for embryos undergoing the early stages of osteogenesis even slightly acidic solutions can be problematic. To circumvent this problem I have modified the acid-free protocol of Walker and Kimmel (2007) that was designed for zebrafish embryos and successfully stained embryos as early as Anolis stage 9, shortly after the first cartilage appears. In this one day protocol alcian blue is dissolved at a high concentration using heat and dissolved into a working stock that is applied to the embryos.

Fluorescent Imaging and Fluorescent Labeling of Fresh Material

Skull of an A. cristatellus juvenile photographed with fluorescent light, no stains, labels, or dyes.

The protocols described above are end-point protocols; once you prepare the material there isn’t much more that you can do with it beyond pure morphological examination. But what if you want to use the material to examine gene expression, histology, or another experimental procedure? For each of these it may be necessary to have a record of skeletal morphology prior to the destructive procedure. I use fluorescent imaging of fresh material to circumvent this issue. Collagen has relatively strong autofluorescence due to the cross linking of collagen fibrils. Collagen autofluorescence can be picked up with many standard blue filters equipped to modern fluorescent microscopes. Using this trick I acquire high quality photographs of the skull with virtually no time of preparation allowing me to relate various cellular and molecular parameters to skull shape and size.

There are also several fluorescent calcium chelators that can be used to label skeletal elements in anoles. The three most commonly used labels are calcein (green), alizarin complexone (red), and tetracycline (blue) (but be warned that I had several early experiments ruined by unexpected antibiotics being given to crickets by the vendor). Staggering administration of different labels allows for a real-time measurement of growth rates for different skeletal elements, as is commonly performed in studies of fracture repair. The only caveat to this method is determining the appropriate time between pulses because the skeleton needs grow enough that the labels are distinct but, given enough time, the first signal will be remodeled out of the bone.

Pulse labeling of the anole skeleton with calcium chelators. The distance between green and red lables is the amount of bone growth that occured over 30 days.

This is just a sample of the ways to visualize the anole skeleton. I have not touched on technologies such as CT scanning or other 3D methods even though they are being used in the anole community. It would be great to hear about other people’s experience with photographic tricks and specimen preparation techniques (I once witnessed Steve Poe allow ants to skeletonize a lizard that had died in the field). With that I wish you all a happy October.


Dingerkus, G., and L.D. Uhler. 1977. Enzyme clearing of alcian blue stained whole small vertebrates for demonstration of cartilage. Stain Technol. 52 (4): 229-232.

Hanken, J. & Wassersug, R. 1981. The visible skeleton: a new double-stain technique reveals the nature of the ‘hard’ tissues. Funct. Photogr. 16, 22–26.

Taylor,W. R.&Dyke,G. C.V. 1985. Revised procedures for staining and clearing small fishes and other vertebrates for bone and cartilage study. Cybium 9, 107–119.

Walker, M. B. &Kimmel, C. B. 2007. A two-color acid-free cartilage and bone stain for zebrafish larvae. Biotechnic. Histochem. 82, 23–28.

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2 Responses to Glowing Green Lizard Heads: Tips and Tricks of Skeletal Preparation

  1. Cool! I’ve always wanted to study skeletal structures. 🙂


  2. Pingback: Anolis – now in 3D! |

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