The Journal of Plastination
The official publication of the International Society for Plastination
ISSN 2311 -7761
IN THIS ISSUE:
3-D Reconstruction of the Ethmoidal Arteries of the Medial Orbital Wall Using Biodur® E12 – p 5
Silicone Cast in situ: A Technique to Demonstrate the Arterial Supply of the Female Reproductive Organs of an African Lion (Panthera leo) – p 11
Recycling Histopathology Solvents: A Funding Source for Plastination – p16
Comparative Staining Methods With Room Temperature Plastination (15 - 18ºC) of Brain Specimens, Using Biodur® S10 / S3 – p 21
Establishing a Plastination Laboratory at the College of Veterinary Medicine, University of Basra, Iraq – p 30
Volume 26 (2);i December 2014
The Journal of Plastination
ISSN 2311-7761 The official publication of the International Society for Plastination
Editorial Board:
Renu Dhingra Philip J. Adds New Delhi, India Editor-in-Chief Division of Biomedical Sciences (Anatomy) Geoffrey D. Guttman St. George’s, University of London Fort Worth, TX USA London, UK
M.S.A. Kumar Robert W. Henry North Grafton, MA USA Associate Editor Department of Comparative Medicine Rafael Latorre College of Veterinary Medicine Murcia, Spain Knoxville, Tennessee, USA
Scott Lozanoff Selcuk Tunali Honolulu, HI USA Assistant Editor Department of Anatomy Ameed Raoof. Hacettepe University Faculty of Medicine Ann Arbor, MI USA Ankara, Turkey Mircea-Constantin Sora Vienna, Austria Executive: Carlos Baptista, President Hong Jin Sui Rafael Latorre, Vice-President Dalian, China Selcuk Tunali, Secretary Joshua Lopez, Treasurer Carlos Baptista
Toledo, OH USA
Instructions for Authors
Manuscripts and figures intended for publication in The Journal of Plastination should be sent via e-mail attachment to: [email protected]. Manuscript preparation guidelines are on the last two pages of this issue.
i The Journal of Plastination 26(2):1 (2014)
Journal of Plastination Volume 26 (2); December 2014
Contents
Letter from the President, Carlos. A. C. Baptista 2
Letter from the Editor, Philip J. Adds 4
3-D Reconstruction of the Ethmoidal Arteries of the Medial Orbital Wall Using 5 Biodur® E12, P. J. Adds, A. Al-Rekabi
Silicone Cast in situ: A Technique to Demonstrate the Arterial Supply of the 11 Female Reproductive Organs of an African Lion (Panthera leo), M.J. Hartman, H.B. Groenewald
Recycling Histopathology Solvents: A Funding Source for Plastination, J. 16 Hawkes, M.S.A. Kumar
Comparative Staining Methods With Room Temperature Plastination (15 - 21 18°C) of Brain Specimens, Using BiodurTM S10 / S3, M. S. Mooncey, M. Gill Sagoo
Establishing a Plastination Laboratory at the College of Veterinary Medicine, 30 University of Basra – Iraq, A. A. Sawad, F. S. Al-Asadi
The 18th International Conference on Plastination, Pereira, Colombia 34
Instructions for Authors 35
The Journal of Plastination 26(2):2 (2014)
LETTER FROM THE PRESIDENT Dear Fellow Plastinators: Since our meeting in Saint Petersburg last summer, several changes have taken place within our society and with the Journal of Plastination. These are exciting times for the ISP and the Journal. I would like to take this opportunity to share with you the good news.
The Society membership continues to grow. I would like to extend a warm welcome to our new members. Thank you for joining the ISP. The sources for recruiting new members have been workshops on plastination. Currently we have three workshops available: Murcia, Dalian and Toledo. Each of these three locations covers a distinct geographical area: Europe, Asia and North America, respectively. I am grateful for the knowledge and dedication of Rafael Latorre, Bob Henry and Hong Jin Sui, who provide a constant source of information used Carlos A. C. Baptista, MD, PhD for training in the workshops.
The interest in plastination has been growing rapidly. This is especially true in Brazil and Africa. In Brazil there are at least a dozen plastination labs currently being built. In the past three years I have been invited to minister theoretical courses on plastination in Brazil. The attendance at these courses is growing quickly. Last September I attended the Brazilian Congress of Anatomy in the city of Curitiba, and there were more than 150 participants in a plastination course. For this reason the time has come for a practical course on plastination in Brazil. Professor Athelson Bittencourt, from the Federal University of Espirito Santo, is organizing The 11th Interim Conference on Plastination that will take place in the city of Vitoria from July 13-16, 2015.
In 2016 we will be heading to Colombia for the 18th International Conference on Plastination and our biennial meeting of the ISP. Start now to plan your trip to the city of Pereira, Colombia. The meeting is being organized by Dr. Ricardo Jimenez. It will be an extraordinary meeting. Please refer to page 34 of this issue for an announcement about the meeting.
During the ISP Business meeting in Saint Petersburg, it was suggested that we create a student membership category. To qualify for this type of membership, a student must be sponsored by an ISP member in good standing. Please nominate your student to be a member of the ISP. There is no cost associated with this type of membership.
I am pleased to announce that there were several nominations for the position of Council. The elections will be held in January 2015. The new governance of the
The Journal of Plastination 26(2):3 (2014)
ISP will be composed of the President, Vice-President, immediate Past-President, Secretary, Treasurer and four Council members. I would like to encourage you to become actively involved in our society.
I am delighted to announce that a new design for the Plastination Index is up and running on the ISP website. The Index has been updated thanks to Gilles Grondin who came out of his retirement to help with the task of updating the index. The changes are not yet complete. Soon you will be able to search the database for articles. The Index will serve as a more robust source of information to our readers and authors.
Finally, as mentioned in his Letter from the Editor, Philip Adds has set a target for 2015 for the indexing of the Journal of Plastination in PubMed. This indexation will be an important landmark for our Journal. My hope is that more articles will be submitted for publication because of the journal Impact Factor that will come with the indexing. I am very proud of our journal. If you perform a search in Google Scholar you will notice that all the issues of our journal are available. The Journal of Plastination was the idea of Harmon Bickley (the “father” of the journal) and has been, for almost 30 years, a shining source of information for scientists and plastinators. More importantly, the indexing and impact factor are signs of the commitment of the Journal to maintain quality and its international reputation for excellence.
Happy New Year!
With my warmest regards from Toledo, Ohio, USA
Yours Sincerely
Carlos A. C. Baptista President
The Journal of Plastination 26(2):4 (2014)
LETTER FROM THE EDITOR
Dear Readers,
As you can see, we are continuing the tradition of this Journal in bringing out two issues a year, and it gives me great pleasure to introduce the second issue of 2014. I hope that you will find these very varied papers of interest. In this issue we look back to the 17th ICP in St Petersburg, with papers from presentations that were given there, and we also look forward to 2016 with a preview of the 18th ICP to be held in Pereira, Colombia.
I would like to take this opportunity to thank the members of the editorial board for their vital contribution to the Journal, by acting as reviewers for the
manuscripts that are submitted for publication. Of course, the Journal cannot Philip J. Adds, MSc, FIBMS survive without high-quality papers to publish, and we cannot publish without peer review by suitably qualified and experienced reviewers. Our Editorial Board members work incredibly hard on behalf of the Journal and I would like to thank them on behalf of our readers.
But I would also like to offer an invitation to any of our readers who would also like to be involved in reviewing manuscripts and joining the Editorial Board. If you would be willing to be a reviewer, can you please email me at [email protected], including brief details of your area of expertise? Needless to say, your contribution would be very much appreciated, and you would have the satisfaction of helping to shape the Journal as we head towards the exciting challenges of 2015. With the support of the President of the ISP, Carlos Baptista, I have set the targets for 2015 of getting the Journal of Plastination indexed to PubMed, obtaining its Impact Factor.
We cannot achieve these goals without the continued support of our readers, and without continued submission of your manuscripts. Thank you for your support in the past; I look forward with confidence to the future.
With best wishes for the New Year,
Phil Adds
Editor, The Journal of Plastination
The Journal of Plastination 26(2):5-10 (2014)
ORIGINAL 3-D Reconstruction of the Ethmoidal Arteries of the
ORIGINAL RESEARCHORIGINAL RESEARCH Medial Orbital Wall Using Biodur® E12 ARTICLE
Philip J. Adds ABSTRACT: Objectives Ahmad Al-Rekabi The medial wall of the orbit is reported to contain anterior and posterior ethmoidal foramina, through which pass branches of the ophthalmic artery. These arteries are a potential source of
Division of Biomedical bleeding during surgical procedures involving the medial orbital wall. However, recent research Sciences (Anatomy) has revealed variable numbers of accessory ethmoidal foramina, which have also been shown to St George's, University transmit vascular structures, making intraorbital surgery unpredictable and potentially hazardous. of London This study aims to elucidate the branching pattern of the arterial supply of the medial orbital wall, London, UK particularly in cases of multiple ethmoidal foramina. Materials and Methods Orbits were retrieved from cadavers donated for anatomical examination. Red silicone was injected into the ophthalmic artery via the internal carotid. The medial orbital wall was then
dissected from contiguous craniofacial structures and embedded in Biodur® Epoxy E12 resin.
Sections of 0.3 mm thickness were cut with a slow speed diamond saw, stained with Miller’s stain for elastin and then photographed with a digital camera. Three-dimensional reconstructions were carried out using WinSURF software. Results The optical qualities of the epoxy resin blocks were excellent, though this was not always the case with the individual sections. However, in the stained sections, the arteries were clearly visible. Using WinSURF, the outlines of the branches of the ethmoidal arteries and the bone lining the medial wall of the orbit were delineated. A three-dimensional model of the pattern of arterial branching was created. Conclusion Surgeons operating along the medial wall of the orbit need to be aware that multiple branches of the ethmoidal artery may be encountered. Three-dimensional reconstructions of the branching pattern give a clearer understanding of the blood supply to the medial wall. Work is on-going to map the variations in the branching of the ophthalmic artery. . KEY WORDS: ethmoidal arteries; ethmoid foramen; plastination; epoxy; 3-D reconstruction
Correspondence to: PJ Adds, Biomedical Sciences (Anatomy), St George’s University of London, Cranmer Terrace, London SW17 0RE UK. Tel: +44 (0) 208 725 5208, email [email protected]
Introduction sphenoid posteriorly, and pierced by two foramina (the anterior and posterior ethmoidal foramina) that lie along The medial wall of the orbit and its arteries are of much the fronto-ethmoidal suture line (Fig. 1). The bone of the interest to ophthalmic and oculoplastic surgeons, who ethmoid is particularly thin (0.2 – 0.4 mm in thickness) may need to carry out procedures involving the medial and is consequently very vulnerable to traumatic or orbital wall in cases of trauma, severe epistaxis or iatrogenic fracture. The lamina papyracea so formed thyroid eye disease. It is vital, therefore, that the surgeon makes up the majority of the surface of the medial orbital operating in this region has access to a full and accurate wall (Dutton, 2011). description of the vascular structures that they are likely to encounter. The anterior and posterior ethmoidal foramina are important landmarks during surgical exploration along Conventional wisdom describes the medial orbital wall the medial orbital wall, and their relative positions are as being formed by contributions from the ethmoid, often remembered by the “24-12-6” rule. This describes maxilla and lacrimal bones, with the body of the
6 Adds and Al-Rekabi the distance in millimetres between the anterior lacrimal The ethmoidal foramina transmit important crest and the anterior ethmoidal foramen, from the neurovascular structures. The anterior and posterior anterior ethmoidal foramen to the posterior ethmoidal ethmoidal arteries, branches from the ophthalmic artery, foramen and finally from the posterior ethmoidal foramen pass through the anterior and posterior foramina to the optic canal (Abed et al., 2012). respectively. The ophthalmic artery is the first branch from the internal carotid artery as it enters the middle cranial fossa (Fig.2). The nasociliary nerve, a branch of the ophthalmic division of the trigeminal, gives off anterior and (variably) posterior ethmoidal branches that accompany the arteries through the respective foramina to supply the ethmoidal air cells, anterior cranial cavity and (via the cribriform plate), the nasal cavity (Dutton, 2011).
It is well known that the position, and indeed number, of the ethmoidal foramina is likely to vary between individuals, and recently the degree of variation has been assessed in both Asian and Caucasian orbits. Takahashi et al. (2011) examined 54 Japanese orbits and reported either one or two extra accessory ethmoidal foramina in 18 orbits (33.3 %), while Abed et al. (2012), in a study of 47 Caucasian orbits, reported
variations (including the first report in the literature of Figure 1. Skull showing the medial wall of the orbit. quintuple foramina) in 21 orbits (44%), and furthermore Arrowheads indicate anterior and posterior ethmoidal suggested that the navigational aide memoire should be foramina, * indicates the optic canal. A single accessory revised to “26-17-7”. foramen can be seen between the anterior and posterior foramina (arrow). The question remained to be answered whether these accessory foramina, occurring in nearly half the Caucasian and a third of the Asian population were likely to be of significance. Are these accessory foramina merely defects in the bony wall of the orbit, or do they, like the anterior and posterior foramina, also transmit vascular (and possibly neural) structures? Anecdotal evidence from ENT and ophthalmic surgeons suggests that unexpected bleeding is often encountered, and a histological study carried out by the authors using a dissecting microscope suggests that, not only are there many more accessory foramina that previously suspected (8 were identified in one individual), but they all transmit vascular structures (data not shown). The text-book image of the medial wall, then, is clearly in need of revision. The distribution and branching pattern of the ethmoidal arteries in one-third to one-half of the population is likely to be significantly more complex than previously suspected.
In this study, we aimed to reconstruct the 3-D branching Figure 2. Conventional depiction of the anterior and pattern of the ethmoidal arteries using WinSURF posterior ethmoidal arteries passing through their software, (www.akuaware.com, Kailua, HI, USA), by respective foramina. Adapted from Bartleby.com. embedding the medial orbital wall in Biodur® E12 resin
3-D Reconstruction 7 and cutting thin sections which were then photographed. Dehydration and impregnation The method used was adapted from the standard The dissected specimen was dehydrated by immersion published E12 ultra-thin epoxy technique described by Sora (2007). in 100% acetone (VWR) pre-cooled to -20° and stored in a sealed container in the freezer. The acetone was Materials and Methods replaced daily for three days with fresh 100% pre-cooled acetone, after which the container was brought out of the Specimen preparation freezer and allowed to equilibrate to room temperature before the specimen was transferred to fresh acetone at Pairs of orbits were retrieved from fresh-frozen cadaveric room temperature to dissolve the fat. heads in the Dissecting Room at St George’s School of Medicine, London, UK. The cadavers had all undergone For impregnation, a mixture of fresh Biodur® serological testing and appropriate consent had been E12/E6/E600 was prepared using the ratio by volume of obtained prior to death. Prior to further dissection, the 100/50/0.2, and mixed thoroughly. The dissected orbit inferior (proximal) end of the internal carotid artery was was submerged in the mixed resin and placed in a clamped, and saline was pumped through the superior vacuum oven at 30° C (Heraeus VT 6130 M), where it end using a syringe and cannula to flush the arteries. was allowed to equilibrate overnight. The pressure was Low-viscosity red silicone resin (Biodur® KEM 06) was then lowered (i.e. the vacuum was increased) in stages then injected into the ophthalmic artery via the internal as follows: 40 cm Hg (550 Mbar), 32 cm Hg (425 Mbar), carotid, using a 1 ml syringe with a wide-bore cannula. 24 cm Hg (320 Mbar), 16 cm Hg (210 Mbar) for 45 minutes to 1 hour at each stage, before the temperature The skull section was then immersed in 4% formalin for was raised to 60° C for the final 2 stages: 8 cm Hg (100 one week to fix the tissues before further dissection was Mbar), 2 cm Hg (25 Mbar). The temperature was raised carried out. The specimen was removed from the for the final two stages in order to decrease the viscosity formalin, drained and rinsed under running tap water. of the resin and thereby facilitate penetration of the resin The skin was removed from the lateral aspect of the into the specimen. Using this protocol, vacuum skull, then using a hand-held oscillating saw (de Soutter impregnation can be completed in one day following the Medical), the lateral half of the orbits was removed and overnight incubation. the two orbits were split down the mid-line. The globe, lateral ocular adnexa and periorbital fat were removed Embedding, sectioning and staining by careful dissection to expose the medial extra-ocular muscles (superior oblique and medial rectus) and the The specimen was removed from the vacuum and ophthalmic artery and its branches (Fig. 3). transferred to a mold for embedding. The molds were cut out of blocks of Styrofoam, lined with kitchen foil (Fig. 4); small plastic food containers are also ideal for this purpose. If Styrofoam is used, great care must be taken to avoid overfilling, as spillage of the resin will dissolve the Styrofoam. The same resin mixture that was used for the impregnation stage can be used for embedding; it is not necessary to prepare a fresh mixture since the impregnation stage is so short. The mold was then placed in the oven at 65° C for 5 days to harden. It may be helpful to indicate anterior/posterior and medial/lateral on the mould with a permanent marker before placing in the oven.
Before sectioning, the block can be trimmed if necessary Figure 3. Left orbit section (viewed from lateral side); ica, using the oscillating saw. The trimmed block was then internal carotid artery; oa, ophthalmic artery; on, optic clamped in a Buehler Isomet slow-speed diamond saw, nerve. with a 0.4 x 127 mm blade (Fig. 5), lubricated with “Cool 2” coolant /lubricating fluid (Buehler). Serial sections of 0.3 mm thickness were cut, and the sections were
8 Adds and Al-Rekabi washed with distilled water, dried and placed on a fibre- Visibility of the vascular structures can be improved by optic light source (Schott) and photographed with a staining the sections. Epoxy-embedded thin sections can Nikon D3100 DSLR fitted with a Sigma 105mm F2.8 be stained directly, without the need to re-hydrate the macro lens (Fig. 6). specimens before staining. To enhance the visibility of the sectioned arteries in this study, a modified staining procedure with Miller’s stain for elastin was used:
1. Acidified potassium permanganate 10 mins
2. Rinse in tap water 5 mins
3. Decolorize with oxalic acid 5 mins
4. Wash with distilled water
5. Rinse in 95% alcohol
Figure 4. Molds were cut from blocks of Styrofoam and 6. Stain in Miller’s stain 2 hours lined with plastic foil. 7. Wash in 95% alcohol
8. Counterstain with picro-sirius red 5 mins
9. Blot dry
10. Mount with immersion oil for imaging.
(Modified from: http://www.ihcworld.com.)
Reconstruction was carried out using WinSURF software. Objects of interest (the ophthalmic artery and its ethmoidal branches and the bone of the lamina papyracea) were identified and highlighted on the images of sequential sections.
Results Figure 5. Sections were cut with a Buehler Isomet slow- speed diamond saw. The speed setting was set to 3.0. The specimens were thoroughly impregnated despite the shortened time scale. The optical qualities of the epoxy resin block were excellent (Fig. 7a), though this was not always the case with the individual sections where saw- marks and discolouration affected the appearance of some of the sections (Fig. 7b). It is helpful to use frequent changes of lubricant, since the lubricant bath soon becomes clogged with residue from sawing and leads to sections with poor optical quality. Approximately 30 slices were obtained from each specimen.
Staining with Miller’s stain for elastin enhanced the visibility of the arteries by highlighting the artery walls seen in section (Fig. 8), facilitating the 3-D reconstruction.
Figure 6. Sections were placed on a fibre-optic light Three-dimensional reconstruction was carried out using source and photographed with a Sigma 105mm F2.8 WinSURF 4.2 software (Moody and Lozanoff, 1997). macro lens macro lens mounted on a Nikon DSLR camera. The bone of the medial wall was easily identifiable in the
3-D Reconstruction 9 sections, and the visibility of the artery sections was enhanced by histological staining with Miller’s stain for elastin. A 3-D reconstruction was created, showing the medial wall, the course of the main branch of the ophthalmic artery, and the anterior and posterior ethmoidal arteries (Fig. 9).
Figure 7. a) Embedded specimen being cut, showing the Figure 9. Three-dimensional reconstruction of the medial exceptionally good optical qualities of the epoxy block. wall arteries (left orbit) showing the anterior and posterior The anterior ethmoidal artery can be clearly seen ethmoidal arteries branching off from the ophthalmic branching from the ophthalmic artery and passing through artery and penetrating the wall of the lamina papyracea. the anterior ethmoidal foramen; b) cut section before The anterior ethmoidal artery can be seen continuing staining showing saw marks and discolouration. superiorly and becoming enlarged as it does so.
Discussion
The reconstructed images appeared relatively well defined and bore a close resemblance to the embedded specimens. The reconstructed structures, which included the medial wall and the ophthalmic artery and its branches, can then be displayed individually or as a whole interactive model that can be rotated in 3D space and viewed in the WinSURF viewer (Fig. 9). Additionally, various features such as the transparency, colour, animation and a variety of other options can be adjusted to facilitate the visualization of the complex orbital anatomy. The resulting 3-D rendered images display details that are likely to be of considerable interest to oculoplastic or ophthalmic surgeons, permitting them to view the structures from all angles and to view their spatial relationships.
It is necessary to cut the sections have as thin as Figure 8. Magnified image of section after staining with possible (0.3mm) in order to maximise the amount of modified Miller’s staining technique for elastin. Three information and enable an accurate reconstruction of arteries are shown in transverse section (arrows), plus a such a small specimen. The unavoidable loss of material tortuous length of artery running in the same plane as the from the thickness of the blade (0.4 mm) meant that the section (arrowhead); * indicates the medial rectus muscle; total thickness of each cut was 0.7 mm, so that, red colouration shows collagen within the bone of the medial wall. inevitably, some information was lost with every cut. This disrupted the continuity of the segments, although with reasonable deduction the problem could be solved, and the 3-D reconstructions were found to give a good graphical representation of the branching pattern of the arteries. However, it may be the case that very small 10 Adds and Al-Rekabi branches of the artery can be completely lost between Moody D & Lozanoff S. 1997: SURFdriver: A practical sections. computer program for generating three-dimensional models of anatomical structures using a PowerMac. Clin Although the E12 process is time-consuming, it was Anat 11: 132. found to produce sections that preserved considerable details of blood vessels, soft tissue and bone. The Sora M-C. 2007: Epoxy plastination of biological tissue: structure and spatial relationship of the tissues is not E12 ultra-thin technique. J Int Soc Plastination 22: 40-45 altered in the process. The optical qualities of the Takahashi Y, Kakizaki H, Nakano T. 2011: Accessory embedded blocks are excellent, and even though this ethmoidal foramina: an anatomical study. Ophthal Plast was not always the case for the individual sections, it Reconstr Surg 27: 125-127. was still possible to identify the structures of interest.
Another advantage of the E12 ultra-thin technique is that it does not require decalcification of bone, and histological staining can be carried out directly on the cut sections without the need to take the sections to water beforehand, as is the case in wax-embedded sections.
In conclusion, the E12 method is an effective method for visualising details of small areas of anatomical interest. Of course, in the case of the ethmoidal arteries, where there has been found to be variation in more than 30% of individuals, it will be necessary to carry out multiple reconstructions on a large sample of orbits in order to visualise the variations in the pattern. Now that the embedding and sectioning technique has been refined, work is on-going to build up a catalogue of ethmoidal artery variants.
Acknowledgements
The authors are grateful to the donors and their families, without whose generosity this work would not have been possible. We also gratefully acknowledge the help of Ray Moss and Maria McGlyn, The Image Resource Facility, St George’s, University of London, for their invaluable assistance with the histology.
References
Abed SF, Shams P, Shen S, Adds PJ. Uddin, JM. 2012: A cadaveric study of ethmoidal foramina variation and its surgical significance in Caucasians. Br J Ophthalmol 96: 118-121.
Dutton JJ. 2011: Atlas of clinical and surgical orbital anatomy 2nd ed. Elsevier Saunders p 16-17, 87-88. http://www.bartleby.com/107/illus514.html (accessed 29/07/2014) http://www.ihcworld.com/_protocols/special_stains/miller %27s_elastic_ellis.htm (accessed 29/07/2014)
The Journal of Plastination 26(2):11-15 (2014)
Silicone cast in situ: A technique to demonstrate the arterial supply of the female reproductive organs of an TECHNICAL BRIEF BRIEFTECHNICAL African lion (Panthera leo)
M.J. Hartman1 ABSTRACT: A technique to demonstrate the arterial supply of the female reproductive organs of the African lion (Panthera leo) is described. A 122 kg, one year old nulliparous lioness was used. H.B. Groenewald2 A tin-condensation based, room temperature vulcanization silicone with a durometer shore 30 A
hardness silicone, coloured with red pigment was used to fill the reproductive arterial supply in Department of situ via the abdominal aorta. After the silicone polymer-mix cured, the organs were removed and Anatomy & Physiology macerated yielding a flexible arterial cast of the female reproductive organs. The cast endured University of Pretoria handling well and showed good flexibility. In situ casting has the advantage of minimal leakage Onderstepoort since no vessels are cut in the system to be studied. Republic of South Africa
KEY WORDS: African lion; anatomy; arterial supply;silicone cast
Correspondence to: 1 Department of Companion Animal Clinical Studies, Faculty of Veterinary Science, University of Pretoria, Private Bag X04, Onderstepoort 0110, Republic of South Africa. email [email protected] 2 Department of Anatomy and Physiology, Faculty of Veterinary Science, University of Pretoria, Private Bag X04, Onderstepoort 0110, Republic of South Africa.
Introduction of the human heart, tracheobronchial tree and brain ventricles was done (Aultman et al., 2003). In this study, The surplus of African lions on lion farms in South Africa a process describing four stages was advocated that has created a need for population control. No included harvesting and preparation of the organs, description of the vascular supply of the female African silicone injection, curing and maceration. The sequence lion’s (Panthera leo) reproductive organs was found in in which plastination chemicals are added can improve the literature and its accurate anatomical detail had to be stability of the silicone impregnation mixture at room established. Cast production, to aid anatomical temperature (Glover, 2004) and cold-temperature description, dates to the late fifteenth century. At this techniques have been used commonly over the past 20 time Leonardo da Vinci used wax to inject the ventricles years (De Jong and Henry, 2007). Silicone casting of of the sheep brain (Huard, 1968). Since then various the uterine lumen of a mare has also been reported products have been used to aid in anatomical (Chaurasia and Nayak, 2009) and the preservation of descriptions. The preparation of organ silicone casts to organs using an alkyd resin has been used in various study the trachea and bronchi (Henry, 1992a) and its species (Ari and Çinaroğlu, 2011). associated vasculature (Henry, 1992b) and the chambers of the heart and its great vessels (Henry et al., In order to perform laparoscopic sterilization of the 1998) has been done since 1992. In 1995 a resin cast African lioness (Panthera leo) a sound knowledge of the with E20 red was used to study anatomical cavities morphology and arterial supply of the female (Pretorius and Geyer, 1995) and the comparative use of reproductive tract is required. A morphological study of gelatine and silicone in brain ventricles was done in the splanchnology and topography of the female 2000 (Grondin, 2000). Resin, latex and acrylic casts reproductive organs of the African lion has already been have been illustrated in a Veterinary textbook (König and performed by the authors on three cadaveric specimens Liebich, 2004). (Hartman and Groenewald, 2012) however detailed knowledge of the arterial supply was lacking. In situ Improvements on existing techniques have been vascular silicone casting was chosen for the purpose of evaluated recently. A more comprehensive study to studying the female lion’s reproductive blood supply prior enhance the quality and durability of silicone specimens to performing any surgical procedure on this organ
12 Hartman and Groenewald system. A local company in South Africa was sourced to and ventral to the aorta, the aorta was approached from supply silicone materials, red pigment and equipment the left and was ligated immediately cranial to the renal (Advanced Materials Technology (Pty) Ltd, 2010). arteries in order to secure preservation bilaterally of the origins of the ovarian arteries. The caudal mesenteric Materials and Methods artery was ligated where it was readily accessible in the mesocolon to prevent silicone from being lost to the A one year and one month old captive bred nulliparous arterial supply to the large intestine. No other dissection lioness weighing 122.0 kg was used. She was initially was done in order not to lacerate any blood vessels immobilized with 360 mg of a tiletamine/zolazepam which would allow extravasation of silicone product. The combination (Zoletil®, Virbac, South Africa) administered project was approved by the University of Pretoria via a 3ml remote injection dart (DanInject®, South Animal Use and Care Committee and Research Africa). An 18G intravenous catheter (Jelco®, Smiths Committee (protocol number v038-09). Due to the Medical) was placed in a lateral saphenous vein and an ethical sensitivity of wildlife research the use of larger additional 140 mg of Zoletil was given intravenously. The number specimens was not feasible. lioness was then transported to the Onderstepoort Veterinary Academic Hospital (OVAH) under constant Preparation of the product supervision. Mold Max 30 RTV silicone, durometer shore 30 A Upon arrival at the OVAH a second 14G intravenous hardness, viscosity 25000cps (Advanced Materials catheter (Jelco®, Smiths medical) was placed in one of Technology (Pty) Ltd, 2010, from Smooth-On, USA) the cephalic veins and 300mg of propofol (Propofol 1% which is a tin/condensation cure (rather than Fresenius, Fresenius Kabi, South Africa) given platinum/addition cure) was obtained. Five hundred intravenously. The lioness was intubated and maintained grams of polymer were measured into a 1 kg plastic on isoflurane (Isofor, Safeline Pharmaceuticals Pty Ltd, bucket and 5% by weight silicone thinning fluid was Florida, South Africa). A third 14G intravenous catheter added to decrease the viscosity nearly 50% (13800cps) was paced in the external jugular vein and heparin to enhance injection. To create a red color, 15 ml of red sodium 5000IU/ml (Fresenius 5ml/vial) at a dose of 75 silicone pigment was added and mixed for 5 minutes; Units/kg (1.8ml) injected. 10% (50 grams) parts by weight of MM30 STD catalyst Procedure was then added to the thinned and colored silicone and stirred thoroughly for 5 minutes. Once the catalyst is The jugular vein was used to flush the entire added, curing will begin within 60 minutes. This silicone cardiovascular system through with 3L Ringer’s lactate. preparation was placed into a 5L plastic bucket in After approximately 2L of Ringer’s lactate had run in a preparation for degassing. The bucket with the thinned, metal trocar and cannula was placed in the contralateral coloured, polymer-mix was placed in a vacuum chamber carotid artery by means of a cut down technique. While and a vacuum of 0.86 bar (28mm Hg) applied. As full under anaesthesia the lioness was exsanguinated. vacuum was approached, air bubbles rose to the surface Euthanasia was therefore effected without recovery from from within the polymer. This trapped air expanded the anaesthesia, while the patient was still in plane two of volume of the mix ~ 4X, rising almost to the top of the anaesthesia. bucket and then spontaneously collapsed. Vacuum was released and then immediately applied a second time. An incision was made through the abdominal wall at the During the second vacuum cycle, the silicone did not rise lateral border of the rectus abdominis muscle. Two to the same extent. The silicone preparation was poured additional incisions extending from this paramedian slowly into a 400 ml twin-chamber cartridge from a one incision to the lateral border of the epaxial muscles metre height to further eliminate pre-existing air bubbles. directly caudal to the thirteenth rib as well as cranial to The dual chamber cartridge can be used to mix two the tensor fasciae latae muscle were completed. The different solutions, however in our study the silicone was lateral abdominal wall was reflected dorsally using this already prepared. Two plungers were inserted into the technique bilaterally. Since this was a fresh specimen cartridge without their black plastic seals to allow excess without rigor mortis it was possible to reflect all the air to escape. The seals were inserted as soon as layers of the lateral abdominal wall dorsally in one layer. silicone appeared through the escape ports. The Because the caudal vena cava is situated to the right
Silicone cast in situ 13 cartridge was inserted into the dispensing handgun and The cadaver was stored overnight at 23°C which is ideal the static mixing nozzle was screwed on to the tip of the for curing. Approximately 400 ml of silicone-mix was cartridge (Fig. 1). The static mixing nozzle was filled used. with silicone-mix to expel air from the nozzle. The total silicone preparation time was 35 minutes. Maceration
After 21 hours of curing the silicone in a normal position, the organ system was removed from the cadaver. Ostectomies of the pelvic floor were performed at the lateral borders of the obturator foramina and through the lateral border of the ramus of the ischium to remove the floor of the pelvic cavity and the contents of the pelvic cavity and abdominal reproductive organs. The intra- pelvic arteries including the entire length of bilateral internal pudendal arteries were removed with the pelvic and reproductive viscera. The caudal gluteal artery was severed soon after the internal pudendal artery branched off. After removing the reproductive tract, the tissue block was placed into 10% potassium hydroxide (KOH) in distilled water. After six days of maceration (Fig. 3) the KOH was decanted and replaced with fresh 10% Figure 1. Silicone handgun with twin chamber cartridge KOH. However, this new solution of KOH was made loaded with red pigmented silicone. with tap water instead of distilled water. This new solution was ineffective due to precipitation of the KOH The aorta was transected immediately caudal to the and resulted in the solution becoming milky and ligature that had been applied earlier, and the caudal ineffective. A new KOH solution was constituted using part of the abdominal aorta was secured to the tip of the distilled water. This solution once again was effective dispenser of the silicone gun using two 0 nylon ligatures and showed no signs of precipitation. After two more (Fig. 2). Prepared, thinned and coloured silicone was weeks the KOH was replaced with fresh solution. At this injected by hand pressure until the silicone appeared in stage (three weeks after the start of the maceration an incision made in the plantar aspect of the metatarsus. process) all the tissues were macerated apart from The injection device was left attached to the aorta to tissue around the vulva area which was originally very retain silicone intravascularly during the curing process. bulky.
Figure 3. Maceration process day six. Figure 2. Silicone gun static mixing nozzle secured to the abdominal aorta ready for injection. After another two weeks of maceration (five weeks in total) some gelatine-like tissue was still present in the area of the vulva. However at this stage it was decided
14 Hartman and Groenewald to place the casts in a running water bath to rinse off the Discussion remaining tissue. This had good results; the remaining tissue disintegrated into smaller pieces through the The value of studying the arterial supply to an organ hydrodynamic forces of the running tap water within one system in species of unknown anatomy using silicone hour. In areas with tissue remnants, gentle digital casting in situ prior to surgical exploration provides manipulation was applied and together with running valuable information for the surgeon. water remaining tissue was removed. The casts were We found that silicone can be used to a produce arterial boiled in water for thirty minutes until all tissue remnants casts of accurate anatomic detail similar to other were denatured and removed. The casts were then techniques (König and Liebich, 2004). This is a study placed in 6% hydrogen peroxide for two hours. Some describing the use of a silicone product to study arterial very small remnants had to be gently teased off with supply to an organ system situated in a complex forceps. The casts were left in a running water bath anatomical location as used in other studies (Smodlaka overnight to yield a completely macerated cast of the et al., 2008, 2009) in contrast to the use of loose organs arterial system. The arteries were then allowed to re- (Henry, 1992b; Aultman et al., 2003). arrange themelves in their natural position in a solution of glycerine, formalin and water which allowed the cast The arterial casts were severely entangled by the to be suspended inside a perspex container built to size. running water in the water bath. It was challenging and time consuming to re-arrange the arteries in their natural Results orientation after this process. Leaving the casts in a This process yielded a silicone cast of good quality in running water bath might be regarded as an terms of detail, flexibility and anatomical accuracy (Fig. unnecessary step in the authors’ opinion. However once 4). The casts endured handling well due to good the casts were disentangled the silicone casts tended to flexibility and appeared to provide a true replica of the adapt their own spatial arrangement by virtue of the arterial supply of the female reproductive organs of the memory in the silicone over time. The use of tap water African lion (Panthera leo). in preparation of the KOH solution should be avoided and distilled water is essential. A firmer product such as resin might not result in this complication.
In this study a platinum / addition cure silicone was first used which suffered cure inhibition and did not cure properly inside the arteries. Cure inhibition can result on a variety of surfaces, sulphur being the biggest culprit, and biological tissues being the cause in our study. Tin / condensation cure silicones lose some moisture over time, shrink slightly, dry out and then become easier to tear. This process can take 1 - 6 years depending on the quality of the silicone (MM30 has a typical life of 5 years) (Advanced Materials Technology (Pty) Ltd., 2010) when after this point the silicone becomes more delicate to handle. Platinum / addition cure silicones however do
not lose any moisture so the shrinkage is negligible; this Figure 4. Red pigmented silicone cast depicting the arterial supply to the reproductive organs of the African silicone will have a much longer shelf life once cured. lioness. Aorta (AO), external iliac artery (EI), internal iliac Current reports of platinum silicone moulds over 20 artery (II), median sacral artery (MS), ovarian artery (O), years old exist and they would probably last a lifetime uterine artery (U), caudal gluteal artery (CG), internal (Advanced Materials Technology (Pty) Ltd., 2010). pudendal artery (IP), vaginal artery (V), caudal vesicular artery (CV), urethral artery (UR), caudal rectal artery (CR), This technique provided the ability to fill the entire ventral perineal artery (VP), clitoral artery (C). arterial supply of the female reproductive organs in situ similar to the use of silicone in loose specimens (Aultman et al., 2003). It was possible to depict the spatial arrangement of the arteries of this visceral and
Silicone cast in situ 15 intra-pelvic organ system in its natural position by then S10/S15 technique and products. J Int Soc Plastination allowing the cast to cure inside the carcass. It was 22: 2-14. possible to remove the organ structure from a complex location in the body, complete the maceration process Glover R. 2004: Silicone plastination, room temperature and yield a cast reflecting the arterial anatomy. Care methodology: Basic techniques, applications and has to be taken during dissection of the internal benefits for the interested user. J Int Soc Plastination pudendal artery due to its complex anatomical location. 19: 7. This method prevents leakage of silicone from severed Grondin G, Sianothai A, Orly R. 2000: In situ ventricular vessels should the organs be removed prior to casting, casts of S10 plastinated human brains. J Int Soc and it could possibly be used for other organ Plastination 15: 18-21. structures.The cast was of good quality in terms of detail and anatomical accuracy with the main arteries Hartman MJ, Groenewald HB. 2013: Morphology of the infiltrated. The cast endured handling well and showed female reproductive organs of the African lion (Panthera good flexibility. Silicone can be used in abdominal leo). Acta Zool-Stockholm 94: 437–446. organs in situ to illustrate, study and anatomically describe arterial supply. The arterial supply to the Hartman MJ, Monnet E, Kirberger RM. 2013: reproductive organs of the African lioness is similar to Laparoscopic Sterilization of the African Lioness that of the domestic cat and information obtained from (Panthera leo). Vet Surg 42: 559–564. this silicone study could subsequently be used during laparoscopic ovariectomy and salpingectomy of the Henry RW. 1992a: Silicone tracheobroncial casts. J Int African lioness (Hartman et al., 2013). The results from Soc Plastination 6: 38-40. this technique were also used in a subsequent study to Henry RW. 1992b: Silicone pulmonary vascular casts compare the efficacy of direct observation and trans- with attached tracheobronchial casts. J Int Soc illumination in describing the arterial supply to the Plastination 6: 41-44. reproductive organs of the African lioness. Henry RW, Daniel GB, Reed RB. 1998: Silicone castings Acknowledgements of the chambers of the heart and the great vessels. J Int The authors thank Mr. Paul Carnall of Advanced Soc Plastination 13: 17-19. Materials Technology (Pty) Ltd., Kempton Park, South Huard P. 1968. Leonardo da Vinci. Dessins anatomiques Africa for supplying equipment and technical assistance. (anatomie artistique, descriptive et fonctionelle). Paris: References Roger Dacosta.
Advanced Materials Technology (Pty) Ltd., 2010. Mold König HE, Liebich HG. 2004: Veterinary anatomy of Max 30. Kempton Park, South Africa. domestic mammals. Schattauer, Stuttgart, New York.
Ari HH, Çinaroğlu S. 2011: A new approach to Pretorius WF, Geyer HJ. 1995: The use of E20 red resin preservation of some organs using alkyd resin. Res Vet for casting anatomical cavities. J Int Soc Plastination 9: Sci 90: 16-19. 37.
Aultman A, Blythe J, Sowder H, Trotter R, Raoof A. Smodlaka H, Henry RW, Schumacher J. 2008: 2003: Enhancing the value of organ silicone casts in Macroscopic anatomy of the heart of the ringed seal human gross anatomy education. J Int Soc Plastination (Phoca hispida). Anat Histol Embryol 37: 30-35. 18: 9-13. Smodlaka H, Henry RW, Reed RB. 2009: Macroscopic Chaurasia S, Nayak V. 2009: Silicone corrosion cast of anatomy of the ringed seal [Pusa (phoca) hispida] lower uterus of mare for museum. Vet Pract 10: 179-180. respiratory system Anat Histol Embryol 38: 177-183
De Jong K, Henry RW. 2007: Silicone plastination of biological tissue: Cold-temperature technique - Biodur
The Journal of Plastination 26(2):16-20 (2014)
TECHNICAL BRIEF Recycling Histopathology Solvents: A Funding Source for
TECHNICAL BRIEFTECHNICAL Plastination
ABSTRACT: The plastination process produces large quantities of waste acetone contaminated Jack Hawkes with water and lipids dissolved in the solvent. Disposal creates environmental pollution and makes the plastination process more expensive. Recycling the acetone is the ideal solution. Like M.S.A. Kumar most plastination facilities, our plastination unit incorporates acetone recycling equipment. We describe how we have used the excess capacity of the recycling unit to help our hospital’s Biomedical Sciences histopathology section to recycle their used alcohol and Pro-Par Clearant (a xylene substitute). As
Cummings School of a result, they save some $7,000 a year (that were allocated towards purchasing new reagents), a Veterinary Medicine part of which funds our plastination facility. The recycling plant is environmentally responsible and Tufts University in line with the University’s ‘green’ initiatives. As of early 2014, we have recycled approximately North Grafton, MA 6624 L (1,750 gallons) of solvent for the hospital at $US3.3/L (12.50/gal), for a total income of almost $22,000.
KEY WORDS: acetone; alcohol; histopathology; recycling; solvent
Correspondence to: Dr. Jack Hawkes, Biomedical Sciences Department, Cummings School of Veterinary Medicine at Tufts University, 200 Westboro Rd., North Grafton, Ma. 01536 Tel. +001 508 887 4674; Fax: +001 508 839 8787; E-mail: [email protected]
Introduction have recycled approximately 6624 L (1750 gallons) of solvent for the hospital at $3.3/L (12.50/gal), for a total The Cummings School of Veterinary Medicine’s income of almost $22,000. We typically recycle between plastination program was initiated in 2007, with the twenty and twenty-five gallons of histopathology solvents primary goal of reducing animal usage in our anatomy a month, in addition to processing roughly fifty-five teaching program. The facility includes an acetone gallons of acetone for our lab needs. In this paper, we recycler to distill the acetone waste that is a byproduct of describe the recycling process involved in distilling the plastination process. It was soon realized that the histopathology solvents on the campus and the recycling unit could have wider use for recycling organic monetary benefit. solvents generated by various laboratories on the campus. We describe here a mutually beneficial On the other hand, such a program will not work without agreement with our school hospital’s histopathology a reasonable commitment on the part of both parties section to recycle their used alcohol and Pro-Par toward making the program work. Problems like clearant (a xylene substitute). The process involves contamination will inevitably crop up, and it’s important picking up used solvent from the histopathology to have procedures in place to deal with them. The department, recycling it, and selling it back to them for success of our program is due in large part to the half of the cost of new solvents (including what they institution’s support of recycling and innovation. would pay for shipping). They saved approximately $7,000 in 2013, and half these revenues (from resale to There are important physical limitations to recycling. the histopathology lab) help fund the plastination Most importantly, alcohol is an azeotrope, a liquid whose program. In addition, the university saves the costs that concentration can only be changed to a certain point by would be involved in disposing of used solvents (which distillation, because the liquid and the vapor it produces may equal 50% of the solvent’s purchase price(Taylor have the same composition. In the case of ethanol, for 2014)). Recycling solvents is environmentally friendly, as example, pure water of course boils at 100°C; pure it reduces the consumption of petrochemicals, as well as ethanol boils at 78.3°C; but a mixture of 95.63% ethanol the gasoline that would have been used to transport and 4.37% water (by weight) boils at a lower them (the xylene substitute is produced 1500 km from temperature,78.2°C, so by conventional distillation, our campus). The combination of environmental ethanol can only be recycled to about 95% purity. Higher sensitivity and fiscal responsibility fostered by the purity (100%) alcohol is produced on an industrial scale recycling program has also won recognition from Tufts either through addition of an entraining agent like University as a ‘green’ initiative. As of early 2014, we
Recycling Histopathology Solvents 17 benzene, or by removing the water with a desiccant (molecular sieves).
Also, alcohol is sensitive to contamination by xylene or xylene substitutes. Alcohol which contains even small amounts of xylene or xylene substitutes cannot be completely purified by recycling, because these compounds cannot be completely removed. The mixture of water, alcohol and xylene, for example, forms a complicated azeotropic mixture with each of the three isomers (ortho, meta and para) of xylene (Fele et al., 2000)
Materials and Methods
The Histopathology Process Figure 1. Overview of tissue embedding and staining. Fixed tissue goes into two changes of formalin (upper In order to understand histopathology solvent recycling, left), is dehydrated through six changes of alcohol, and it may be helpful to review the process of tissue embedded in paraffin (bottom left). The resulting block is preparation in the histopathology lab (Fig. 1). The first sliced, slices are mounted on slides, and slices are cleared of paraffin (upper right) and stained (lower right). stages in tissue preparation are similar to the process of Solvent baths marked with an asterisk are recycled. plastination: tissue is first fixed and then dehydrated in alcohol prior to embedding in paraffin. The fixed tissue sample goes through two changes of fresh formalin, then Recycling Regulations increasing concentrations of alcohol through 100%, finally through two changes of xylene substitute before An initial step involved in recycling solvents is to make being embedded in paraffin. The resulting block is sure that the process meets the requirements of sectioned and mounted on a slide. At this point, the regulatory bodies. Anyone contemplating recycling process is reversed (xylene replaced with alcohol) so alcohol, in particular ethanol, should verify (and that the tissue section may be stained with water-based document) the legal requirements of their activities. In dyes. The slide is then cover-slipped. the United States the Bureau of Alcohol, Tobacco and Firearms, a division of the Department of Justice, is the Key points: There are two primary solvents used for appropriate regulatory body. Denatured alcohol is histopathology: alcohol and xylene (or xylene ethanol to which methanol or isopropanol (or both) has substitutes). Alcohol is miscible in water and removes been added to make it unfit for human consumption. water from the tissue specimen. Xylene is miscible in Since these alcohols cannot be removed from ethanol, both alcohol and paraffin, and thus aids paraffin to distillation should not pose a legal problem. Ethanol is infiltrate the tissue. Xylene is not miscible in water. When alcohol which is fit for human consumption, and which water is added to an alcohol-xylene solution (even one requires obtaining a tax stamp for purchase. The ATF containing a minute quantity of xylene), the xylene first agency informed us that distillation of dirty ethanol for causes optical distortion (cloudiness), then comes out of which a tax stamp had been purchased would be solution. If xylene remains in the final stages of the slide- allowable under their rules, since the purpose of making process it can cause stain spotting on the slide, distillation would be simply to clean it and to return it to ruining it. The consequences of solvent contamination its as-purchased state. It goes without saying that liquids are serious, and precautions against it must be taken. that have been distilled in a unit that has been used for plastination solvents are unfit for human consumption. However, anyone contemplating ethanol recycling would be well-advised to get written clarification of the law from the appropriate regulatory body before proceeding.
Companies that engage in commercial solvent recycling are heavily regulated by a variety of agencies. For this 18 Hawkes and Kumar reason we do not recycle any solvents for clients outside product/distillate is discarded. Next xylene is distilled the university system. (primarily to separate xylene from paraffin dissolved in it), and the product/xylene is saved. Since alcohol is very Equipment susceptible to contamination by xylene (and xylene substitutes), for the first two years of operation two A Procycler A (Fig. 2) (B/R Instruments, Easton, MD, batches of acetone were run between Pro-Par and USA), was purchased in 2007 for $13,400 and has been alcohol to ensure that there was no residual Pro-Par in operating daily for seven years, with only a few minor the recycler. A two-column distillation unit, which has a problems (which B/R Instruments have been extremely separate boiler and column for alcohol and xylene, helpful in resolving). prevents this problem.
Discussion
Quality Control
Because some histopath samples (i.e. biopsies) are irreplaceable, it is important to work closely with the clients of recycled solvent, and to discuss how to handle problems with solvent quality before they happen. Quality control of recycled solvents is critical. Even small amounts of Pro-Par in recycled alcohol, for example, can cause spotting on slides when they are stained, ruining the biopsy sample.
Last year Pro-Par contamination of our alcohol was a problem. Pro-Par is soluble in fat. Investigation revealed that the acetone being distilled between Pro-Par and alcohol (to clean the unit) was very high in fat content. The fat residue which remained in the boiler absorbed Pro-Par which then contaminated the alcohol during its recycling period. As a result of this incident, we purchased a separate boiler dedicated to alcohol recycling (at a cost of $2,250), although this step is probably only financially justifiable for larger programs. We also instituted rigorous quality control measures, which we now recommend to anyone contemplating recycling histopathology solvents; more limited quality control measures will probably benefit those who only Figure 2. B/R Instruments Solvent recycler, in an recycle acetone, by monitoring quality and banking explosion-proof enclosure. samples in case of future problems.
When the recycler was purchased, we did not foresee It is important to maintain a daily record of the recycling recycling xylene substitutes, so we purchased a single- activity. We record the history of each batch of solvents column recycler, at a saving of some $5,000. Generally, we recycle. Quality analysis of the recycled solvent is xylene/alcohol recyclers sold have two boilers and two also important. We record the specific gravity of each columns, one for each solvent, while alcohol (or batch of solvent we run, and bank samples of all the acetone) recyclers have only one column. A single solvents we recycle. Every container of recycled solvent column recycler may be used for xylene. However, is marked with its batch number, so if there turns out to since xylene’s boiling point is higher than that of water, be a problem with it, the banked sample can be recycling xylene is a two-stage process with our single qualitatively analyzed to pinpoint the problem. column recycler. First, water and alcohol (which have lower boiling points) are distilled off, and the
Recycling Histopathology Solvents 19 Results percentage of ethanol). It should be noted that the alcohol we distill, as previously noted, has been Specific Gravity denatured. It consists of approximately 85% ethanol (S.G. 0.789), approximately 5% methanol (S.G. 0.792), Our primary tool for analysis is the hydrometer (for approximately 5% isopropanol (S.G. 0.786), and at least alcohol, one calibrated to read in proof). The specific 5% water. For simplicity’s sake, and because we are gravity of every batch of histopath solvents we run (and primarily concerned with consistency from batch to a sampling of the acetone batches we run, as well) is batch, we assume our alcohol to be composed entirely recorded. Ideally, these tests would be run at a of ethanol. Note that if ethanol is denatured with equal standardized temperature (normally 20° C), but because volumes of methanol and isopropanol the resulting it is quite cumbersome to run these tests at standardized specific gravity is the same as for pure ethanol, since temperatures, we have either found tables or charts of isopropanol’s S.G. is 0.003 less than that of ethanol, the solvents we use at various concentrations and while methanol’s is 0.003 greater. temperatures, or have constructed charts of our previous results (to use as a rough guide). Results are thus For acetone, we use a chart (Fig. 3), constructed from obtained considerably more quickly, and possibly more data published by the University of Stirling accurately as well, since results don’t depend on (http://www.thin.stir.ac.uk/tag/drying/), which gives attempting to record a solvent’s specific gravity at the the specific gravities for water contents between 0–11% moment it warms up to the temperature for a hydrometer at three temperatures: 71.6°F, 68°F and 64.4°F. This is calibrated. range covers the temperatures normally found in our The U.S. National Bureau of Standards has published a acetone; for temperatures in between the data points we very complete table, entitled “The True Percents of Proof extrapolate: for example, a hydrometer reading of 0.800 Spirit for any Indicationof the Hydrometer at indicates acetone that is 97% pure at 64.4 F, 96.5% at Temperatures Between 0° and 100° F.” (available at 68F, and 96% at 71.6 F. The acetone from our recycler http://www.ttb.gov/foia/Gauging_Manual_Tables/Ta averages 97% purity. For Pro-Par, we have constructed ble_1.pdf), which allows compensation for any a table of the specific gravities we have recorded at temperature between 0° F and 100° F at any proof from various temperatures in the past; this allows us to 1 to 200 (proof, of course, is simply double the identify any batch which falls outside the norms of what we’ve seen in the past.
% Water in Acetone at Three Ambient Temperatures
0.84
0.83
0.82
0.81
0.8
Specific Gravity sp.gravity at 64.4 F sp. gravity at 68 F sp. gravity at 71.6 F 0.79
0.78
0.77 1 2 3 4 5 6 7 8 9 10 11 % Water in Acetone w/w
Figure 3. Chart of water percent in acetone by specific gravity at three temperatures, constructed from data published by the University of Stirling, Scotland (http://www.thin.stir.ac.uk/tag/drying/). 20 Hawkes and Kumar
Contamination cartridge after each use. We change filters approximately every six months. We carefully check for xylene (or xylene substitute) contamination of every batch of alcohol we recycle. Conclusion Xylene is miscible in alcohol, but not miscible in water. If a few drops of contaminated alcohol are floated on We have found that making use of our recycler’s unused water, the xylene will come out of solution in the alcohol, capacity for recycling solvents for the histopathology and be seen as an oily sheen on the water (this is how department in our school’s hospital has several benefits. we used to test for contamination). At concentrations It converts unused capacity to income, and helps meet lower than those which will force the xylene out of the school’s goals of environmental responsibility. solution, xylene has the effect of disturbing the optical Recycling has taught us a great deal about maximizing qualities of the alcohol/water solution, making it cloudy; the quality of recycling for all our solvents. Finally, in this is a variation of the histological technique for these days of cost-consciousness, the evidence of fiscal checking or determining the purity of alcohol using responsibility provided by this business arrangement xylene (Humason, 1962; Hall, 2001). We have found this helps ensure that we are able to continue our to be the more sensitive visual test. plastination program.
Pro-Par References
Fele, L, Štemberger, N, Grilc, V. 2000: Separation of Finally, there are a couple of tricks we have picked up in water + ethanol + (o-, m-, p-) xylene systems. J Chem recycling the xylene substitute used by our Eng Data 45: 784-791. histopathology lab. Manufactured by Anatech, Ltd. (Battle Creek, MI, USA; http://www.anatechltdusa.com/), Hall, J, 2001: A simple, rapid method for measuring the Pro-Par is a xylene substitute (technically, it is a percentage of water in alcohols used for dehydrating propylene glycol ether) marketed for, among other tissues. Biotech & Histochem 76: 41- 42 things, its ability to be recycled. The distillate of Pro-Par has a very strong and objectionable fishy odor; for this Humason, G, 1962: Animal Tissue Techniques. San reason, the distillate is filtered through activated Francisco: W.H. Freeman, p. 32. charcoal, which removes the odor. Pro-Par attacks many plastics, as well as cured silicone adhesive. For this Taylor, J. 2014: Cost Benefit Analysis furnished by reason we use refillable filter cartridges (Catalog No. James Taylor, B/R Instruments, personal communication 9007T53, McMaster-Carr, NJ, USA), and dispose of the 2/21/14.
The Journal of Plastination 26(2):21-29 (2014)
Comparative Staining Methods With Room Temperature ORIGINAL Plastination (15 - 18ºC) of Brain Specimens, Using RESEARCH BiodurTM S10 / S3 RESEARCHORIGINAL
ABSTRACT: Plastinated brain specimens are easy to handle, long-lasting, and can be used as Mumtaz S. Mooncey effective tools in neuroanatomy teaching. This study compares four different staining methods to differentiate between the gray and the white matter of brain specimens. It further investigates an Mandeep Gill Sagoo alternative and economical method of room temperature plastination of the stained specimens.
St. George’s, A total of 10 formalin-fixed brain specimens were obtained from St. George’s University of London University of London anatomy dissection room. The tissue was used in accordance with the Human Tissue Act (2004). Cranmer Terrace The brain specimens were cut into sagittal or coronal slices, and then stained. The plastination London, UK process involves fixation, dehydration, impregnation and curing. Dehydration was performed at - 30°C, over 3 weeks, using acetone of increasing concentrations. Vacuum impregnation with BiodurTM S10/S3 and curing with S6 were carried out at room temperature (15-18°C). The size of the brain specimens was measured at 3 stages; after staining/before dehydration, after impregnation and after curing. The ability of the stains to withstand the plastination procedure was also assessed.
The Roberts staining method was found to be the most effective because it showed the clearest differentiation between the gray and white matter. The color produced by all staining methods was maintained following completion of plastination. The average shrinkage of the stained brain specimens after plastination was acceptable (12.80% in length, and 9.95% in width), and there was minimal effect on the overall appearance of the specimens. This study further extends the potential applications of the staining method combined with a cost-effective room-temperature plastination method.
KEY WORDS: BiodurTM S10/S3; brain specimens; low cost plastination; room temperature impregnation; staining.
Correspondence to: Mandeep Gill Sagoo, Work address: Biomedical Sciences (Anatomy), St. George’s, University of London, Cranmer Terrace, London, UK. SW17 0RE. Email: [email protected]
Introduction created resources (Weiglein, 1996; Holladay and Hudson, 1989; Cook and Dawson, 1996; Baeres et al., Plastination is a unique technique to prevent decay and 2001; Lozanoff et al., 2003). decomposition of valuable cadaveric specimens. It is a process of removing water and fat from cadaveric In this study, the brain specimens were firstly stained specimens, and replacing them with polymers through and then plastinated. The gray matter largely consists of dehydration, impregnation and curing processes (von neuronal cell bodies, and white matter consists of glial Hagens, 1986). cells, myelinated axons and lipids (Patestas and Gartner, 2006). In staining methods, the cell bodies are Plastinated specimens can be extremely useful teaching expected to take up the stain, leaving the white matter tools, especially in neuroanatomy. Plastination increases unstained. According to the studies by LeMasurier durability and longevity of fragile wet brain specimens. It (1935) and Suriyaprapadilok and Withyachumnarkul has been described in the literature how plastinated (1997), the phenol component of the stain combines with specimens enable students to have a more real hands- lipids of the white matter, therefore forming a protective on learning experience, in comparison to artificially 22 Mooncey and Sagoo film which prevents the white matter from taking up the using a rotary meat slicer. The remaining brain stain. According to Mulligan (1931), the chemicals which specimens had been previously sliced and stored in 50% make up the stain penetrate the grey and white matter ethanol solution for two months. This produced a total of differently. 37 brain slices.
In this study, the following four staining methods were The brain slices were washed in tap water for one hour, used; Mulligan, tannic acid Mulligan, LeMasurier and and then immersed in 50% ethanol for one week with Roberts (Gregg, 1975; LeMasurier, 1935; Roberts and daily agitation, to wash out the embalming chemicals. Hanaway, 1969). This was to enable comparison of This was followed by washing the slices in running tap different staining techniques and to find the optimum water for one hour. method that differentiates between gray and white matter most effectively. The specimens were stained using either Mulligan’s staining method, tannic acid staining method, Plastination is often performed in specialized LeMasurier’s staining method or Roberts’ staining laboratories at low temperatures between -15°C and - method (for protocols, see appendix). 25°C (as described by Tianszhong and Jingren, 1998; de Jong and Henry, 2007). However, in the present For plastination, the materials used were acetone (Anala study, the plastination (forced impregnation) of the R Normapur, VWR), -30°C laboratory freezer, vacuum 3 stained brain specimens was performed at a room pump (rotary vane vacuum pump, 6m /h), polymers S10, TM temperature of +15 to +18°C. The reduction in size S3 and S6 (Biodur Germany), acetonometer, grids for (width and length) of the brain specimens at the separating brain slices, air-tight containers and a following stages: before dehydration, after impregnation, plastination chamber. and after curing, was assessed. The plastination procedure: Moreover, the study assessed the ability of the stains to withstand the plastination procedure. Although staining Dehydration of brain specimens can be effective, previous studies o The stained brain slices were pre-cooled to +4 C in an have shown the color and intensity of the stain have a air-tight container, for 24 hours. This was done to avoid tendency to fade with time (Gregg, 1975). Baeres and temperature shock and potential shrinkage of the brain Møller (2001) have reported that Mulligan staining does o slices, when submerged in pre-cooled acetone at -30 C. not fade on plastination. In this study, we investigated The acetone volume used was 10 times greater than the the ability of Mulligan, tannic acid, LeMasurier and specimens to be dehydrated. The water from the brain Roberts stains to withstand plastination. specimens was replaced with acetone of gradually The authors believe that this technique could extend the increasing concentrations (94-100%), over the course of potential applications of staining and room temperature 3 weeks. The concentration of acetone was monitored plastination, along with the advantage of simplicity of for verification of complete dehydration. Ideally, modified set-up (Sagoo and Adds, 2013). freezers should be used for the dehydration process to prevent risk of explosion; however, in this study, due to Materials and methods lack of resources, air-tight containers were used in standard laboratory freezers during dehydration. The brain specimens were obtained from 10 formalin- Baptista et al. (1992) reported that maintaining acetone fixed cadavers, average age 72.9 years (range 71 to 91 vapor below zero degrees Fahrenheit (-17.78°C) is yrs.). The cadavers were embalmed with 10% formalin, essential to prevent the vapors reaching flammable 10% polyethylene glycol, 5% phenol and 75% ethanol in levels. the Dissecting Room of St. George’s, University of London. Impregnation
Consent for anatomical examination and imaging had The dehydrated specimens were removed from the TM been given under the Human Tissue Act (2004). Out of acetone bath and immersed in pre-cooled Biodur 10 brains, two brains were cut into sagittal slices and one into coronal slices of approximately 1cm thickness,
Comparative Staining Methods 23
S10/S3 (100:1) at -30oC. The silicone level was 2-4 cm were taken to establish the degree of shrinkage. In higher than the level of the specimens to facilitate addition, photographs were taken at each step. A paired impregnation. These specimens were then placed in the T-test (one-tailed) was performed on the lengths and vacuum chamber to equilibrate for 24 hours at room widths of the brain specimens, using Graph Pad Prism 4 temperature (15-18oC). The pressure was then reduced software. gradually over an average period of six days, to replace the acetone in the specimens with silicone polymer. The Staining comparison: pressure was measured using an analogue pressure For staining color comparison, a method for grading the gauge installed with the impregnation chamber, and stained brain specimens was devised (See Grading gradually lowered until the final pressure of 5mmHg was Table 1) to investigate the difference in staining. It was reached. The impregnated specimens were then performed visually and the differentiation between the removed from the polymer and drained on a paper towel gray and white matter in the specimens was graded as for a period of 2 - 3 days. follows: Curing 10 - poor differentiation. It refers to excessive BiodurTM S6 polymer was used for curing. The mixing of a stain in the gray and white matter specimens were cured in a closed chamber where which leads to no or minimal demarcation. BiodurTM S6 was placed in a petri dish to help cross link 11 - moderate differentiation. It refers to staining the polymer chains in the specimens. This was done of the gray matter with a moderate mixing in the over a period of three weeks. white matter. Statistical Analysis: 12 - good differentiation. It refers to staining of Measurements of the specimens were taken with a steel the gray matter with no or minimal mixing to rule (millimeters) at three stages of the plastination clearly demarcate it from the white matter. process: after staining/before dehydration, after impregnation, and after curing. These measurements
Table 1: SCORING TABLE – Table showing the different scoring levels for the colors produced by staining the brain slices.
24 Mooncey and Sagoo
Results ruby-red-stained gray matter and the pink stained white matter. From the slices stained by Mulligan’s method, 2 Staining: out of 7 (29%) were stained effectively; however, relatively, there was a moderate differentiation between The effects of different staining methods on the brain the gray and white matter, which were both different slices were compared (Table 2). The most effective stain shades of blue. The LeMasurier method stained only 1 resulted in relatively clearest differentiation between the out of 8 (12%) of the slices, with relatively poor gray and white matter, and the strongest color intensity. differentiation between the gray and white matter. The Overall, the Roberts method stained 13 out of 18 (72%) tannic acid method did not stain any of the slices in this slices effectively, with good contrast and color intensity. study effectively. There was often a burgundy line (Fig. 1) between the
Table 2: RESULTS – Images of stained specimens at different stages of the plastination process – Before dehydration, after impregnation, and after curing.
Comparative Staining Methods 25
Plastination – measurements: mean shrinkage was 12.80% (length) and 9.95% (width) (P < 0.05) (Table 3). Despite the shrinkage, the Measurements of each specimen were taken before and structural details were preserved after curing and the after these stages of the process: before stain withstood the process of plastination. The average dehydration/after staining, after impregnation, and after impregnation time for the room temperature plastination curing. was approximately six days, and the overall process took approximately 8 weeks. From analysis of the raw data, there was a decrease in the size of the brain specimens after impregnation. The
Table 3: Statistical Analysis before dehydration and after impregnation – Table showing the lengths and widths of brain slices before dehydration and after impregnation, including calculations of the mean, median, standard deviation, and percentage changes.
Widths Lengths Before After Before After dehydration impregnation dehydration impregnation
Mean 8.87 7.98 6.91 6.03
Number of slices 33 33 33 33
Median 6.90 6.40 8.20 6.70
Standard Deviation 4.14 3.79 3.02 2.72
Percentage 9.95 12.80
Discussion In comparison to the low temperature plastination technique (de Jong and Henry, 2007), the room It has been found that 1 cm thick slices tolerate handling temperature plastination method used in this study without framing (Ulfig and Wuttke, 1990). In this study, produced results of a similar standard, with shrinkage Roberts’ method was found to be the best method to levels of approximately 10-12%. This is similar to the differentiate between the gray and white matter. A levels of shrinkage reported by Suriyaprapadilok and demarcating burgundy line was found between the Withyachumnarnkul (1997) for plastination of stained grayand white matter. Although the reason for this line is brain slices (approximately 10%). The overall unclear, it contributed towards the effectiveness of the appearance of the specimens was maintained and the stain. The tannic acid method was found to be ineffective level of shrinkage was acceptable for neuroanatomy and the slices did not pick up the stain. The modified teaching. Furthermore, in wet specimens, the stains Mulligan’s and LeMasurier’s methods were of usually fade unless kept in the dark (Gregg, 1975); intermediate effectiveness. The staining of the brain however, in this study, the plastinated slices retained specimens from these methods was inconsistent, their color and have thus far shown no tendency to fade. resulting in relatively poor differentiation between the gray and white matter. In previous studies at St. Hence, the staining and plastination methods used in George’s, Mulligan stain was found to be effective in this study were found to be capable of producing good staining the slices (Sagoo and Adds, 2013); however, in results with lower financial set-up costs. Adoption of the this study it was not as effective. The inconsistency in technique by more universities and institutions could staining could be due to unknown pathologies that could advance neuroscience teaching, with additional financial have affected the brains, or differences in storage and and practical benefits. sectioning of the specimens. 26 Mooncey and Sagoo
Limitations The other staining methods used in this study were adapted from the initial Mulligan’s method, by varying A number of factors such as variation in the age of the the solutions and timings accordingly. brain specimens and daily room temperature changes may have affected the results of the study. Additionally, studies suggest that if there was any damage to the white matter of the specimens, then the stain would be unable to differentiate between the two types of matter, due to the damaged area of tissue. Thus, good preservation of the brain is very important in order to reveal anatomical details (Roberts and Hanaway, 1969). No initial tests were performed to determine any observer variability for grading the stained tissue. For future studies, piloted and validated grading methods will be used to yield more accurate grading. No measurements were taken to establish the shrinkage of gray matter in comparison to white; however, no obvious distortion was found. This comparative shrinkage Mulligan’s – before dehydration between gray and white matter could be investigated in future experiments.
Acknowledgements
The authors would like to thank Mr. Philip J. Adds for his support and guidance. We are most grateful to those individuals who donated their bodies and brains for research purposes, without whom this study would not have been possible.
Appendix
Staining protocols Mulligan’s – after impregnation Mulligan’s staining method is derived from Tompsett’s modified Mulligan staining procedure (Tompsett, 1956). Tannic acid staining method gives a gray-black color to Mulligan’s method shows effective contrast, by uptake of the grey matter, and produces a long-lasting stain blue color by the gray matter, and no staining of the (adapted from Gregg, 1975). The steps are as follows: white matter (adapted from Tompsett, 1956 and Edwards, 1959). The steps are as follows: 1. Submerge in Mulligan’s solution (as above) for 4 minutes; wash with ice water for 10 seconds. 1. Submerge in Mulligan’s solution (5 g phenol crystals, 0.5 g copper sulfate, 0.125 ml 0.1 N 2. Transfer to 0.4% tannic acid water solution for 1 hydrochloric acid and 100 ml distilled water) at minute, and then wash in running tap water for 1 60° C for 5 minutes; wash with running tap water minute. for 10 seconds. 3. Transfer to 0.08% ferric ammonium sulfate for 2. Transfer to 2% aqueous ferric chloride for 1 10 seconds, and then wash in running tap water minute, and then wash with running tap water for for 1 hour. Store the specimens immersed in tap 2 minutes water until needed.
3. Transfer to 1% potassium ferrocyanide for 4 minutes followed by washing for 1 hour in running tap water.
Comparative Staining Methods 27
3. Transfer to 1% potassium ferrocyanide for 3 minutes followed by 1 hour washing in running tap water. Store the specimens immersed in tap water until needed.
Tannic acid – before dehydration
LeMasurier’s – before dehydration
Tannic acid – after impregnation
LeMasurier’s – after impregnation
Tannic acid – after curing
LeMasurier’s staining method creates a sharp contrast by producing a bright blue colored stain in the gray LeMasurier’s – after curing matter, which is effective and long-lasting (adapted from LeMasurier, 1935). The steps are as follows:
1. Submerge in Mulligan solution (as above) for 2 minutes; wash with ice water for 1 minute. 2. Transfer to 1% ferric chloride for 2 minutes, and then wash in running tap water for 5 minutes. 28 Mooncey and Sagoo
Roberts’ staining method stains the gray matter red- References brown in color, using potassium ferrocyanide (adapted from Roberts, 1969). The steps are as follows: Baeres FMM, Møller M. 2001: Plastination of dissected brain specimens and Mulligan-stained sections of the 1. Submerge in Mulligan solution (as above) for 6 human brain. Eur J Morphol 39: 307-311. minutes; wash with running tap water for 5 minutes. Baeres FMM, Wamberg J, Møller M. 2000: Preparation of plastinated specimens of the human central nervous 2. Transfer to 2% potassium ferrocyanide for 1 system for use in teaching of medical and dental minute, followed by washing in running tap water for 1 hour. Store the specimens in air-tight students, 10th Int Conf Plast, Saint-Etienne, France, container until needed. 2000. Abstract in J Int Soc Plastination 16: 34-35.
Baptista CAC, Bellm P, Plagge MS, Valigosky M. 1992: The use of explosion proof freezers in plastination: Are they really necessary? J Int Soc Plastination 6: 34-37.
Cook P, Dawson B. 1996: Plastination methods used in Auckland, New Zealand, J Int Soc Plastination 10: 32- 33.
De Jong K, Henry RW. 2007: Silicone plastination of biological tissue: cold temperature technique – Biodur™ S10/S15 technique and products. J Int Soc Plastination 22: 2-14.
Robert’s – before dehydration Human Tissue Act 2004. UK: Department of Health, 2004 (11/2004). Available from: URL:
http://www.legislation.gov.uk/ukpga/2004/30/pdfs/ukpga _20040030_en.pdf
Edwards JJ, Edwards MJ. 1959: Medical museum technology. London: Oxford Univ. Press, p 87-88.
Gregg RV. 1975: Tannic acid-iron alum reaction: stain of choice for macroscopic sections of brain to be embedded in plastic. Stain Technol 50: 87-91.
Holladay SD, Hudson, LC. 1989: Use of plastinated Robert’s – after impregnation brains in teaching neuroanatomy at the North Carolina State University, College of Veterinary Medicine. J Int Soc Plastination 3: 15-17.
LeMasurier HE. 1935: Simple method of staining macroscopic brain sections. Arch Neuro Psychiatr 34: 1065-1067.
Lozanoff S, Lozanoff BK, Sora MC, Rosenheimer J, Keep MF, Tregear J, Saland L, Jacobs J, Saiki S, Alverson D. 2003: Anatomy and the access grid: exploiting plastinated brain sections for use in distributed medical education. Anat Rec B New Anat: 270: 30-37.
Robert’s – after curing
Comparative Staining Methods 29
Mulligan JH. 1931: A method of staining the brain for Comparison between Different Staining methods. J Int macroscopic study. J Anat 65: 468-472. Soc Plastination 12: 27-32.
Patestas M, Gartner LP. 2006: A textbook of Tianzhong Z, Jingren L, Kerming Z. 1998: Plastination at neuroanatomy. Oxford, UK: Blackwell Science Ltd., p 7- room temperature. J Int Soc Plastination. 13: 21-25. 399. Tompsett DH. 1956: Anatomical techniques. Edinburgh Roberts M, Hanaway J. 1969: Preparation of brain slices and London: E&S. Livingstone, Ltd. for macroscopic study by the copper sulfate phenol- ferrocyanide technique. Stain Technol 44: 143-146. Ulfig N, Wuttke M. 1990: Plastination of stained sections of the human brain. Anat Anz 170: 309-312. Sagoo MG, Adds PJ. 2013: Low temperature dehydration and room temperature impregnation of brain Von Hagens G. 1986: Heidelberg plastination folder: nd slices using Biodur TM S10/S3. J Int Soc Plastination 25: Collection of all technical leaflets for plastination. 2 ed. 3-8. Heidelberg, Anatomisches Institut, Universität Heidelberg. Suriyaprapadilok L, Withyachumnarnkul B. 1997: Plastination of stained sections of the human brain: Weiglein AH. 1996: Preparing and using S-10 and P-35 brain slices. J Int Soc Plastination 10: 22-25. The Journal of Plastination 26(2):30- 33 (2014)
SHORT ESTABLISHING A PLASTINATION LABORATORY AT THE COMMUNICATION
SHORT COMMUNICATIONSHORT COLLEGE OF VETERINARY MEDICINE, UNIVERSITY OF BASRA, IRAQ
ABSTRACT: Plastination is a laboratory preservation technique applied to specimens so that Alaa A. Sawad they can be used as models in the study of anatomy, for both research and education. This technique reduces the exposure to harmful gases and toxic fumes, and provides fixed, non- Fawzi S. Al-Asadi perishable samples at a low cost. The technique uses simple equipment and laboratory devices that can be obtained locally with low costs. Department of Anatomy College of Veterinary The plastination laboratory in The College of Veterinary Medicine at the University of Basra, Iraq, Medicine, University of was designed and built to use the standard S10 method of plastination. The different ways of Basra, Basra, Iraq financing the laboratory, and the technical equipment and supplies necessary for the establishment of the laboratory will be described with the initial results of this experiment.
KEY WORDS: acetone; freezer; impregnation; specimens
Correspondence to: Dr. Alaa A.Sawad, College of veterinary Medicine, University of Basrah, Basrah, Iraq; Email [email protected]
Introduction The main chemical used in the plastination process is acetone, which is used in large amounts and can be very The plastination technique was created by Dr. Gunther dangerous if not handled correctly. Acetone has a flash von Hagens in Heidelberg, Germany, in 1978 as a point of -18 °C. Cooling acetone in the freezer to -25° C unique way to preserve specimens and tissues. This reduces this potential hazard (Gubbins, 1990). process replaces the fat and water in biological tissues with polymeric materials which are subsequently At The College of Veterinary Medicine at the University hardened, to produce dry, odorless and non-perishable of Basra, plastinated anatomical sections have been specimens (Weiglein, 1997). prepared to aid in teaching anatomy. The plastinated specimens are non-toxic, dry, can be used in every The plastination technique requires careful handling of climate, and demonstrate the exact anatomical details. chemicals to avoid injury from inhaled fumes or contact with the skin. The compounds used have a potential Setting up the laboratory irritating or allergic effect, therefore adequate ventilation, impervious gloves and safety glasses must be used The plastination laboratory was designed according to (Holladay et al., 2001). In recent years, plastination has the Technical Leaflets for Plastination from Heidelberg revolutionized the way students can study anatomy and (Von Hagens, 1986), and with the technical support and has become an important means of preservation of cooperation of the plastination laboratory at the organs (Latorre et al., 2007; Riederer, 2013). University of Zagazig (Egypt). Financial support was obtained from the College of Veterinary Medicine, A plastination laboratory design often involves many University of Basra, to increase the integrity of technical and structural facility problems. This can result laboratories and to encourage new technologies in in economic problems that may delay the possibility of teaching anatomy. obtaining plastinated anatomical specimens in a short period of time (Reina-de la Torre et al., 2004).
The main concern in designing a plastination laboratory is to control the potential danger for fire or explosion.
Setting Up a Plastination Laboratory 31
The design of the plastination laboratory was We started to produce plastinated specimens in October, implemented to initiate the S10 standard method, which 2013, and during this time of operation, we have includes four steps: fixation, dehydration, impregnation succeeded in producing about 30 different plastinated and curing. specimens (Fig. 6).
(1) In fixation 10% formaldehyde solution was used for Conclusion 5-10 days; The plastination laboratory at the College of Veterinary (2) Dehydration was done by acetone (3.5 dollars per Medicine, University of Basra, has produced good litre, made in South Africa) at -5°C, which was changed quality plastinated specimens since October 2013, which 4-6 times according to the size of specimen. To obtain are used as anatomical models for education and for complete dehydration, the volume of the acetone was 5- studying veterinary anatomy. Examples of specimens 10 times larger than the volume of the specimens. An produced are: a dissected ostrich heart, a series of acetonometer was used to make sure the dehydration dissections of buffalo brains, a dissected sheep’s kidney, was complete. the muscles of the eye of an ox, and a dissected fish.
(3) During impregnation, the specimen was placed in In the future, our ambition is to increase the number and liquid silicone resin in a vacuum chamber (Fig. 1). quality of our plastinated specimens. We believe that the Vacuum was applied slowly, the acetone was made to plastination technique facilitates the connection of boil and leave the cells, allowing the polymers to students with anatomy, thus our experience with this penetrate. technique has been positive.
(4) Curing was the final step, in which the specimen was Suggestions for the use of plastination in medical placed in an enclosed chamber containing the curing education would include hiring highly trained technicians agent S6, for 5- 6 days for full hardening (Fig. 2). The (at least one full-time) to ensure the continued plastination laboratory used a modified commercial production of high quality samples and safe handling of freezer for impregnation and dehydration, with the solvents and chemicals. compressor and thermoregulation isolated outside of the freezer. The freezer box joints were treated with rubber silicone to avoid an acetone vapor explosion (Fig. 3). Four small holes for passing the vacuum lines were made in the body of the freezer and connected to the vacuum pump (ALUE ve115 rotary type 220-240V, 50/60 Hz, single phase). The vacuum pump was turned on and off every 3 hours by using electronic timers (Fig. 4). We used stainless steel containers (16 litres capacity) for dehydration and impregnation, with a galvanized metal basket. During forced impregnation, the sample was immersed in a polymer solution (Biodur S10 +S3) and placed in a vacuum chamber.
The plastination laboratory was designed to be near the dissecting room with an area of 24 square meters, containing two freezers, one for dehydration and the other for forced impregnation (Fig. 5). It contains a water Figure 1. Modified vacuum chambers sink and drain, as well as a good ventilation system using exhaust fans with a capacity of 100 cubic meters/hour. All electrical switches in the room are isolated by rubber silicone. There are two storage rooms for chemicals and equipment. 32 Sawad and Al-Asadi
Figure 5. Basra plastination laboratory
Figure 2. Hardening Chamber
Figure 3. Modified commercial freezer
Figure 6. Plastinated specimens produced in the laboratory
Figure 4. Vacuum pump with the rubber tubes and valves
Setting Up a Plastination Laboratory 33
References 2004: Setting up a plastination laboratory at the Faculty of Medicine of the Autonomous University of Barcelona. Gubbins RBG. 1990: Design of a plastination laboratory. Eur J Anat 8: 1-6. J Int Soc Plastination 4:24-27. Riederer BM. 2013: Plastination and its importance in Hernández F, Gil F, López O, Ayala MD, Ramírez teaching anatomy. Critical points for long-term G, Vázquez JM, Arencibia A, Henry RW. 2007: How preservation of human tissue. J Anat 224:1-7. useful is plastination in learning anatomy? J Vet Med Educ 34:172-176. Von Hagens, G. 1986: Heidelberg plastination folder: Collection of technical leaflets for plastination. 2nd Ed. Holladay SD, Blaylock BL, Smith BJ. 2001: Risk factors Heidelberg: Anatomisches Institut 1, Universität associated with plastination: I. Chemical toxicity Heidelberg. considerations. J Int Soc Plastination 16: 9-13. Weiglein AH. 1997: Plastination in the neurosciences. Latorre RM, García-Sanz MP, Moreno M, Reina-De La Acta Anat 158:6-9. Torre F, Rodriguez-Baeza A, Domenech-Mateu JM. 34- The Journal of Plastination 26(2)
The 18th International Conference on Plastination Pereira, Colombia July, 2016
Program Director: Dr. Ricardo Jimenez Fundacion Universitaria Autonoma de las Americas Department of Medical Education Avenida Sur 98 - 56 Pereira, Risaralda 660001 / CO
Email Address: [email protected] Phone Number: 57 311 330 10 16
The Journal of Plastination 26(2):35-37 (2014)
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All manuscripts must be submitted to the Editorial Office Manuscript preparation via the e-mail: [email protected]. If you experience any problems or need further information, Cover Letter please contact Philip J. Adds, [email protected]. The cover letter should include a statement of authorship, notification of conflicts of interest, ethical Authors must have an e-mail address at which they may adherence, and any financial disclosures. be reached. Cover letters may be addressed to the Editor-in-Chief, Journal of Plastination. Necessary Files for Submission Include: Cover letter Manuscript Manuscript (including references and figure legends) The manuscript should consist of subdivisions in the Table(s) (when appropriate) following sequence: Figure(s) (when appropriate) Title Page Copyright Release Form (after acceptance) Abstract with keywords Text Note: The above items should be prepared as separate Introduction files. Each file must contain a file extension (.doc, tif, jpg, Materials and methods eps). Results File formats appropriate for text and table Discussion submissions: Microsoft Word References File formats appropriate for figure submissions: TIFF, Figure Legends JPEG (JPG) and EPS Title Page Categories of submissions: The first page of the manuscript should include: Articles published in Journal of Plastination are grouped Title of paper into general article types (listed below). Final designation Each author’s name of a manuscript’s article type is determined by the Institution from which paper emanated, with city, EDITOR. state, and postal code. Each affiliation should be Original Research – Plastination listed as a separate entity, with a superscript number Original Research – preservation that links it to the individual author. Education Case reports Technical brief notes Review - by invitation only Legacy – institutions and people Correspondence 36 The Journal of Plastination 26(2) (2014)
For example: References to published works, abstracts and books 1 2 1 S. D. HOLLADAY *, B. L. BLAYLOCK and B. J. SMITH must include all that are relevant and necessary to 1Department of Biomedical Sciences and Pathobiology, the manuscript. Virginia Maryland Regional College of Veterinary Citations in the text should be in parentheses and Medicine, Virginia Polytechnic Institute and State listed chronologically; e.g. (Bickley et al., 1981; von University, Blacksburg, VA 24061-0442, USA. Hagens, 1985; Henry and Haynes, 1989) except when 2College of Pharmacy and Health Sciences, University of the authors name is part of a sentence; e.g. "…von Louisiana at Monroe, Monroe, LA 71209, USA. Hagens (1985) reported that…" When references are made to more than one paper by the same author Corresponding Author’s name, address, telephone published in the same year, designate each citation and telefax numbers, and e-mail address. as 1999 a, b, c, etc. For example: Literature cited may only include the publications, *Correspondence to: Dr Shane D. HOLLADAY, Department which are cited in the text. References are to be of Biomedical Sciences and Pathobiology, Virginia listed alphabetically using abbreviated journal names Maryland Regional College of Veterinary Medicine, according to Index Medicus. Page numbers of the Virginia Polytechnic Institute and State University, citation must be included. Blacksburg, VA 24061-0442, USA. Tel.: +001 404 739 Examples of the reference style are as follows: 6403; Fax.: +001 404 739 6492; E-mail: For a journal article: [email protected] Bickley HC, von Hagens G, Townsend FM. 1981: An improved method for preserving of teaching It is the corresponding author’s responsibility to notify the specimens. Arch Pathol Lab Med 105:674-676. Editorial Office of changes of address. Only the For a book section: corresponding author should communicate with the Henry R, Haynes C. 1989: The urinary system. In: Editorial office for matters regarding each manuscript. Henry R, editor. An atlas and guide to the dissection of the pony, 4th ed. Edina, MN: Alpha Editions, p 8- Abstract & Key Words: 17. The abstract should be no longer than 250 words. It For other publications: should contain a description of the objectives, materials Von Hagens G. 1985: Heidelberg plastination folder: and methods, results, and conclusions. The abstract Collection of technical leaflets for plastination. should include a section on technique/technical Heidelberg: Anatomiches Institut 1, Universität development if the paper is significantly technical in Heidelberg, p 16-33. nature. The abstract must be written in complete sentences and be intelligible without reference to the Figure legends rest of the paper. No references should be used in the Legends for all figures should be brief, specific and abstract. not be a substitute listing for the result section, and
appear on a separate page at the end of the On the same page, list, in alphabetical order, five Key manuscript, following the list of references. Words that reflect the content of the manuscript. Consult Legends must be numbered consecutively as they the Medical Subject Headings for appropriate key words. first appear in the textAll symbols or abbreviations Key words should be set in lower case (except for appearing in any figure must be defined in the essential capitals), separated by a semicolon and bolded. legend.
Text Tables The body of the text should be written using American All tables must be cited in the text and have titles. English spelling. Table titles should be complete but brief. Information Where quantities are specified, S.I. units should be used. other than that defining the data should be Equivalent Imperial or U.S. units, if desired, should follow presented as footnotes. in parentheses e.g. 1 Kg (2.2 pounds).
References:
Instructions for Authors 37
Create tables using the table creating and editing Figures should be created, saved and submitted as feature of Microsoft Word. Do not use Excel or either a TIFF, JPEG (JPG) or an EPS file. comparable spreadsheet programs. Line drawings must have a resolution of at least 1200 Each table should be simple and uncomplicated, with dpi, and electronic photographs, scanned images, NO vertical and as few horizontal lines as possible. radiographs, CT and MRI scans must have a Each table is to appear on a separate page and must resolution of at least 300 dpi. include the table title and appropriate column heads. The size of each figure should be at least 8.25 cm / Save each table in a separate word document file and 3.25 inches (one-column width) or 16 cm / 6 inches upload individually, like figures. (two-column width). Do not embed tables within the body of the Magnification must be recorded and have a “scale manuscript. bar” in the photo. Since reproduction of illustrations is costly, authors should limit the number of figures Figures to those which adequately present the findings, and All figures must be cited in the text and must have add to the understanding of the manuscript. legends. Figures that are submitted in color must be published Each figure should be attached as a separate file and in color. Authors are responsible for the costs of any labeled with the appropriate number. color reproductions. Contact the editor for details.