The Journal of Plastination 25(1):1-30 (2013) 9

The Journal of Plastination The official publication of the International Society for Plastination

IN THIS ISSUE: Low-temperature Dehydration and Roomtemperature Impregnation of Brain Slices Using Biodur TM S10/S3 – p 3 P35 Plastination: Experiences with Delayed Impregnation – p 9 Injection Plastination: A Low-Tech, Inexpensive Method for Silicone Preservation of Small Vertebrates – p 13 Demonstration of Systolic and Diastolic Phases of the Cardiac Cycle in a Plastinated Human Heart - p 18 Three Dimensional Reconstruction of a Female Pelvis using Plastinated Cross-sections – p 22

Volume 25 (1); July 2013

The Journal of Plastination 25(1):1-30 (2013)

The Journal of Plastination ISSN 1090-2171 The official publication of the International Society for Plastination

Editorial Board: Renu Dhingra New Delhi, India

Geoffrey D. Guttman Scranton, PA USA

M.S.A. Kumar North Grafton, MA USA Rafael Latorre Mucia, Spain

Scott Lozanoff Honolulu, HI USA

Ameed Raoof. Ann Arbor, MI USA

Mircea-Constantin Sora Vienna, Austria Hong Jin Sui Dalian, China

Instructions for Authors

Carlos A. C. Baptista. M.D., Ph.D., MPH Interim Editor Department of Neuroscience University of Toledo, College of Medicine Toledo, Ohio, USA Robert W. Henry Associate Editor Department of Comparative Medicine College of Veterinary Medicine Knoxville, Tennessee, USA

Selcuk Tunali, MD, Ph.D. Assistant Editor Department of Anatomy Hacettepe University Faculty of Medicine Ankara, Turkey

Philip J. Adds, MSc. FIBMS Assistant Editor Division of Biomedical Sciences (Anatomy) St. George’s, University of London London, UK Executive: Carlos Baptista, President Rafael Latorre, Vice-President Christoph von Horst, Secretary Ameed Raoof, Treasurer

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

The Journal of Plastination 25(1):1-30 (2013)

Journal of Plastination

Volume 25 (1); July 2013

Contents Letter from the President, Carlos. A. C. Baptista

2

Low-temperature Dehydration and Room- temperature Impregnation of Brain Slices Using Biodur TM S10/S3, Mandeep Gill Sagoo, Philip J Adds

3

P35 Plastination: Experiences with Delayed Impregnation, M. Üzel, A.H. Weiglein

9

Injection Plastination: A Low-Tech, Inexpensive Method for Silicone Preservation of Small Vertebrates, Shawnda L. Kumro, Ashton V. Crocker, Randy L. Powell

12

Demonstration of Systolic and Diastolic Phases of the Cardiac Cycle in a Plastinated Human Heart, A. Raoof, L. Marchese, A. Marchese, A. Wischmeyer

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Three Dimensional Reconstruction of a Female Pelvis Using Plastinated Crosssections - Using Plastination for 3D Reconstruction, M. C. Sora, R. Jilavu, P. Matusz

22

17th International Conference on Plastination

28

Instructions for Authors

29

The Journal of Plastination 25(1): 2 (2013) 2

LETTER FROM THE PRESIDENT

Dear Friends and Plastinators: It is with great pleasure that I present to you Volume 25, issue 1 of the Journal of Plastination. I would like to express my appreciation and my thanks to Dr. Ming Zhang for the initial organization of this issue. Issue 2 of the Journal of Plastination is already being organized and it is my hope it will be released in December 2013. One year from now, July 2014, we will be in St Petersburg, Russia for the 17th International Conference on Plastination. Our host, Dimitry Starchik, is organizing a superb scientific program and a spectacular social program as well. I encourage you to mark your calendars and attend this outstanding conference.

Carlos A. C. Baptista, MD, PhD

Once again we did not have an interested party to host the Interim Meeting this year. In 2011, I had the same difficulty in finding someone to host the event. I was delighted to host that meeting and hoped to have another laboratory host the meeting in 2013. The tradition has been that the Interim Meeting is held in United States and is comprised of a workshop and platform sessions. I am making an appeal to those knowledgeable ISP members in the U.S. to genuinely consider the possibility of hosting the next Interim Meeting. These biennial meetings foster collaboration among our membership, in addition to cultivating and supporting our newer constituents. I would be happy to provide direction and supportive planning documents to assist the hosting laboratory. I am delighted to report that the ISP membership is growing fast with several new laboratories sprouting in Brazil, Africa, Asia, Europe and US. There are now several workshops being organized and presented to supply the demand for learning the technique. The workshop in Murcia, organized by Rafael Latorre, has been a beacon of knowledge in plastination for many years. In 2013, Honjin Sui hosted his workshop in Dalian, China, and Carlos Baptista and Robert Henry hosted a workshop in Toledo, Ohio, USA. I find this increased activity exciting and a positive sign of a deeper interest in plastination throughout the world. I look forward to working with you all to continue this effort. Sincerely

Carlos A. C. Baptista President of the International Society for Plastination

The Journal of Plastination 25(1):3-8 (2013)

Mandeep Gill Sagoo* Philip J. Adds Division of Biomedical Sciences (Anatomy) St George's, University of London London, UK

Low-temperature dehydration and room-temperature impregnation of brain slices using Biodur TM S10/S3 ABSTRACT: The standard method for plastination with Biodur TM S10/S3 involves lowtemperature dehydration in a volatile intermediary solvent followed by forced impregnation under vacuum at -15°C. However, some institutions have been reluctant to install low-temperature impregnation equipment because of health and safety and cost considerations. The aim of this study is to investigate a low-budget and simple to set up room temperature plastination procedure to prepare neuroanatomy teaching resources. Previous studies at St George’s, University of London have shown that a low-temperature dehydration/ room temperature impregnation protocol TM for Biodur S10/S3 can produce results comparable, if not equal, to the standard method. Fiftyfour formaldehyde-fixed brain slices were dehydrated in acetone at -30° C and vacuum impregnated at room temperature. Twenty slices were stained with Mulligan’s stain before plastination. The slices were measured before dehydration and after impregnation to monitor shrinkage. Shrinkage was acceptable (6.99% in lengths and 6.19% in widths) in both stained and unstained slices, and did not detract from the appearance of the slices. The stain has thus far not faded on exposure to light. Therefore, this procedure can be used to plastinate brain slices with quality comparable to low temperature plastination, which further extends the potential applications of room-temperature plastination. KEY WORDS: low cost plastination; brain slices; Mulligan staining; teaching resource; TM Biodur S10/S3 *Correspondence to: Mandeep Sagoo, Biomedical Sciences (Anatomy), St. George’s University of London, Cranmer Terrace, Tooting, London, SW170RE Email: [email protected]

Introduction: Plastination, developed by von Hagens in Heidelberg, Germany, is a unique technique of obtaining valuable, dry, odorless and nontoxic biological specimens (von Hagens, 1986; von Hagens et al., 1987). Since 1990, plastinated specimens have been used to teach anatomy and neuroanatomy, and these specimens are a valuable supplement for learning 3-dimensional anatomy, for computer assisted modules and for selfdirected anatomy trails (Ulfig and Wuttke, 1990; Purinton, 1991; Weiglein, 1993; Olry and Grondin, 1994; Côté et al., 1995; Szarvas, Szaras, Groscurth, 1995; Weiglein, 1997a; Baeres, Wamberg, Møller, 2001; Lozanoff et al., 2003). Moreover, these specimens are durable, easy to use and do not drip on students’ handbooks (Holladay and Hudson, 1989). For enhancing neuroanatomy teaching at St. George’s, University of London, we have produced plastinated brain slices using a low-temperature dehydration/ room temperature impregnation method (Adds, 2008) which differs from the standard method (de Jong and Henry,

2007) in that forced impregnation is carried out at room temperature. Mulligan staining is a well-established staining method for gray matter, however the stain soon fades from wet specimens unless kept in the dark (Baeres and Møller, 2001). In this study, we assess the ability of Mulligan’s stain to withstand dehydration and impregnation during plastination, and its subsequent resistance to fading in the cured specimen. The advantage of the method described here is that it is a low budget and simple to set-up room-temperature procedure which produces high quality teaching resources. Materials and methods: The materials used were acetone (Anala R Normapur, VWR), grids for separating brain slices (BiodurTM), -30° C laboratory freezer, air-tight containers, vacuum pump (rotary vane vacuum pump, 6m3/h) and plastination TM chamber, polymers S10, S3 and S6 (Biodur Germany).

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4 Sagoo et al. The brains were obtained from seven formaldehydefixed cadavers (embalmed with 10% formaldehyde, 10% polyethylene glycol, 5% phenol and 75% ethanol) from the Dissecting Room of St. George’s, University of London. Consent for anatomical examination and imaging had been given under the Human Tissue Act (2004). The brains were cut into sections approximately 1cm thick using a rotary meat slicer, giving a total of 54 transverse and coronal slices. The length and width of each slice was measured with a steel ruler before dehydration and after impregnation. Twenty randomly selected slices were stained with Mulligan’s stain prior to plastination. The brain slices were washed in tap water for three hours, and then immersed in 50% ethanol for a week with daily agitation, to wash out the embalming chemicals. This step was repeated twice, using fresh 50% ethanol. This was followed by washing the slices in running tap water for one hour. Dehydration The Mulligan-stained (see below for method) and unstained slices were pre-cooled to +4˚C then submerged in pre-cooled 100% acetone in an air-tight container at -30˚C. The volume of acetone used was 10 times that of the tissue to be dehydrated. The acetone was replaced weekly for three weeks. The concentration of acetone was monitored with an acetonometer until it stabilised at 99% for verification of complete dehydration. Impregnation The dehydrated brain slices were immersed in BiodurTM S10/S3 (100:1). The S10/S3 mixture had been pre-cooled to -30° C. The immersed slices were then placed in the vacuum chamber to equilibrate for 24 hours at room temperature (15 - 18°C). After 24 hours the vacuum pump was started and the pressure was reduced gradually. At every point where a few acetone bubbles were seen on the surface of silicone, it was left at that pressure to equilibrate for 2-3 hrs. Pressure was regulated by adjusting the shutoff and bypass valves to permit stabilization of the pressure at certain levels. The pressure was not lowered if

bubbles were still rising. In this way, the pressure was gradually reduced over a period of approximately 9 days until the final pressure of ~5 mm Hg was reached. The change in pressure was measured with an analogue pressure gauge installed with the impregnation chamber. The pressure in the chamber was lowered by approximately one third per day. Care was taken to ensure that during impregnation the level of silicone was maintained no more than 2-4 cm higher than the level of brain slices in order to facilitate vaporisation of acetone. The brain slices were then allowed to equilibrate for 24 hours before removal. The S10/S3 mixture was stored in the freezer at -30°C when not in use to delay thickening of the mixture and to allow reuse. Curing The slices were removed from the polymer and drained on a paper towel for 2-3 days. The slices were placed in a sealed plastic tank containing BiodurTM S6 which was placed in a glass Petri dish at room temperature for 2-3 weeks to crosslink the polymer chains. The S6 was replenished as necessary. Mulligan staining The staining method was derived from Tompsett’s modified Mulligan-staining procedure (Tompsett, 1956). In this procedure the timing is critical as the white matter tends to picks up the stain. 1. Submerge in Mulligan solution (5 g phenol crystals, 0.5 g copper sulphate, 0.125 ml 0.1 N hydrochloric acid and 100 ml distilled water) at 60˚C for 5 minutes; wash with running tap water for 10 seconds. 2. Transfer to 2% aqueous ferric chloride for 1 minute, and then wash in running water for 2 minutes. 3. Transfer to 1% potassium ferrocyanide for 4 minutes followed by overnight washing in running water. Statistical Analysis: A paired T-test (one-tailed) was performed on the lengths and widths of 54 brain slices before dehydration and after impregnation using Graph Pad Prism 4 software.

Brain Impregnation 5 Results: The mean shrinkage was 6.99 % (length) and 6.19% (width) (P .01 (ns) > .01 (ns)

Percentage % 6.99 6.19

Shrinkage was approximately 7% from a combination of low temperature dehydration and room temperature impregnation, with and without stain, which was around 3% less than Suriyaprapadilok and Withyachumnarnkul (1997) report for low temperature impregnation of brain slices. Therefore, it demonstrates that, contrary to expectations, S10/S3 impregnation of brain material can be accomplished satisfactorily at room temperature. In the current economic climate, this protocol has the added advantage of reduced capital costs; however, it is vital to use air-tight containers in the freezer to avoid the risk of explosion. It is necessary to maintain acetone vapor below zero degrees Fahrenheit (-18° C) to prevent the vapors reaching flammable level (Baptista, Bellm, Plagge, Valigosky, 1992). To conclude, this report suggests an alternative, achievable and relatively low-cost method of plastination of neuroanatomy specimens. Work is ongoing to investigate further applications of this approach.

6 Sagoo et al. Limitations: Shrinkage after dehydration was not observed to analyze the effect of dehydration. Moreover, no measurements were taken to establish the shrinkage of grey matter in comparison to white; however, no obvious distortion was found. Acknowledgements: The authors would like to thank Ms. Lynda Jane Phillipson for her help in staining the brain slices.

Figure 3: Horizontal, Mulligan-stained section of the cerebrum. The basal ganglia are clearly seen, together with the internal capsule and caudatolenticular bridges of gray matter. Figure 1: Horizontal, plastinated section of unstained cerebellum showing dentate nuclei

Figure 2: Horizontal, plastinated section of unstained cerebrum showing caudate nuclei, internal capsules and lentiform nucleus

Figure 4: Horizontal, Mulligan-stained section of the cerebellum and rostral part of the pons. The dentate nucleus and the fibrae pontis transversae are seen

Brain Impregnation 7

Figure 7: Post-plastination: horizontal, Mulligan stained and plastinated section of cerebellum showing dentate nuclei . The staining is unaffected by the plastination process.

Figure 5: Horizontal, Mulligan stained section of cerebellum and midbrain showing substantia nigra.

References:

Figure 6: Pre-plastination: horizontal, Mulligan stained section of cerebellum showing dentate nuclei. (Staining timing and the quality of the specimen is crucial to prevent the color leaking into the white matter.)

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Adds PJ. 2008: A low-temperature dehydration/ room-temperature impregnation protocol for brain tissue using Biodur S10/S3. J Int Soc Plastination 23:41.

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Baeres FMM and Møller M. 2001: Plastination of dissected brain specimens and Mulligan-stained sections of the human brain. Eur J Morphol, 39 (5):307-311.

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Baeres FMM, Wamberg J, Møller M. 2001: Preparation of Plastinated Specimens of the Human Central Nervous System for Use in Teaching of Medical and Dental Students. 10th Int Conf Plast, Saint-Etienne, France, 2000. Abstract in J Int Soc Plastination 16: 34-35.

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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 (1): 34-37.

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Côté ME, Veilleux F, Christin MJ, Fortin MJ, Olry R. 1995: Plastination: a new approach to the teaching of topographical anatomy. Chiropractic Centennial Foundation, Washington, DC, USA.

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de Jong K and Henry RW. 2007: Silicone plastination of biological tissue: cold temperature technique – Biodur™ S10/S15 technique and products. J Int Soc Plastination 22:2-14.

8 Sagoo et al. •

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Holladay SD, Hudson LC. 1989: Use of plastinated brains in teaching neuroanatomy at the North Carolina State University, College of Veterinary Medicine. J Int Soc Plastination 3 (1): 15-17. 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 270B (1):30-37. Olry R and Grondin G. 1994: Plastination in chiropractic teaching: critical analysis and place of plastinated specimens in anatomical pedagogics, 7th International Conference on Plastination, Graz, Austria. J Int Soc Plastination 1995, 9(1): 21. Purinton PT. 1991: Plastinated brains used with computer assisted learning modules for teaching veterinary neuroanatomy laboratories. J Int Soc Plastination 5 (1): 16-19. Suriyaprapadilok L and Withyachumnarnkul B. 1997: Plastination of stained sections of the human brain: comparison between different staining methods. J Int Soc Plastination 12 (1): 27-32. Szarvas B, Szaras L, Groscurth P. 1995: Use of plastinated brain sections for medical education. J

Int Soc Plastination 9 (1): 23-24. •

Tompsett DH. 1956: Anatomical Techniques, Edinburgh and London: E.&S. Livingstone Ltd., p 225-230.

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Ulfig N. 1990: Staining of human fetal and adult brain slices combined with subsequent plastination. J Int Soc Plastination 4 (1): 33-38.

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Ulfig N, Wuttke M. 1990: Plastination of stained sections of the human brain. Anat Anz 170 (5): 309-312.

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von Hagens G. 1986: Heidelberg plastination folder. Collection of all technical leaflets for nd plastination. 2 ed. Heidelberg, Anatomische Institut, Universitat, Hiedelberg .

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von Hagens G, Tiedemann K and Kriz W. 1987: The current potential of plastination. Anat Embryol 175:411-421.

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Weiglein A. 1993: Plastinated brain-specimens in the anatomical curriculum at Graz University. J Int Soc Plastination 7 (1): 3-7 . Weiglein AH. 1997a: Plastination in the neurosciences. Acta Anat 158: 6-9

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The Journal of Plastination 25(1): 9-11 (2013)

M. Üzel* Department of Anatomy, Cerrahpasa Medical Faculty, Istanbul University Turkey A.H. Weiglein Institute of Anatomy Medical University of Graz Graz, Austria

P35 Plastination: Experiences with delayed impregnation ABSTRACT: During an educational demonstration of the P35 technique, brain slices which had been immersed in P35 resin and stored in a cold room (5° Celsius) for approximately two years were used. The resin was very viscous and it was difficult to remove the steel basket containing the brain slices from the container of resin. There were technical difficulties during the manipulation of the slices: slices were brittle and fragile, filter paper spacers were stuck to the specimens, curing had begun where the slice touched the grid and gel-like resin remnants were stuck on the metal grids. Despite the very long immersion period and the problems encountered, the final specimen was satisfactory from an optical point of view.

KEY WORDS: plastination; P35; second immersion; impregnation *Correspondence to: M. Uzel E-mail: [email protected]

Introduction: The well-known classic P35 method is the most used basic technique for brain slice plastination. It is used to obtain semitransparent brain slices, and yields excellent gray-white matter distinction. Its main steps include: fixation with formaldehyde, slicing and placing on stainless steel grids, flushing & precooling to +5°C, two dehydration baths of 2-4 days, two immersion baths (1 day each), forced impregnation, casting in double glass chambers, light curing, heat curing, and finishing (Weiglein, 1996; Weber et al., 2007). By completing all of these steps, the final product is a beautiful, durable brain slice helpful for studying sectional anatomy of the brain. In this case report, the optical quality of the end product, as well as the technical experience after a very long duration (two years instead of one day) in the second immersion bath of the P35 method are shared. Technical case report During an educational demonstration of the P35 technique at the Institute of Anatomy, Medical University of Graz, Graz, Austria, the brain slices that were to be used were placed in the second immersion bath (P35/A9 mixture) approximately two years previously and were stored in the immersion bath during this period at +5°C. The normal methodology would be to commence impregnation after the slices have been in the second

bath for 1 day. Therefore, it was logical to continue the plastination process with 24 hours of forced impregnation of the brain slices which were submerged in this two-year-old resin-hardener (P35-A9) mixture. After the vacuum/impregnation was completed, the slices were removed from the basket/polymer and positioned on the glass, and a double glass chamber was constructed. The chambers were filled with fresh resin mix (P35/A9), the slices were positioned with a wire, and the slices were then cured using UV-A light and heat. Observations Problems were observed from the beginning. After storage and vacuum application, the resin from the immersion bath was too viscous and it was very difficult to remove the steel basket of impregnated slices from the resin. After a struggle, the basket was removed. The slices were rigid, fragile, brittle and difficult to handle (unfortunately some of the slices broke into pieces) (Fig. 1). The filter papers between the slices were often united with the slices. With a lot of effort, most of the papers were removed from the slices. Portions of the slices had started to cure, and gel-like resin remnants were stuck to the metal grids (Fig. 2). Also, the grids left marks on some of the specimens (Fig. 3). Because of their fragility and the increased viscosity of the resin, it was difficult to put the slices onto the glass for construction of the double glass chambers without

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10 Üzel et al. breaking them. After assembling the double glass chambers, the chambers were filled with fresh P35/A9 mixture and the slice position was adjusted with a wire. The hardening procedure was the usual UV-A light-heat combination. Some finished specimens had excavations and traces of filter paper on them. Despite the very long immersion period (two years) and the problems encountered, the final specimens were optically satisfactory (Fig. 4). Discussion The duration of the second immersion step of the P35 method is normally one day. In this case, the specimens were forgotten and unintentionally remained in the second immersion bath (in the cold room) for two years. Because of this very long second immersion period, problems were encountered during the remaining P35 procedure. The problems (i.e. brittle, fragile, and partially cured specimens and gel-like resin sticking on the metal grids) encountered during the handling of the specimens were probably due to the very long (two years) exposure time of the specimens to acetone (von Hagens, 1986) and the increased resin viscosity. The acetone or length of time of the resin and catalyst being mixed likely caused a reaction to change the polymer into a gel. In conclusion, the time for production of brain slices with the P35 method can be extended up to several months by storing the slices in a cold immersion bath. However, too long a duration should be avoided because the slices start to cure at the regions where they are directly in contact with the steel grid, and viscosity of the resin mix increases markedly. These findings suggest that in the P35 method, brain slices can stay in the second immersion bath for months without having any decrease in optical quality of the final specimen. On the other hand, changes in the physical properties of the resin should be expected which will result in mechanical problems that plastinators should be prepared to deal with.

Figure 1: P35 broken impregnated brain slices.

Figure 2: Remnants of (P35/A9) resin mix on the metal grids.

Figure 3: Cured P35 brain slice.

Delayed impregnation 11

References • von Hagens G, 1985: Heidelberg Plastination Folder, 2nd English Edition: Collection of all technical leaflets for plastination, pg. 3:9-10. Biodur Products, D-69126 Heidelberg, Germany . • Weiglein AH, 1996: Preparing and using S-10 and P-35 brain slices. J Int Soc Plastination 10(1): 22-25. • Weber W, A Weiglein, R Latorre, RW Henry, 2007: Polyester plastination of biological tissue: P35 technique, J Int Soc Plastination 22: 50-58.

Figure 4: Grid marks on finished specimen.

The Journal of Plastination 25(1): 12-17 (2013)

Shawnda L. Kumro Ashton V. Crocker Randy L. Powell* Department of Biological & Health Sciences, Texas A & M University Texas, USA

Injection Plastination: A Low-Tech, Inexpensive Method for Silicone Preservation of Small Vertebrates ABSTRACT: The plastination process using vacuum impregnation replaces tissue fluids with curable polymers and results in dry, non-toxic specimens. We detail a method to produce high quality plastinated specimens using an injection impregnation process. This alternate, nonvacuum method is very low-tech and has minimal start-up costs. We were able to successfully use this technique on small vertebrates ranging from 1 to 700 grams. The plastinated specimens were life-like and the natural contours of the animals were maintained. Dissection revealed polymer had penetrated throughout the viscera and deep muscles. In addition, internal morphology including major muscle groups retained their shape with no apparent shrinkage.

KEY WORDS: alternate process; inexpensive; injection; plastination; low-tech; small vertebrates *Correspondence to: Randy Powell E-mail: [email protected]

Introduction: Plastination is a multi-step process in which tissue fluids are replaced by curable polymers; e.g., silicone, epoxy, polyester (Bickley et al., 1981; Pashaei, 2010). The results are dry, durable specimens (entire/partial body or individual organs) that look more life-like. Because plastinated specimens are non-toxic, they can be freely handled and examined without gloves. Consequently, the incorporation of plastinated specimens for teaching and research purposes has been adopted by medical and veterinary schools worldwide (Dawson et al., 1990; Cook, 1996; Mansor, 1996; Correia et al., 1998; Peris, 2000; Zhong et al., 2000). The fundamentals of the plastination process were originally developed and documented by Gunther von Hagens over 30 years ago (von Hagens et al., 1987; see Pashaei, 2010 for a more detailed history). While the basic principles of plastination have remained somewhat consistent since its initial introduction (i.e., fixation, dehydration, forced vacuum impregnation, and curing), there have been numerous developments, alternate techniques, and improvements in the process. These developments include: new polymers that permit ambient (room) temperature plastination (Tianzhong et al., 1998; Henry, 2007; Raoof et al., 2007), progress in sheet and ultra thin sheet plastination (Latorre et al., 2004; Sora et al., 2004; Latorre and Henry, 2007; Sora, 2007), staining and re-coloring of plastinated tissue (Riepertinger and Heuckendorf, 1993; Suriyaprapadilok

and Withyachumnarnkul, 1997; An and Zhang, 1999; Mendez, 2008; Steinke et al., 2008a), and methods to produce light-weight plastinated specimens (Henry and Nel, 1993; Steinke et al., 2008b). Integrating plastinated specimens into a teaching environment has been shown to provide positive educational outcomes in the classroom (Latorre et al., 2011). The characteristics of plastinated specimens (life-like feel, no odor, no toxicity) allow students the benefit of greater tactile interaction with the specimens which may result in an increased time spent with the material. Moreover, incorporation of plastinated specimens in teaching labs has been shown as an excellent method to improve the “hands-on experience” (see Dawson et al., 1990, for a student survey of responses to plastinated specimens). Similarly, the use of plastinated specimens for exhibition and display in museums has been demonstrated as equally valuable. Unfortunately, start-up costs for the production of plastinated specimens using traditional methods may be prohibitive for small labs and institutions with limited budgets and space. Furthermore, traditional plastination methods require special safety consideration and equipment, such as explosion hazards and adequate exhaust venting (Henry and Nel, 1993). Plastinated specimens (entire/partial body forms and individual organs) are available for purchase from a few companies; however, availability is limited to a small number of species, and the specimens can be quite expensive.

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Injection Plastination 13 Vacuum impregnation, which allows for the penetration and saturation of curable polymers into tissue, has been the hallmark step in plastination. While injection (i.e., into deep tissues on large specimens or into specialized structures) has been suggested to improve silicone perfusion during vacuum impregnation (Henry and Nel, 1993; Sivrev et al., 1997), it has not been reported for use exclusively as a mechanism for polymer introduction. Although large organisms may not be good candidates, smaller specimens, with less tissue mass, can be thoroughly saturated with polymer using injection alone. Plastination of small organisms has received little attention, and scant information has been reported (see Asadi and Mahmodzadeh, 2004). Heretofore, no one has reported on injection impregnation for small, wholebody specimens. Our goals were to produce high quality plastinated specimens using an alternate, non-vacuum impregnation process, and to minimize start-up costs. Materials and methods: Fixation: Specimens were fixed in 10% formalin (3.7% absolute formaldehyde) by injection using oral and cloacal injection sites with 21G-18G needles. Following injection, specimens were positioned for hardening and submerged in 10% formalin at room temperature for 24 to 48 hours. After fixation, specimens were rinsed thoroughly in tap water to remove any excess, unbound formalin. Dehydration: Specimens were submerged in three consecutive 100% acetone baths at room temperature with a 1:10 specimen/acetone ratio. In addition, the specimens were flushed through the previous injection sites (using a syringe and 21G needle) with 100% acetone initially and between baths. Each bath lasted two to five days, and the specimen/acetone mixture was agitated once each day. Acetone concentration was monitored with a hydrometer to determine the progression of specimen dehydration. Dehydration was considered satisfactory when the final acetone bath reached a specific gravity of 0.80. Silicone impregnation: Specimens were removed from the final acetone bath, and a silicone polymer mixture was immediately applied to the entire surface of the specimens using a brush. Following surface application, specimens were injected with silicone polymer mixture (via previous injection sites, penetrating into body cavities and deep tissue) with 23G and 21G needles and 1 to 5 ml syringes. Larger specimens that required more silicone (e.g., large rodents), were injected using 50 ml

syringes and 18G or 15G needles. The amount of polymer injected into the specimens varied between 1 and 25 ml per injection site depending on the size and species of the organism. The objective was to force in as much silicone as possible (until it began to leak out), while maintaining the original morphology of the specimen. The silicone polymer mixture was composed of NCS10 and the cross-linker, NCS6 (North Carolina products) at a ratio of 90:10. The NCS10/6 mixture remains stable at room temperature (Henry, 2007) and has a very low viscosity (easily injected through 23G needles with a 1 ml syringe). After the initial surface application and injection, specimens were wrapped in a layer of paper towels followed by an external layer of thin plastic wrap (to slow acetone evaporation). Specimens were inspected daily, injected with additional NCS10/6 mixture (to replace evaporating acetone), and re-wrapped in clean paper towels and plastic wrap. Daily injection with the NCS10/6 mixture continued until no acetone odor was detected on the paper towels or the specimen (approx. 3 to 15 days). After acetone evaporation was complete, the specimens were injected with NCS10/6/3 mixture of 90% NCS10/6 and 10% NCS3 (catalyst: North Carolina products), followed by a thin surface application (with a small brush) of NCS3 on the entire specimen. Mammalian specimens required blotting to remove excess NCS10/6 from the fur. The catalyst (NCS3) was then applied directly to the skin at various points using a 1ml plastic syringe to limit polymerization in the fur. The specimens were covered with thin plastic wrap. Daily inspection and additional NCS10/6/3 mixture injection was repeated as necessary for three to five days. Curing: Specimens were unwrapped, wiped clean of any silicone polymer that may have leaked out, and placed on paper towels in a small aquarium. Approximately 1ml of NCS5 (chain extender: North Carolina products) was placed in an open (60 X15mm) cell culture dish and a glass cover was placed over the aquarium to maintain an atmosphere of NCS5 vapor. Each subsequent day, chain extender (NCS5) was replaced, the specimens were wiped clean of any uncured silicone polymer that may have leaked, and specimens were inspected for any signs of distortion (shrinkage of the tail, legs or abdominal areas). If necessary, additional NCS10/6/3 polymer mixture was injected at or near any distorted areas. Excess polymer was periodically brushed from the pelage of mammals during the entire curing time.

14 Kumro et al. Specimens were maintained in the NCS5 vapor chamber for final curing for 20 to 30 days. Results Numerous vertebrate species that included an assortment of amphibians, reptiles, and mammals were plastinated. Specimens ranged in size from a 1.0g lizard to a 700g snake. The plastinated specimens were odorless, their surfaces were dry to the touch, and they exhibited a small degree of flexibility. In general, the plastinated specimens were life-like and the natural contours of the animals were maintained (Figs. 1, 2, 3). Gross dissection was performed on several specimens. Silicone had penetrated throughout the viscera and the internal morphology was maintained, as major organs remained in approximate position (Figs. 4, 5). The internal structures were odorless and dry with negligible shrinkage. Superficial, as well as deep muscles appeared thoroughly plastinated and major muscle groups retained their shape with little to no reduction in size (Fig. 6). Histological examination revealed polymer perfusion into the tissues of the internal organs and muscles, and that polymer had filled the interstitial as well as intracellular spaces. Discussion We argue that the success of the injection method is based on the physical properties and interactions of animal integument, acetone, and silicone polymer. Injection of silicone polymer results in an increased internal pressure that is contained by the integument of the specimen. The semi-permeability of the integument allows the passage of acetone molecules, while retaining the larger, more viscous silicone molecules. The much heavier silicone polymer (~27,000 g mol-1) readily -1 displaces the lighter acetone molecules (58.08 g mol ), causing movement of acetone through the integument, mouth, and cloaca. Acetone rapidly evaporates at room temperature; however, silicone polymer has an extremely low vapor pressure and negligible evaporation. In addition, the degree of keratinization of the integument results in varying rates of evaporation due to differences in permeability. These differences necessitate special measures for less keratinized specimens, such as amphibians, which require rapid coating with NCS10/6 and wrapping, to slow acetone evaporation. It should be emphasized that premature evaporation of acetone causes pronounced shrinkage and stiffness in specimens and prevents adequate injection of silicone polymer resulting in plastinated specimens of poorer quality.

The results demonstrate that high-quality plastinated specimens can be produced using only injection impregnation. This alternate, non-vacuum process is very low-tech and has minimal start-up costs. Injection plastination will allow small laboratories on limited budgets the ability to process specimens for teaching collections and display. Although we were able to use this technique successfully on a 700g animal, there are no doubt limitations to the upper size of specimens that can be successfully processed with this method. It should be emphasized that to produce high-quality plastinated specimens, it is important to be consistent with daily inspection and processing of specimens; especially in controlling the rate of acetone evaporation and adequate injection of silicone polymer. As with traditional plastination techniques, there is a small learning curve and some degree of skill and art involved. Our preliminary findings also indicate that small animal plastinated specimens are better teaching models in taxonomic lab courses and are preferred over traditional fluid-preserved and dry prepared specimens. Acknowledgements The following people were helpful in providing suggestions, helpful information, and/or feedback: Dr. Robert W. Henry, College of Veterinary Medicine, The University of Tennessee; Faculty, staff, and students of mammalogy and vertebrate zoology from the Department of Biological and Health Sciences, Texas A&M University-Kingsville.

Injection Plastination 15 life-like appearance.

Figure 1: Various species of mammals, amphibians and reptiles were plastinated using the injection method. In general, natural contours of the animals were maintained. a) Western Diamondback Rattlesnake, Crotalus atrox, b) Texas Coralsnake, Micrurus tener, c) Hispid Cotton Rat, Sigmodon hispidus, d) Mexican Spiny Pocket Mouse, Liomys irroratus, e) Yellow Mud Turtle, Kinosternon flavescens, f) Brazilian Free-tailed Bat, Tadarida brasiliensis, g) Cane Toad, Rhinella marina, h) Texas Toad, Anaxyrus speciosus, i) Prairie Skink, Plestiodon septentrionalis.

Figure 2: Texas Coralsnake (Micrurus tener). This specimen displayed excellent color retention.

Figure 3: Cane Toad (Rhinella marina). The natural contours of the animal were well maintained. Glass eyes were installed on this specimen to enhance its

Figure 4: Gulf Coast Toad (Bufo nebulifer). The specimen shows polymer penetration of the viscera and preservation of organs and internal morphology. a, b) liver, c) stomach, d, e) egg mass

Figure 5: Hispid Cotton Rat (Sigmodon hispidus). The specimen shows polymer penetration of the viscera and preservation of internal morphology. a) heart, b) liver, c) cecu, d) small intestine, e) spleen.

16 Kumro et al.

Figure 6. Hispid Cotton Rat (Sigmodon hispidus). The muscles of this specimen are thoroughly plastinated and muscle groups retained morphology.

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Latorre R, Henry RW. 2007: Polyester plastination of biological tissue: P40 technique for body slices. J Int Soc Plastination 22: 69-77.

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Latorre RM, García-Sanz MP, Moreno M, Hernández F, Gil F, López O, Ayala MD, Ramírez G, Vázquez JM, Arencibia A, Henry RW. 2011: How useful is plastination in learning anatomy? J Vet Med Ed 34: 172-176.

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Mansor O. 1996: Use of plastinated specimens in a medical school with a fully integrated curriculum. J Int Soc Plastination 11: 16-17.

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Mendez BA, Romero RL, Trigo FJ, Henry RW, Candanosa AE. 2008: Evaluation of imidazole for color reactivation of pathological specimens of domestic animals. J Int Soc Plastination 23: 17-24.

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Pashaei S. 2010: A brief review on the history, methods and applications of plastination. Int J Morphol 28: 1075-1079.

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Peris KJ. 2000: Plastination technology for biomedical research and studies in Kenya. J Int Soc Plastination 15: 4-9.

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Raoof A, Henry RW, Reed RB. 2007: Silicone plastination of biological tissue: Room temperature technique Dow™/Corcoran technique and products. J Int Soc Plastination 22: 21-25.

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An P, Zhang M. 1999: A technique for preserving the subarachnoid space and its contents in a natural state with different colours. J Int Soc Plastination 14(1): 12-17.

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Asadi MH, Mahmodzadeh A. 2004: Ascaris plastination through S10 techniques. J Int Soc Plastination 19: 20-2.

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Bickley HC, von Hagens G, Townsend, FM. 1981: An improved method for the preservation of teaching specimens. Arch Pathol Lab Med 105: 674-6.

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Cook P. 1996: Plastination as a clinically based teaching aid at the University of Auckland. J Int Soc Plastination 11: 22.

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Riepertinger A, Heuckendorf E. 1993: E 20 colorinjection and plastination of the brain. J Int Soc Plastination 7: 8-12.

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Correia JAP, Prinz RAD, Benevides de Freitas EC, Pezzi LHA. 1998: Labeling and storing plastinated specimens-An experience from Universidade Federal Do Rio De Janeiro. J Int Soc Plastination 13(2): 17-20.

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Sivrev D, Kayriakov J, Trifonov Z, Djelebov D, Atanasov M. 1997: Combined plastination methods for preparation of improved ophthalmologic teaching models. J Int Soc Plastination 12(2): 12-14.

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Dawson TP, James RS, Williams GT. 1990: Silicone plastinated pathology specimens and their teaching potential. J Pathol 162: 265-72.

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Sora MC. 2007: Epoxy plastination of biological tissue: E12 ultra-thin technique. J Int Soc Plastination 22: 40-45.

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Henry RW. 2007: Silicone plastination of biological tissue: Room temperature technique North Carolina technique and products. J Int Soc Plastination 22: 26-30.

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Sora MC, Strobl B, Radu J. 2004: High temperature E12 plastination to produce ultra-thin sheets. J Int Soc Plastination 19: 22-25.

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Steinke H, Rabi S, Saito T. 2008a: Staining body slices before and after plastination. Eur J Anat 12: 51-55.

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Steinke H, Rabi S, Saito T, Sawutti A, Miyaki T, Itoh M, Spanel-Borowski K. 2008b: Light-weight plastination. Ann Anat 190: 428-431.

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Suriyaprapadilok L, Withyachumnarnkul B. 1997: Plastination of stained sections of the human brain:

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Henry RW, Nel PPC. 1993: Forced impregnation for the standard S10 method. J Int Soc Plastination 7: 27-31. Latorre R, Arencibia A, Gil F, Rivero M, Ramirez G, Vaquez-Auto JM, Henry RW. 2004: Sheet plastination with polyester: An alternative for all tissues. J Int Soc Plastination 19: 33-39.

Injection Plastination 17 Comparison between different staining methods. J Int Soc Plastination 12(1): 27-32. •

Tianzhong Z, Jingren L, Kerming Z. 1998: Plastination at room temperature. J Int Soc Plastination 13(2): 21-25.

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von Hagens G, Tiedemann K, Kriz W. 1987: The current potential of plastination. Anat Embryol 175: 411–21.

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Zhong ZT, Xuegui Y, Ling C, Jingren L. 2000: The history of plastination in China. J Int Soc Plastination 15: 25-29.

The Journal of Plastination 25(1): 18-21 (2013)

A. Raoof* L. Marchese A. Marchese A. Wischmeyer Division of Anatomical Sciences, Office of Medical Education The University of Michigan Ann Arbor USA

Demonstration of Systolic and Diastolic Phases of the Cardiac Cycle in a Plastinated Human Heart ABSTRACT: Plastination has enhanced the way students study human gross anatomy by providing them with three-dimensional specimens that they can hold and manipulate. These specimens allow students to learn gross anatomy, especially difficult areas, more efficiently. However, the intricacies of organ function in life are is still difficult to understand from dissected specimens. At the University of Michigan Medical School, innovative approaches to enhance the quality of plastinated specimens have been implemented to demonstrate complex anatomical features. The heart is a particularly difficult organ for students to,visualise because of the unique changes it undergoes during systolic and diastolic phases of the cardiac cycle. The aim was to develop a plastinated heart model that demonstrates how cardiac valves function during the systolic and diastolic phases. Five hearts were collected from cadavers, dissected and plastinated. Various incisions in the heart were made to reveal the cardiac valves. Corks, sutures, and hinges were used to position and to hold the valves in place either in its contracted state (systole) (2 out of 5 hearts) or in its relaxed state (diastole) (3 out of 5 hearts). A pilot survey was administered to get students’ feedback on these plastinated models. The results indicate that a majority of students favor this novel animated model as it displays both systolic and diastolic phases while keeping superficial structures of the heart intact. KEY WORDS: plastination, heart, valves, systole, diastole *Correspondence to: Dr. Ameed Raoof, Office of Medical Education, The University of Michigan Medical School, 3740 Med. Sci. II Bldg., Ann Arbor, MI, 48109-5608, USA; E-mail: [email protected].

Introduction:

Materials and methods:

The advent of the plastination process in gross anatomy laboratories provided students with a learning tool that transforms two-dimensional textbook images into threedimensional models that can be examined and manipulated (von Hagens et al., 1987). One of the most challenging organs to understand for medical students is the human heart, with its complex physiology and unique two-stroke mechanism. Knowledge of the basic anatomy and physiology of the heart is an essential component of any medical or health science curriculum. Demonstrating how heart valves open and close during systolic and diastolic phases of a cardiac cycle is difficult with cadaveric specimens. The aim of this study was to develop a plastinated heart model that demonstrates how cardiac valves function during the systolic and diastolic phases. Plastinated models of hearts in both systole and diastole would allow students to identify each valve and its role in each phase of the heartbeat, as well as provide them with an intact three-dimensional tool for learning the fine details of heart anatomy and how its structure is related to its function.

Five embalmed human heart specimens were harvested from cadavers donated to the University of Michigan’s Anatomical Donations Program, age ranged between 70-80 years (average 75). The hearts were kept in a water bath throughout the duration of the dissection to preserve their hydration and to enhance clot removal from the heart. At first the pericardium was removed, as well as fat from around veins and arteries. To remove the superior aspect of the heart, the semilunar cusps were located and an incision was made just above the pulmonary and aortic valves. This incision was extended around the heart superior to the coronary sulcus for a full transverse section of the atria and aorta and pulmonary trunk. To mimic the contracted state of the heart (systole), the mitral and tricuspid valves were sutured closed while the aortic and pulmonary semilunar valves were positioned open using corks of a suitable size in two of the hearts.. The other three hearts were used to demonstrate the heart in its relaxed state (diastole). The mitral and tricuspid valves were positioned open using corks and the aortic and pulmonary semilunar valves were sutured closed. After the heart valves were prepared, the two portions of the

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Systolic and Diastolic Phases 19 heart were sutured to align and preserve the original external anatomy (Raoof, 2001). After flushing the hearts with water, they were dehydrated in cold acetone. The prepared hearts were plastinated using the room temperature method (Raoof 2001, Raoof et al., 2007). The impregnation polymer was mixed with 8% cross-linker (CR 22- Dow Corning). After forced impregnation, all sutures and corks were removed. A small brass hinge was attached with pins and glue to keep the basal and apical portions of the specimens as one unit. The hinges were attached with sutures, fine screws, and rubber silicone (Figure 2). After the hinge had been securely attached to the heart, the specimen was cured using a catalyst (CT 32- Dow Corning) (Raoof, 2001, Raoof et al., 2007). The impregnated and cured hearts were used by undergraduate students in Human Anatomy. The course introduced the students to the basic concepts of systemic anatomy and included visits to the lab where pertinent plastinated specimens are displayed. To evaluate student perception after using the plastinated hearts, a pilot survey was developed and administered to these 199 undergraduate students. Questionnaires were distributed and students were asked to express their opinions to three questions on a 5-point Likert scale. The questions were (1) “whether the heart specimens have been useful in demonstrating structural relationships”, (2) “whether the specimens have been beneficial to learning the heart’s anatomy”, and (3) “whether the specimens have been beneficial to understanding function in correlation with structure”. Data from the survey was compiled and analyzed using Microsoft Excel. Results Careful preparation of the hearts permitted production of heart specimens for demonstration of systole and diastole of the cardiac cycle while preserving the relationship of superficial anatomy of the heart (Figure 3). The valves were misshaped during closure and opening but could be made anatomically correct prior to hardening/application of the catalyst. The value of these models for undergraduate study of heart anatomy and physiology was assessed by conducting a pilot survey among undergraduate anatomy students. Students were allowed to spend time studying and manipulating the heart, after which they were asked if they felt a model of this type was beneficial

to learning anatomy. Sixty out of 199 students responded to the survey. A majority of students felt that these specimens were useful in demonstrating structural relationships (Figure 4) and were helpful to their understanding of details of heart anatomy (Figure 5) and how structure correlates with function (Figure 6). Discussion and Conclusion Metal hinges were used to create a simple animated model of the heart that would permit students not only to study the external features of the heart, but also observe the internal structures and to gain a better understanding of the correlation between valve function and blood flow during various stages of the cardiac cycle. These systolic and diastolic models help students comprehend the sequence of events that make up a cardiac cycle. Technically, it is essential to modify valve contours which are misshaped due to the pressure of the cork and/or suture to their final perceived anatomical shape after silicone impregnation and before adding the catalyst. During preparation of the cardiac valves an incision was made above the pulmonary and aortic valves to view the semilunar valves. The incision around the heart superior to the coronary sulcus permitted the view of the atrioventricular valves. After plastination the two portions of the heart were sutured to align and preserve the original external anatomy (Figure 1). Baptista and Conran, 1989, prepared the cardiac valves prior to dehydration by anchoring the leaflets of the right and left atrioventricular valves with small pieces of moistened cotton introduced through a small incision in the right and left ventricles. Small pieces of cotton were also positioned in the cusps of the semilunar valves. After plastination, the incision was widened reviewing the internal structures of the ventricles such as chordae tendineae and papillary muscles, and their relationship with the cardiac valves. Gomez et al., 2011, in an attempt to correlate plastinated heart slices with echocardiographic images, used 13 dog hearts fixed by dilation. The plastinated slices corresponded accurately with the echocardiographic images revealing the internal anatomy with great details. The dilated cavities of the plastinates (due to fixation) made comparison with echocardiographic images of systole difficult. A current project will further enhance understanding of heart anatomy. The heart is being sectioned along a sagittal plane through the septum and a second hinge attached, giving another view into the heart. Hinges are rather obtrusive, therefore, magnets will be used instead of hinges.

20 Raoof et al.

Figure 4: Student response, pilot survey, question 1 - whether the heart specimens have been useful in demonstrating structural relationships. Figure 1: Left specimen: heart sutured at the pulmonary trunk. Right specimen: base of the heart transected from the apex.

Figure 5: Student response, pilot survey, question 2 - whether the specimens have been beneficial to learning the heart’s anatomy.

Figure 2: Transected human heart with hinge

Figure 6: Student response, pilot survey, question 3 - whether the specimens have been beneficial to understanding function in correlation with structure.

Figure 3: Plastinated human heart displays the systolic (left) and diastolic (right) forms.

Systolic and Diastolic Phases 21 References •

Baptista, C.A.C. and Conran P.B.1989: Plastination of the heart: Preparation for the study of the cardiac valves. J Int. Soc. Plastination 3: 3-7

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Gomez A., Del Palacio J. F., Latorre R., Henry R. W., Sarria R., Arbors O. L. 2011: Plastinated heart slices aid echocardiographic interpretation in the dog. Vet Radiol Ultrasound 53 (2): 197-203

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Raoof, A. 2001: Using a room-temperature plastination technique in assessing prenatal changes in the human spinal cord. J Int Soc Plastination 16: 5-8

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Raoof A, Henry RW, Reed RB 2007: Silicone plastination of biological tissue: room temperature TM plastination technique - Dow /Corcoran technique and products. J Int Soc Plastination. 22: 21-25.

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von Hagens G, Tiedemann K, Kriz W. 1987: The current potential of plastination. Anat Embryol 175(4): 411-421.

The Journal of Plastination 25(1): 22-27 (2013)

Mircea-Constantin Sora* Radu Jilavu Plastination Laboratory Center for Anatomy and Cell Biology Medical University of Vienna, Austria Petru Matusz Anatomical Department University of Medicine and Pharmacy "Victor Babes" Timisoara, Romania

Three dimensional reconstruction of a female pelvis using plastinated cross-sections - Using Plastination for 3D Reconstruction ABSTRACT: In this study, a three-dimensional (3-D) model of the pelvis was built based on thin slice plastination cross-sections of the adult female pelvis and 3-D reconstruction technology. A female pelvis was obtained, MRI scanned, plastinated, sectioned and subjected to 3-D computerized reconstruction using the WinSURF modeling system (SURFdriver Software). Qualitative observations revealed that the morphological features of the model were consistent with those displayed by typical cadaveric specimens, whilst closer morphometric analysis indicated that the model did not significantly differ from a sample of cadaveric specimens. This conclusion proves that the utilization of plastinates for generating tissue sections can be successfully integrated into 3-D computerized modeling . A better understanding of pelvic floor anatomy is relevant to gynaecologists, radiologists, surgeons, urologists, physical therapists and all professionals who take care of women with pelvic floor dysfunction. The objective of this study was to describe the method of developing this computerized model of the human female pelvis using plastinated slices. It is a method which could be applied to reconstruct any desired region of the human body.

KEY WORDS: female, pelvic floor, levator ani muscle, anatomy, plastination, 3-D reconstruction *Correspondence to: Mircea-Constantin Sora, M.D., PhD Centre for Anatomy and Cell Biology Plastination and Topographic Anatomy, Währingerstr. 13/ 3 A-1090 Wien, Austria E-mail: [email protected]

Introduction: The pelvic floor has a complex spatial structure, knowledge of which is essential when assessing pathologies in this area (Contouris,1988; Wiliams,1995). Women, for the most part, undergo pelvic floor examinations for urinary incontinence or prolapse of the internal genitalia or of the urinary bladder (Goodrich, 1993; Lienemann, 1997). The advent of lateral urethrocystography (UCG) and colpocystourethrography has made it possible to visualize and objectively assess pathological changes in the urinary bladder and urethra, whereas pathological changes in the pelvic floor and of the supportive structures of connective tissue can at best only be evaluated indirectly. The pelvic floor muscles serve as a barrier for perianal fistulas and abscesses, and knowledge of the shape of the pelvic floor muscles, required to assess any perianal fistulas and their extent, is gained using MR imaging. The detailed anatomical and morphological information provided by sheet plastination can be added to this MR image information, which has a poorer detail resolution.

The improved properties of plastinated specimens are mainly accounted for by the superior qualities of curable polymers. In the plastination technique, tissue water and lipids are replaced by cured polymers. The class of polymer used determines the mechanical (flexible or firm) and optical (opaque or transparent) properties of the specimen. Plastinated specimens are dry, odorless, and durable; they even retain structural details down to the histological level. Today, 30 years after its introduction (v. Hagens, 1977), plastination is being applied in more than 250 departments of anatomy, pathology, forensic science and biology all over the world. In research, the technique of sheet plastination allows the arrangement of all tissue-components to be studied in their undisturbed context. This is of major interest in the borderline area between gross anatomy and histology with respect to muscular and connective tissue patterns. Epoxy resins are used to produce transparent body or organ slices, which, for research purposes, allow the study of the topography of all body

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Using Plastination for 3D Reconstruction 23 structures in an uncollapsed and non-dislocated state. In addition, the specimens are useful in advanced training programs in sectional topography, resident training in CT and NMR (de Barros, 2001; Lazanoff, 2003; Thomas, 2004). Pelvic floor dysfunction, which includes urinary and fecal incontinence as well as pelvic organ prolapse, is a highly prevalent condition in women. Ten percent of all women undergo at least one operation to treat pelvic floor dysfunction during their lifetime (Mant, 1997). The annual cost of treating urinary incontinence in Austria alone is estimated at 48 million Euros annually. However, little is known about specific pelvic floor pathomorphology and even less about pathophysiology as it relates to pelvic floor dysfunction. DeLancey et al., (2003) used Magnetic Resonance Imaging (MRI) to investigate levator ani muscle damage, and Lien et al., (2004) constructed a computerized model in order to determine the stretch forces that exceed the forces which muscle tissue can usually sustain. In our study, a three-dimensional (3-D) model of the pelvis was built for the first time based on thin slice plastination crosssections of the adult female pelvis and 3-D reconstruction technology. In order to investigate this topic, we need to know the orientation of the levator ani muscle and the interaction of muscles, bones, connective tissue and pelvic organs. Sections of a plastinated pelvis can help us to understand the levator ani architecture, and as the relation of muscles, fascia, organs and bones can be studied perfectly it is also well suited for a 3-D computerized reconstruction. This offers the opportunity to compare the use of identical physical and virtual models for the development of a 3-D anatomical computer model based system and its interactive manipulation.

Materials and methods: The female human pelvis used for this study was removed from a fresh unfixed cadaver. A large specimen block, containing the pelvis, was cut into the desired sized blocks for impregnation with the maximum size of 250mm x 150mm x 150mm. High-field, highresolution magnetic resonance imaging provides high soft-tissue contrast that allows imaging of the regular female pelvic floor anatomy to a level almost equal to plastinated macroscopic anatomic cross-sections. In this study macroscopic cadaveric cross-sections of the

regular female pelvic floor anatomy are compared and correlated in-vitro with high resolution MR imaging of the same orientation. The cadaveric pelvis block was cut and adjusted to fit the standard Bruker magnetic resonance head coil (dimensions: 140x90x100 mm). Wooden markers were installed as landmarks to allow correct orientation and slice selection. The scanned pelvis block was frozen at -80°C for one week and then plastinated following the standard ultrathin E12 slice plastination method. (An and Zhang, 1999; Johnson, 2000; Lane, 1990; Sora, 2007 a). Freeze substitution is the standard dehydration procedure for plastination, and shrinkage is minimized when cold acetone is used. The tissue block was placed into a -25°C freezer and then submerged in cold (-25°C) technical quality 100% acetone for dehydration. Methylene chloride was used for the degreasing and then impregnation was performed using the following epoxy mixture (Biodur E12, Rathausstr.18, 69126 Heidelberg, Germany): E12 (resin)/ E6 (hardener)/ E600 (accelerator) (Sora et al., 2007a). Once impregnation had been completed, the tissue block was removed from the vacuum chamber and inserted into a mold constructed of Styrofoam and lined with polyethylene foil. The mold containing the impregnated specimen and resin-mix was then placed in a 65°C oven for four days to harden the resin-mix. The tissue/resin block was cooled to room temperature and the mold removed. Before sawing, three colored plastic sticks were placed inside the block as markers. Using a contact point diamond blade saw, Exact 310 CP (Exact Apparatebau GmbH, Norderstedt, Germany) the hardened E12 block was cut into 1.6 ± 0.26 mm slices. Between each tissue slice, the width of the saw blade (0.4 mm), was lost. Finally the caudal surfaces of the plastinated slices were scanned into a computer using an EPSON GT-10000+ Color Image Scanner. In every scan we included a ruler as a calibration marker and used the UTHSCSA IMAGE TOOL v.2.0 for Windows software (The University of Texas Health Science Center in San Antonio) for measurements. Scanned images of the tissue slices were loaded into WinSURF (http://www.surfdriver.com) and traced from the monitor. The following features, defined as objects, were used in the reconstruction: pelvic girdle, levator ani muscle, obturator internus muscle, rectum, urinary bladder, urethra and the vagina. Each object was traced and numbered accordingly. Afterwards, the reconstruction was rendered, visualized, and qualitatively checked for surface discontinuities by

24 Sora et al. rotating the model. Furthermore, WinSURF offers a measuring tool to record height, width, and depth measurements after rendering the model. Minimum requirements for the software are extremely modest and include a 200 MHz processor, Windows 2000, Windows XP, Windows 7, 24 MB of free available systems RAM (64 MB recommended), 50 MB of available disk space, 1024 x 768, 16-bit color display, CD ROM drive and 3 1/2” floppy or 100 MB Zip Drive, and a mouse or compatible pointing device; all of which are included in standard PC equipment. Results Sectional plastination showed the structures of the pelvic floor muscles and their relationship to adjacent structures with a resolution down to microscopic level (Fig. 1, 2). This method allowed assessment of the course of muscle fibers. Diagnostically adequate visualization of the pelvic floor was achieved by MR imaging with a pixel size of 0.89 x 0.89 mm and soft tissue contrast. By segmentation of the outlines, it was possible to create a 3-D model which consisted of the levator ani muscle, the vagina, the urethra, the rectum, the obturator internus muscles, and the pelvic bones. The anatomical structures of the pelvis could be easily identified and the borders could be traced rapidly and reliably. The thin plastinated slices displayed the nerves, muscles, vessels and bones of the pelvic region distinctly . Once scanned and loaded into WinSURF, edge detection was used to quickly collect tissue borders or contours. The generated 3-D pelvis model displays a morphology corresponding qualitatively to the actual cadaver specimen (Fig. 3). The reconstructed images appeared well-defined, especially the spatial positions and complicated relationships of contiguous structures of the female pelvis. All reconstructed structures can be displayed in groups or as a whole and interactively rotated in 3-D space. Various features such as transparency control, individual object selection, animation and a variety of manipulation modes facilitate visualization of the complex pelvic anatomy (Fig. 4). Discussion The pelvic floor has a complex spatial structure, of which only parts are visualized on sectional images (Fritsch, 1995; Fritsch, 2004; Fröhlich, 1997;). It is, however, necessary to correctly relate the visualized part to the

entire structure in order to properly assess pathologies. A 3-D model in which the positions of the imaging plane of interest are shown clearly improves the vividness of depiction. Additional information, such as the course of muscle fibers or connections of the muscles with each other for a given imaging plane, was gained from the plastinated sections. In contrast to anatomical preparation, the structures and spatial relationships of the tissues were not altered by plastination . Thin plastinated slices of 1.6 mm are essential if an accurate 3-D reconstruction is desired (Qiu, 2003; Sha, 2001; Sora, 2002; Sora, 2007 b). Although the process of plastination extends the time and effort required to generate images for analysis, considerable detail is provided and the reconstructed pelvic model exhibits the bones and surrounding soft tissue. Various groups had previously investigated the female pelvic floor by computed tomography (CT), magnetic resonance (MRI) or ultrasound (US), but none had evaluated it by plastination. Transparent body slices are used for research purposes because they allow the study of body structures in a non-collapsed and non-dislocated state. Sectional plastination anatomy also confers a major advantage since the decalcification of bony tissue is not necessary, and spatial relationships are retained between contiguous features of differing composition. Thus, topography between bones and contiguous soft tissues is retained without additional chemical manipulation. Computer models and animations of anatomical features are becoming increasingly attractive as a means to communicate complex spatial relationships and concepts effectively (Dev, 2002). Although many educational animations are based on artistic renderings (Gould, 2001), more recent applications use virtual representations derived from actual cadaveric material (Neider, 2000; Lozanoff, 2003). A logical advantage of these models is that they provide a greater sense of realism, which increases the amount of perceived information. In the future one can envision anatomy lectures consisting of only 3-D models without 2-D images (Trelease, 2002). One of the standard interventions in the therapy of urinary incontinence is the tension-free vaginal tape (TVT) intervention (de Leval, 2003). We used the reconstructed pelvis to simulate a TVT placement (Fig. 5). This model will furthermore provide data to calculate stretch ratios and other biomechanical tissue properties.

Using Plastination for 3D Reconstruction 25 The system described here relies on relatively inexpensive hardware, including a scanner and computer. The WinSURF reconstruction package, from SURFdriver Software © (surface reconstruction driver), was developed expressly for use in three-dimensional anatomical reconstruction and is a simple icon-driven system (Moody and Lozanoff, 1998).

MOI – Obturator internus R – Rectum S – Pubic symphysis V – Vagina U – Urethra

One major problem that occurs with existing anatomical databases is the low resolution for smaller anatomical structures. Plastination can provide a useful alternative for generating anatomical databases. Moreover, plastinates are significantly easier to cut, stain, and handle compared to fresh-frozen tissue, since they are significantly more durable due to the silicone infiltrate. Although the female pelvis reconstruction presented here did not appear to be affected by a loss of information (due to tissue loss between slices), further testing will be required to examine this issue. The capability to reconstruct individual and combined images of the pelvic structures, view them from all surgical angles, and allow for accurate measurement of their spatial relationships provides important guidance for surgeons. The reconstructed model can also be used for residency education, testing unusual surgical techniques and for the development of new surgical approaches. The 3-D model of the female pelvis presented in this paper provides a stereoscopic view to study the adjacent relationship and arrangement of respective pelvis sections.

Figure 1: Transparent cross-section of the female pelvis. a. a plastinated cross-section and b. comparable MRI image. G – Gluteus maximus FI – Ischiorectal fossa MLA – Levator ani MOE – Obturator externus

Figure 2: Plastinated cross-section at the level of the hip joint. A – Acetabulum C – Coxal bone M – Marker MOI - Obturator internus MLA – Levator ani R – Rectum S – Pubic symphysis V - Vagina VU – Urinary bladder

Figure 3: A 3-D reconstruction of the female pelvis. a. View of the levator ani muscle (red) from caudo-frontal. b. View from the right side with removed pelvic bones. c.

26 Sora et al. View from cranial into the pelvis. MLA – Levator ani MOI - Obturator internus R – Rectum V – Vagina U – Urethra VU – Urinary bladder

References •

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Figure 4: Front view of the 3-D reconstruction with transparent bony structures. MLA – Levator ani MOI – Obturator internus R – Rectum V – Vagina U – Urethra VU – Urinary bladder

Figure 5: Reconstructed pelvis with a tension-free vaginal tape (TVT), after removal of the right side of the bony structures. MLA – Levator ani MOI – Obturator internus R – Rectum TVT – Tension-free vaginal tape V – Vagina U – Urethra VU – Urinary bladder

Using Plastination for 3D Reconstruction 27 •

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The Journal of Plastination 25(1): 28 (2013)

17th International Conference on Plastination St. Petersburg, Russia

The Journal of Plastination 25(1): 29 (2013)

Journal of Plastination

Instructions for Authors (Revised January 2013)

JOURNAL OF PLASTINATION is owned and controlled by the International Society for Plastination (ISP).

responsibility to obtain permission to reproduce illustrations, tables and figures from other publications.

Goals - The Journal of Plastination (ISSN 10902171) is to provide a medium for the publication of scientific papers dealing with all aspects of plastination and preservation of biological specimens.

Copyright Transfer Form may be downloaded from http://www.journal.plastination.org/downloads/copyright.pdf. After the form is completed and signed by all the authors, it should be submitted to the Editorial Office ([email protected]) as a pdf or jpeg file via an email attachment.

Submission Guidelines All manuscripts must be submitted to the Editorial Office via the e-mail: [email protected]. If you experience any problems or need further information, please contact either Dr. Carlos Baptista, [email protected], or Dr Selcuk Tunali, [email protected]. Authors must have an e-mail address at which they may be reached. Necessary Files for Submission Include: • Cover letter • Manuscript (including references and legends) • Table(s) (when appropriate) • Figure(s) (when appropriate) • Copyright Release Form (after acceptance)

figure

Note: The above items should be prepared as separate files. Each file must contain a file extension (.doc, tif, jpg, eps). • File formats appropriate for text and table submissions: Microsoft Word • File formats appropriate for figure submissions: TIFF, JPEG (JPG) and EPS Categories of submissions: Articles published in Journal of Plastination are grouped into general article types (listed below). Final designation of a manuscript’s article type is determined by the EDITOR. • Original Research – Plastination • Original Research – preservation • Education • Case reports • Technical brief notes • Review - by invitation only • Legacy – institutions and people • Correspondence • Editorial Acceptance of a submission implies the transfer of copyright from the authors to the publisher. It is the author's

Manuscript preparation Cover Letter The cover letter should include a statement of authorship, notification of conflicts of interest, ethical adherence, and any financial disclosures. Cover letters may be addressed to the Editor-in-Chief, Journal of Plastination. Manuscript The manuscript should consist of subdivisions in the following sequence: Title Page Abstract with keywords Text Introduction Materials and methods Results Discussion References Figure Legends Title Page The first page of the manuscript should include: • Title of paper • Each author’s name • Institution from which paper emanated, with city, state, and postal code. Each affiliation should be listed as a separate entity, with a superscript number that links it to the individual author. For example: S. D. HOLLADAY1*, B. L. BLAYLOCK2 and B. J. SMITH1 1 Department of Biomedical Sciences and Pathobiology, Virginia Maryland Regional College of Veterinary Medicine, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061-0442, USA. 2 College of Pharmacy and Health Sciences, University of Louisiana at Monroe, Monroe, LA 71209, USA. • Corresponding Author’s name, address, telephone and telefax numbers, and e-mail address.

The Journal of Plastination 25(1):30 For example: *Correspondence to: Dr Shane D. HOLLADAY, Department of Biomedical Sciences and Pathobiology, Virginia Maryland Regional College of Veterinary Medicine, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061-0442, USA. Tel.: +001 404 739 6403; Fax.: +001 404 739 6492; E-mail: [email protected] It is the corresponding author’s responsibility to notify the Editorial Office of changes of address. Only the corresponding author should communicate with the Editorial office for matters regarding each manuscript. Abstract & Key Words: The abstract should be no longer than 250 words. It should contain a description of the objectives, materials and methods, results, and conclusions. The abstract should include a section on technique/technical development if the paper is significantly technical in nature. The abstract must be written in complete sentences and be intelligible without reference to the rest of the paper. No references should be used in the abstract. On the same page, list, in alphabetical order, five Key Words that reflect the content of the manuscript. Consult the Medical Subject Headings for appropriate key words. Key words should be set in lower case (except for essential capitals), separated by a semicolon and bolded. References: • References to published works, abstracts and books must include all that are relevant and necessary to the manuscript. • Citations in the text should be in parentheses and listed chronologically; e.g. (Bickley et al., 1981; von Hagens, 1985; Henry and Haynes, 1989) except when the authors name is part of a sentence; e.g. "…von Hagens (1985) reported that…" When references are made to more than one paper by the same author published in the same year, designate each citation as 1999 a, b, c, etc. • Literature cited may only include the publications, which are cited in the text. References are to be listed alphabetically using abbreviated journal names according to Index Medicus. Page numbers of the citation must be included. • Examples of the reference style are as follows: • For a journal article: Bickley HC, von Hagens G, Townsend FM. 1981: An improved method for preserving of teaching specimens. Arch Pathol Lab Med 105:674-676. • For a book section: Henry R, Haynes C. 1989: The urinary system. In: Henry R, editor. An atlas and guide to the dissection of the pony, 4th ed. Edina, MN: Alpha Editions, p 8-17.

• For other publications: Von Hagens G. 1985: Heidelberg plastination folder: Collection of technical leaflets for plastination. Heidelberg: Anatomiches Institut 1, Universität Heidelberg, p 16-33. Figure legends • Legends for all figures should be brief, specific and not be a substitute listing for the result section, and appear on a separate page at the end of the manuscript, following the list of references. • Legends must be numbered consecutively as they first appear in the text. • All symbols or abbreviations appearing in any figure must be defined in the legend. Tables • All tables must be cited in the text and have titles. Table titles should be complete but brief. Information other than that defining the data should be presented as footnotes. • Create tables using the table creating and editing feature of Microsoft Word. Do not use Excel or comparable spreadsheet programs. • Each table should be simple and uncomplicated, with NO vertical and as few horizontal lines as possible. • Each table is to appear on a separate page and must include the table title and appropriate column heads. • Save each table in a separate word document file and upload individually, like figures. • Do not embed tables within the body of the manuscript. Figures • All figures must be cited in the text and must have legends. • Each figure should be attached as a separate file and labeled with the appropriate number. • Figures should be created, saved and submitted as either a TIFF, JPEG (JPG) or an EPS file. • Line drawings must have a resolution of at least 1200 dpi, and electronic photographs, scanned images, radiographs, CT and MRI scans must have a resolution of at least 300 dpi. • The size of each figure should be at least 8.25 cm / 3.25 inches (one-column width) or 16 cm / 6 inches (twocolumn width). • Magnification must be recorded and have a “scale bar” in the photo. Since reproduction of illustrations is costly, authors should limit the number of figures to those which adequately present the findings, and add to the understanding of the manuscript. • Figures that are submitted in color must be published in color. Authors are responsible for the costs of any color reproductions. Contact the editor for details.