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The Journal of Plastination 25(1):1-30 (2013)9The Journal of PlastinationThe official publication of the International Society for PlastinationIN THIS ISSUE:Low-temperatureDehydration and RoomtemperatureImpregnationof Brain Slices UsingBiodur TM S10/S3 – p 3P35 Plastination:Experiences with DelayedImpregnation – p 9Injection Plastination:A Low-Tech, InexpensiveMethod for SiliconePreservation of SmallVertebrates – p 13Demonstration of Systolicand Diastolic Phases ofthe Cardiac Cycle in aPlastinated HumanHeart - p 18Three DimensionalReconstruction of a FemalePelvis using PlastinatedCross-sections – p 22Volume 25 (1); July 2013

The Journal of Plastination 25(1):1-30 (2013)Journal of Plastination Volume 25 (1); July 2013ContentsLetter from the President, Carlos. A. C. Baptista 2Low-temperature Dehydration and Room- temperature Impregnation of BrainSlices Using Biodur TM S10/S3, Mandeep Gill Sagoo, Philip J AddsP35 Plastination: Experiences with Delayed Impregnation, M. Üzel, A.H.WeigleinInjection Plastination: A Low-Tech, Inexpensive Method for SiliconePreservation of Small Vertebrates, Shawnda L. Kumro, Ashton V. Crocker,Randy L. PowellDemonstration of Systolic and Diastolic Phases of the Cardiac Cycle in aPlastinated Human Heart, A. Raoof, L. Marchese, A. Marchese, A. WischmeyerThree Dimensional Reconstruction of a Female Pelvis Using Plastinated Crosssections- Using Plastination for 3D Reconstruction, M. C. Sora, R. Jilavu, P.Matusz3912182217th International Conference on Plastination 28Instructions for Authors 29

The Journal of Plastination 25(1):3-8 (2013)ORIGINALRESEARCHARTICLEMandeep Gill Sagoo*Philip J. AddsDivision of BiomedicalSciences (Anatomy)St George's, Universityof LondonLondon, UKLow-temperature dehydration and room-temperatureimpregnation of brain slices using Biodur TM S10/S3ABSTRACT: The standard method for plastination with Biodur TM S10/S3 involves lowtemperaturedehydration in a volatile intermediary solvent followed by forced impregnation undervacuum at -15°C. However, some institutions have been reluctant to install low-temperatureimpregnation equipment because of health and safety and cost considerations. The aim of thisstudy is to investigate a low-budget and simple to set up room temperature plastination procedureto prepare neuroanatomy teaching resources. Previous studies at St George’s, University ofLondon have shown that a low-temperature dehydration/ room temperature impregnation protocolfor Biodur TM S10/S3 can produce results comparable, if not equal, to the standard method. Fiftyfourformaldehyde-fixed brain slices were dehydrated in acetone at -30° C and vacuumimpregnated at room temperature. Twenty slices were stained with Mulligan’s stain beforeplastination. The slices were measured before dehydration and after impregnation to monitorshrinkage. Shrinkage was acceptable (6.99% in lengths and 6.19% in widths) in both stained andunstained slices, and did not detract from the appearance of the slices. The stain has thus far notfaded on exposure to light. Therefore, this procedure can be used to plastinate brain slices withquality comparable to low temperature plastination, which further extends the potentialapplications of room-temperature plastination.ORIGINAL RESEARCHKEY WORDS: low cost plastination; brain slices; Mulligan staining; teaching resource;Biodur TM S10/S3*Correspondence to: Mandeep Sagoo, Biomedical Sciences (Anatomy), St. George’s Universityof London, Cranmer Terrace, Tooting, London, SW170REEmail: msagoo@sgul.ac.ukIntroduction:Plastination, developed by von Hagens in Heidelberg,Germany, is a unique technique of obtaining valuable,dry, odorless and nontoxic biological specimens (vonHagens, 1986; von Hagens et al., 1987). Since 1990,plastinated specimens have been used to teachanatomy and neuroanatomy, and these specimens area valuable supplement for learning 3-dimensionalanatomy, for computer assisted modules and for selfdirectedanatomy 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, thesespecimens are durable, easy to use and do not drip onstudents’ handbooks (Holladay and Hudson, 1989).For enhancing neuroanatomy teaching at St. George’s,University of London, we have produced plastinatedbrain slices using a low-temperature dehydration/ roomtemperature impregnation method (Adds, 2008) whichdiffers from the standard method (de Jong and Henry,2007) in that forced impregnation is carried out at roomtemperature. Mulligan staining is a well-establishedstaining method for gray matter, however the stainsoon fades from wet specimens unless kept in the dark(Baeres and Møller, 2001).In this study, we assess the ability of Mulligan’s stain towithstand dehydration and impregnation duringplastination, and its subsequent resistance to fading inthe cured specimen. The advantage of the methoddescribed here is that it is a low budget and simple toset-up room-temperature procedure which produceshigh quality teaching resources.Materials and methods:The materials used were acetone (Anala R Normapur,VWR), grids for separating brain slices (Biodur TM ), -30°C laboratory freezer, air-tight containers, vacuum pump(rotary vane vacuum pump, 6m3/h) and plastinationchamber, polymers S10, S3 and S6 (Biodur TMGermany).

4 Sagoo et al.The brains were obtained from seven formaldehydefixedcadavers (embalmed with 10% formaldehyde,10% polyethylene glycol, 5% phenol and 75% ethanol)from the Dissecting Room of St. George’s, Universityof London.Consent for anatomical examination and imaging hadbeen given under the Human Tissue Act (2004). Thebrains were cut into sections approximately 1cm thickusing a rotary meat slicer, giving a total of 54transverse and coronal slices. The length and width ofeach slice was measured with a steel ruler beforedehydration and after impregnation. Twenty randomlyselected slices were stained with Mulligan’s stain priorto plastination.The brain slices were washed in tap water for threehours, and then immersed in 50% ethanol for a weekwith daily agitation, to wash out the embalmingchemicals. This step was repeated twice, using fresh50% ethanol. This was followed by washing the slicesin running tap water for one hour.DehydrationThe Mulligan-stained (see below for method) andunstained slices were pre-cooled to +4˚C thensubmerged in pre-cooled 100% acetone in an air-tightcontainer at -30˚C. The volume of acetone used was10 times that of the tissue to be dehydrated. Theacetone was replaced weekly for three weeks. Theconcentration of acetone was monitored with anacetonometer until it stabilised at 99% for verification ofcomplete dehydration.ImpregnationThe dehydrated brain slices were immersed inBiodur TM S10/S3 (100:1). The S10/S3 mixture hadbeen pre-cooled to -30° C. The immersed slices werethen placed in the vacuum chamber to equilibrate for24 hours at room temperature (15 - 18°C). After 24hours the vacuum pump was started and the pressurewas reduced gradually. At every point where a fewacetone 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 andbypass valves to permit stabilization of the pressure atcertain levels. The pressure was not lowered ifbubbles were still rising. In this way, the pressure wasgradually reduced over a period of approximately 9days until the final pressure of ~5 mm Hg was reached.The change in pressure was measured with ananalogue pressure gauge installed with theimpregnation chamber. The pressure in the chamberwas lowered by approximately one third per day. Carewas taken to ensure that during impregnation the levelof silicone was maintained no more than 2-4 cm higherthan the level of brain slices in order to facilitatevaporisation of acetone. The brain slices were thenallowed to equilibrate for 24 hours before removal. TheS10/S3 mixture was stored in the freezer at -30°Cwhen not in use to delay thickening of the mixture andto allow reuse.CuringThe slices were removed from the polymer and drainedon a paper towel for 2-3 days. The slices were placedin a sealed plastic tank containing Biodur TM S6 whichwas placed in a glass Petri dish at room temperaturefor 2-3 weeks to crosslink the polymer chains. The S6was replenished as necessary.Mulligan stainingThe staining method was derived from Tompsett’smodified Mulligan-staining procedure (Tompsett,1956). In this procedure the timing is critical as thewhite 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 acidand 100 ml distilled water) at 60˚C for 5 minutes; washwith 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 minutesfollowed by overnight washing in running water.Statistical Analysis:A paired T-test (one-tailed) was performed on thelengths and widths of 54 brain slices beforedehydration and after impregnation using Graph PadPrism 4 software.

Brain Impregnation 5Results:The mean shrinkage was 6.99 % (length) and 6.19%(width) (P .01 (ns) 6.99Width 11.15±0.36 10.46±0.37 > .01 (ns) 6.19(ns) – not significantand the stain did not fade with time on exposure todaylight (Figs 3 – 7). After fixation, the total timetaken was 8 weeks and 2 days (sectioning andstaining - 2 days, dehydration - 3 weeks, forcedimpregnation - average 9 days to impregnate and 2days to equilibrate, curing – average 2 days to drainand 3 weeks to cure, and tagging - 1 day).Discussion:We have found that using low-temperaturedehydration and room-temperature impregnation ofbrain slices gives very good results with BiodurS10/S3. Shrinkage was comparable to that achievedduring low-temperature impregnation, and theMulligan’s staining was robust enough to withstanddehydration and impregnation. Suriyaprapadilok andWithyachumnarnkul (1997) have previously performedplastination of Mulligan-stained thin brain slices (withthickness 4-6 mm) which were later framed withplastic plates. We have found that thicker slices(approximately 1 cm) can tolerate handling withoutframing (as suggested by Ulfig, 1990; Ulfig andWuttke, 1990). Furthermore, in wet brain specimens,Mulligan’s stain soon fades unless kept in the dark(Baeres and Møller, 2001) whereas the plastinatedslices retain their color and have thus far shown notendency to fade.Shrinkage was approximately 7% from a combinationof low temperature dehydration and room temperatureimpregnation, with and without stain, which wasaround 3% less than Suriyaprapadilok andWithyachumnarnkul (1997) report for low temperatureimpregnation of brain slices. Therefore, itdemonstrates that, contrary to expectations, S10/S3impregnation of brain material can be accomplishedsatisfactorily at room temperature. In the currenteconomic climate, this protocol has the addedadvantage of reduced capital costs; however, it is vitalto use air-tight containers in the freezer to avoid therisk of explosion. It is necessary to maintain acetonevapor below zero degrees Fahrenheit (-18° C) toprevent the vapors reaching flammable level(Baptista, Bellm, Plagge, Valigosky, 1992).To conclude, this report suggests an alternative,achievable and relatively low-cost method ofplastination of neuroanatomy specimens. Work is ongoingto investigate further applications of thisapproach.

6 Sagoo et al.Limitations:Shrinkage after dehydration was not observed toanalyze the effect of dehydration. Moreover, nomeasurements were taken to establish the shrinkageof grey matter in comparison to white; however, noobvious distortion was found.Acknowledgements:The authors would like to thank Ms. Lynda JanePhillipson for her help in staining the brain slices.Figure 3: Horizontal, Mulligan-stained section ofthe cerebrum. The basal ganglia are clearly seen,together with the internal capsule andcaudatolenticular bridges of gray matter.Figure 1: Horizontal, plastinated section of unstainedcerebellum showing dentate nucleiFigure 2: Horizontal, plastinated section of unstainedcerebrum showing caudate nuclei, internal capsules andlentiform nucleusFigure 4: Horizontal, Mulligan-stained section ofthe cerebellum and rostral part of the pons. Thedentate nucleus and the fibrae pontis transversaeare seen

Brain Impregnation 7Figure 5: Horizontal, Mulligan stained section ofcerebellum and midbrain showing substantianigra.Figure 7: Post-plastination: horizontal, Mulliganstained and plastinated section of cerebellumshowing dentate nuclei . The staining is unaffected bythe plastination process.References:• Adds PJ. 2008: A low-temperature dehydration/room-temperature impregnation protocol for braintissue using Biodur S10/S3. J Int Soc Plastination23:41.• Baeres FMM and Møller M. 2001: Plastination ofdissected brain specimens and Mulligan-stainedsections of the human brain. Eur J Morphol, 39(5):307-311.Figure 6: Pre-plastination: horizontal, Mulliganstained section of cerebellum showing dentatenuclei. (Staining timing and the quality of thespecimen is crucial to prevent the color leakinginto the white matter.)• Baeres FMM, Wamberg J, Møller M. 2001:Preparation of Plastinated Specimens of theHuman Central Nervous System for Use inTeaching of Medical and Dental Students. 10th IntConf Plast, Saint-Etienne, France, 2000. Abstractin J Int Soc Plastination 16: 34-35.• Baptista CAC, Bellm P, Plagge MS, Valigosky M.1992: The use of explosion proof freezers inplastination: Are they really necessary? J Int SocPlastination 6 (1): 34-37.• Côté ME, Veilleux F, Christin MJ, Fortin MJ, OlryR. 1995: Plastination: a new approach to theteaching of topographical anatomy. ChiropracticCentennial Foundation, Washington, DC, USA.• de Jong K and Henry RW. 2007: Siliconeplastination of biological tissue: cold temperaturetechnique – Biodur S10/S15 technique andproducts. J Int Soc Plastination 22:2-14.

8 Sagoo et al.• Holladay SD, Hudson LC. 1989: Use ofplastinated brains in teaching neuroanatomy atthe North Carolina State University, College ofVeterinary Medicine. J Int Soc Plastination 3 (1):15-17.• Lozanoff S, Lozanoff BK, Sora MC, RosenheimerJ, Keep MF, Tregear J, Saland L, Jacobs J, SaikiS, Alverson D. 2003: Anatomy and the accessgrid: exploiting plastinated brain sections for usein distributed medical education. Anat Rec 270B(1):30-37.• Olry R and Grondin G. 1994: Plastination inchiropractic teaching: critical analysis and place ofplastinated 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 withcomputer assisted learning modules for teachingveterinary neuroanatomy laboratories. J Int SocPlastination 5 (1): 16-19.• Suriyaprapadilok L and Withyachumnarnkul B.1997: Plastination of stained sections of thehuman brain: comparison between differentstaining methods. J Int Soc Plastination 12 (1):27-32.Int Soc Plastination 9 (1): 23-24.• Tompsett DH. 1956: Anatomical Techniques,Edinburgh and London: E.&S. Livingstone Ltd., p225-230.• Ulfig N. 1990: Staining of human fetal and adultbrain slices combined with subsequentplastination. J Int Soc Plastination 4 (1): 33-38.• Ulfig N, Wuttke M. 1990: Plastination of stainedsections of the human brain. Anat Anz 170 (5):309-312.• von Hagens G. 1986: Heidelberg plastinationfolder. Collection of all technical leaflets forplastination. 2 nd ed. Heidelberg, AnatomischeInstitut, Universitat, Hiedelberg .• von Hagens G, Tiedemann K and Kriz W. 1987:The current potential of plastination. Anat Embryol175:411-421.• Weiglein A. 1993: Plastinated brain-specimens inthe anatomical curriculum at Graz University. J IntSoc Plastination 7 (1): 3-7.• Weiglein AH. 1997a: Plastination in the neurosciences.Acta Anat 158: 6-9• Szarvas B, Szaras L, Groscurth P. 1995: Use ofplastinated brain sections for medical education. J

The Journal of Plastination 25(1): 9-11 (2013)ORIGINALRESEARCHARTICLEM. Üzel*Department ofAnatomy, CerrahpasaMedical Faculty,Istanbul UniversityTurkeyA.H. WeigleinInstitute of AnatomyMedical Universityof GrazGraz, AustriaP35 Plastination: Experiences with delayed impregnationABSTRACT: During an educational demonstration of the P35 technique, brain slices which hadbeen immersed in P35 resin and stored in a cold room (5° Celsius) for approximately two yearswere used. The resin was very viscous and it was difficult to remove the steel basket containingthe brain slices from the container of resin. There were technical difficulties during themanipulation of the slices: slices were brittle and fragile, filter paper spacers were stuck to thespecimens, curing had begun where the slice touched the grid and gel-like resin remnants werestuck 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: muzel@istanbul.edu.trORIGINAL RESEARCHIntroduction:The well-known classic P35 method is the most usedbasic technique for brain slice plastination. It is used toobtain semitransparent brain slices, and yields excellentgray-white matter distinction. Its main steps include:fixation with formaldehyde, slicing and placing onstainless steel grids, flushing & precooling to +5°C, twodehydration baths of 2-4 days, two immersion baths (1day each), forced impregnation, casting in double glasschambers, light curing, heat curing, and finishing(Weiglein, 1996; Weber et al., 2007). By completing allof these steps, the final product is a beautiful, durablebrain slice helpful for studying sectional anatomy of thebrain.In this case report, the optical quality of the end product,as well as the technical experience after a very longduration (two years instead of one day) in the secondimmersion bath of the P35 method are shared.Technical case reportDuring an educational demonstration of the P35technique at the Institute of Anatomy, Medical Universityof Graz, Graz, Austria, the brain slices that were to beused were placed in the second immersion bath (P35/A9mixture) approximately two years previously and werestored in the immersion bath during this period at +5°C.The normal methodology would be to commenceimpregnation after the slices have been in the secondbath for 1 day. Therefore, it was logical to continue theplastination process with 24 hours of forcedimpregnation of the brain slices which were submergedin this two-year-old resin-hardener (P35-A9) mixture.After the vacuum/impregnation was completed, theslices were removed from the basket/polymer andpositioned on the glass, and a double glass chamberwas constructed. The chambers were filled with freshresin mix (P35/A9), the slices were positioned with awire, and the slices were then cured using UV-A lightand heat.ObservationsProblems were observed from the beginning. Afterstorage and vacuum application, the resin from theimmersion bath was too viscous and it was very difficultto remove the steel basket of impregnated slices fromthe 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 oftenunited with the slices. With a lot of effort, most of thepapers were removed from the slices. Portions of theslices had started to cure, and gel-like resin remnantswere stuck to the metal grids (Fig. 2). Also, the grids leftmarks on some of the specimens (Fig. 3). Because oftheir fragility and the increased viscosity of the resin, itwas difficult to put the slices onto the glass forconstruction of the double glass chambers without

10 Üzel et al.breaking them. After assembling the double glasschambers, the chambers were filled with fresh P35/A9mixture and the slice position was adjusted with a wire.The hardening procedure was the usual UV-A light-heatcombination. Some finished specimens had excavationsand traces of filter paper on them. Despite the very longimmersion period (two years) and the problemsencountered, the final specimens were opticallysatisfactory (Fig. 4).DiscussionThe duration of the second immersion step of the P35method is normally one day. In this case, the specimenswere forgotten and unintentionally remained in thesecond immersion bath (in the cold room) for two years.Because of this very long second immersion period,problems were encountered during the remaining P35procedure. The problems (i.e. brittle, fragile, and partiallycured specimens and gel-like resin sticking on the metalgrids) encountered during the handling of the specimenswere probably due to the very long (two years) exposuretime of the specimens to acetone (von Hagens, 1986)and the increased resin viscosity. The acetone or lengthof time of the resin and catalyst being mixed likelycaused a reaction to change the polymer into a gel.In conclusion, the time for production of brain slices withthe P35 method can be extended up to several monthsby storing the slices in a cold immersion bath. However,too long a duration should be avoided because the slicesstart to cure at the regions where they are directly incontact with the steel grid, and viscosity of the resin mixincreases markedly. These findings suggest that in theP35 method, brain slices can stay in the secondimmersion bath for months without having any decreasein optical quality of the final specimen. On the otherhand, changes in the physical properties of the resinshould be expected which will result in mechanicalproblems that plastinators should be prepared to dealwith.Figure 1: P35 broken impregnated brain slices.Figure 2: Remnants of (P35/A9) resin mix on the metalgrids.Figure 3: Cured P35 brain slice.

Delayed impregnation 11References• von Hagens G, 1985: Heidelberg PlastinationFolder, 2nd English Edition: Collection of alltechnical leaflets for plastination, pg. 3:9-10. BiodurProducts, D-69126 Heidelberg, Germany.• Weiglein AH, 1996: Preparing and using S-10 andP-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: P35technique, J Int Soc Plastination 22: 50-58.Figure 4: Grid marks on finished specimen.

The Journal of Plastination 25(1): 12-17 (2013)ORIGINALRESEARCHARTICLEShawnda L. KumroAshton V. CrockerRandy L. Powell*Department ofBiological & HealthSciences, TexasA & M UniversityTexas, USAInjection Plastination: A Low-Tech, Inexpensive Methodfor Silicone Preservation of Small VertebratesABSTRACT: The plastination process using vacuum impregnation replaces tissue fluids withcurable polymers and results in dry, non-toxic specimens. We detail a method to produce highquality plastinated specimens using an injection impregnation process. This alternate, nonvacuummethod is very low-tech and has minimal start-up costs. We were able to successfullyuse this technique on small vertebrates ranging from 1 to 700 grams. The plastinated specimenswere life-like and the natural contours of the animals were maintained. Dissection revealedpolymer had penetrated throughout the viscera and deep muscles. In addition, internalmorphology including major muscle groups retained their shape with no apparent shrinkage.KEY WORDS: alternate process; inexpensive; injection; plastination; low-tech; smallvertebrates*Correspondence to: Randy Powell E-mail: randy.powell@tamuk.eduORIGINAL RESEARCHIntroduction:Plastination is a multi-step process in which tissue fluidsare replaced by curable polymers; e.g., silicone, epoxy,polyester (Bickley et al., 1981; Pashaei, 2010). Theresults are dry, durable specimens (entire/partial body orindividual organs) that look more life-like. Becauseplastinated specimens are non-toxic, they can be freelyhandled and examined without gloves. Consequently,the incorporation of plastinated specimens for teachingand research purposes has been adopted by medicaland 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 wereoriginally developed and documented by Gunther vonHagens over 30 years ago (von Hagens et al., 1987; seePashaei, 2010 for a more detailed history). While thebasic principles of plastination have remained somewhatconsistent since its initial introduction (i.e., fixation,dehydration, forced vacuum impregnation, and curing),there have been numerous developments, alternatetechniques, and improvements in the process. Thesedevelopments include: new polymers that permitambient (room) temperature plastination (Tianzhong etal., 1998; Henry, 2007; Raoof et al., 2007), progress insheet 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; Suriyaprapadilokand Withyachumnarnkul, 1997; An and Zhang, 1999;Mendez, 2008; Steinke et al., 2008a), and methods toproduce light-weight plastinated specimens (Henry andNel, 1993; Steinke et al., 2008b).Integrating plastinated specimens into a teachingenvironment has been shown to provide positiveeducational outcomes in the classroom (Latorre et al.,2011). The characteristics of plastinated specimens(life-like feel, no odor, no toxicity) allow students thebenefit of greater tactile interaction with the specimenswhich may result in an increased time spent with thematerial. Moreover, incorporation of plastinatedspecimens in teaching labs has been shown as anexcellent method to improve the “hands-on experience”(see Dawson et al., 1990, for a student survey ofresponses to plastinated specimens). Similarly, the useof plastinated specimens for exhibition and display inmuseums has been demonstrated as equally valuable.Unfortunately, start-up costs for the production ofplastinated specimens using traditional methods may beprohibitive for small labs and institutions with limitedbudgets and space. Furthermore, traditional plastinationmethods require special safety consideration andequipment, such as explosion hazards and adequateexhaust venting (Henry and Nel, 1993). Plastinatedspecimens (entire/partial body forms and individualorgans) are available for purchase from a fewcompanies; however, availability is limited to a smallnumber of species, and the specimens can be quiteexpensive.

Injection Plastination 13Vacuum impregnation, which allows for the penetrationand saturation of curable polymers into tissue, has beenthe hallmark step in plastination. While injection (i.e.,into deep tissues on large specimens or into specializedstructures) has been suggested to improve siliconeperfusion during vacuum impregnation (Henry and Nel,1993; Sivrev et al., 1997), it has not been reported foruse exclusively as a mechanism for polymerintroduction. Although large organisms may not be goodcandidates, smaller specimens, with less tissue mass,can be thoroughly saturated with polymer using injectionalone. Plastination of small organisms has received littleattention, and scant information has been reported (seeAsadi and Mahmodzadeh, 2004). Heretofore, no onehas reported on injection impregnation for small, wholebodyspecimens. Our goals were to produce high qualityplastinated specimens using an alternate, non-vacuumimpregnation 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 andcloacal injection sites with 21G-18G needles. Followinginjection, specimens were positioned for hardening andsubmerged in 10% formalin at room temperature for 24to 48 hours. After fixation, specimens were rinsedthoroughly in tap water to remove any excess, unboundformalin.Dehydration: Specimens were submerged in threeconsecutive 100% acetone baths at room temperaturewith a 1:10 specimen/acetone ratio. In addition, thespecimens were flushed through the previous injectionsites (using a syringe and 21G needle) with 100%acetone initially and between baths. Each bath lastedtwo to five days, and the specimen/acetone mixture wasagitated once each day. Acetone concentration wasmonitored with a hydrometer to determine theprogression of specimen dehydration. Dehydration wasconsidered satisfactory when the final acetone bathreached a specific gravity of 0.80.Silicone impregnation: Specimens were removed fromthe final acetone bath, and a silicone polymer mixturewas immediately applied to the entire surface of thespecimens using a brush. Following surface application,specimens were injected with silicone polymer mixture(via previous injection sites, penetrating into bodycavities and deep tissue) with 23G and 21G needles and1 to 5 ml syringes. Larger specimens that required moresilicone (e.g., large rodents), were injected using 50 mlsyringes and 18G or 15G needles. The amount ofpolymer injected into the specimens varied between 1and 25 ml per injection site depending on the size andspecies of the organism. The objective was to force inas much silicone as possible (until it began to leak out),while maintaining the original morphology of thespecimen. The silicone polymer mixture was composedof NCS10 and the cross-linker, NCS6 (North Carolinaproducts) at a ratio of 90:10. The NCS10/6 mixtureremains stable at room temperature (Henry, 2007) andhas a very low viscosity (easily injected through 23Gneedles with a 1 ml syringe). After the initial surfaceapplication and injection, specimens were wrapped in alayer of paper towels followed by an external layer of thinplastic wrap (to slow acetone evaporation).Specimens were inspected daily, injected with additionalNCS10/6 mixture (to replace evaporating acetone), andre-wrapped in clean paper towels and plastic wrap.Daily injection with the NCS10/6 mixture continued untilno acetone odor was detected on the paper towels orthe specimen (approx. 3 to 15 days). After acetoneevaporation was complete, the specimens were injectedwith NCS10/6/3 mixture of 90% NCS10/6 and 10%NCS3 (catalyst: North Carolina products), followed by athin surface application (with a small brush) of NCS3 onthe entire specimen. Mammalian specimens requiredblotting to remove excess NCS10/6 from the fur. Thecatalyst (NCS3) was then applied directly to the skin atvarious points using a 1ml plastic syringe to limitpolymerization in the fur. The specimens were coveredwith thin plastic wrap. Daily inspection and additionalNCS10/6/3 mixture injection was repeated as necessaryfor three to five days.Curing: Specimens were unwrapped, wiped clean of anysilicone polymer that may have leaked out, and placedon paper towels in a small aquarium. Approximately 1mlof NCS5 (chain extender: North Carolina products) wasplaced in an open (60 X15mm) cell culture dish and aglass cover was placed over the aquarium to maintainan atmosphere of NCS5 vapor. Each subsequent day,chain extender (NCS5) was replaced, the specimenswere wiped clean of any uncured silicone polymer thatmay have leaked, and specimens were inspected for anysigns of distortion (shrinkage of the tail, legs orabdominal areas). If necessary, additional NCS10/6/3polymer mixture was injected at or near any distortedareas. Excess polymer was periodically brushed fromthe pelage of mammals during the entire curing time.

14 Kumro et al.Specimens were maintained in the NCS5 vapor chamberfor final curing for 20 to 30 days.ResultsNumerous vertebrate species that included anassortment of amphibians, reptiles, and mammals wereplastinated. Specimens ranged in size from a 1.0g lizardto a 700g snake. The plastinated specimens wereodorless, their surfaces were dry to the touch, and theyexhibited a small degree of flexibility. In general, theplastinated specimens were life-like and the naturalcontours of the animals were maintained (Figs. 1, 2, 3).Gross dissection was performed on several specimens.Silicone had penetrated throughout the viscera and theinternal morphology was maintained, as major organsremained in approximate position (Figs. 4, 5). Theinternal structures were odorless and dry with negligibleshrinkage. Superficial, as well as deep musclesappeared thoroughly plastinated and major musclegroups retained their shape with little to no reduction insize (Fig. 6). Histological examination revealed polymerperfusion into the tissues of the internal organs andmuscles, and that polymer had filled the interstitial aswell as intracellular spaces.DiscussionWe argue that the success of the injection method isbased on the physical properties and interactions ofanimal integument, acetone, and silicone polymer.Injection of silicone polymer results in an increasedinternal pressure that is contained by the integument ofthe specimen. The semi-permeability of the integumentallows the passage of acetone molecules, while retainingthe larger, more viscous silicone molecules. The muchheavier silicone polymer (~27,000 g mol -1 ) readilydisplaces the lighter acetone molecules (58.08 g mol -1 ),causing movement of acetone through the integument,mouth, and cloaca. Acetone rapidly evaporates at roomtemperature; however, silicone polymer has anextremely low vapor pressure and negligibleevaporation. In addition, the degree of keratinization ofthe integument results in varying rates of evaporationdue to differences in permeability. These differencesnecessitate special measures for less keratinizedspecimens, such as amphibians, which require rapidcoating with NCS10/6 and wrapping, to slow acetoneevaporation. It should be emphasized that prematureevaporation of acetone causes pronounced shrinkageand stiffness in specimens and prevents adequateinjection of silicone polymer resulting in plastinatedspecimens of poorer quality.The results demonstrate that high-quality plastinatedspecimens can be produced using only injectionimpregnation. This alternate, non-vacuum process isvery low-tech and has minimal start-up costs. Injectionplastination will allow small laboratories on limitedbudgets the ability to process specimens for teachingcollections and display. Although we were able to usethis technique successfully on a 700g animal, there areno doubt limitations to the upper size of specimens thatcan be successfully processed with this method. Itshould be emphasized that to produce high-qualityplastinated specimens, it is important to be consistentwith daily inspection and processing of specimens;especially in controlling the rate of acetone evaporationand adequate injection of silicone polymer. As withtraditional plastination techniques, there is a smalllearning curve and some degree of skill and art involved.Our preliminary findings also indicate that small animalplastinated specimens are better teaching models intaxonomic lab courses and are preferred over traditionalfluid-preserved and dry prepared specimens.AcknowledgementsThe following people were helpful in providingsuggestions, helpful information, and/or feedback: Dr.Robert W. Henry, College of Veterinary Medicine, TheUniversity of Tennessee; Faculty, staff, and students ofmammalogy and vertebrate zoology from theDepartment of Biological and Health Sciences, TexasA&M University-Kingsville.

Injection Plastination 15life-like appearance.Figure 1: Various species of mammals, amphibians andreptiles were plastinated using the injection method. Ingeneral, 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, Kinosternonflavescens, f) Brazilian Free-tailed Bat, Tadaridabrasiliensis, g) Cane Toad, Rhinella marina, h) Texas Toad,Anaxyrus speciosus, i) Prairie Skink, Plestiodonseptentrionalis.Figure 4: Gulf Coast Toad (Bufo nebulifer). Thespecimen shows polymer penetration of the visceraand preservation of organs and internal morphology. a,b) liver, c) stomach, d, e) egg massFigure 2: Texas Coralsnake (Micrurus tener). Thisspecimen displayed excellent color retention.Figure 5: Hispid Cotton Rat (Sigmodon hispidus). Thespecimen shows polymer penetration of the viscera andpreservation of internal morphology. a) heart, b) liver, c)cecu, d) small intestine, e) spleen.Figure 3: Cane Toad (Rhinella marina). The naturalcontours of the animal were well maintained. Glasseyes were installed on this specimen to enhance its

16 Kumro et al.• Latorre R, Henry RW. 2007: Polyester plastination ofbiological tissue: P40 technique for body slices. J IntSoc Plastination 22: 69-77.• Latorre RM, García-Sanz MP, Moreno M,Hernández F, Gil F, López O, Ayala MD, Ramírez G,Vázquez JM, Arencibia A, Henry RW. 2011: Howuseful is plastination in learning anatomy? J Vet MedEd 34: 172-176.Figure 6. Hispid Cotton Rat (Sigmodon hispidus). Themuscles of this specimen are thoroughly plastinatedand muscle groups retained morphology.References• An P, Zhang M. 1999: A technique for preserving thesubarachnoid space and its contents in a naturalstate with different colours. J Int Soc Plastination14(1): 12-17.• Asadi MH, Mahmodzadeh A. 2004: Ascarisplastination through S10 techniques. J Int SocPlastination 19: 20-2.• Bickley HC, von Hagens G, Townsend, FM. 1981:An improved method for the preservation of teachingspecimens. Arch Pathol Lab Med 105: 674-6.• Cook P. 1996: Plastination as a clinically basedteaching aid at the University of Auckland. J Int SocPlastination 11: 22.• Correia JAP, Prinz RAD, Benevides de Freitas EC,Pezzi LHA. 1998: Labeling and storing plastinatedspecimens-An experience from UniversidadeFederal Do Rio De Janeiro. J Int Soc Plastination13(2): 17-20.• Dawson TP, James RS, Williams GT. 1990: Siliconeplastinated pathology specimens and their teachingpotential. J Pathol 162: 265-72.• Henry RW. 2007: Silicone plastination of biologicaltissue: Room temperature technique North Carolinatechnique and products. J Int Soc Plastination 22:26-30.• Henry RW, Nel PPC. 1993: Forced impregnation forthe 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: Sheetplastination with polyester: An alternative for alltissues. J Int Soc Plastination 19: 33-39.• Mansor O. 1996: Use of plastinated specimens in amedical school with a fully integrated curriculum. JInt Soc Plastination 11: 16-17.• Mendez BA, Romero RL, Trigo FJ, Henry RW,Candanosa AE. 2008: Evaluation of imidazole forcolor reactivation of pathological specimens ofdomestic animals. J Int Soc Plastination 23: 17-24.• Pashaei S. 2010: A brief review on the history,methods and applications of plastination. Int JMorphol 28: 1075-1079.• Peris KJ. 2000: Plastination technology forbiomedical research and studies in Kenya. J Int SocPlastination 15: 4-9.• Raoof A, Henry RW, Reed RB. 2007: Siliconeplastination of biological tissue: Room temperaturetechnique Dow/Corcoran technique and products.J Int Soc Plastination 22: 21-25.• Riepertinger A, Heuckendorf E. 1993: E 20 colorinjectionand plastination of the brain. J Int SocPlastination 7: 8-12.• Sivrev D, Kayriakov J, Trifonov Z, Djelebov D,Atanasov M. 1997: Combined plastination methodsfor preparation of improved ophthalmologic teachingmodels. J Int Soc Plastination 12(2): 12-14.• Sora MC. 2007: Epoxy plastination of biologicaltissue: E12 ultra-thin technique. J Int SocPlastination 22: 40-45.• Sora MC, Strobl B, Radu J. 2004: High temperatureE12 plastination to produce ultra-thin sheets. J IntSoc Plastination 19: 22-25.• Steinke H, Rabi S, Saito T. 2008a: Staining bodyslices before and after plastination. Eur J Anat 12:51-55.• Steinke H, Rabi S, Saito T, Sawutti A, Miyaki T, ItohM, Spanel-Borowski K. 2008b: Light-weightplastination. Ann Anat 190: 428-431.• Suriyaprapadilok L, Withyachumnarnkul B. 1997:Plastination of stained sections of the human brain:

Injection Plastination 17Comparison between different staining methods. JInt Soc Plastination 12(1): 27-32.• Tianzhong Z, Jingren L, Kerming Z. 1998:Plastination at room temperature. J Int SocPlastination 13(2): 21-25.• von Hagens G, Tiedemann K, Kriz W. 1987: Thecurrent potential of plastination. Anat Embryol 175:411–21.• Zhong ZT, Xuegui Y, Ling C, Jingren L. 2000: Thehistory of plastination in China. J Int Soc Plastination15: 25-29.

ORIGINALRESEARCHARTICLEA. Raoof*L. MarcheseA. MarcheseA. WischmeyerDivision of AnatomicalSciences, Office ofMedical EducationThe University ofMichigan Ann ArborUSAThe Journal of Plastination 25(1): 18-21 (2013)Demonstration of Systolic and Diastolic Phases of theCardiac Cycle in a Plastinated Human HeartABSTRACT: Plastination has enhanced the way students study human gross anatomy byproviding them with three-dimensional specimens that they can hold and manipulate. Thesespecimens 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 dissectedspecimens. At the University of Michigan Medical School, innovative approaches to enhance thequality of plastinated specimens have been implemented to demonstrate complex anatomicalfeatures. The heart is a particularly difficult organ for students to,visualise because of the uniquechanges it undergoes during systolic and diastolic phases of the cardiac cycle. The aim was todevelop a plastinated heart model that demonstrates how cardiac valves function during thesystolic and diastolic phases. Five hearts were collected from cadavers, dissected andplastinated. 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 contractedstate (systole) (2 out of 5 hearts) or in its relaxed state (diastole) (3 out of 5 hearts). A pilot surveywas administered to get students’ feedback on these plastinated models. The results indicatethat a majority of students favor this novel animated model as it displays both systolic anddiastolic 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 MichiganMedical School, 3740 Med. Sci. II Bldg., Ann Arbor, MI, 48109-5608, USA;E-mail: ameedr@umich.edu.ORIGINAL RESEARCHIntroduction:Materials and methods:The advent of the plastination process in gross anatomylaboratories provided students with a learning tool thattransforms two-dimensional textbook images into threedimensionalmodels that can be examined andmanipulated (von Hagens et al., 1987). One of the mostchallenging organs to understand for medical students isthe human heart, with its complex physiology and uniquetwo-stroke mechanism. Knowledge of the basicanatomy and physiology of the heart is an essentialcomponent of any medical or health science curriculum.Demonstrating how heart valves open and close duringsystolic and diastolic phases of a cardiac cycle is difficultwith cadaveric specimens. The aim of this study was todevelop a plastinated heart model that demonstrateshow cardiac valves function during the systolic anddiastolic phases. Plastinated models of hearts in bothsystole and diastole would allow students to identifyeach valve and its role in each phase of the heartbeat,as well as provide them with an intact three-dimensionaltool for learning the fine details of heart anatomy andhow its structure is related to its function.Five embalmed human heart specimens were harvestedfrom cadavers donated to the University of Michigan’sAnatomical Donations Program, age ranged between70-80 years (average 75). The hearts were kept in awater bath throughout the duration of the dissection topreserve their hydration and to enhance clot removalfrom the heart. At first the pericardium was removed, aswell as fat from around veins and arteries.To remove the superior aspect of the heart, thesemilunar cusps were located and an incision was madejust above the pulmonary and aortic valves. Thisincision was extended around the heart superior to thecoronary sulcus for a full transverse section of the atriaand aorta and pulmonary trunk. To mimic the contractedstate of the heart (systole), the mitral and tricuspidvalves were sutured closed while the aortic andpulmonary semilunar valves were positioned open usingcorks of a suitable size in two of the hearts.. The otherthree hearts were used to demonstrate the heart in itsrelaxed state (diastole). The mitral and tricuspid valveswere positioned open using corks and the aortic andpulmonary semilunar valves were sutured closed. Afterthe heart valves were prepared, the two portions of the

Systolic and Diastolic Phases 19heart were sutured to align and preserve the originalexternal anatomy (Raoof, 2001).After flushing the hearts with water, they weredehydrated in cold acetone. The prepared hearts wereplastinated using the room temperature method (Raoof2001, Raoof et al., 2007). The impregnation polymerwas mixed with 8% cross-linker (CR 22- Dow Corning).After forced impregnation, all sutures and corks wereremoved. A small brass hinge was attached with pinsand glue to keep the basal and apical portions of thespecimens as one unit. The hinges were attached withsutures, 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- DowCorning) (Raoof, 2001, Raoof et al., 2007).The impregnated and cured hearts were used byundergraduate students in Human Anatomy. The courseintroduced the students to the basic concepts ofsystemic anatomy and included visits to the lab wherepertinent plastinated specimens are displayed. Toevaluate student perception after using the plastinatedhearts, a pilot survey was developed and administeredto these 199 undergraduate students. Questionnaireswere distributed and students were asked to expresstheir opinions to three questions on a 5-point Likertscale. The questions were (1) “whether the heartspecimens have been useful in demonstrating structuralrelationships”, (2) “whether the specimens have beenbeneficial to learning the heart’s anatomy”, and (3)“whether the specimens have been beneficial tounderstanding function in correlation with structure”.Data from the survey was compiled and analyzed usingMicrosoft Excel.ResultsCareful preparation of the hearts permitted production ofheart specimens for demonstration of systole anddiastole of the cardiac cycle while preserving therelationship of superficial anatomy of the heart(Figure 3). The valves were misshaped during closureand opening but could be made anatomically correctprior to hardening/application of the catalyst.The value of these models for undergraduate study ofheart anatomy and physiology was assessed byconducting a pilot survey among undergraduateanatomy students. Students were allowed to spend timestudying and manipulating the heart, after which theywere asked if they felt a model of this type was beneficialto learning anatomy. Sixty out of 199 studentsresponded to the survey. A majority of students felt thatthese specimens were useful in demonstrating structuralrelationships (Figure 4) and were helpful to theirunderstanding of details of heart anatomy (Figure 5) andhow structure correlates with function (Figure 6).Discussion and ConclusionMetal hinges were used to create a simple animatedmodel of the heart that would permit students not only tostudy the external features of the heart, but also observethe internal structures and to gain a better understandingof the correlation between valve function and blood flowduring various stages of the cardiac cycle. Thesesystolic and diastolic models help students comprehendthe sequence of events that make up a cardiac cycle.Technically, it is essential to modify valve contours whichare misshaped due to the pressure of the cork and/orsuture to their final perceived anatomical shape aftersilicone impregnation and before adding the catalyst.During preparation of the cardiac valves an incision wasmade above the pulmonary and aortic valves to view thesemilunar valves. The incision around the heart superiorto the coronary sulcus permitted the view of theatrioventricular valves. After plastination the twoportions of the heart were sutured to align and preservethe original external anatomy (Figure 1).Baptista and Conran, 1989, prepared the cardiac valvesprior to dehydration by anchoring the leaflets of the rightand left atrioventricular valves with small pieces ofmoistened cotton introduced through a small incision inthe right and left ventricles. Small pieces of cotton werealso positioned in the cusps of the semilunar valves.After plastination, the incision was widened reviewingthe internal structures of the ventricles such as chordaetendineae and papillary muscles, and their relationshipwith the cardiac valves. Gomez et al., 2011, in anattempt to correlate plastinated heart slices withechocardiographic images, used 13 dog hearts fixed bydilation. The plastinated slices corresponded accuratelywith the echocardiographic images revealing the internalanatomy with great details. The dilated cavities of theplastinates (due to fixation) made comparison withechocardiographic images of systole difficult.A current project will further enhance understanding ofheart anatomy. The heart is being sectioned along asagittal plane through the septum and a second hingeattached, giving another view into the heart. Hinges arerather obtrusive, therefore, magnets will be used insteadof hinges.

20 Raoof et al.Figure 4: Student response, pilot survey, question 1 - whetherthe heart specimens have been useful in demonstratingstructural relationships.Figure 1: Left specimen: heart sutured at the pulmonarytrunk. Right specimen: base of the heart transected fromthe apex.Figure 5: Student response, pilot survey, question 2 - whetherthe specimens have been beneficial to learning the heart’sanatomy.Figure 2: Transected human heart with hingeFigure 6: Student response, pilot survey, question 3 - whetherthe specimens have been beneficial to understanding functionin correlation with structure.Figure 3: Plastinated human heart displays the systolic (left)and diastolic (right) forms.

Systolic and Diastolic Phases 21References• Baptista, C.A.C. and Conran P.B.1989: Plastinationof the heart: Preparation for the study of the cardiacvalves. J Int. Soc. Plastination 3: 3-7• Gomez A., Del Palacio J. F., Latorre R., Henry R.W., Sarria R., Arbors O. L. 2011: Plastinated heartslices aid echocardiographic interpretation in thedog. Vet Radiol Ultrasound 53 (2): 197-203• Raoof A, Henry RW, Reed RB 2007: Siliconeplastination of biological tissue: room temperatureplastination technique - Dow TM /Corcoran techniqueand products. J Int Soc Plastination. 22: 21-25.• von Hagens G, Tiedemann K, Kriz W. 1987: Thecurrent potential of plastination. Anat Embryol175(4): 411-421.• Raoof, A. 2001: Using a room-temperatureplastination technique in assessing prenatalchanges in the human spinal cord. J Int SocPlastination 16: 5-8

ORIGINALRESEARCHARTICLEMircea-ConstantinSora*Radu JilavuPlastinationLaboratoryCenter forAnatomy and CellBiologyMedical Universityof Vienna, AustriaPetru MatuszAnatomical DepartmentUniversity of Medicineand Pharmacy "VictorBabes" Timisoara,RomaniaThe Journal of Plastination 25(1): 22-27 (2013)Three dimensional reconstruction of a female pelvis usingplastinated cross-sections - Using Plastination for 3DReconstructionABSTRACT: In this study, a three-dimensional (3-D) model of the pelvis was built based on thinslice plastination cross-sections of the adult female pelvis and 3-D reconstruction technology. Afemale pelvis was obtained, MRI scanned, plastinated, sectioned and subjected to 3-Dcomputerized reconstruction using the WinSURF modeling system (SURFdriver Software).Qualitative observations revealed that the morphological features of the model were consistentwith those displayed by typical cadaveric specimens, whilst closer morphometric analysisindicated that the model did not significantly differ from a sample of cadaveric specimens. Thisconclusion proves that the utilization of plastinates for generating tissue sections can besuccessfully integrated into 3-D computerized modeling . A better understanding of pelvic flooranatomy is relevant to gynaecologists, radiologists, surgeons, urologists, physical therapists andall professionals who take care of women with pelvic floor dysfunction. The objective of this studywas to describe the method of developing this computerized model of the human female pelvisusing plastinated slices. It is a method which could be applied to reconstruct any desired regionof 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 BiologyPlastination and Topographic Anatomy, Währingerstr. 13/ 3 A-1090 Wien, AustriaE-mail: mircea-constantin.sora@meduniwien.ac.atORIGINAL RESEARCHIntroduction:The pelvic floor has a complex spatial structure,knowledge of which is essential when assessingpathologies in this area (Contouris,1988; Wiliams,1995).Women, for the most part, undergo pelvic floor examinationsfor urinary incontinence or prolapse of theinternal genitalia or of the urinary bladder (Goodrich,1993; Lienemann, 1997). The advent of lateralurethrocystography (UCG) and colpocystourethrographyhas made it possible to visualize and objectively assesspathological changes in the urinary bladder and urethra,whereas pathological changes in the pelvic floor and ofthe supportive structures of connective tissue can at bestonly be evaluated indirectly. The pelvic floor musclesserve 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 anatomicaland morphological information provided by sheetplastination can be added to this MR image information,which has a poorer detail resolution.The improved properties of plastinated specimens aremainly accounted for by the superior qualities of curablepolymers. In the plastination technique, tissue water andlipids are replaced by cured polymers. The class ofpolymer used determines the mechanical (flexible orfirm) and optical (opaque or transparent) properties ofthe specimen. Plastinated specimens are dry, odorless,and durable; they even retain structural details down tothe histological level. Today, 30 years after itsintroduction (v. Hagens, 1977), plastination is beingapplied in more than 250 departments of anatomy,pathology, forensic science and biology all over theworld. In research, the technique of sheet plastinationallows the arrangement of all tissue-components to bestudied in their undisturbed context. This is of majorinterest in the borderline area between gross anatomyand histology with respect to muscular and connectivetissue patterns. Epoxy resins are used to producetransparent body or organ slices, which, for researchpurposes, allow the study of the topography of all body

Using Plastination for 3D Reconstruction 23structures in an uncollapsed and non-dislocated state.In addition, the specimens are useful in advancedtraining programs in sectional topography, residenttraining in CT and NMR (de Barros, 2001; Lazanoff,2003; Thomas, 2004).Pelvic floor dysfunction, which includes urinary and fecalincontinence as well as pelvic organ prolapse, is a highlyprevalent condition in women. Ten percent of all womenundergo at least one operation to treat pelvic floordysfunction during their lifetime (Mant, 1997). Theannual cost of treating urinary incontinence in Austriaalone is estimated at 48 million Euros annually.However, little is known about specific pelvic floorpathomorphology and even less about pathophysiologyas it relates to pelvic floor dysfunction. DeLancey et al.,(2003) used Magnetic Resonance Imaging (MRI) toinvestigate levator ani muscle damage, and Lien et al.,(2004) constructed a computerized model in order todetermine the stretch forces that exceed the forceswhich muscle tissue can usually sustain. In our study, athree-dimensional (3-D) model of the pelvis was built forthe first time based on thin slice plastination crosssectionsof the adult female pelvis and 3-Dreconstruction technology.In order to investigate this topic, we need to know theorientation of the levator ani muscle and the interactionof muscles, bones, connective tissue and pelvic organs.Sections of a plastinated pelvis can help us tounderstand the levator ani architecture, and as therelation of muscles, fascia, organs and bones can bestudied perfectly it is also well suited for a 3-Dcomputerized reconstruction. This offers the opportunityto compare the use of identical physical and virtualmodels for the development of a 3-D anatomicalcomputer model based system and its interactivemanipulation.Materials and methods:The female human pelvis used for this study wasremoved from a fresh unfixed cadaver. A largespecimen block, containing the pelvis, was cut into thedesired sized blocks for impregnation with the maximumsize of 250mm x 150mm x 150mm. High-field, highresolutionmagnetic resonance imaging provides highsoft-tissue contrast that allows imaging of the regularfemale pelvic floor anatomy to a level almost equal toplastinated macroscopic anatomic cross-sections. Inthis study macroscopic cadaveric cross-sections of theregular female pelvic floor anatomy are compared andcorrelated in-vitro with high resolution MR imaging of thesame orientation. The cadaveric pelvis block was cutand adjusted to fit the standard Bruker magneticresonance head coil (dimensions: 140x90x100 mm).Wooden markers were installed as landmarks to allowcorrect orientation and slice selection.The scanned pelvis block was frozen at -80°C for oneweek and then plastinated following the standard ultrathinE12 slice plastination method. (An and Zhang, 1999;Johnson, 2000; Lane, 1990; Sora, 2007 a).Freeze substitution is the standard dehydrationprocedure for plastination, and shrinkage is minimizedwhen cold acetone is used. The tissue block was placedinto a -25°C freezer and then submerged in cold (-25°C)technical quality 100% acetone for dehydration.Methylene chloride was used for the degreasing andthen impregnation was performed using the followingepoxy mixture (Biodur E12, Rathausstr.18, 69126Heidelberg, Germany): E12 (resin)/ E6 (hardener)/ E600(accelerator) (Sora et al., 2007a). Once impregnationhad been completed, the tissue block was removed fromthe vacuum chamber and inserted into a moldconstructed of Styrofoam and lined with polyethylenefoil. The mold containing the impregnated specimen andresin-mix was then placed in a 65°C oven for four daysto harden the resin-mix. The tissue/resin block wascooled to room temperature and the mold removed.Before sawing, three colored plastic sticks were placedinside the block as markers. Using a contact pointdiamond blade saw, Exact 310 CP (Exact ApparatebauGmbH, Norderstedt, Germany) the hardened E12 blockwas cut into 1.6 ± 0.26 mm slices. Between each tissueslice, the width of the saw blade (0.4 mm), was lost.Finally the caudal surfaces of the plastinated slices werescanned into a computer using an EPSON GT-10000+Color Image Scanner. In every scan we included a ruleras a calibration marker and used the UTHSCSA IMAGETOOL v.2.0 for Windows software (The University ofTexas Health Science Center in San Antonio) formeasurements. Scanned images of the tissue sliceswere 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 internusmuscle, 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 ameasuring tool to record height, width, and depthmeasurements after rendering the model. Minimumrequirements for the software are extremely modest andinclude a 200 MHz processor, Windows 2000, WindowsXP, 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 31/2” floppy or 100 MB Zip Drive, and a mouse orcompatible pointing device; all of which are included instandard PC equipment.ResultsSectional plastination showed the structures of the pelvicfloor muscles and their relationship to adjacentstructures with a resolution down to microscopic level(Fig. 1, 2). This method allowed assessment of thecourse of muscle fibers. Diagnostically adequatevisualization of the pelvic floor was achieved by MRimaging with a pixel size of 0.89 x 0.89 mm and softtissue contrast.By segmentation of the outlines, it was possible to createa 3-D model which consisted of the levator ani muscle,the vagina, the urethra, the rectum, the obturatorinternus muscles, and the pelvic bones. The anatomicalstructures of the pelvis could be easily identified and theborders could be traced rapidly and reliably. The thinplastinated slices displayed the nerves, muscles, vesselsand bones of the pelvic region distinctly . Once scannedand loaded into WinSURF, edge detection was used toquickly collect tissue borders or contours. Thegenerated 3-D pelvis model displays a morphologycorresponding qualitatively to the actual cadaverspecimen (Fig. 3).The reconstructed images appeared well-defined,especially the spatial positions and complicatedrelationships of contiguous structures of the femalepelvis. All reconstructed structures can be displayed ingroups or as a whole and interactively rotated in 3-Dspace. Various features such as transparency control,individual object selection, animation and a variety ofmanipulation modes facilitate visualization of thecomplex pelvic anatomy (Fig. 4).DiscussionThe pelvic floor has a complex spatial structure, of whichonly parts are visualized on sectional images (Fritsch,1995; Fritsch, 2004; Fröhlich, 1997;). It is, however,necessary to correctly relate the visualized part to theentire structure in order to properly assess pathologies.A 3-D model in which the positions of the imaging planeof interest are shown clearly improves the vividness ofdepiction. Additional information, such as the course ofmuscle fibers or connections of the muscles with eachother for a given imaging plane, was gained from theplastinated sections. In contrast to anatomicalpreparation, the structures and spatial relationships ofthe tissues were not altered by plastination .Thin plastinated slices of 1.6 mm are essential if anaccurate 3-D reconstruction is desired (Qiu, 2003; Sha,2001; Sora, 2002; Sora, 2007 b). Although the processof plastination extends the time and effort required togenerate images for analysis, considerable detail isprovided and the reconstructed pelvic model exhibits thebones and surrounding soft tissue. Various groups hadpreviously investigated the female pelvic floor bycomputed tomography (CT), magnetic resonance (MRI)or ultrasound (US), but none had evaluated it byplastination. Transparent body slices are used forresearch purposes because they allow the study of bodystructures in a non-collapsed and non-dislocated state.Sectional plastination anatomy also confers a majoradvantage since the decalcification of bony tissue is notnecessary, and spatial relationships are retainedbetween contiguous features of differing composition.Thus, topography between bones and contiguous softtissues is retained without additional chemicalmanipulation.Computer models and animations of anatomical featuresare becoming increasingly attractive as a means tocommunicate complex spatial relationships andconcepts effectively (Dev, 2002). Although manyeducational animations are based on artistic renderings(Gould, 2001), more recent applications use virtualrepresentations derived from actual cadaveric material(Neider, 2000; Lozanoff, 2003). A logical advantage ofthese models is that they provide a greater sense ofrealism, which increases the amount of perceivedinformation. In the future one can envision anatomylectures consisting of only 3-D models without 2-Dimages (Trelease, 2002).One of the standard interventions in the therapy ofurinary incontinence is the tension-free vaginal tape(TVT) intervention (de Leval, 2003). We used thereconstructed pelvis to simulate a TVT placement (Fig.5). This model will furthermore provide data to calculatestretch ratios and other biomechanical tissue properties.

Using Plastination for 3D Reconstruction 25The system described here relies on relativelyinexpensive hardware, including a scanner andcomputer. The WinSURF reconstruction package, fromSURFdriver Software © (surface reconstruction driver),was developed expressly for use in three-dimensionalanatomical reconstruction and is a simple icon-drivensystem (Moody and Lozanoff, 1998).MOI – Obturator internusR – RectumS – Pubic symphysisV – VaginaU – UrethraOne major problem that occurs with existing anatomicaldatabases is the low resolution for smaller anatomicalstructures. Plastination can provide a useful alternativefor generating anatomical databases. Moreover,plastinates are significantly easier to cut, stain, andhandle compared to fresh-frozen tissue, since they aresignificantly more durable due to the silicone infiltrate.Although the female pelvis reconstruction presentedhere did not appear to be affected by a loss ofinformation (due to tissue loss between slices), furthertesting will be required to examine this issue. Thecapability to reconstruct individual and combined imagesof the pelvic structures, view them from all surgicalangles, and allow for accurate measurement of theirspatial relationships provides important guidance forsurgeons. The reconstructed model can also be usedfor residency education, testing unusual surgicaltechniques and for the development of new surgicalapproaches. The 3-D model of the female pelvispresented in this paper provides a stereoscopic view tostudy the adjacent relationship and arrangement ofrespective pelvis sections.Figure 2: Plastinated cross-section at the level of the hipjoint.A – AcetabulumC – Coxal boneM – MarkerMOI - Obturator internusMLA – Levator aniR – RectumS – Pubic symphysisV - VaginaVU – Urinary bladderFigure 1: Transparent cross-section of the female pelvis.a. a plastinated cross-section and b. comparable MRIimage.G – Gluteus maximusFI – Ischiorectal fossaMLA – Levator aniMOE – Obturator externusFigure 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 aniMOI - Obturator internusR – RectumV – VaginaU – UrethraVU – Urinary bladderReferences• An PC, Zhang M. 1999: A technique for Preservingthe Subarachnoid Space and its Contents in aNatural State with Different Colours. J Int SocPlastination 14: 12-17.• Contouris N (1988) The human levator ani muscle.Advances in andrology. Carl Schirren Symposium,Diesbach Verlag, p 159-165.• de Barros N, Junqueira Rodrigues C, JunqueiraRodrigues A Jr, de Negri Germano MA, Guido CerriG, 2001: The value of teaching sectional anatomy toimprove CT scan interpretation. Clin Anat 14 (1):36-41.Figure 4: Front view of the 3-D reconstruction withtransparent bony structures.MLA – Levator aniMOI – Obturator internusR – RectumV – VaginaU – UrethraVU – Urinary bladder• DeLancey JO, Kearney R, Chou Q, Speights S,Binno S. 2003: The appearance of levator animuscle abnormalities in magnetic resonance imagesafter vaginal delivery. Obstet Gynecol 101(1):46-53.• Dev P, Montgomery K, Senger S, Heinrichs WL,Srivastava S, Waldron K. 2002: Simulated medicallearning environments on the internet. J Amer MedInformatics Assoc 9: 437-447.• Fritsch H, Hötzinger H. 1995: Tomographicalanatomy of the pelvis, visceral pelvic connectivetissue, and its compartments. Clin Anat 8(1): 17-24.• Fritsch H, Lienemann A, Brenner E, Ludwikowski B.2004: Clinical anatomy of the pelvic floor. Adv AnatEmbryol Cell Biol 175: III-IX, 1-64.• Fröhlich B, Hötzinger H, Fritsch H. 1997:Tomographical anatomy of the pelvis, pelvic floor,and related structures. Clin Anat 10(4): 223-30.Figure 5: Reconstructed pelvis with a tension-free vaginaltape (TVT), after removal of the right side of the bonystructures.MLA – Levator aniMOI – Obturator internusR – RectumTVT – Tension-free vaginal tapeV – VaginaU – UrethraVU – Urinary bladder• Goodrich MA, Webb MJ, King BF, Bampton AE,Campeau NG, Riederer SJ 1993: Magneticresonance imaging of pelvic floor relaxation:dynamic analysis and evaluation of patients beforeand after surgical repair. Gynecol Obstet 82: 883-891.• Gould D. 2001: The brachial plexus: developmentaland assessment of a computer based learning tool.Med Educ Online 6: 1-7.

Using Plastination for 3D Reconstruction 27• Johnson G, Zhang M, Barnett R. 2000: AComparison between epoxy resin slices andhistology sections in the study of spinal connectivetissue structure. J Int Soc Plastination 15(1): 10-13.• de Leval J. 2003: Novel surgical technique for thetreatment of female stress urinary incontinence:transobturator vaginal tape inside-out. Eur Urol44(6): 724-730.Lane A. 1990: Sectional anatomy: standardizedmethodology. J Int Soc Plastination 4: 16-22.• Lien KC, Mooney B, DeLancey JO, Ashton-Miller JA.2004: Levator ani muscle stretch induced bysimulated vaginal birth. Obstet Gynecol 103: 31-40.• Lienemann A, Anthuber C, Baron A, Kohz P, ReiserM. 1997: Dynamic MR colpocystorectographyassessing pelvic- floor descent. Eur Radiol 7: 1309-1317.• Lozanoff S, Lozanoff B K, Sora MC, RosenheimerJ,.Keep MF, J Tregear, Saland L, Jacobs J, Saiki S,Alverson D. 2003: Anatomy and the access grid:exploiting plastinated brain sections for use indistributed medical education. Anat Rec 270: 30-37.• Mant J, Painter R, Vessey M. 1997: Epidemiology ofgenital prolapse: observations from the OxfordFamily Planning Association study. Br J ObstetGynaecol 104(5): 579-85.• Moody D, Lozanoff S, 1998: SURFdriver a practicalcomputer program for generatingthree-dimensionalmodels of anatomical structures using a PowerMac.Clin Anat 11: 133• Neider GL, Scott JN, Anderson MD. 2000: UsingQuickTime virtual reality objects in computerassistedinstruction of gross anatomy: Yorick the VRskull. Clin Anat 13: 287-293.• Qiu MG, Zhang SX, Liu ZJ, Tan LW, Wang YS,Deng JH, Tang ZS. 2003: Plastination andcomputerized 3-D reconstruction of the temporalbone. Clin Anat 16: 300-303.• Sha Y, Zhang SX, Liu ZJ, Tan LW, Wu XY, Wan YS,Deng JH, Tang ZS. 2001: Computerized 3-Dreconstructionsof the ligaments of the lateral aspectof ankle and subtalar joints. Surg Radiol Anat 23:111-114.• Sora MC, Strobl B, Forster-Streffleur S, Staykov D.2002: Optic nerve compression analyzed by usingplastination. Surg Radiol Anat 24: 205-208.• Sora M.C., 2007 a: Epoxy plastination of biologicaltissue: E12 ultra-thin technique. J Int SocPlastination, 22: 40-45.• Sora M.C., Genser-Strobl B., Radu J., Lozanoff S.2007 b: Three-dimensional reconstruction of theankle by means of ultrathin slice plastination. ClinAnat. 20(2): 196-200.• Trelease RB. 2002: “Anatomical informatics:millenial perspectives on a newer frontier.” Anat Rec269: 224-235.• Williams PL 1995: Muscles of the pelvis. In: Gray'sanatomy, 38th edn. Churchill-Livingstone, London, p831-835.• von Hagens G 1977: Patents: DBPat 27 10 147(1978) Brit Pat 1 558 802 (1984) Brit Pat 22 082 041(1978) Belg Pat 863.949 (1978) RSA Pat 78/1330(1980) Austr. Pat 360 272 (1980) US Pat 4, 205.059(1981) US Pat 4, 244, 992 (1981) US Pat 4, 278,701 (1982) US-Pat 4, 320, 157:

17th International Conference on PlastinationSt. Petersburg, RussiaThe Journal of Plastination 25(1): 28 (2013)

The Journal of Plastination 25(1): 29 (2013)Journal of PlastinationInstructions for Authors(Revised January 2013)JOURNAL OF PLASTINATION is owned and controlledby the International Society for Plastination (ISP).Goals - The Journal of Plastination (ISSN 1090-2171) is to provide a medium for the publication of scientificpapers dealing with all aspects of plastination andpreservation of biological specimens.Submission GuidelinesAll manuscripts must be submitted to the Editorial Office viathe e-mail: carlos.baptista@utoledo.edu. If you experienceany problems or need further information, please contacteither Dr. Carlos Baptista, carlos.baptista@utoledo.edu, orDr Selcuk Tunali, tunali@hacettepe.edu.tr.Authors must have an e-mail address at which they may bereached.Necessary Files for Submission Include:• Cover letter• Manuscript (including references and figurelegends)• Table(s) (when appropriate)• Figure(s) (when appropriate)• Copyright Release Form (after acceptance)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 tablesubmissions: Microsoft Word• File formats appropriate for figure submissions:TIFF, JPEG (JPG) and EPSCategories of submissions:Articles published in Journal of Plastination are grouped intogeneral article types (listed below). Final designation of amanuscript’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• EditorialAcceptance of a submission implies the transfer ofcopyright from the authors to the publisher. It is the author'sresponsibility to obtain permission to reproduce illustrations,tables and figures from other publications.Copyright Transfer Form may be downloaded fromhttp://www.journal.plastination.org/downloads/copyright.pdf.After the form is completed and signed by all the authors, itshould be submitted to the Editorial Office(carlos.baptista@utoledo.edu) as a pdf or jpeg file via an e-mail attachment.Manuscript preparationCover LetterThe cover letter should include a statement of authorship,notification of conflicts of interest, ethical adherence, andany financial disclosures.Cover letters may be addressed to the Editor-in-Chief,Journal of Plastination.ManuscriptThe manuscript should consist of subdivisions in thefollowing sequence:Title PageAbstract with keywordsTextIntroductionMaterials and methodsResultsDiscussionReferencesFigure LegendsTitle PageThe 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 aseparate entity, with a superscript number that links it to theindividual author.For example:S. D. HOLLADAY 1 *, B. L. BLAYLOCK 2 and B. J. SMITH 11 Department of Biomedical Sciences and Pathobiology,Virginia Maryland Regional College of VeterinaryMedicine, Virginia Polytechnic Institute and StateUniversity, Blacksburg, VA 24061-0442, USA.2 College of Pharmacy and Health Sciences, University ofLouisiana at Monroe, Monroe, LA 71209, USA.• Corresponding Author’s name, address, telephoneand telefax numbers, and e-mail address.

The Journal of Plastination 25(1):30For example:*Correspondence to: Dr Shane D. HOLLADAY, Departmentof Biomedical Sciences and Pathobiology, VirginiaMaryland Regional College of Veterinary Medicine,Virginia Polytechnic Institute and State University,Blacksburg, VA 24061-0442, USA. Tel.: +001 404 7396403; Fax.: +001 404 739 6492; E-mail:journal93@plastina.orgIt is the corresponding author’s responsibility to notify theEditorial Office of changes of address. Only thecorresponding author should communicate with theEditorial office for matters regarding each manuscript.Abstract & Key Words:The abstract should be no longer than 250 words. It shouldcontain a description of the objectives, materials andmethods, results, and conclusions. The abstract shouldinclude a section on technique/technical development if thepaper is significantly technical in nature. The abstract mustbe written in complete sentences and be intelligible withoutreference to the rest of the paper. No references should beused in the abstract.On the same page, list, in alphabetical order, five KeyWords that reflect the content of the manuscript. Consult theMedical Subject Headings for appropriate key words. Keywords should be set in lower case (except for essentialcapitals), separated by a semicolon and bolded.References:• References to published works, abstracts and booksmust include all that are relevant and necessary to themanuscript.• Citations in the text should be in parentheses andlisted chronologically; e.g. (Bickley et al., 1981; vonHagens, 1985; Henry and Haynes, 1989) except when theauthors name is part of a sentence; e.g. "…von Hagens(1985) reported that…" When references are made to morethan one paper by the same author published in the sameyear, 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 listedalphabetically using abbreviated journal names according toIndex Medicus. Page numbers of the citation must beincluded.• Examples of the reference style are as follows:• For a journal article:Bickley HC, von Hagens G, Townsend FM. 1981: Animproved 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: HenryR, editor. An atlas and guide to the dissection of the pony,4 th 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 andnot be a substitute listing for the result section, and appearon a separate page at the end of the manuscript, followingthe list of references.• Legends must be numbered consecutively as theyfirst appear in the text.• All symbols or abbreviations appearing in anyfigure 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 otherthan that defining the data should be presented as footnotes.• Create tables using the table creating and editingfeature of Microsoft Word. Do not use Excel or comparablespreadsheet 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 mustinclude the table title and appropriate column heads.• Save each table in a separate word document fileand upload individually, like figures.• Do not embed tables within the body of themanuscript.Figures• All figures must be cited in the text and must havelegends.• Each figure should be attached as a separate fileand labeled with the appropriate number.• Figures should be created, saved and submitted aseither a TIFF, JPEG (JPG) or an EPS file.• Line drawings must have a resolution of at least1200 dpi, and electronic photographs, scanned images,radiographs, CT and MRI scans must have a resolution of atleast 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 (twocolumnwidth).• Magnification must be recorded and have a “scalebar” in the photo. Since reproduction of illustrations iscostly, authors should limit the number of figures to thosewhich adequately present the findings, and add to theunderstanding of the manuscript.• Figures that are submitted in color must bepublished in color. Authors are responsible for the costs ofany color reproductions. Contact the editor for details.