The official publication of the International Society for Plastination

The Journal of Plastination

I SSN 2 311 -77 61

Analysis of Radio Frequency

Identification Tagging of

Biological Specimens – p 6

Pitcher Plant Plastination:

Preserving Botanical Specimens

For Education And Display – p15

Updated Protocol for the Hoffen

P45 Sheet Plastination

Technique – p22

Tissue shrinkage after P45

plastination – p26

IN THIS ISSUE:

Volume 31 (2); December 2019

Pitcher Plant Before Plastination

The Journal of Plastination 31 (2):1 (2019)

The Journal of Plastination

ISSN 2311-7761 ISSN 2311-777X online The official publication of the International Society for Plastination

Editorial Board:

Rafael Latorre Murcia, Spain

Scott Lozanoff Honolulu, HI USA

Ameed Raoof. Ann Arbor, MI USA

Mircea-Constantin Sora Vienna, Austria

Hong Jin Sui Dalian, China

Carlos Baptista Toledo, OH USA

Philip J. Adds Editor-in-Chief Institute of Medical and Biomedical Education (Anatomy) St. George’s, University of London London, UK

Robert W. Henry Associate Editor Department of Comparative Medicine College of Veterinary Medicine Knoxville, Tennessee, USA

Selcuk Tunali Assistant Editor Department of Anatomy Hacettepe University Faculty of Medicine Ankara, Turkey

Executive Committee: Rafael Latorre, President Dmitry Starchik, Vice-President Selcuk Tunali, Secretary Carlos Baptista, Treasurer

Instructions for Authors

Manuscripts and figures intended for publication in The Journal of Plastination should be sent via e-mail attachment to: editor.plastination@gmail.com. Manuscript preparation guidelines are on the last four pages of this issue.

The Journal of Plastination 31 (2):1 (2019)

Journal of Plastination Volume 31 (2); December 2019

Contents

Letter from the President, Rafael Latorre 2

Letter from the Editor, Philip J. Adds 4

Analysis of Radio Frequency Identification Tagging of Biological Specimens Prior to Plastination; G.R. Vandezande, R.A. Wamble, J.A. Huggins, J.R. Kerfoot, and M.G. Bolyard

6

Pitcher Plant Plastination: Preserving Botanical Specimens For Education And Display; Michal R. Golos, Anne-Kristin Lenz, Rafael O. Moreno Tortolero, Sean Davis, Ulrike Bauer

15

Updated Protocol for the Hoffen P45 Sheet Plastination Technique; Okoye, Chukwuemeka Samuel, Hong-Jin Sui

22

Tissue shrinkage after P45 plastination; Okoye, Chukwuemeka Samuel, Dou Ya-Ru, Sui Hong-Jin

26

Instructions for Authors 31

The Journal of Plastination 31 (2):2 (2019)

LETTER FROM

THE PRESIDENT

Letter from the President of the International Society for Plastination

Dear Friends and Plastinators:

It is with great pleasure that I present to you Volume 31, Issue 2, of the Journal of

Plastination. I want to thank the authors of the papers who have chosen our journal to

publish their results. These articles allow us to know more details about specific

protocols with very interesting applications in our laboratories. I would like to thank

the reviewers for taking their time to review the manuscripts.

This issue presents four remarkable papers. The first paper, Analysis of Radio

Frequency Identification Tagging of biological specimens prior to plastination, from Dr.

Vandezande et al. shows how radio frequency identification tags are a feasible and

low-cost option for the identification and management of plastinated biological

specimens. This study demonstrates the possibility of embedding these tags in

advance to have the option of tracking and monitoring samples during the

plastination process. The second study, Pitcher plant plastination: preserving botanical

specimens for education and display, from Dr. Golos et al., contemplates one of the

most difficult applications, the plastination of plants. It offers a method to preserve

the trapping leaf of a carnivorous pitcher plant in its natural shape and coloration for

long-term display. The last two papers, from Dr. Okoye, are about P45 plastination

technique. One of them is an Updated protocol for the Hoffen P45 sheet plastination

technique. With this protocol the plastinated sections obtained are semi-transparent,

durable slices with a clear delineation of the tissue morphology including the

connective tissue. The other P45 paper, Tissue shrinkage after P45 plastination,

present physical properties of P45 plastinated specimens, with shrinkage between

6.39 and 19.86 %.

As most of you know, the XX International Conference on Plastination

(https://www.icp2020chile.com ) will be held in Temuco, Chile, next July. The host is

Prof. Nicolás Ottone and his team from Universidad de La Frontera. The dates are July

20-24th, 2020. As president of the ISP I would like you all to become actively involved

in this conference, sending communications and attending it personally. It will be a

great opportunity to share new experiences about innovation and to establish future

collaborations for the advancement of plastination. I hope I can meet all of you in

Temuco.

I should remind that to attend this Congress, the General Assembly of the ISP

approved three travel grants for postgraduate students, working in plastination, and

members of the International Society for Plastination.

Rafael Latorre, DVM, PhD

The Journal of Plastination 31(2):3 (2019)

I would like to welcome all new members of the International Society for Plastination

and to invite all of you to participate in the Journal of Plastination. Please, share with

us your results, your expertise in plastination and other anatomical techniques.

With best regards from Murcia, Spain

Rafael Latorre

The Journal of Plastination 31 (2):4 (2019)

LETTER FROM THE EDITOR

Scopus Focus: Indexing the Journal of Plastination

Dear Colleagues,

As you can see from the screenshot below, The Journal of Plastination is now indexed on

Scopus, Elsevier’s abstract and citation database, covering around 36,377 titles from

approximately 11,678 publishers. A search by ISSN (23117761) brings up seven records, from

The Journal of Plastination 31(1) (Fig. 1).

Figure 1. Screenshot of a search for ISSN 2317761 (The Journal of Plastination) on Scopus

In a survey of academic databases, Burnham (2006) concluded that Scopus was “easy to

navigate, even for the novice user”. The facility for searching both backwards and forwards

from a particular citation was considered to be particularly helpful to the researcher, and the

multidisciplinary aspect allows the researcher to easily search outside of his or her discipline.

This is a very important advance for the Journal, and means, of course, that articles

published in the Journal (from 2109 onwards, unfortunately it is not retroactive) should now

reach a much wider readership amongst academic researchers. It is hugely disappointing

that one can read scholarly articles on plastination in high-profile peer-reviewed journals, in

which The Journal of Plastination, (or its predecessor The Journal of the International Society

for Plastination), surely the gold standard source for papers on the techniques and research

potential of plastination, are rarely cited . A PubMed search for papers with ‘plastination’ in

either the title or abstract, published since 1st January 2018, yielded 5 papers: four beautiful

research papers, and one paper on anatomical learning resources in Korea (Chung and

Chung, 2018; Lui et al., 2018; Thorpe Lowis et al., 2018; Kumar et al., 2019; Xu et al., 2020).

What is striking is that out of a combined total of 145 references listed, the Journal of

Plastination (or its predecessor) are cited only three times. It is to be hoped that with

inclusion in Scopus will come greater visibility, and hence more citations, giving the Journal

the academic weight that it may appear to lack now.

Unfortunately, Scopus is not free, nor is it available at all institutions. We are currently

Philip J. Adds, MSc, FAS, FFIBMS, SFHEA

The Journal of Plastination 31(2):5 (2019)

investigating the possibility of the International Society for Plastination taking out a

subscription, so that members of the ISP can have access – and hopefully help to further the

reach of the Journal. The Journal is also currently listed in Google Scholar, though coverage

appears to be incomplete and rather random. An application for inclusion in the Web of

Science database has been submitted, and I hope to update readers in the next issue,

although, as we have seen, evaluation can be a very slow process. If we can get the journal

listed on Scopus and Web of Science, we will have a much stronger case for inclusion in

PubMed.

In this issue we present papers demonstrating the wide reach of plastination. Xu et al. and

Okoye et al. report on aspects of the new P45 sheet plastination method from Hoffen in

China; Golos et al. describe plastination of the pitcher plant – it is unusual to see reports of

plastination of non-animal specimens, so this paper is particularly welcome. We also publish

research from Vandezande et al. on a novel method of identifying specimens with radio

frequency identification tags, during and after, the plastination process. This application is

particularly relevant during batch plastination of human organs from different individuals, as

it is legal requirement in most countries to be able to accurately identify and track donated

tissues and organs. We look forward to further reports from their team.

With best wishes,

Philip J Adds Editor-in-Chief References

Burnham JF. 2006: Scopus database: a review. Biomed Digit Libr 3:1: https://doi.org/10.1186/1742-5581-3-1

Chung BS, Chung MS. 2018: Homepage to distribute the anatomy learning contents including Visible Korean products, comics, and books. Anat Cell Biol 51: 7–13. https://doi.org/10.5115/acb.2018.51.1.7

Kumar N, Solanki JB, Shil P, Patel DC, Meneka R, Chaurasia S. 2019: Dry preservation of Toxocara vitulorum by plastination technique. Vet World 12:1428-1433. doi: 10.14202/vetworld.2019.1428-1433. PMID: 31749577; PMCID: PMC6813614.

Liu P, Li C, Zheng N, Yuan X, Zhou Y, Chun P, Chi Y, Gilmore C, Yu S, Sui H. 2018: The myodural bridges' existence in the sperm whale. PLoS One 13: e0200260. doi: 10.1371/journal.pone.0200260. PMID: 29985953; PMCID: PMC6037366.

Thorpe Lowis CG, Xu Z, Zhang M. 2018: Visualisation of facet joint recesses of the cadaveric spine: a micro-CT and sheet plastination study. BMJ Open Sport Exerc Med 4:e000338. doi: 10.1136/bmjsem-2017-000338. PMID: 29527323; PMCID: PMC5841519.

Xu Z, Mei B, Liu M, Tu L, Zhang H, Zhang M. 2020: Fibrous configuration of the fascia iliaca compartment: An epoxy sheet plastination and confocal microscopy study. Sci Rep 10:1548. doi: 10.1038/s41598-020-58519-0. PMID: 32005916; PMCID: PMC6994512.

The Journal of Plastination 31(2): 6-14 (2019)

ORIGINAL RESEARCH

Analysis of Radio Frequency Identification Tagging of Biological Specimens Prior to Plastination

G.R. Vandezande

R.A. Wamble

J.A. Huggins

J.R. Kerfoot

M.G. Bolyard.

Department of Biology

Union University

Jackson, TN 38305, USA

ABSTRACT:

The utilization of plastinated specimens has increased significantly in anatomy

instruction, providing self-directed aids for student inquiry. Due to increasing access to

plastinated specimens, low-cost identification systems are being developed for the

monitoring of their usage, handling, and distribution. Radio frequency identification

(RFID) technology has been used in the healthcare field and recent plastination studies

for its automated identification and tracking of multiple artifacts. This study

demonstrates a streamlined tag system for biological specimens subjected to the S10

cold-temperature plastination process. Four commercially available RFID tag types

were selected and embedded in biological specimens, prior to the plastination process.

The results indicated that the four types of tags selected are reliable and have the ability

to sustain lengthy periods of time in the harsh plastination conditions. Embedding RFID

tags in varying tissue types represents a successful small-scale study for seamless

tracking of anatomical specimens. Comparison of the four RFID tag types reveals that

there was no significant difference in the composition and successful performance

following the plastination process. This study further demonstrates that RFID tags are a

feasible and low-cost option for the identification and management of plastinated

biological specimens.

KEY WORDS: anatomical specimens; healthcare field; radio frequency identification;

RFID tags; student inquiry * Correspondence to: Dr. Mark G. Bolyard, Department of Biology, Union University, Jackson, TN 38305, USA. Tel.: +1 731-661-6586; E-mail: mbolyard@uu.edu

Introduction

In response to the heightened demand for health care

professionals, the integration of enhanced anatomy

curricula has been implemented in North American

universities. Experiential learning within the anatomy

field has become increasingly difficult, due largely to the

inability to find and collect biological specimen resources

(Marks, 1996; Older, 2004; Lockwood and Roberts

2007). For decades, anatomists have turned to tissue

preservation to produce high quality, long lasting

specimens through plastination, which relies on the

physical replacement of water and lipids with curable

polymer. Although plastination requires chemicals to fix

the specimens and dehydrate the tissue, specimens do

not require long-term immersion or storage in chemicals.

Cadaveric remains are essential for academic instruction

and research, but can be costly and difficult to maintain.

Although plastination equipment and materials can also

be costly, its utilization allows for permanent

preservation of remains, in addition to alleviating the

residual costs of fixing additives or repurchasing (von

Hagens et. al., 1987). One challenge for plastination is

tracking and monitoring specimens during and after the

plastination process. Tracking specimens through the

plastination process can be difficult because of the

limited access during procedural steps, in addition to

plastinating indistinguishable specimens (e.g. organs

from several donated remains). Likewise, monitoring

specimen access following plastination is extremely

important due to plastination expenses, in addition to

ensuring the integrity and respectful handling of donated

remains (Schmitt et al., 2014). Plastinated specimens

Analysis of Specimen RF Identification Tagging - 7

RFID Tag Type Manufacturer Read Range

(Fixed Reader)

Read Range

(Handheld

Reader)

Material

Compatibility

Dimensions

OMNI-ID FIT 400P Omni-ID, Rochester, NY,

US

3.5 m (11.5 ft) 1.75 m (5.7 ft) Plastic Assets 17.6 x 7.1 x 4.1 mm

XERAFY DASH-IN

XS

Xerafy, Dallas, TX, US 2.0 m (6.6 ft) 1.5 m (5.0 ft) Metal Assets 12.3 x 3 x 2.2 mm

OMNI-ID EXO 200 Omni-ID, Rochester, NY,

US

2.0 m (6.6 ft) 1.0 m (3.28 ft) Plastic Assets 14.5 x 12 x 5.4 mm

OMNI-ID EXO 400P Omni-ID, Rochester, NY,

US

3.5 m (11.5 ft) 1.75 m (5.7 ft) Plastic Assets 23.5 x 13 x 6.9 mm

Table 1. RFID tag specifications

do not require storage containers and are safe to handle,

therefore, they can be misplaced, lost, or stolen, much

easier than other fixed specimens. Management of

specimens can be improved by utilizing radio frequency

identification (RFID) tracking systems, benefitting body

donation programs, universities with anatomical studies,

and professional health programs (Porzionato et al.,

2012). However, a streamlined system for classification

and tracking has not yet been established (Noël and

Connolly, 2016). A recent study successfully attached

over 300 RFID tags onto biological specimens after the

plastination process. Analysis of these tags revealed the

successful and economically feasible tracking system

tracking abilities of RFID tags (Noël and Connolly, 2016).

Tracking and management of plastinated specimens is

important not only after specimens have been prepared,

but from the time of harvesting. Having the ability to

track assets greatly reduces misidentification.

Implementation of RFID tagging can be done at the time

of organ harvesting or acquisition, however, it was not

known how these RFID tags would fare going through

the rigorous plastination process. RFID tags are

advantageous due to their small, inlaid integration that

protects them from harsh environments, and passive tag

ability requiring no battery source (Hanna and

Pantanowitz, 2015). RFID tags utilize a digitized bar

code, which allows multiple tags to be read

simultaneously without disruption of signal frequency or

directly scanning a visible barcode. A RFID reader sends

a high frequency radio wave powering the passive RFID

tag, activating its integrated circuit. Each tag stores an

electronic product code (EPC) within the integrated

circuit on a memory chip. The EPC number is emitted

from the RFID tag following activation and read by the

network-connected RFID reader. The reader

communicates between the tags and a computer

database, which inventories the EPCs for storage and

tracking purposes. Each RFID tag possesses a unique

EPC that is programmed into the memory chip by the

manufacturer. The factory EPC typically consists of a

unique twenty-four-digit code that is used for specific tag

identification. Standard factory EPCs come in a Hex

format which can only be coded in numbers zero through

nine and the character letters A-F. If a RFID tat

possesses “read and write” capability, the Hex format

may be altered to ASCII. The ASCII format utilizes the

numbers zero through nine as well as letters A-Z. The

RFID tags used for this study possessed the “read and

write” capability, which can be advantageous when

tracking large quantities of specimens.

Plastination requires documentation and close

examination of the specimens throughout the entire

process, particularly if similar tissues or organs from

different donated remains are used (Schmitt et al.,

2014). To overcome this difficulty, specimens in this

study had a unique RFID tag embedded in the tissue,

prior to the plastination process. The objectives of this

project were to successfully embed four unique RFID tag

types into twenty different specimens prior to the S10

plastination process, implement an RFID classification

system for the biological specimens by creating unique,

distinguishable EPCs, and assess the durability and

functionality of the four RFID tag types following

plastination.

Materials and Methods

The durability of four RFID tag types (Table 1) was

assessed following the S10 cold-temperature

plastination method outlined by DeJong and Henry

(2007). The TSL 1128 Bluetooth UHF RFID Reader and

commercially available RFID tags (Omni-ID Fit 400P,

Omni-ID Exo 400P, Omni-ID Exo 200, Xerafy Dash-IN

XS) were purchased from Atlas RFID Solutions

(Birmingham, Alabama, US 35203). Methods for

inserting tags are contingent on the specimens used, but

8 – Bolyard, et al

Figure 1. Insertion of an Omni-ID Exo 200 RFID tag into a Leopard frog (Rana sphenocephala) and scanning of the inserted tag: (a) making a small incision in the specimen, (b) inserting the RFID tag, (c) suturing the incision, (d) reading the tag with the TSL 1128

Bluetooth UHF RFID Reader.

for these experiments, tags were inserted following the

method demonstrated in Figure 1.

The TSL 1128 Bluetooth UHF RFID Reader was

selected to read the RFID tags, as it provides a cost-

effective way to read and write EPC UHF transponders.

The TSL 1128 is capable of communicating with a wide

variety of host Bluetooth devices, such as iOS or

Android phones/tablets, while being compatible with a

variety of database software. The operating frequency

range of 902-982 MHz was selected in order to

successfully activate and read the RFID tags under

study. The maximum read distance of this reader was 4

meters, which was well within the desired range.

Due to the harsh conditions, careful consideration was

given to deciding which RFID tags were most capable of

surviving the plastination process. The RFID tags were

required to operate uniformly in the range of 860 to 960

MHz and contain Alien H3 IC, providing large user

memory (96 EPC-bits, extensible to 480 bits) and

enhanced IC security. Manufacturer specification,

including thermal operating ranges (-20°C to 85°C),

thermal and chemical cycling durability, signal output,

overall size, and cost effectiveness were also

considered. Furthermore, the selected tags had a

maximum readable distance of less than 4 meters

(short-range RFID tags), which are optimal for the TSL

1128 UHF RFID Reader. Tags were tested individually

for performance and the ability to emit readable EPC’s

prior to embedding into the specimens to be plastinated.

Each RFID tag arrived with a unique EPC from the

manufacturer. The factory specific EPC consisted of a

unique twenty-four-digit code that is used for individual

tag identification. For logistic purposes, modification was

required to enable faster tracking and better

categorization of plastinated specimens. The TSL 1128

UHF RFID handheld reader and the RFID Tag Finder

iOS application (v 1.0.8.2449) from Technology

Solutions UK LTD was used to rewrite the factory EPC’s.

Utilizing the read and write capabilities of the TSL 1128,

the twenty-four-digit HEX code was converted to a

twelve-digit ASCII format. Specimens were given an

eight-digit abbreviation for classification. Subsequently,

all codex EPCs were used to track and categorize the

plastinated specimens.

The cold-temperature BiodurTM S10/S15 plastination

technique outlined by DeJong and Henry (2007) was

used in this study. A silicone polymer mix (impregnation

mixture) was previously prepared consisting of the S10

silicone polymer, S3 catalyst and chain extender. Mixing

prior to impregnation allows the silicone molecule

elongation to start, resulting in longer silicone chain

length and a more viscous impregnation mixture. The

impregnation mixture was stored in a freezer at -15° C to

retard chain elongation until the impregnation phase.

Embalmed (formalin- fixed) specimens were placed in 50

% ethanol for seven days, removing excess embalming

fluids prior to the plastination process (DeJong and

Henry, 2007).

After tag assessment and S10 cold-temperature

plastination preparations, 20 tags (5 of each type) were

embedded into 20 formalin-fixed specimens (Table 2).

Because the tags differ slightly in size, tags were

selected to match each specimen based on the size of

the specimen compared to the tag. After insertion of the

tags, several specimens required suturing to seal

openings. The specimens were then placed in a flushing

water bath for 7 days to remove any residual formalin

(DeJong and Henry, 2007).

The specimens were then placed in a chemical-resistant

receptacle and submerged in a 90% acetone bath. The

receptacle was placed in a freezer (-15° C) for 8 days.

An acetonometer was used to determine the acetone

Analysis of Specimen RF Identification Tagging - 9

Specimen Binomial Classification RFID Placement Site Abbreviation for Codex

Pigfish Orthopriatis chrysoptera Abdominal cavity PigFish1

Fetal Pig Sus scrofa domesticus Superior femoral region (Right) FetalPg1, FetalPg2

Axolotl Ambystoma mexicanum Abdominal cavity Axolotl1

Southern Flying Squirrel Glaucomys volans Superior femoral region (Right) SFSquir1

Bichir Polypetrus retropinnis Abdominal cavity Bichir01

Southern Brook Lamprey Ichthyomyzon gagei Abdominal cavity SBLampe1

Mole Salamander Ambystoma talpoideum Abdominal cavity MSalama1

Shrew Cryoptotis parva Superior femoral region (Right) Shrew001

Atlantic NeedleFish Strongyura marina Abdominal cavity ANeedle1

Long-tailed Weasel Mustela frenata Superior femoral region (Right) LtWease1

Grey Squirrel Sciursus carolinensis Superior femoral region (Right) GSquir01

Leopard Frog Rana sphenocephala Abdominal cavity LeoFrog1

White Footed Mouse Peromyscus leucopus Superior femoral region (Right) Wfmouse1

Mink Mustela vison Superior femoral region (Right) Mink0001

Eastern Box Turtle Terrapene carolina Superior to Right leg, Ventral to

Carapace

EBTurtl1, EBTurtl2

Alligator Alligator mississippiensis Superior femoral region (Right) Alligat1, Alligat2, Alligat3

Table 2. Specimens used for plastination, RFID tag placement sites, and edited Codex for each specimen. Codex abbreviations must be 8 alphanumeric characters.

concentrations for the next 45 days of specimen

submersion as the concentration was gradually

increased to 99% (DeJong and Henry, 2007). The

receptacle containing the specimens was removed from

the freezer and placed at room temperature for 6 days to

undergo the defatting process. The gradual increase in

solvent temperature initiates defatting, removing excess

fat/lipids, resulting in increased tissue permeability,

allowing for better polymer penetration and distribution,

and producing a more life-like and durable specimen.

Monitoring was conducted by observing the color of the

acetone solvent that the specimens were submerged in.

The transition in the color of the acetone from clear to

yellowish/brown color signified the defatting process is

complete (DeJong and Henry, 2007).

After the defatting process, specimens were immediately

placed in a freezer containing a vacuum chamber

designed to withstand one atmosphere decrease in

pressure. This chamber contained the S10 silicone

polymer, S3 catalyst and chain extender. Specimens

were left for one day to stabilize at -14° C. The vacuum

pump was turned on and the pressure was reduced to

22 cm Hg. The pressure was slowly lowered over the

next 14 days to 4 cm Hg, resulting in the removal of

acetone vapor from the specimens, allowing for the

forced impregnation of the S10 polymer mix. Pressure

was further reduced to 1 cm Hg (0.013 atm), resulting in

further gas removal, for the next 20 days. On day 36,

gas release ceased, signifying the completion of forced

impregnation. The pressure was slowly returned to

normal atmospheric pressure and left to equalize for 2

days until the specimens were removed (DeJong and

Henry, 2007).

The specimens were then placed out at room

temperature for 19 days, allowing excess impregnation

mix to drain, and silicone molecules to undergo chain

extension, linking their S3 extender portions together

during the pre-curing process. The pre-curing phase was

extended to

30 days to yield more pliable, flexible specimens. Finally,

specimens were placed in a chemical-resistant curing

chamber, containing a desiccant and the S6 cross-linker

during the gas curing phase, resulting in the connection

of adjacent silicone molecule chains (DeJong and Henry,

2007).

During the five-month plastination process, the RFID

tags were assessed for readability after the following

plastination steps: flushing, dehydration, impregnation,

and curing. Each specimen containing an RFID tag was

10 – Bolyard, et al

Tag Type Signal Strength

X Difference(cm) S difference(cm) t- Value V- value df P-Value

Omni Fit 400 100% -6.0 2.65 5.071 4 0.0071* 75% -11.0 7.11 3.461 4 0.0258* 50% -28.6 8.29 7.710 4 0.0015* Farthest 116.8 32.63 8.004 4 0.0013*

Omni Exo 200 100% 0.0 0.00 0.000 4 NA 75% 0.8 1.30 1.500 4 0.2652 50% -0.4 3.29 0.272 4 0.799 Farthest -11.8 16.30 1.619 4 0.1808

Omni Exo 400 100% -1.8 1.48 2.714 4 0.0533* 75% -1.2 3.11 0.862 4 0.4375 50% -8.6 16.02 1.2000 4 0.2963 Farthest -16.8 68.04 0.552 4 0.6103

Xerfy 100% 0.0 0.00 0.000 4 NA 75% -0.2 0.84 4.000 4 0.7728 50% -0.2 0.84 0.535 4 0.6213 Farthest -0.8 2.68 0.667 4 0.5415

Table 3. Statistical analyses of signal strength differences before and after plastination, including means

(�̅�difference), standard deviations (S), and sample statistics (paired samples t-test [t] or Wilcoxon signed- ranks test [v]) with associated P- less than 0.05.

scanned to ensure each tag would emit a readable EPC

to the TSL 1128 Bluetooth UHF RFID Reader (Fig. 1d).

RFID tag read distances were measured at 100%, 75%,

and 50% signal strength, as well as the farthest distance

to a readable signal, both before and after plastination.

Statistical analyses were performed on the mean

differences of the distances before and after plastination

for each RFID tag used, regardless of the sample tested.

Normally distributed data were analyzed using a paired

t-test, otherwise a Wilcoxon signed ranks test was used.

All analyses were performed at an -level of 0.05 using

R statistical software.

Results

Prior to insertion and plastination, all tags provided

readable EPCs at distances up to 160 mm, although the

Omni-ID tags (Fit 400P, Exo 400P, Exo 200) provided

readable EPCs at greater distances than the Xerafy

Dash-IN tags (see data in the Appendix). All twenty

RFID tags emitted an accurate and readable EPC

following flushing. Four of the Xerafy Dash-IN tags, and

one Omni-ID Fit 400P tag did not emit an accurate and

readable EPC following dehydration. However, following

impregnation and curing, all twenty RFID tags once

again emitted an accurate and readable EPC. After read

range capabilities were collected at 100%, 75%, and

50% signal strength, as well as the farthest distance to a

readable signal, average distances and standard

deviations were calculated (Table 3).

F.2 Omni-ID Fit 400P Tag Analysis

Differences in means before and after plastination were

significantly different among all read strengths, and for

the greatest distance to a readable signal (p <0.05;

Table 3), indicating that, in this study, plastination had a

significant impact on the function of these RFID tags.

However, even after plastination these tags had the

second highest mean distance to a readable signal

(Appendix), indicating that they were still very useful for

this application.

F.3 Xerafy-IN XS RFID Tag Analysis

No significant difference was detected in mean

differences across read strengths, indicating that the

plastination process did not have a significant impact on

the function of these tags (Table 3), although these tags

had the lowest mean distance to a readable signal of the

tags tested following plastination (Appendix).

F.4 Omni-ID Exo 200 RFID Tag Analysis

No significant difference was detected in mean

differences across read strengths, indicating that the

plastination process did not have a significant impact on

the function of these tags (Table 3), although these tags

had the second lowest mean distance to a readable

signal of the tags tested following plastination

(Appendix).

F.5 Omni-ID Exo 400P RFID Tag Analysis

Differences in distances at 100% signal strength

approached a statistically significant difference before

Analysis of Specimen RF Identification Tagging - 11

and after plastination (p = 0.0533; Table 3), and

additional testing may indicate significance, but no

significant difference was detected among any of the

other signal strengths. These tags had the highest mean

distance to a readable signal of the tags tested following

plastination (Appendix).

Discussion

This study demonstrates a cost-effective approach to

tagging specimens prior to the plastination process.

Tagging these specimens with RFID sensors prior to

plastination provides an efficient tracking system that

maintains correct identification and categorization

through the entire preservation process, and throughout

the life of the specimen. It is also more straightforward to

insert the tags prior to plastination rather than after the

process is complete. The twenty RFID tags emitted

readable EPCs after flushing. However, four of the

Xerafy Dash-IN XS and one of the Omni-ID Fit 400P

RFID tags did not produce EPCs following the

dehydration step. This was likely due to diminished read

range capabilities while being submerged in preservation

fluid, which was also observed by Noël and Connolly,

(2016). The four tags were unable to be scanned at

close enough distances for detection due to the solvent’s

corrosive effects on the scanner. In order to prevent

damage, and maintain the integrity of the handheld

scanner, plastinated samples were scanned at a

distance of 30 cm (1 ft). Although five tags were unable

to be read after defatting, all twenty RFID tags emitted

accurate and readable EPCs following impregnation and

curing. The 4 commercially-available RFID tag types

tested provide a standard for large-scale tagging of

biological specimens undergoing the plastination

process. The twenty RFID tags used in this study will

continue to be monitored for any long-term disruption or

malfunction after being exposed to the harsh conditions

of plastination.

With regard to the function of the four RFID tags relative

to each other, there are several factors to consider. First,

there is the loss of signal at certain points during the

plastination process when using the Xerafy Dash-IN XS

and the Omni-ID Fit 400P. Second, the Omni-ID Fit

400P showed statistically significant differences in

signals before and after plastination. Third, when

considering the differences in the mean of the farthest

signal detected after plastination, the order of distances

(from the Appendix) is Omni Exo 400 (77.8 cm), Omni

Fit 400 (49.4 cm), Omni Exo 200 (28.2 cm), and Xerafy

(8 cm). Therefore, selection of the appropriate RFID tag

will be based on particular applications of the

researchers.

Future investigation could explore optimal depths for

implantation of RFID tags, using multiple samples of the

same tissue, and integrating all four RFID types into the

same sample. These could allow for differences to be

calculated among specimens of varying tissues,

providing useful data for optimal tag choice regarding

individual tissues. Also, multiple RFID tag types

embedded in a single specimen could provide a much

closer comparative analysis between RFID tag types.

Due to RFID capabilities and technology, close proximity

of multiple tags does not have an effect on surrounding

RFID tag signals. Additional RFID tags, such as the Fit

220 HT (Atlas RFID), should also be evaluated. It is also

our hope to compare the effectiveness of this wider

range of RFID tags in cold- vs room-temperature

plastination systems, as each temperature system is

beneficial for different types of specimens.

As universities continue to enhance their anatomy

curricula to aid in the development of aspiring healthcare

providers (Marks, 1996; Older, 2004; Lockwood and

Roberts 2007) methodologies for specimen

management continue to be important. Demonstrating

the distinct characteristics of each anatomical specimen

continues to be pedagogically important. Plastination

provides a means to turn these valuable, perishable

specimens into non-perishing, reusable teaching tools.

These tissues are also accurate in terms of structure and

approximate in terms of color, are chemical free, odor

free, maintenance free, safe for handling, and are able to

retain integrity/clarity throughout the specimens

handling. Protecting donated remains and costly

anatomical specimens is essential (Schmitt et al., 2014).

Implementation of RFID tagging promotes proper

handling and care, due to its monitoring and tracking

capabilities, potentially saving plastinated resources as

well as funding for medical universities or programs.

With RFID technology, monitoring and tracking

difficulties become alleviated, allowing maximum

confidence that specimens will sustain the quality of

preservation (Wakefield, 2007). It may also be possible

to use RFID tags with greater information storage

capacity to provide educational information in addition to

identifiers. The data provided in this study demonstrates

the possibility of embedding RFID tags within various

biological tissues, categorizing plastinated specimens,

tracking and monitoring samples during the plastination

12 – Bolyard, et al

process, and managing the usage of valuable

plastinates following plastination. Furthermore, this study

promotes a stronger relationship with body donation and

medical programs. Plastination continues to serve as a

valuable tool for the future for anatomical research and

education, and can be enhanced through the

implementation of RFID technology.

Acknowledgements

The authors wish to acknowledge Tyler Lockard and the

Atlas RFID team for dedicating their time, resources, and

support throughout this project. We also appreciate the

work of Drs. Marc Lockett and Michael Schiebout in

editing Mr. Vandezande’s original Masters manuscript,

Dr. Micah Fern for assistance with photography, and Ms.

Anna Laura Livingston for assistance with manuscript

and table preparation.

References

DeJong K, Henry RW. 2007: Silicone plastination of

biological tissue: Cold-temperature technique BiodurTM

S10/S15 technique and products. J Int Soc Plastination

22:2-14.

Hanna MG, Pantanowitz L. 2015: Bar coding and

tracking in pathology. Surg Pathol Clin 8:123–135.

Lockwood AM, Roberts AM. 2007: The anatomy

demonstrator of the future: An examination of the role of

the medically-qualified anatomy demonstrator in the

context of tomorrow’s doctors and modernizing medical

careers. Clin Anat 20:455–459.

Marks SC Jr. 1996: Information technology, medical

education, and anatomy for the twenty- first century. Clin

Anat 9:343–348.

Noël G, PJC, Connolly CC. 2016: Monitoring the use of

anatomical teaching material using low-cost radio

frequency identification system: A comprehensive

assessment. Anat Sci Educ 9:197–202.

doi:10.1002/ase.1575.

Older J. 2004: Anatomy: A must for teaching the next

generation. Surgeon 2:79-90.

Porzionato A, Macchi V, Stecco C, Mazzi A, Rambaldo

A, Sarasin G, Parenti A, Scipioni A, De Caro R. 2012:

Quality management of body donation program at the

University of Padova. Anat Sci Educ 5:264–272.

Schmitt B, Wacker C, Ikemoto L, Meyers FJ, Pomeroy C.

2014: A transparent oversight policy for human

anatomical specimen management: The University of

California, Davis experience. Acad Med 89:410–414.

Von Hagens G, Tiedemann K, Kriz W. 1987: The current

potential of plastination. Anat Embryol (Berl) 175:411–

421.

Wakefield D. 2007: The future of medical museums:

Threatened but not extinct. Med J Aust 187:380–381.

Analysis of Specimen RF Identification Tagging - 13

Appendix

Omni-ID Fit 400P

Before Embedding

Specimen 100%(cm) 75%(cm) 50%(cm) Farthest Signal(cm)

U318PigFish1 8 20 50 180

U318FetalPg1 9 21 48 170

U318FetalPg2 7 11 44 158

U318Axolotl1 5 14 39 160

U318SFSquir1 8 17 40 163

Mean: 7.4 16.6 44.2 166.2

Following Plastination

U318Pigfish1 1 8 22 50

U318FetalPg1 0 0 6 10

U318FetalPg2 5 10 20 85

U318Axolotl1 0 4 10 60

U318SFSquir1 1 6 20 42

Mean: 1.4 5.6 15.6 49.4

Xerafy Dash-iN XS

Before Embedding

Specimen 100%(cm) 75%(cm) 50%(cm) Farthest Signal(cm)

U318Bichir01 0 0 2 8

U318SBLampe1 0 0 1 4

U318MSalama1 0 1 3 11

U318Shrew001 0 1 2 12

U318ANeedle1 0 0 2 9

Mean: 0 0.4 2 8.8

Following Plastination

U318Bichir01 0 1 3 10

U318SBLampe1 0 0 1 5

U318MSalama1 0 0 2 10

U318Shrew001 0 0 2 7

U318ANeedle1 0 0 1 8

Mean: 0 0.2 1.8 8

14 – Bolyard, et al

Omni-ID Exo 200

Before Embedding

Specimen 100%(cm) 75%(cm) 50%(cm) Farthest Signal (cm)

U318LtWease1 0 8 20 46

U318GSquir01 0 1 9 50

U318LeoFrog1 0 1 6 32

U318Wfmouse1 0 1 9 34

U318Mink0001 1 7 16 38

Mean: 0.2 3.6 12 40

Following Plastination

U318LtWease1 0 7 15 21

U318GSquir01 0 3 8 25

U318LeoFrog1 0 1 10 45

U318Wfmouse1 0 3 10 30

U318Mink0001 1 8 15 20

Mean: 0.2 4.4 11.6 28.2

Omni-ID Exo 400P

Before Embedding

Specimen 100%(cm) 75%(cm) 50%(cm) Farthest Signal(cm)

U318Alligat1 4 10 23 50

U318Alligat2 3 8 20 120

U318Alligat3 2 11 36 98

U318EBTurtl1 5 10 40 100

U318EBTurtl2 3 6 34 105

Mean: 3.4 9 30.6 94.6

Following Plastination

U318Alligat1 2 12 38 132

U318Alligat2 3 10 20 90

U318Alligat3 1 6 22 115

U318EBTurtl1 1 8 20 36

U318EBTurtl2 1 3 10 16

Mean: 1.6 7.8 22 77.8

The Journal of Plastination 31(2): 15-21 (2019)

ORIGINAL RESEARCH

Pitcher Plant Plastination: Preserving Botanical Specimens For Education And Display

Michal R. Golos1§*

Anne-Kristin Lenz1§*

R. O. Moreno Tortolero2

Sean Davis2

Ulrike Bauer1

1 School of Biological

Sciences, University of

Bristol, Bristol, UK

2 School of Chemistry,

University of Bristol, Bristol,

UK

§ These authors contri-

buted equally to this work.

ABSTRACT:

The lifelike preservation of three-dimensional plant material poses particular challenges,

and there is still no established method for it. The aim of the present study was to develop

a method to preserve the trapping leaf of a carnivorous pitcher plant in its natural shape

and coloration for long-term display in a public exhibition. Fresh pitchers were subjected to

one of the following preservation methods: freeze-drying, coating in PDMS, and

plastination. The resulting specimens were then compared against fresh and air-dried

material. Plastination was found to be superior to the other preservation methods in

yielding lifelike specimens for display. In particular, plastinates retained their shape better

and exhibited no obvious shrinkage. However, the process altered the coloration

significantly due to the loss of chlorophyll and mobilisation of anthocyanins (red–blue

pigments) during the dehydration and impregnation stages. Exposure of the finished

plastinated specimen to bright light also caused it to turn brown over a period of several

weeks. Further work is needed to refine the procedures for plastination of botanical

material. In particular, a method should be sought for fixing chlorophyll and other plant

pigments. These issues notwithstanding, plastination shows promise as a 3D preservation

method to supplement herbarium material and educational displays.

KEY WORDS: S10 method; room temperature plastination; freeze drying; PDMS coating; color retention

* Correspondence to: Michal R. Golos (michal.golos@bristol.ac.uk) and Anne-KristinLenz (anne-kristin.lenz@bristol.ac.uk), School of Biological Sciences, University of Bristol, 24 Tyndall Avenue, Bristol, BS8 1TQ, UK.

Introduction

The importance of lifelike biological specimens as

teaching tools has long been recognized. Preservation

methods vary depending on the type of specimen. While

vertebrates are typically stuffed (Péquignot, 2006) and

invertebrates are either air dried and pinned, preserved

in ethanol, or critical-point dried (Huber, 1998; Quicke et

al., 1999), plant material is generally pressed and dried

(Miller and Nyberg, 1955; MacFarlane, 1985). While this

method yields durable and easy-to-store specimens, it is

not well suited to highly three-dimensional organs such

as pitcher plant traps (e.g. genus Nepenthes and

Sarracenia) or complex flowers such as those of orchids

or pipe vines (genus Aristolochia). Nepenthes pitchers

are hollow, cup-shaped leaves (Cheek and Jebb, 2001;

Clarke, 2001) specialized to capture and digest

predominantly invertebrate prey (Moran and Clarke,

2010). Each pitcher is connected to the main leaf blade

via a thin tendril (Fig. 1A). Other distinctive features are

a pair of ventral ‘wings’, the collar-shaped pitcher rim

(peristome), and a lid shielding the opening from rain

(Clarke, 2001). Pressing dramatically alters these

distinctive geometries, thereby obscuring taxonomically

relevant morphological information, and potentially

producing herbarium specimens of limited use for the

identification of fresh plant material (Shivas, 1983; Lamb,

1989; Clarke and Moran, 2011).

Historically, these limitations spurred the production of

botanical models and replicas, as perhaps best

exemplified by the Blaschka “glass flowers” at Harvard,

which number more than 4000 and are known for their

craftsmanship and general scientific accuracy (Parke,

1983; McNally and Buschini, 1993). But even so, certain

botanical inaccuracies have been noted (Rossi-Wilcox,

2008, 2015). Wax was also used to create lifelike

replicas – a giant Nepenthes rajah pitcher was

‘preserved’ in this manner at the Royal Botanic Gardens,

Kew (Nelson, 1991). While these techniques have their

16 - Golos, et al

Figure 1. Pitchers of Nepenthes × hookeriana before and after preservation. A) Fresh pitcher. Labelled parts: Tendril, Lid, Peristome (pitcher rim), Wings, Pitcher Cup, Leaf Blade. B) Pitcher plastinated with the refined room temperature method. C) Final exhibit on display at Cambridge University Library, March 2019. D) Closeup of the same exhibit in September 2019. Note the color change of the pitcher and leaf which is likely due to bright light exposure, and the simulated pool of pitcher fluid with cockroach prey. Photographs A–C by MR Golos and A-K Lenz; D by Rachel Sawicki (Conservator,

Cambridge University Library).

merits, it is desirable to preserve real botanical material

wherever possible.

Previous efforts to preserve pitcher plant traps in their

three-dimensional form varied in outcome. Shivas (1983)

freeze-dried pitchers at −50°C under vacuum, achieving

good shape but only partial color retention. Stewart

(2008) employed a method which, though described as

such, was not true freeze-drying, as it involved placing

the pitcher in an ordinary freezer at standard pressure.

The method resulted in complete loss of coloration as

well as lid and wing curling, as the specimens thawed

and then dried following removal from the freezer. It was

also a lengthy process, taking four to eight weeks

(Stewart, 2008). Other methods include preservation

with glycerin and encasing in blocks of acrylic. However,

the former gives an unnaturally dark, oily appearance,

while latter is a challenging material to work with

(Stewart, 2008) and has the obvious disadvantage of

precluding close examination.

Shanos (1985) and Stewart (2008) described a method

of desiccating pitcher plants and other carnivorous

plants by covering them in silica gel in a sealed

container. The pressure exerted by the silica beads on

all sides maintained the shape and orientation of delicate

parts such as the lid and wings and prevented shriveling

in thin-walled species. But the dehydrated specimens

were very fragile (Shanos, 1985; Stewart, 2008) and

sensitive to moisture, and required carefully monitored

storage conditions. Protective coatings, while increasing

durability, will alter both texture and optical appearance.

The aim of the present study was to develop a

preservation method that could yield a lifelike, three-

dimensional pitcher plant specimen for display at an

exhibition of the Cambridge Philosophical Society. In

particular, we explored the potential of plastination, a

method that is unrivalled in the preservation of three-

dimensional vertebrate specimens, from organs to whole

bodies (von Hagens et al. 1987). Recently, this method

was successfully applied to mushrooms (Diz et al.,

2004a, 2004b; Looney and Henry, 2014; Henry et al.,

2016). Plant plastination, however, remains considerably

more obscure: while brief mentions are scattered across

various publications (Shama Sundar, 2010; Henry et al.,

2016) and even appear in the earliest patents related to

the technique (von Hagens, 1978, 1980), to our

knowledge it has not previously been detailed in the

academic literature.

Materials and Methods

Plant material

Pitcher plants (Nepenthes × hookeriana) were grown in

a climate-controlled chamber on a 12-hour photoperiod

with an average daytime temperature of 30°C and 60%

humidity, and a night-time temperature of 24°C and 80%

humidity. Initial trials to compare different preservation

methods were performed on eight pitchers, each

approximately 10 cm tall. For the final display, we

plastinated a pitcher of 24 cm height. Pitchers were cut

from plants, and subjected to one of the below-described

preservation methods within 30 minutes. The pitchers

were handled by their tendrils whenever possible as this

was deemed the part most resistant to mechanical

damage. One pitcher was left to air-dry at room

temperature for comparison.

Freeze-drying

A total of four pitchers were freeze-dried for 48 hours

using a FreeZone 1L benchtop freeze-drying system

(Labconco Corp.). Prior to freeze-drying, each pitcher

was subjected to one of the following treatments: 1)

submersion in liquid nitrogen (−196°C) until bubble

formation ceased; 2) submersion in 20% ethylene glycol

solution for several days followed by submersion in liquid

nitrogen; 3) freezing at −80°C; and 4) freezing at −20°C.

Preserving Botanical Specimens - 17

Table 1. Time, temperature and pressure parameters for

the refined plastination method for pitcher plants.

These temperatures were chosen as they are readily

achievable in common labs. The rationale behind rapid

cooling with liquid nitrogen was to produce amorphous

ice, thereby minimizing crystal formation and associated

tissue damage.

PDMS coating

A single freshly cut pitcher was submerged in absolute

ethanol and kept at −20°C to dehydrate the specimen.

After a week it was removed from the freezer,

submerged in polydimethylsiloxane (PDMS; Sigma

Aldrich) and, using a nylon string, hung upside down in

an oven at 40°C for three days to cure the coating.

Plastination

In total, four pitchers were plastinated. For the first, we

followed the room temperature method for plastination

as described by Henry (2007b) and Looney and Henry

(2014). This method comprises three main steps:

dehydration, impregnation, and curing. In order to

improve the result, we slightly modified this method for

the remaining three specimens by omitting the initial

acclimation step during the impregnation phase (Table

1). Both acetone (used in the standard method) and

ethanol were trialed as dehydrating fluids. While both

were found to mobilize green (chlorophyll) and red

pigments (anthocyanins) to some degree, acetone

appeared to have a less severe effect and was therefore

used for all further plastination work.

The first pitcher was dehydrated in 98% acetone at

−20°C for seven days, then transferred to a polymer mix

of ten parts S10 polymer and one part S6 cross-linker

(Biodur), and allowed to equilibrate overnight. It was

then placed in a vacuum chamber (Vacuum Oven

Digital; Fistreem International) with an attached

diaphragm vacuum pump (Rotavac Vario Pumping Unit;

Heidolph Instruments) and the pressure rapidly reduced

to 300 mbar. Over the following 48 hours the pressure

was incrementally decreased to 10 mbar, where it was

held for an additional 24 hours to ensure complete

replacement of the acetone by the polymer mix. After

that, the pressure was increased back to atmospheric

pressure within a minute before the pitcher was removed

from the polymer bath, hung upside down, and left to

drain for several hours.

The specimen was then sprayed with S3 catalyst

(Biodur) on its inner and outer surfaces and wrapped in

cling film to create a saturated environment in which the

vaporized catalyst could more efficiently diffuse into the

specimen. The catalyst treatment was repeated three

times over the course of five days. Paper towels were

used to wipe away excess catalyst from the outer

surface of the specimen.

When plastinating subsequent pitchers, the acclimation

step between dehydration and impregnation was

omitted, to improve the result for the thin plant material.

Further refinements to the curing step were made when

plastinating the large pitcher for the exhibition. The

catalyst was sprayed only on the inside of the pitcher as

well as on the undersides of the lid and leaf. Spraying

only one side of each surface in this way was sufficient

to initiate the desired chain reaction while minimizing the

exposed areas that might retain a liquid layer of excess

catalyst following curing. When wrapping in cling film,

particular care was taken not to deform the fragile lid and

wings; a loose roll of cling film was placed between the

wings to help maintain their shape. Finally, the large

pitcher was partially filled with polymer mix to simulate a

pool of pitcher fluid, complete with insect ‘prey’ (Fig. 1D).

This also helped maintain the structural integrity of the

exhibit upon curing.

Evaluation

The shape and color of the specimens resulting from

different preservation methods were monitored and

recorded over a total of four months (one month for the

exhibit pitcher). A qualitative assessment of the changes

was made by visual comparison against fresh pitchers.

At the end of this period, images of all specimens were

taken. All specimens were kept at 20°C in a room with

low-intensity artificial lights. Additionally, the exhibit

pitcher was monitored for the first few weeks of the

exhibition, during which time it was kept in a Perspex

18 - Golos, et al

Figure 2. Comparison of Nepenthes × hookeriana pitchers subjected to different preservation methods. A) Fresh pitcher. B–E) Freeze dried after (B) plunge-freezing in liquid nitrogen (−196°C), (C) submerging in ethylene glycol for several days and plunge-freezing in liquid nitrogen (−196°C), (D) freezing at −80°C, and (E) freezing at −20°C. F) Dehydrated in ethanol, then PDMS coated. G) Plastinated at room temperature following the original method. H) Plastinated at room temperature using our refined method (omitting the acclimation step after dehydration). I) Air dried without further treatment.

Photographs A–I by M. Golos and AK. Lenz.

Figure 3. A) Air-dried pitcher, showing shriveled tendril (top arrow) and wings (bottom arrow). Note the uneven surface of the pitcher body resulting from shrinkage, which nonetheless showed excellent color retention. B) Pitcher freeze dried at −80°C, showing buckling of peristome surface (top arrow; inset) and pitcher wing deformation (bottom arrow). While the pitcher body did not contract significantly, ‘bleeding’ of red pigments was obvious. C) PDMS-coated pitcher, showing severe drying artefacts affecting the peristome (top arrow) and tendril (bottom arrow). Color retention was very poor, though

the wings were surprisingly well preserved.

box and exposed to natural light (Fig. 1C), and examined

after the conclusion of the six-month exhibition (Fig. 1D).

Results

None of the tested methods was able to deliver a fully

lifelike result, but plastination with the aforementioned

refinements yielded the best specimen for display (Fig.

2). While some methods excelled at preserving color,

others retained the original shape better. In comparison

to a fresh sample (Fig. 2A), all drying approaches (air

drying and freeze drying with and without prior

treatment) resulted in some degree of shrinkage, and

specimens became brittle. Particularly obvious

deformations were curled-up wings, shrunken pitcher

rims and tendrils, and downward folding of the lid (Fig.

3).

Shrinkage was most severe in the air-dried pitcher (Figs.

2I, 3A), but color retention was better than for any other

method. The liquid nitrogen-treated, freeze-dried pitcher

(Fig. 2B) shrank less but developed multiple long cracks

across the pitcher cup. Color retention was good.

Ethylene glycol treatment (Fig. 2C) reduced the

cracking, but the resulting specimen gradually lost its

natural color and turned brown over the course of

several weeks. Pitchers frozen at −80°C (Fig. 2D) and

−20°C (Fig. 2E) showed significant shrinking of the

pitcher rim (Fig. 3B) but otherwise good shape retention,

particularly those treated at −80°C. Cracks did not occur

with this method; however, color retention was poorer

with increasing temperature, and red pigments leached

out into adjacent tissues.

PDMS coating (Fig. 2F) did not cause cracking and

achieved generally good shape retention. The delicate

wings were preserved well in their natural position;

however, the pitcher rim shrank more than in all other

preservation methods apart from air drying (Fig. 3C).

Coating with PDMS preserved the red pigments well, but

led to a complete loss of green color. In addition, the

lower half of the pitcher turned brown during the coating

process. In contrast to all other methods, the PDMS

coating also resulted in an unnaturally glossy

appearance.

Plastination using the standard room temperature

method (Fig. 2G) led to slight shrinkage, with the rim and

wings affected most severely. Omitting the overnight

acclimation phase after the transition from acetone to

polymer mix (Fig. 2H) eliminated this shrinkage almost

entirely. However, both plastination methods caused

severe discoloration. The green chlorophyll was lost

during dehydration in acetone, and the red pigments

moved within the tissue, which led to a red-colored

pitcher with increased saturation. In comparison to the

dried pitcher, the plastinated samples were less brittle,

but the wings in particular remained fragile as they

consist of very thin tissue. Stuffing cling film between the

Preserving Botanical Specimens - 19

wings prevented them from rolling inwards and improved

the result in the final exhibit (Fig. 1B).

Apart from the bleaching of green to near-white, the final

plastinated exhibit (Fig. 1B) appeared lifelike. However,

in contrast to previous plastinated pitchers that were

kept under low-light conditions at 20°C, the final exhibit

gradually turned brown over the six-month course of the

exhibition (Fig. 1D). This might be a result of the

exposure to stronger lighting or higher temperatures in

comparison to our storage conditions. To disentangle the

relative contributions of light and temperature, we kept

three plastinated pitchers in different environmental

conditions for a period of two weeks and recorded the

temperature continuously. The results suggest that high

light intensity rather than increased temperature

contributed to the discoloration observed in the exhibit.

Discussion

We successfully adapted and applied the plastination

method for tissue preservation to a three-dimensional

plant organ for the first time. Because the tissue of our

specimen was not more than a few millimeters thick, the

procedure was much faster than that for e.g. zoological

specimens or mushrooms (the room-temperature

process for the latter taking around three weeks; Looney

and Henry, 2014). The entire plastination process took

only two weeks. Our refined method yielded a

reasonably robust specimen with good shape retention

and intense red pigmentation. In terms of 3D shape

preservation, plastination proved superior to air drying,

freeze drying and PDMS coating. However, freeze

drying at both −196°C and −80°C achieved better color

preservation, especially of the green pigments. Both

plastination and freeze-drying therefore excelled in

different aspects of preservation. Since Nepenthes

pitcher coloration is highly variable in nature

(McPherson, 2009), exact color preservation was

deemed less important than natural shape retention in

an educational context, and the refined plastination

method was used for the final exhibit.

Several additional modifications could and should be

trialed to further improve the results of both methods.

For freeze drying, the use of a dry ice-acetone cooling

bath could help to maintain the temperature of the

sample stable at −78°C. This might help to eliminate

shrinkage. For plastination, fixation of the plant tissue in

glutaraldehyde or FAA (formaldehyde, acetic acid, and

ethanol) could be trialed. In addition, an acetone dilution

series could be tested as a milder alternative to direct

transfer into 98% acetone. Cold temperature plastination

(de Jong and Henry, 2007; Henry, 2007a; Looney and

Henry, 2014) could also lead to improved results.

For use in a public exhibition environment, light-induced

discoloration was a significant problem. The color

change only became apparent after two to three weeks.

Therefore, the procedure is not currently suitable for the

preparation of long-term exhibits with exposure to strong

natural or artificial lighting, and further research is

necessary to understand the effects of light and

temperature more thoroughly. Nevertheless, plastination

can be useful for educational purposes whenever it is

not possible to have a live plant, particularly for

structures whose three-dimensional shape is essential

for explaining their biological function, as for carnivorous

plant traps, kettle trap flowers, or flowers with moving

parts that play an essential role in their pollination

biology.

Plastinated pitchers could also potentially serve a

scientific function and be deposited alongside type

material in herbaria. Under adequate cool and dark

storage conditions, browning should be reduced or

prevented entirely. Plastinates could supplement

pressed specimens in the same way as “wet”, alcohol-

preserved specimens. By providing vital information

about 3D structure, plastinated specimens could

facilitate taxonomic identification and enhance functional

understanding. Future work should investigate the

applicability of this method to other plant species and

tissues.

Conclusion

Room-temperature plastination was successfully

adapted and used to preserve the three-dimensional

pitcher trap of a carnivorous plant (Nepenthes ×

hookeriana). The resulting specimen (Fig. 1B–D) was

displayed at the Cambridge Philosophical Society

exhibition (University Library, Cambridge) from March to

September 2019 (Dean, 2019). Color changes were

observed during both the dehydration and impregnation

steps of the plastination process, and significant

browning occurred post-curing, most likely as a result of

light exposure. Further refinements are needed to better

preserve the natural pigmentation. Other methods such

as drying, freeze-drying and PDMS coating obtained

inferior results with regard to shape preservation, but

freeze-drying at low temperatures (−196° C and −80° C)

yielded better color retention.

20 - Golos, et al

Acknowledgements

We thank Christopher Burgess, Rachel Sawicki and the

rest of the team at Cambridge University Library for their

tremendous work in putting together the pitcher plant

display, which was seen by approximately 56,000

visitors during the six months of the exhibition.

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context: a revised protocol for the preservation of

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de Jong K, Henry RW. 2007: Silicone plastination of

biological tissue: cold-temperature technique – Biodur™

S10/S15 technique and products. J Int Soc Plastination

22: 2–14.

Diz A, Martinez-Galisteo A, Berlango J, Conde-Pérez A.

2004a: Some aspects on fungi plastination. Murcia,

Spain: 12th Int Conf Plastination. (Abstract in J Int Soc

Plastination 19: 55–56.)

Diz A, Martinez-Galisteo A, Sanchez-Rodriguez M,

Conde-Pérez A. 2004b: Plastination of fungi as an aid in

teaching botanic classification. Murcia, Spain: 12th Int

Conf Plastination. (Abstract in J Int Soc Plastination 19:

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von Hagens G. 1978: DE patent 2710147: Preserving

human, animal or plant specimens - by impregnation

with a polymerisable plastic material without affecting

outline. European Patent Office. Available from:

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o?CC=DE&NR=2710147

von Hagens G. 1980: US patent 4205059: Animal and

vegetal tissues permanently preserved by synthetic resin

impregnation. Google Patents. Available from:

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von Hagens G, Tiedemann K, Kriz W. 1987: The current

potential of plastination. Anat Embryol (Berl) 175(4):

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Henry RW. 2007a: Silicone plastination of biological

tissue: cold temperature technique – North Carolina

technique and products. J Int Soc Plastination 22: 15–

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tissue: room-temperature technique – North Carolina

technique and products. J Int Soc Plastination 22: 26–

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The Journal of Plastination 31(2): 22-25 (2019)

TECHNICAL REPORT

Updated Protocol for the Hoffen P45 Sheet Plastination Technique

Okoye, Chukwuemeka

Samuel1

Hong-Jin Sui 1, 2

*

1 Department of Anatomy,

Dalian Medical University,

Dalian, P. R. China

2 Dalian Hoffen Bio-

Technique Co. Ltd., Dalian,

P. R. China

ABSTRACT:

Plastination is a method of biological preservation. Sheet plastination has relevance in

education and research. Both epoxy and polyester sheet plastination are currently used.

P45 is a polyester plastination technique. P45 sheet plastination produces an intact,

semi-transparent anatomical structure, with well highlighted connective tissue. The

technology comes at a low cost; it’s easy to produce, and easy to handle. We present

here an updated protocol for the P45 polyester plastination technique.

KEY WORDS: plastination; sheet plastination; polyester method; body slices; polyester resin; P45 technique * Correspondence to: Hong-Jin Sui. Tel.: +8613904287577; Email: suihj@hotmail.com

Introduction

Plastination is simply the process of replacing water and

lipid molecules from biological tissues with curable

polymer. Sheet plastination was developed for the

preservation of body slices, and has been extensively

used over the past decade as a teaching and research

tool (Thomas & Steinke, 2004; Sora & Gender-Strobl,

2007; Ottone et al., 2018). The E12 method is a sheet

plastination technique that uses Epoxy resin for

impregnation, while P35 and P40 sheet plastination

utilize polyester for impregnation (Latorre et al., 2004).

The E12, P35, and P40 sheet plastination methods were

invented by Gunther von Hagens (von Hagens et al.,

1987). The P45 sheet plastination method uses

polyester for impregnation, and was invented by Hong-

Jin Sui in 2003, and patented in China in 2006 (Sui,

2006). All of these sheet plastination techniques utilize

forced impregnation and casting between glass plates.

The techniques for the different sheet plastination

methods are similar. However, E12, P35 and P40 use a

flat chamber technique, while P45 uses an open vertical

chamber technique. Another difference in technique is in

the curing process: unlike the other methods, the P45

technique utilizes warm water for curing. This article

presents an update of the P45 slice plastination

technique.

Materials and Methods

The methods have been previously described by Gao et

al. (2006) and Sui & Henry (2007). The P45 plastination

technique can be divided into four basic steps: specimen

preparation and slicing, dehydration, impregnation, and

curing.

Specimen preparation and slicing

Equipment needed for specimen preparation and slicing:

bandsaw, grids and screen, wood or metal box of

appropriate size (see below).

Procedure for specimen preparation and slicing

Fresh or fixed tissue specimens can be processed.

Formalin fixation may reduce or eliminate any potential

biohazard, but may affect the color of the specimen.

Nevertheless, fixing the specimens can be done before

or after slicing.

The specimen is first frozen in an ultra-cold deep freeze

at -70° C for about two days to two weeks, depending on

the size of the specimen. Freezing the specimen allows

for easy and stable slicing of the specimen. Before

freezing, any undesirable tissue on the surface of the

specimen can be removed, for example trimming of the

hair, etc. After freezing, the specimen is further

enhanced for slicing by embedding it in a polyurethane

block.

Updated Protocol for Hoffan P45 Technique - 23

Procedure for embedding in polyurethane

An appropriate wooden or metal box is used. The box is

first lined with a plastic sheet for protection against the

polyurethane. The specimen is positioned appropriately

in the middle of the box, allowing a gap at the perimeter.

The cutting line should be marked, with the specimen

placed in the correct position, depending on the type of

section desired, i.e. coronal, sagittal, or horizontal, etc.

The polyurethane mixture is poured into the box, around

the specimen. The polyurethane is then allowed to rise,

foam and solidify. The specimen is then ready to be

sectioned.

Slicing Excess polyurethane is first trimmed off. The specimen

is then cut into smaller sections of about 2-3 mm

thickness, by first setting the guide stop of the bandsaw,

and then slicing the specimen. The tissue lost as tissue

dust between adjacent slices due to the blade thickness

is approximately 1 mm thick. A bandsaw with

appropriate blade thickness, and saw teeth size and

inter-distance should be utilized. We use a 3/4 teeth

blade, which means there is a larger tooth after each

three smaller one. The size of the selected blade

depends upon the size of the specimen: the bigger the

size of specimen, the bigger the teeth.

Slices are placed on a polyethylene grid, with a cotton

fiber screen. The grid should be acetone resistant. The

sawdust on the sliced sections is removed with stream of

gently running water, or by carefully scraping it off with a

blunt knife. It is not necessary to keep the slices frozen

during this stage. The grids with their washed slices are

stacked and then tied with twine to hold each stack as a

unit. The stack should be as small and portable as

possible for easy transfer during the dehydration step.

The stacked unit is then transferred into the first cold

acetone (-25° C) bath or a fixative bath.

Fixation and Bleaching This is an optional step. The slices can be fixed in

formaldehyde for one or two weeks at room temperature

by submerging the stacked unit in a 10% formalin bath.

After completing the fixation process, excess

formaldehyde is removed from the tissue slices by

rinsing in cold running water overnight. Slices can then

be immersed in 5% hydrogen peroxide (bleach)

overnight, to improve tissue color brightness and

transparency. Subsequently, excess bleach is washed

off by rinsing in running water for one hour or more.

Bleaching is suggested for any tissue that is dark in

color.

Dehydration

The dehydration process is performed by using the

freeze substitution method. Dehydration using acetone

also performs a second function of degreasing. A

preliminary step is to pre-cool the slices to 5° C in order

to prevent ice crystal formation, and minimize shrinkage

upon placement into the cold acetone. The stacked unit

of slices is first submerged in a bath of 100% acetone at

-25° C for one week. The stacked unit is then transferred

into another fresh acetone bath at -15° C for one week.

Finally, the stacked unit is transferred into a bath of

100% acetone at room temperature for another week.

From the first to the third change of acetone, the

concentration of acetone in the acetone bath is

monitored each day using an acetonometer. Once the

concentration of acetone remains stable for three

consecutive days, the stacked unit is ready to be moved

on to the next dehydrating solution.

If more transparency of fat is desired, the dehydrated

slices may be placed into methylene chloride

(dichloromethane) and monitored daily until the desired

degreasing is achieved.

Forced impregnation

Impregnation equipment: Vacuum chamber with a

transparent lid, vacuum pump, vacuum tubing and fine

adjustment needle-valves, vacuum gauge, and Bennert

mercury or digital manometer.

Forced impregnation in this step involves the

replacement of acetone with P45 polyester resin. This is

based on a difference of vapor pressure of acetone and

P45 polyester resin.

Constructing the casting chamber

The casting chamber used in this step is an open vertical

chamber. The top part of the chamber remains open

(while the bottom and two sides of the casting chamber

are clamped), and this chamber is then placed vertically

in the vacuum chamber, i.e. standing on its bottom end.

The open vertical chamber is constructed from two

plates of 5 mm tempered glass, flexible 4 mm latex

tubing, and several large fold-back clamps.

The tubing is placed between the two tempered glass

sheets, round the margins of the sheets (except for the

top edge). The glass sheets and tubing are clamped

together around the perimeter of the bottom and sides of

the glass using the fold-back clamps (Fig 1). A piece of

hardened P45 is placed at the bottom of the chamber to

24 – Okoye, Sui

Figure 1. An open vertical chamber with sagittal head slice ready for impregnation

Figure 2. A. Water bath with chambers undergoing curing; B. Close-up of an open vertical chamber with

a specimen undergoing curing in the water bath

prevent the specimen slice from touching the latex

tubing.

Preparing the P45 polyester mixture

The impregnation resin mixture is prepared as follows:

1000 ml P45 resin (Hoffen polyester, China) is mixed

with 10 g of P45A, 30 ml P45B, and 5 g of P45C. The

P45A and P45C are plasticizers, and P45B is a

hardener. The P45 resin is mixed just before

constructing the open vertical chamber, since it thickens

over time. Refrigeration of the P45 resin mixture will

retard its thickening, and can be used to store the mixed

resin for subsequent use.

Procedure for forced impregnation of the slices

The slices are removed from the final acetone bath and

placed into the open vertical chambers. The chambers

are then filled with the P45 polyester resin mixture

(Hoffen polyester, China) using a customized funnel.

After pouring the P45 resin mixture into the open vertical

chamber, air bubbles can be manually removed from the

casting chambers using a 1 mm stainless steel wire. The

open vertical chambers are then placed upright into the

vacuum chamber for impregnation at room temperature.

The absolute pressure in the vacuum chamber is

gradually decreased to 0 mmHg. Bubbles are slowly

released from the tissue slices. The vacuum is

maintained at 0 mmHg until bubbling ceases. The

bubbling activity occurring during this step can be

monitored through the transparent glass lid of the

vacuum chamber. The duration of the impregnation step

is usually around eight hours.

Curing

After impregnation, the pressure in the vacuum chamber

is released, and the casting chambers are transferred to

a curing chamber. The slices in the open vertical

chamber may need to be aligned properly, and any

residual trapped air bubbles should be removed. Both

these procedures are done with the 1 mm stainless wire.

The curing chamber is a warm water bath maintained at

40o C, with a small attached circulatory pump which

equilibrates the temperature of the water in the water

bath, since water is a poor conductor of heat. The

casting chambers are kept in the warm water bath for

three days (Fig 2).

Finishing

After curing, the open vertical chambers are removed

from the water bath and allowed to cool to room

temperature. The chamber is then dismantled by

removing the clamps, tubing, and glass. The P45 slice is

then taken out and wrapped with a plastic sheet or

lightweight foil, for protection against scratches. A

Updated Protocol for Hoffan P45 Technique - 25

bandsaw is used to trim off excess cured resin, and to

give the P45 slice the desired shape. A wood sander is

used to smooth the edges of the slice. Following

sanding, the slice is wrapped in a new plastic sheet or

foil, to avoid scratches on the surface of the slice. The

P45 plastinated sheet is now ready for use or storage.

Results

The P45 sections are semi-transparent, durable slices

with a clear delineation of the tissue morphology

including the connective tissues. Shrinkage is 2-8%

(Okoye et al., 2019), and the refractive index is 1.49.

Discussion

Most sheet plastination uses the flat chamber technique.

Instead of a flat chamber, the casting chamber utilized in

the P45 technique is an open vertical chamber, and it is

a potential time saver. In the other sheet plastination

techniques, such as E12, P35, and P40, the resin for

impregnation is replaced with fresh resin before curing

(Latorre & Henry, 2007; Weber et al., 2007; Ottone et al.,

2018). However, in the P45 technique, the same resin

and casting chamber that is used for impregnation is

also used for curing. This also means that the amount of

resin used in this technique is minimal. Furthermore,

curing is performed in a warm water bath, and this saves

on the amount of energy used in the plastination

process, reduces monitoring of the process, makes

complex equipment unnecessary, and saves time, since

no dismantling or mounting is required at the curing step.

Like other polyester techniques, the P45 sections are not

just embedded in the resin, but the slices are

incorporated as part of a single cured sheet of P45

polyester resin. Thus, the P45 technique is not

complicated, and requires less time and equipment. The

P45 sections show good anatomical details. The soft

tissues, connective tissue, and myofascial fibers of the

P45 plastinated specimens are clearly defined.

References

Gao H, Liu J, Yu S, Sui HJ. 2006: A new polyester

technique for sheet plastination. J Int Soc Plastination

21:7-10.

Latorre R, Henry RW. 2007: Polyester plastination of

biological tissue: P40 technique for body slices. J Int Soc

Plastination 22:69-77.

Okoye CS, Dou Y-R, Sui HJ. 2019: Tissue shrinkage

after P45 plastination. J Plastination 31(2): 25-33.

Ottone NE, Baptista CA, Latorre R, Bianchi HF, Del Sol

M, Fuentes R. 2018: E12 sheet plastination: techniques

and applications. Clin Anat, 31(5):742-756.

Sora M-C, Gender-Strobl B. 2007: The sectional

anatomy of the carpal tunnel and its related

neurovascular structures studied by using plastination.

Eur J Neurol 12:380-384.

Sui HJ. 2006: Sheet plastination of biological tissue and

its production method. China patent, Patent No. ZL 03 1

34109.8.

Sui HJ, Henry RW. 2007: Polyester plastination of

biological tissue: Hoffen P45 technique. J Int Soc

Plastination 22: 78-81.

Thomas M, Steinke H. 2004: Thin layer plastination of

the shoulder. Clin Sports Med Int 1:9-15.

von Hagens G, Tiedemann K, Kriz W. 1987. The current

potential of plastination. Anat Embryol 175:411-421.

The Journal of Plastination 31(2): 26-30 (2019)

ORIGINAL RESEARCH

Tissue shrinkage after P45 plastination

Okoye, Chukwuemeka

Samuel1

Dou Ya-Ru1

Sui Hong-Jin 1, 2

1Department of Anatomy,

Dalian Medical University,

Dalian, P. R. China

2Dalian Hoffen Bio-

Technique Co. Ltd., Dalian

.

ABSTRACT: Plastination is a method of preserving biological tissues with a curable

polymer. Sheet plastination is a method of preparing plastinated tissue slices for

education and research. Both epoxy and polyester sheet plastination are currently used.

P45 sheet plastination produces an intact, semi-transparent anatomical structure, with

well highlighted connective tissue. There is little information in the literature regarding

the physical properties of P45 plastinated specimens. Shrinkage during plastination is to

be expected. In this study we present data on the shrinkage of the following P45

plastinated tissue slices: eight head and neck sections, five thoracic sections, eight

abdominal sections, five pelvic sections, and three arm sections. The standard P45

protocol was followed, and a digital image of the specimens was taken before and after

the plastination process. Analysis of the images showed that shrinkage varied between

6.39 (±3.9) % for cerebral cortex, and 19.86 (±1.68) % for lung tissue.

KEY WORDS: sheet plastination; polyester method; body slices; polyester resin;

P45 technique; tissue shrinkage * Correspondence to: Sui Hong-Jin. Tel.: +8613904287577; Email: suihj@hotmail.com

Introduction

Plastination is simply the process of substituting water

and lipid molecules from biological tissues with curable

polymer. It is considered a major improvement in the

preservation of biological specimens (Riederer, 2014;

McRae et al., 2015). Plastinated specimens pose no

health hazards (Henry et al., 1997; Sivrev, 2012).

Sheet plastination has been used over the decades as a

teaching and research tool. In teaching, for instance, the

study of topographical anatomy is helpful for interpreting

MRI and CT images (Thomas, 2004), and slice

plastinates aid in the appreciation and understanding of

sectional anatomy in biomedical images. Sheet

plastination also plays essential roles in clinical anatomy

research, it has been used in the study of joints, body

cavities and spaces, bones, neurovascular structures,

body ligaments, muscles, organs, and 3D computational

reconstruction of anatomical structures (Sora & Genser-

Strobl, 2007). The usage of sheet plastinated specimens

in teaching and research is due to the fact that sheet

plastination presents the body structures in a non-

collapsed and non-dislocated form (Sora et al., 2002).

P45 sheet plastination is a polyester resin plastination

method used to preserve biological tissues. P45 plasti-

nated sheets are semi-transparent slice sections, with

the internal structures of the specimen clearly revealed

(Gao et al., 2006).

Shrinkage of biological specimens during plastination is

expected. The shrinkage values can be used to validate

both morphometric and 3D reconstruction

measurements. This study addresses tissue shrinkage

after P45 sheet plastination.

Materials аnd Methods

Two cadaveric specimens were used for this study, and

approval for the study was given by the Department of

Anatomy, Dalian Medical University. The following

sections were obtained from the specimens: eight head

and neck sections, five thoracic sections, eight

abdominal sections, five pelvic sections, and three arm

sections.

P45 sheet plastination procedure

A detailed P45 sheet plastination procedure has been

documented by Sui and Henry (2007), and Okoye and

Sui (2019). The cadavers were formalin-fixed prior to

processing. The specimens were first frozen in an ultra-

cold deep freezer at -70° C for about two weeks.

Tissue Shrinkage After P45 Plastination - 27

Figure 1. Pelvic section before the P45 plastination

process (A) and after P45 plastination (B).

Slicing

The specimens were embedded in polyurethane and

then sectioned using a bandsaw with a blade thickness

of 0.3 mm, and a cutting speed of 40 m/s. The section

thickness was 2 mm. Between adjacent slices, tissue

lost as tissue dust while sawing was approximately 1

mm thick (Fig. 1a).

The sliced body sections were placed on polyethylene

grid with a cotton fibre screen. The sawdust on the sliced

sections was removed with small stream of gently

running water. The grids with their washed slices were

then stacked and tied with twine to hold each stack as a

unit. The stacked units were then transferred into the

first cold acetone (-25° C) bath

Bleaching

The stacked units were immersed in 5% hydrogen

peroxide overnight to improve tissue color brightness

and transparency. Subsequently, excess bleach was

washed off by washing the slices in running water for

one hour or more.

Dehydration

The stacked units of slices were firstly precooled to 5° C

in order to avoid the formation of ice crystals and

shrinkage, before being submerged in a 100% acetone

bath at -25° C for one week. The stacked units were

then transferred into another fresh acetone bath at -15°

C for one week. Finally, the stacked units were

transferred into 100% acetone at room temperature for

another week.

Preparing the P45 polyester mixture

The impregnation resin mixture was prepared as follows:

1000 ml P45 resin (Hoffen polyester, China) was mixed

with 10 g of P45A, 30 ml P45B and 5 g of P45C. P45A

and P45C are plasticizers and P45B is a hardener. The

open vertical chamber was then prepared.

Building the casting chamber

The casting chamber used in this technique is an open

vertical chamber. The top part of the chamber is left

open, while the bottom and two sides of the casting

chamber are clamped. The open vertical chamber was

placed vertically in the vacuum chamber, i.e. standing on

its bottom end. The chamber was constructed from two

plates of 5 mm tempered glass, flexible 4 mm latex

tubing, and several large fold-back clamps.

The tubing was intercalated between the two tempered

glass sheets, and the glass and tubing were then

clamped together using fold back clamps on three sides,

leaving the top open.

Forced impregnation of the slices

The body sections were removed from the final acetone

bath and placed into the open vertical chambers. The

chambers were then filled with P45 polyester resin

mixture (Hoffen polyester, China) using a customized

funnel at the top open part of the chamber.

The vertical chambers, with the top part are left open,

were then placed upright into a vacuum chamber about

one metre deep. The vacuum chamber was sealed, and

the absolute pressure was progressively decreased to

20 mm Hg, 10 mm Hg, 5 mm Hg, and 0 mm Hg,

maintaining slow bubble production and release. The

pressure was maintained at 0 mm Hg until bubbling

ceased. The impregnation process was performed at

room temperature and was completed in approximately

8 hours.

28 – Sui, et al

Figure 2. A thoracic section uploaded into the Image J

software before the plastination process (A), and after

P45 plastination (B). The dotted line is the tracing line

used by the software for measuring the superior surface

area of the organ

Curing

After impregnation, the open vertical chambers were

transferred to a water bath at 40° C for curing, for three

days.

Finishing

After curing, the open vertical chambers were removed

from the water bath and dismantled. The P45 slice were

wrapped with a plastic sheet or light-weight foil.

Measurement of the tissues

A calibrated photographic documentation of the slice

sections was taken before tissue slicing and after P45

plastination (i.e. after curing) (Fig. 1b). The photographs

were uploaded to Image J software (Image J 1.52i), and

the surface area of each organ was measured (Fig. 2).

The brain, kidney, liver, muscles and spleen were

measured. The muscles measured were the gluteus

maximus and the triceps. The cerebral cortex of the

brain was measured on the brain sections.

Measurements of each organ before slicing and after

curing were documented, and the percentage shrinkage

was calculated.

Results

The P45 sections were in good condition, the slices were

semi-transparent and the connective tissues and organs

in each sheet were all intact. The measurements of the

organs before and after P45 plastination, and the

percentage shrinkage of the tissues are presented in

Table 1.

Liver Brain Kidney Muscle Lung Spleen

No. of slices

measured

5 8 5 5 5 5

Area of superior

surface before

plastination

(cm2)

493.92 524.98 83.38 61.21 453.890 143.21

Area of superior

surface after

plastination

(cm2)

443.59 491.40 76.49 57.09 363.75 127.96

Average

percentage (%)

shrinkage ±SD ±

10.2

3.5

±

6.39

3.9 ±

8.26

2.8

6.73

±

3.61

±

19.86

1.68 ±

10.64

6.35

Table 1 Surface area and average shrinkage of different tissues before and after P45 sheet plastination.

Tissue Shrinkage After P45 Plastination - 29

Figure 3. Graphical representation of the percentage

tissue shrinkage

The lung tissue showed the highest mean percentage

shrinkage value, and the brain tissue the lowest mean

percentage shrinkage (Fig. 3).

Discussion

Shrinkage leads to the reduction of the actual

measurement of the tissue. Thus, the organ shrinkage

value is useful in determining accurate dimensions of

plastinated specimens especially in morphometric

measurements and 3D image reconstructions.

Studies on the shrinkage of thin E12 and standard E12

body sections and also P40 brain slices have been

reported. While E12 and P40 are popular sheet

plastination techniques, the P45 technique is not

currently widely used. However, the shrinkage of this

technique has until now not been reported (Sora et al.,

2002; Sora et al.,

2015).

Several factors can affect tissue shrinkage. Two factors

have been strongly advocated: (1) The dehydrating

temperature affects shrinkage, though dehydrating at

low temperatures (-25° to +5°C) will reduce shrinkage

(von Hagens, 1985, Brown et al., 2002), though the

tissue transparency will be negatively affected because

of minimal defatting (Cook and Al-Ali, 1997). This can be

tackled by increasing the degreasing time (Sora et al.,

2002), but this might also impact on the shrinkage of the

processed tissue.

(2) The shrinkage of the impregnating resin itself may

also affect tissue shrinkage during plastination.

In this study, the sections were measured only before

and after P45 plastination, and did not include shrinkage

during dehydration or impregnation. Taking the

specimens out for measurement during the plastination

process can impact negatively on the shrinkage of the

plastinated section.

The measurement carried out was a bi-dimensional

measurement of the length & width of the organs. The

different P45 organ sections had different shrinkage

rates. The mean percentage reduction of the brain,

kidney and muscles in this study were below 10%, while

the spleen, lungs and liver were above 10%. As

expected, the lung shrinkage was greater than 10%. The

shrinkage of the lung sections was maximal, while the

shrinkage of the brain sections was minimal. This may

be attributed to the properties of the organs – though the

brain is soft, its cells are densely packed. The lungs, on

the other hand, are spongy and have certain elastic

properties. Thus, in addition to dehydration and

shrinkage of the resin, the properties of the organ or

tissue being plastinated may impact its shrinkage.

The percentage shrinkage of brain specimens in this

technique was similar to that found in the P40 technique

for brain tissue (Sora et al., 1999) but differs for kidneys

(Pereira-Sampaio et al., 2011). In the P45 technique, the

bleaching procedure is performed before dehydration,

while in the P40 process, dehydration is performed

before bleaching. The duration of plastination using

polyester resin (P40, P45) is relatively short when

compared to their epoxy counterpart. However, P45

requires no UV light for curing, unlike the P40 technique.

Morphometric and 3D reconstruction measurements

from P45 sections should take into consideration the

shrinkage of the tissues, and also the tissue loss while

sawing the specimens. The values ascertained in the

present study can provide a useful estimate of shrinkage

in future studies.

References

Brown MA, Reed RB, Henry RW. 2002: Effects of

dehydration mediums and temperature on total

dehydration time and tissue shrinkage. J Int Soc

Plastination 17:28-33.

Cook P, Al-Ali S. 1997: Submacroscopic interpretation of

human sectional anatomy using plastinated E12

sections. J Int Soc Plastination 12(2):17-27 .

30 – Sui, et al

Gao H, Liu J, Yu S, Sui H. 2006: A new polyester

technique for sheet plastination. J Int Soc Plastination

21:7-10.

Henry RW, Janick L, Henry C. 1997: Specimen

preparation for silicone plastination. J Int Soc

Plastination 12(1):13-7.

McRae KE, Davies G, Easteal R, Smith GN. 2015:

Creation of plastinated placentas as a novel teaching

resource for medical education in obstetrics and

gynecology. Placenta 36:1045-1051.

Okoye CS, Sui H-J. 2019: Updated protocol of Hoffen

P45 sheet plastination technique. J Plastination 31(2):

19-24

Pereira-Sampaio MA, Marques-Sampaio BP, Sampaio

FJ, Henry RW. 2011: Shrinkage of renal tissue after

impregnation via the cold Biodur plastination technique.

Anat Rec 294:1418-1422.

Riederer BM. 2014: Plastination and its importance in

teaching anatomy. Critical points for long-term

preservation of human tissue. J Anat 224(3):309-315.

Sora MC, Brugger P, Traxler H. 1999: P40 Plastination

of human brain slices: comparison between different

immersion and impregnation conditions. J Int Soc

Plastination 14(1): 22-24.

Sora MC, Brugger PC, Strobl B. 2002: Shrinkage during

E12 Plastination. J Int Soc Plastination 17:23-27.

Sora MC, Genser-Strobl B. 2007: The sectional anatomy

of the carpal tunnel and its related neurovascular

structures studied by using plastination. Eur J Neurol

12(5):380-384

Sora MC, Binder M, Matusz P, Ples H, Sas I. 2015: Slice

plastination and shrinkage. Mater Plast 52(2):186-189.

Sui HJ, Henry RW. 2007: Polyester plastination of

biological tissue: Hoffen P45 technique. J Int Soc

Plastination 22:78-81.

Thomas M, Steinke H. 2004: Thin-layer plastination of

the shoulder. CSMI 1:9-14

Sivrev, D. 2012: Safety and durable P35 and P40

plastination slices of anatomical objects. Conference:

actual questions of theoretical and practical medicine.

Nalchik, Russia 75:109-111.

The Journal of Plastination 31 (2):31 (2019)

Journal of Plastination Instructions for Authors

(Revised July 2017)

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

Goals - The Journal of Plastination (ISSN 1090-2171) aims to provide a medium for the publication of scientific papers dealing with all aspects of plastination and preservation of biological specimens.

Submission Guidelines All manuscripts must be submitted to the Editorial Office via the e-mail: editor.plastination@gmail.com. If you experience any problems or need further information, please contact Philip J. Adds, padds@sgul.ac.uk.

Authors must have an e-mail address at which they may be reached.

Necessary Files for Submission Include:

Cover letter

Manuscript (including references and figure legends)

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

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 (editor.plastination@gmail.com) as a pdf or jpeg file via an e-mail attachment. 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. HOLLADAY

1*, B. L. BLAYLOCK

2 and B. J. SMITH

1

1Department of Biomedical Sciences and Pathobiology,

Virginia Maryland Regional College of Veterinary Medicine, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061-0442, USA. 2College 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.

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: journal93@plastina.org

The Journal of Plastination 31 (2):32 (2019)

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. Text The body of the text should be written using American English spelling. Where quantities are specified, S.I. units should be used. Equivalent Imperial or U.S. units, if desired, should follow in parentheses e.g. 1 Kg (2.2 pounds). 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.

Von Hagens G. 1985: Heidelberg plastination folder: Collection of technical leaflets for plastination. Heidelberg: Anatomiches Institut 1, Universität Heidelberg, p 16-33.

For other publications:

Internet references: Author last name, initial(s). Year: Title of article. URL: Internet address [accessed month, year].

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 (two-column 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.

The Journal of Plastination 31 (2):33 (2019)

Statement of Publication and Research Ethics: This statement is based mainly on the Code of Conduct and Best-Practice Guidelines for Journal Editors (Committee on Publication Ethics, 2011). Responsibilities of the Editor and Editorial Board:

Publication decisions

The editor (in consultation with the Editorial Board where appropriate) is responsible for deciding which of the manuscripts submitted to the Journal of Plastination will be accepted for publication, and into which category of submission they should be placed. The decision will be based solely on the paper's importance, originality and clarity, and the study's validity and its relevance to the scope of the journal. The Editor and Editorial Board will also consider, where appropriate, current legal requirements regarding libel, copyright infringement, and plagiarism.

Confidentiality

The Editor undertakes not to disclose details about any submitted manuscripts to anyone other than the corresponding author, reviewers (and potential reviewers), and the publisher, as appropriate.

Disclosure and conflicts of interest Unpublished materials disclosed in a submitted paper will not be used by the editor or the members of the editorial board for their own research purposes without the author's explicit written consent.

Responsibilities of the Reviewers Contribution to editorial decisions The peer-reviewing process assists the Editor and the Editorial board in making editorial decisions and will also, where appropriate, inform the author of improvements that will, in the opinion of the reviewer, enhance the paper.

Promptness Any selected referee who feels unqualified to review the research reported in a manuscript or knows that its prompt review will be impossible should notify the editor and withdraw from the review process.

Confidentiality

Manuscripts sent for review must be treated by them as confidential documents. They must not be disclosed to or discussed with others unless specifically authorized by the Editor.

Standards of objectivity Reviews must be conducted objectively, without personal criticisms of the author(s). Referees should express their opinions clearly, and justify their comments with examples and supporting arguments.

References and reference citations Reviewers should check that published works cited in the manuscript have also been listed accurately in the References section, and that all references listed have also been correctly cited in the text. Reviewers may also wish to indicate other relevant papers in the literature of which the author(s) may not have been aware. Reviewers will notify the Editor of any substantial similarity or overlap between the manuscript under review and other published papers of which they are aware.

Disclosure and conflict of interest Privileged information or ideas obtained through peer review must be kept confidential and not used for personal advantage. Reviewers should not consider a manuscript in which they have a conflict of interest resulting from competitive, collaborative, or other relationships, or connections with any of the authors, companies, or institutions associated with the manuscript. Any such conflict should be declared to the Editor before agreeing to undertake the review. Duties of the Authors

Reporting standards Authors of original research reports should present an accurate account of the work performed as well as an objective discussion of its significance. Underlying data should be represented accurately in the paper. A paper should contain sufficient detail and references to permit others to replicate the work. Fraudulent or knowingly inaccurate statements constitute unethical behavior and are unacceptable.

Data access and retention Authors may be asked to supply the raw data for their study, and should be prepared to make the data publicly available where appropriate and practicable.

Plagiarism, originality, and acknowledgement of sources

The Journal of Plastination 31 (2):34 (2019)

Authors will submit only entirely original works. The work and/or words of others, where they have been used or quoted, will be appropriately acknowledged and cited.

Multiple, redundant or concurrent publication In general, papers that describe essentially the same research should not be published in more than one journal. Submitting the same paper to more than one journal is considered to be unethical and is unacceptable. Manuscripts that have been published as copyrighted material elsewhere cannot be submitted. Manuscripts that are undergoing the review process should not be resubmitted elsewhere. By submitting a manuscript, the author(s) retain the rights to the published material, although in case of publication, copyright of the published paper passes to the Journal of Plastination.

Authorship of the paper Authorship should be limited to those who have made a significant contribution to the conception, design, execution, or interpretation of the reported study and its subsequent write-up for publication. All those, and only those, who have made significant contributions should be listed as co-authors. The corresponding author must ensure that all contributing co-authors are included in the author list. The corresponding author will also verify that all co-authors have approved the final version of the paper and have agreed to its submission for publication.

Disclosure and conflicts of interest The corresponding author should include a statement disclosing any financial or other substantive conflicts of interest that may be construed to influence the results or interpretation of the manuscript. All sources of financial support for the project should be disclosed. Where there are no conflicts of interest, a statement to that effect should be included.

Fundamental errors in published works When an author subsequently discovers a significant error or inaccuracy in their own published work, it is the author's obligation promptly to notify the Editor of the Journal and to cooperate with the Editor to retract or correct the paper by issuing an erratum.

Research involving human or animal subjects In research involving human subjects, The Journal of Plastination requires that all such studies adhere to the principles of the Declaration of Helsinki. Each manuscript must include details of the a) number of subjects, b) age and sex of the participants, c) inclusion and exclusion criteria, and f) a statement that ethical approval was obtained for the study, and that informed consent was given by the participants. For cadaveric studies, appropriate consent must be in place prior to utilizing the cadavers or specimens. Studies involving experimental animals must conducted in a humane manner and in accordance with relevant guidelines for the care and utilization of laboratory animals. Animal care should be in line with the NIH Guidelines for the Care and Use of Laboratory Animals (NIH, 2015). The manuscript must include a statement that ethical approval of the protocol was obtained. The Journal of Plastination will reject manuscripts if the Editor and/or Editorial Board are not satisfied with the standards of ethical use of animals or data from humans in research. References Committee on Publication Ethics (COPE). (2011, March 7). Code of Conduct and Best-Practice Guidelines for Journal Editors. Retrieved from: https://publicationethics.org/files/Code_of_conduct_for_journal_editors_Mar11.pdf (accessed 5th September 2017) NIH Office of Laboratory Animal Welfare - Public Health Service Policy on Humane Care and Use of Laboratory Animals (NIH, 2015). Retrieved from: https://grants.nih.gov/grants/olaw/references/phspol.htm