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The official publication of the International Society for Plastination
The Journal of Plastination
I SSN 2 311 -77 61
Volume 30 (2); December 2018
Remembering Professor Lance Graham Nash – p5
Remembering the Past While
Looking to the Future: The
First Ten Years of the Journal
of Plastination – p8
Establishing for the First Time
the Use of the Standard S10
Technique for Plastination in
The Sudan – p15
Report of the 19th
International Conference on
Plastination – p19
Abstracts Presented at the
19th International Conference
of the ISP – p20
Minutes of the Business
Meeting of the International
Society of Plastination – p47
Announcement of the 20th
International Conference of
the ISP – p51
IN THIS ISSUE:
In Memoriam
Professor Lance Graham Nash, BSci., MSci., PhD.
(1961 - 2016)
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: [email protected]. Manuscript preparation guidelines are on the last four pages of this issue.
On the Cover: The photograph of Professor Nash originated on Pinterest as saved by The American University of the
Caribbean.
The Journal of Plastination 30(2):1 (2018)
Journal of Plastination Volume 30 (2); December 2018
Contents
Letter from the President, Rafael Latorre 2
Letter from the Editor, Philip J. Adds 4
Remembering Professor Lance Graham Nash; M. Zhang*, D.G. Jones and D.R. Grattan 5
Remembering the Past While Looking to the Future: The First Ten Years of the Journal of Plastination; Philip J. Adds
8
Establishing for the First Time the Use of the Standard S10 Technique for Plastination in The Sudan
15
Report of the 19th International Conference on Plastination; Dalian, China; July 19 -July 22, 2018
19
Abstracts Presented at the 19th International Conference of the ISP; Dalian, China 20
Minutes of the Business Meeting of the International Society of Plastination; July 21, 2018 47
Announcement of the 20th International Conference of the ISP 51
Instructions for Authors 52
The Journal of Plastination 30(1):2 (2018)
LETTER FROM THE
PRESIDENT
Letter from the President of the International Society for Plastination
Dear Friends and Plastinators,
On behalf of the International Society for Plastination (ISP) I would like to thank
all of you who participated in the 19th International Conference on Plastination in
Dalian, China (July 2018). I would like to thank The Dalian Medical University, and
especially the efforts made by Dr. Sui and his local team, to organize this
congress. There were many hours of coordinated work to get everything ready. I
know from experience that organizing this kind of conference, together with a
workshop, takes a lot of effort. The quality of the lectures and communications
was excellent as substantiated in the abstracts contained in this issue of the
Journal of Plastination. Special thanks to Ms. Tanya and the yellow-green
volunteers for their dedication to making everything in this meeting a great
success. Members of the International Society for Plastination had frequent
meetings with the local organization committee to help them develop the best
conference possible. For this reason, I would like to express my special thanks for
the work done by Drs. Carlos Baptista, Robert Henry, Dmitry Starchik and Selcuk
Tunali; without their help this congress would not have been possible. We
learned a lot during the congress, and we established interesting collaborations
for our laboratories in the near future.
In the general meeting of the ISP held on July 21, 2018, Dalian, the new Officers
and Councilors of the International Society for Plastination for the biennial 2018-
2020 were announced: President: Rafael Latorre; Vice-president: Dmitry Starchik;
Secretary: Nicolas Ottone; Treasurer: Carlos Baptista; and Councilors Kees De
Jong, Hong-Jin Sui, Onyemaech Okpara Azu, and Telma Masuko. I would like to
thank Anthony Weinhaus for his support with this election process. Special thanks
to Selcuk Tunali for his tenure as Secretary since 2016. Thanks to Bob Henry, Ming
Zhang and Athelson Bittencourt for serving as councilors since 2016. We greatly
appreciate the time they have dedicated to the ISP. I am sure they will continue
to be involved as active members and we will still have the opportunity to learn
from them. Congratulations to all elected officers, you are the principal part of
our Society during the next two years. Thank you for the opportunity of serving
the ISP as your President for the next two years. I will do my best, and with the
help of all ISP members, especially the board committee, I am sure our Society
will keep growing in both activities and prestige.
We have two years ahead to finish projects we have already started in the ISP and
to begin with new actions to improve our Society. During the general assembly in
Dalian, we presented the following proposals:
Rafael Latorre, DVM, PhD
The Journal of Plastination 30(2):3 (2018)
1. ISP Website:
a. New ISP webpage with a new look and more information about
members, labs, commercials, video tutorials etc.
b. Optimize the information obtained with our ISP survey. A lot of
information can be included in the new ISP website
c. Include the link with the MOOC about S10, P40 and E12 plastination
techniques
d. Announce any course or seminar about plastination around the
world
e. Include a section of new papers about research using plastination as
a tool
2. ISP list:
a. Open to the list any question we receive asking for help or
information about plastination techniques
b. Send to the list topics to discuss around plastination activities
c. Promote new research applications/techniques of plastination
3. Journal of Plastination:
a. Commitment to send papers from the ISP members
b. Commitment to invite to other people to participate in the journal
with original paper/ review
4. Training:
a. Promote at least two new advanced plastination courses in the next
two years
Participants in the general meeting of the ISP also decided on Temuco, Chile, to
host the 20th International Conference on Plastination to be held in 2020. My
best congratulations to Dr. Nicolas Ottone. The other candidate, Honolulu,
Hawaii, was invited to host the 12th Interim Meeting of the ISP in 2019.
I would like to welcome all new members of the ISP and to invite them as well as
the rest of member to participate in the various ongoing activities organized or
hosted by the ISP.
Finally, I want to express my appreciation and gratitude to Dr. Philip Adds, for the
superb job as Editor-in-Chief of the Journal of Plastination and for the continued
organization of the journal.
With the kindest regards from Murcia, Spain
Yours sincerely,
Rafael Latorre
The Journal of Plastination 30(2):4 (2018)
LETTER FROM THE EDITOR
Dear Colleagues,
In the previous issue, I reported that I had submitted an application for The Journal of
Plastination to be indexed on Scopus. Scopus (which is owned by Elsevier) is the largest
abstract and citation database of peer-reviewed literature: scientific journals, books and
conference proceedings, covering over 36,000 titles1.
I am pleased to be able to report that I have received the following response from the
Scopus evaluation team:
Title: Journal of Plastination
ISSN / E-ISSN: 2311-7761 / 2311-777X
Publisher: The International Society for Plastination
The title mentioned above has been evaluated for inclusion in Scopus by the Content
Selection & Advisory Board (CSAB). The review of this title is now complete and the CSAB
has advised that the title will be accepted for inclusion in Scopus. For your information,
the reviewer comments are copied below:
Single blind peer review, yet the journal publishes just one or two issues per year and
belongs to an international scholarly society. Editors and authors are globally dispersed
but are predominantly from the USA. While readability of articles is good, the title is only
fairly cited and the Editor standing is not as high as one would expect it to be, both in
terms of h-index and number of published documents. Further the title is not covered by
any major bibliographic databases which makes accessibility an issue. Would like to
accept the title for its strengths but request re-evaluation in the future to see if
challenges mentioned have been addressed.
Our Source Collection Management department will contact the publisher of this title
within the next three months to initiate the indexing process for Scopus. A content
coverage agreement needs to be in place before we can start adding the content to
Scopus.com. If you are the publisher, please do not send us your content yet unless
requested by our Source Collection Management department.
This is very encouraging, and is a reflection of the high quality of the papers published in
our Journal. In the meantime, I shall continue with our application to the Web of
Science, and hope to re-submit an application to Medline in the near future.
Best wishes,
Philip J Adds Editor-in-Chief References
1. https://www.elsevier.com/en-gb/solutions/scopus
Philip J. Adds, MSc, FIBMS, SSFHEA
The Journal of Plastination 30(2): 5 (2018)
Professor Lance Nash
In Memory Of
Professor Lance Graham Nash, BSci., MSci., PhD.
(1961 - 2016)
Written by: M. Zhang*, D.G. Jones and D.R. Grattan
Department of Anatomy, University of Otago, Dunedin, New Zealand
Corresponding Author: Ming Zhang, MB, MMed, PhD., Department of Anatomy, University of Otago, PO Box 913,
Dunedin 9054, New Zealand. Telephone: 0064 3 479 7378 Fax: 0064 3 479 7254, E-mail:
Professor Lance Nash died on the 14th September 2016 at the age of 55. This tribute is a
celebration of his life and the extraordinary contribution that he made to clinical anatomy
and plastination. We knew Lance for a long time. He completed his BSc, MSc and PhD
degrees in our Department, University of Otago, New Zealand, and then worked at the
American University of the Caribbean in St. Maarten, Netherlands Antilles, and was
promoted to Full Professor in 2011.
Lance devoted his life to clinical anatomy and plastination. Clinical anatomy is a unique
and challenging discipline that requires diverse skills in dissection, an ability to perceive
three-dimensional relationships of anatomical features and their relationship to function,
and other health professional knowledge. Lance was a classically-trained clinical
anatomist with a comprehensive knowledge of the anatomy discipline with skills in the
dissecting room and plastination laboratory. Lance initially trained as a healthcare
professional in Orthotics and Prosthetics, and spent more than a decade working with
medical and paramedical staff in a clinical setting. In this role, he was promoted to
Regional Manager for Orthotic services with the Wellington Capital Health Corporation,
New Zealand.
His PhD work at the University of Otago resulted in high impact publications and was included in the 40th edition of Gray’s
Anatomy. Together with his PhD classmate, Dr Mark Phillips, he established a novel technology: "The use of confocal
microscopy for the examination of E12 sheet plastinated human tissue" (Phillips et al., 2002) and successfully applied it to
the clinical anatomy research on the deep cervical fascia (Nash et al., 2005a; Nash et al., 2005b; Nash et al., 2004; Scali
et al., 2015a; Scali et al., 2015b). Lance's plastination skills were trained by Mr Russell Barnett, a well-known pioneer in
the international plastination society. During his time in Otago, Lance helped Russell to prepare one transverse and one
sagittal sets of the whole cadavers that have been heavily used in both research and undergraduate and postgraduate
teaching in our Department for almost two decades.
Professor Gareth Jones was Head of the Department of Anatomy in Otago when Lance was undertaking his postgraduate
studies. Professor Jones recalls that, since Lance’s background was quite different from so many postgraduate students,
he had to learn to fit in with this new environment. He was older, and had a Maori background, which is not common in
Anatomy. While he sometimes stood out as being different, what shone through was his infectious enthusiasm; rarely
could he be held back. He was very conscious of mapping out new paths for those like him who had had to make their
way up through the ranks. He was a pathfinder in many ways, but rather than see this as a hurdle, he tried to embrace it.
The Journal of Plastination 30(2):6 (2018)
Professor Jones sketched
by Lance
On completion of his postgraduate research training, he took a lecturer position at the American University of the
Caribbean (AUC), a US-accredited medical school on the island of Sint Maarten. He developed extensive experience
teaching all parts of the body and led dissection/sectional anatomy/osteology/radiology laboratory sessions. He
contributed to the histology course. As a successful and highly rated academic he was appointed Chairman of Anatomy,
Embryology and Histology in 2008 and Full Professor in 2011. He initiated and led a number of key reforms in the ACU,
including innovations in audio-visual and IT equipment use in teaching and assessment, improving quality and availability
of cadaveric material through negotiations with a Dutch University and the design and development of a completely new
anatomy facility.
Outside of the classroom and laboratory, Lance was a great team man. He would always
be the first person organizing social events, whether it be small scale gatherings at the
local bar, or large-scale bus trips for team building and morale. He was a genuinely
selfless person, who would offer support and assistance to friends and colleagues in
whatever way he could. He was also an extraordinarily perceptive individual, seemingly
always able to identify when people were having problems, and doing his best to help
them. His sense of humor was somewhat quirky and droll, and not always immediately
appreciated by everyone, but his intention always was to lift the mood of the people
around him.
His attention to detail in clinical anatomy was supported by a keen artistic talent – as a
student, he took to drawing caricatures of his peers to celebrate things like Ph.D.
completions. He was also full of surprises, as shown by his sketch of Professor Jones on
the latter’s birthday (Figure B). He did this entirely of his own volition, and it came as a
total surprise to Professor Jones. It still hangs in Professor Jones’s office as a
reasonable approximation of what he looked like all those years ago! It also reminds him
of an unforgettable character and blossoming academic. He will be missed by all those
that knew him.
Lance and his friends at the 13th ISP conference, Vienna, 2006
The Journal of Plastination 30(2):7 (2018)
References
Bickley HC, von Hagens G, Townsend FM. 1981: An improved method for preserving of teaching specimens. Arch Pathol
Lab Med 105:674-676.
Nash L, Nicholson H, Lee AS, Johnson GM, Zhang M. 2005a: Configuration of the connective tissue in the posterior
atlanto-occipital interspace: a sheet plastination and confocal microscopy study. Spine (Phila Pa 1976) 30:1359-1366.
Nash L, Nicholson HD, Zhang M. 2005b: Does the investing layer of the deep cervical fascia exist? Anesthesiology
103:962-968.
Nash LG, Phillips MN, Nicholson H, Barnett R, Zhang M. 2004: Skin ligaments: regional distribution and variation in
morphology. Clin Anat 17:287-293.
Phillips MN, Nash LG, NBarnett R, Nicholson HD, Zhang M. 2002: The use of confocal microscopy for the examination of
E12 sheet plastinated human tissue. J Int Soc Plastination 17:12-16.
Scali F, Nash LG, Pontell ME. 2015a: Defining the Morphology and Distribution of the Alar Fascia: A Sheet Plastination
Investigation. Ann Otol Rhinol Laryngol 124:814-819.
Scali F, Pontell ME, Nash LG, Enix DE. 2015b: Investigation of meningomyovertebral structures within the upper cervical
epidural space: a sheet plastination study with clinical implications. Spine J 15:2417-2424.
The Journal of Plastination 30(2):8-14 (2018)
Figure 1. Front cover of Volume 1, Number 1, January 1987
Figure 2. a) Canine heart-lung specimen (Henry, 1987);
b) blue whale heart (Miller et al., 2017)
Remembering the Past While Looking to the Future: The First Ten Years of the Journal of Plastination
Written By: Philip J. Adds, Editor-In-Chief, Institute of Medical and Biomedical Education (Anatomy)
St. George’s, University of London, London, UK
The Journal of Plastination, as we know it today, came
into being as the “Journal of the International Society for
Plastination” thirty-one years ago, in January 1987,
under the distinguished Editorship of Dr Harmon Bickley
(Fig 1). The cover, a simple white with red lettering,
featured an axial image of the human abdomen. It is not
clear where this image came from, as there were no
figures at all inside this first issue. However, one of the
papers within it details the procedure for plastination of
whole-body slices with Biodur® S10 or epoxy: “Sectional
anatomy is a valuable approach to the acquisition of an
understanding of body structure. Before our use of
plastinated slices, it had been neglected for many years”
(Lischka & Prihoda, 1987).
The presence of this image on the cover is highly
significant, and demonstrates the importance (and
relevance) of sheet plastination to sectional anatomy
and, hence, to healthcare technology. While CT and MR
were still relatively new in the eighties, they have since
come to dominate medical imaging, and, crucially,
depend on the ability of medical practitioners to interpret
sectional images. Sectional anatomy is now considered
to be an integral part of medical education: even back in
the eighties, the Journal was ahead of the field.
Five papers were published in issue 1, mostly describing
technical aspects of tissue preservation and plastination;
one paper, however, recognized the potential of
plastination in research: “Complete Examination of
Mastectomy Specimens Using Sheet Plastination with
Epoxy Resin”, which had among its authors the inventor
of plastination, Gunther von Hagens (Guhr et al., 1987).
The very first paper in the new journal was by Karine
Oostrom from Utrecht in The Netherlands: “Fixation of
Tissue for Plastination: general principles”, which
discussed different methods of fixation, color
preservation, colour injection, health hazards, and
employee safety (Ostrom, 1987). Ostrom described the
personal protective equipment (PPE) worn in the
plastination lab in Heidelberg: “rubber gloves, plastic
aprons, and goggles or gas masks.” Ostrom continued:
“Those of you who attended the Third International
Conference on
Plastination in San
Antonio will certainly
recall the slide in which
three young ladies
modelled these
fashionable accessories,
and nothing else…The
editor was adamant that
we omit this illustration,
however it would have
served to show that
even fixation can be
fun.” It is good to note
that high editorial
standards were already in place!
The second issue followed in the same year, again with
five papers focussing mainly on technical issues.
Notable authors in issue 2 include Dr Robert Henry, who
contributed a paper on the plastination of hearts and
heart-lung specimens, using canine specimens (Henry,
1987). Thirty years later, Dr Henry was among the
authors of a paper describing heart plastination on a
much larger scale: the salvage and preservation of a
blue-whale heart: “The Challenges of Plastinating a Blue
Whale (Balaenoptera musculus) Heart “(Miller et al.,
2017). It is interesting to compare the image that
accompanied the 1987 paper, (which was in fact the first
anatomical image ever to appear in the Journal), with the
Remembering the Past/Looking to the Future - 9
image of the final specimen that accompanied the 2017
paper (Fig. 2). These images help to show how far
plastination (and the Journal) have come in the last 30
years. Progress indeed.
The author affiliations from Volume 1 show that all
contributing authors were from the early centres of
plastination, one each from Utrecht, Vienna, and
Heidelberg, and the rest from the USA. This narrow
geographical range reflects the limited reach of the
emergent technology of plastination at that time.
The next significant develop in the development of the
journal came 2 years later, in 1989, with the appointment
of Bob Henry as Editor; Harmon Bickley became
Executive Director of the Society. Volume 3 (1989) was
single-issue only, and included, for the first time,
abstracts from meetings of the International Society for
Plastination: the Fourth International Conference (held in
1988 at Macon, Georgia, USA) and the inaugural Interim
Meeting, held a year later in Knoxville, Tennessee.
Volume 3 also heralded the arrival in print of another
significant figure, Dr Carlos Baptista, with two papers
focusing on clinical and applied human anatomy,
“Plastination of the heart: preparation for the study of the
cardiac valves” (Baptista & Conran, 1989), and
“Plastination of the wrist: potential uses in education and
clinical medicine” (Baptista et al., 1989) (Fig. 3), the
latter again focusing on sectional anatomy. As the
authors put it: “These specimens provide an excellent
tool for teaching anatomy and pathology, for patient
education, and potentially as an augentation to MRI
(magnetic resonance imaging) and CT (computer
tomography) analysis” (Baptista et al., 1989).
The next step in the evolution of the Journal came with
Volume 4, another single-issue volume, published in the
fall of 1990. For the first time, an Editorial board
appeared on the first page, consisting of Drs Carlos
Baptista, Harmon Bickley, and P. Tom Purinton. Bob
Henry continued as Editor, and Harmon Bickley as
Executive Director of the Society. Volume 4 contained
the usual eclectic mix of research and technical papers.
Notable among them was a paper by Lane, continuing
the theme of sectional anatomy “Sectional anatomy:
standardized methodology” (Lane, 1990), with an x-ray
image showing levels of axial sections of the body (Fig.
4) and color images of plastinated body sections (Fig. 5).
It is interesting to compare these early papers on
sectional plastinates with the “Visible Human Project”,
which claimed to have revolutionized the study of
anatomy: “The National Library of Medicine (NLM)
introduced the Visible Human Project (VHP) in
Figure 3. a) Heart valve specimen (Baptista & Conran, 1989); b) wrist
section (Baptista et al., 1989)
Figure 4. Levels of sectioning for axial body slices (Lane,
1990)
Figure 5. Images of plastinated body sections
(Lane, 1990)
10 - Adds
Figure 7. Photomicrographs of sheet plastinated human
knees, showing a control and a series of slices showing
degenerative changes (Graf et al., 1992)
November 1994 and in doing so revolutionized our ability
to view and understand human anatomy.”
(https://infocus.nlm.nih.gov/2014/12/31/the-visible-
human-project-at-20/), emphasis added). On the
contrary, it could be claimed that plastination had
already achieved this, nearly a decade earlier!
Another paper in Volume 4 discussed the potential for
using plastinates in the construction of holographic
images (Myers and Bickley, 1990). Holography was
described at the time as “a solution in search of a
problem” (ibid.), because it had thus far fulfilled only a
small part of its potential. Despite speculation about the
potential for holograms in medical education, the
technology never really seemed to take off. Virtual reality
is now the only game in town. Try typing “plastination
hologram” into Google images nowadays, and the only
relevant image likely to appear is Gunther von Hagens
with a hologram of a plastinated couple during
intercourse (Fig. 6).
Volume 6 came out in 1992, another single-issue
volume, which contained a paper from Graf et al., “Early
Morphological Changes in Chondromalacia Patellae in
Humans - Demonstrated With The Plastination Method”,
which described how epoxy sheet plastination was used
on human patellas to investigate, at the microscopic
level, changes in the chondral and subchondral areas. It
was reported, for the first time, that “the origin of the
idiopathic chondromalacia was detected in the
subchondral area and not the cartilage as previously
thought.” These changes had not previously been seen
with arthroscopy, but demonstrated how plastination can
bridge the gap between the macro- and the microscopic,
and shows the application of plastination to both
research and clinical medicine (Fig. 7) (Graf et al.,
1992).
The Journal developed further in 1993, with changes to
the editorial team that were announced in Volume 7. Dr
Robert Henry remained as Editor, but R. Dale Ulmer,
from the College of Medicine, Mobile, Alabama, was
listed as Editor-elect. The Editorial Board remained as
before, but, as evidence of the growing worldwide reach
of plastination, special Journal Correspondents were
listed: for Canada, R. Blake Gubbins (Queen’s
University, Ontario); for Europe, Margit Rokel (St Leon-
Rot, Germany); and for the Far East, Robert Boyes
(Queensland, Australia).
Volume 7 also contained a paper from another author
who was to go on to become a prominent and
distinguished member of the ISP, Andreas Weiglein:
“Plastinated Brain Specimens in the Anatomical
Curriculum at Graz University”. In this paper, Dr Weiglein
described the use of P35 plastinated brain slices in
Figure 6. Gunther von Hagens with a hologram of a couple
during intercourse
(https://www.gettyimages.com.au/detail/news-photo/three-
dimensional-hologram-of-a-plastinated-couple-during-news-
photo/98631343)
Remembering the Past/Looking to the Future - 11
Figure 8. “First known examples of cross-sectional anatomy by
Leonardo da Vinci (1452 – 1519) of the pregnant uterus (a) and the
lower limb (b)“ (Weiglein, 1993)
Figure 9. The original shoulder specimen, E12 plastinated slices and
corresponding MRI, CT and US images (Entius et al., 1993)
neuroanatomy teaching, but he also draws comparisons
between modern-day sectional anatomy and the
anatomical drawings of Leonardo da Vinci (1452 – 1519)
“the first known examples of cross-sectional anatomy”
(Weiglein, 1993) (Figure 8). This was the first paper in
the Journal to discuss the history of anatomy in relation
to modern anatomical techniques.
In the same issue, a paper by Entius et al. brings state of
the art sectional anatomy, medical imaging, and
plastination together. In “A New Positioning Technique
for Comparing Sectional Anatomy of the Shoulder with
Sectional Diagnostic Modalities: Magnetic Resonance
Imaging (MRI), Computed Tomography (CT) and
Ultrasound (US)” the authors describe how skin markers
can be used on anatomical specimens to define planes
of section. After MRI, CT and ultrasound images were
obtained, the specimen was frozen and sectioned at 2
mm thickness. The slices were then plastinated using
the E12 technique, giving sections that exactly matched
the MRI, CT and ultrasound images (Entius et al., 1993)
(Fig. 9).
Volume 8, in 1994, came with a change in the Editorial
team. Dale Ulmer took over as Editor, and Bob Henry
joined the new-look Editorial Board, along with Vincent
DiFabio, Bill Richeimer, and William A. Gardner Jnr.
“Preparation Support”, another innovation, was provided
by Betty Clark and Rosemary Farmer.
Volume 9 (1995), with a suitably sunny cover picture
(Fig. 10), announced the dawn of a new day for
plastination, with the founding of the International
Society for Plastination as an official body. At the historic
4th Biennial Meeting, in Graz, Germany, Bylaws and a
Constitution for the ISP were written and adopted, and a
slate of Officers were proposed and elected, and the
Journal carried, for the first time, Letters from the newly-
elected President, Bob Henry, and the Editor, Dale
Ulmer.
The first-ever Editor’s letter, from Dale Ulmer, carried
this message: “I challenge each plastinator to contribute
one article yearly to our journal and help us advance our
organizational goals. As we learn – we grow.” A
message just as relevant today as it was in 1995.
The 8th International Conference on Plastination (the 5th
Biennial Meeting of the International Society for
Plastination) ventured, for the first time, to the Southern
Hemisphere, to the University of Queensland, Brisbane,
Figure 10. The dawn of a new era: The International Society for
Plastination is now an official body.
12 - Adds
Figure 11. Stained brain slices from Suriyaprapadilok
and Withyachumnarnkul (1997)
in Australia. Confusingly, the cover of Volume 10
featured a photograph showing the Sydney harbor
bridge and the Sydney Opera House. The abstracts from
the meeting were published in this issue, including “The
Use of Silicone Plastinated Specimens for Light and
Electron Microscopy” (Grondin et al., 1996), which
contained the following cryptic message: “if you can find
mistakes in this publication, please consider they are
there for a purpose. We publish something for everyone,
and some people are always looking for mistakes”!
In Volume 11 (1996), an article by Sharon Korbeck
entitled “A Pharaoh’s Farewell: the Making of a Mummy”
was reprinted with the permission of The National
Funeral Directors Association. In his Editor’s Letter, Dale
Ulmer wrote “While this process is not true plastination, it
is, however, a forerunner to the now popular process
that we as Plastinators now use. From time to time, I
believe it is good to examine and see the yester years”.
This passage reminded me of something Craig
Goodmurphy had said at the 15th Biennial Business
Meeting of the International Society for Plastination,
(Honolulu, July 2010): “…the objective of the Journal of
Plastination [should] be expanded to provide a medium
for the publication of scientific papers dealing with all
aspects of preservation of biological specimens including
plastination, sectional anatomy and other anatomical
techniques.” This is a very laudable aim, which had the
backing of the meeting and the Editorial Board. In fact,
as has been shown, the Journal has been doing that all
along, but it is good to be reminded from time to time of
the importance of the history, and range, of anatomical
techniques.
Volume 12 brought the first decade to a close in fine
style. The cover of Issue 1 featured stunning colour
photographs of stained plastinated brain slices (Fig. 11)
from Suriyaprapadilok and Withyachumnarnkul’s (1997)
paper “Plastination of Stained Sections of the Human
Brain: Comparison Between Different Staining Methods”.
Compare this to the cover of Volume 1! There had also
been a change
in the Editorial team, with Gilles Grondin taking over as
Editor, supported by an expanded, international Editorial
Board, of Pamela Arnold, Harmon Bickley, Robert
Henry, Steven Holladay, Larry Janick, Tage N. Kvist,
William Richeimer and Bill Wise from the USA, Russell
Barnett from New Zealand, Régis Olry from Canada, and
Andreas Weiglein, from Austria.
Issue 2 of Volume 12 was published in October 1997,
and included a typically wide-ranging mix of papers,
discussing both cutting-edge research, and the rich
heritage of anatomy. “Submacroscopic Interpretation of
Human Sectional Anatomy Using Plastinated E12
Sections” (Cook and Al-Ali, 1997), expanded on the
possibilities offered by E12 plastination in anatomy
education “The E12 process … has in effect filled a void
in undergraduate teaching. Students are provided with a
clear, unimpeded overview of the planes of the body
seen with a whole section…. providing a firm link
between macroscopic and microscopic anatomy” (Fig.
12).
While Cook and Al-Ali were looking to the future, Olry
and Motomiya (1997) looked back to the Renaissance
and beyond, with their paper “Paolo Mascagni, Ernest
Alexandra Lauth and Marie Philibert Constant Sappey
on the Dissection and Injection of the Lymphatics”. They
describe how the lymphatics were “discovered by
chance, misunderstood for a very long time, … the
subject of much controversy up to the early twentieth
century, when their accurate description was deemed
necessary to promote advances in oncology” (Fig. 13). It
is remarkable that the lymphatic system, though
apparently first described (albeit inaccurately) by
Erasistratus around 250 BC, was a source of continued
controversy up to the 20th century (the role of the
Remembering the Past/Looking to the Future - 13
Figure 12. E12 sections. Clockwise from top left: coronal
head, right eye (magnified), axial thorax section, thoracic
wall (magnified), coronal shoulder section, nasal cavity
(magnified). From Cook and Al-Ali (1997).
Figure 13. Dissected (L) and injected (R) lymphatic vessels
by Rausch, 1665. From Olry and Motomiya (1997)
thymus, for example, was not fully understood until the
1960’s).
That seems a fitting way to conclude this survey of the
first decade of the Journal of Plastination – remembering
the past, while looking to the future. In closing, I would
like to take the opportunity to repeat the words of Dale
Ulmer, in the very first Editor’s letter “I challenge each
plastinator to contribute one article yearly to our journal
and help us advance our organizational goals. As we
learn – we grow”.
References
Baptista CAC, Conran PB: 1989: Plastination of the
heart: preparation for the study of the cardiac valves. J
Int Soc Plastination 3: 3-7
Baptista CAC, Skie M, Yeasting RA, Ebraheim N,
Jackson WT. 1989: Plastination of the wrist: potential
uses in education and clinical medicine. J Int Soc
Plastination 3: 18-21
Cook P, Al-Ali S. 1997: submacroscopic interpretation of
human sectional anatomy using plastinated E12
sections. J Int Soc Plastination 12(2): 17-27
Entius CAC, Kuiper JW, Koops W, de Cast A. 1993: A
new positioning technique for comparing sectional
anatomy of the shoulder with sectional diagnostic
modalities: magnetic resonance imaging (MRI),
computed tomography (CT) and ultrasound (US). J Int
Soc Plastination 7: 23-26
Graf J, Fromm B, Schneider U, Niethard FU. 1992: Early
morphological changes in chondromalacia patellae in
humans demonstrated with the plastination method. J Int
Soc Plastination 6: 25-28
Grondin G, Grondin GG, Talbot BG. 1996: The use of
silicone plastinated specimens for light and electron
microscopy. J Int Soc Plastination 10: 32
Guhr A, Mueller A, Anton H-W, von Hagens G, Bickley
H. 1987: Complete examination of mastectomy
specimens using sheet plastination with epoxy resin. J
Int Soc Plastination 1(1): 23-29
Henry RW. 1987: Plastination of an integral heart-lung
specimen. J Int Soc Plastination 1(2): 20-24
Lane A. 1990: Sectional anatomy: standardized
methodology. J Int Soc Plastination 4: 16-22
Lischka M, Prihoda M. 1987: Establishing and operating
a plastination laboratory at The Institute of Anatomy,
University of Vienna. J Int Soc Plastination 1(1): 12-16
Miller JR, Henry RW, Nader P, Engstrom MD, Iliff S,
Chereminskiy V, von Hagens G. 2017: The challenges of
plastinating a Blue Whale (Balaenoptera musculus)
heart. J Plast 29(2):22-29
14 - Adds
Myers B, Bickely H. 1990: Use of plastinated tissue in
the construction of holograms. J Int Soc Plastination. 4:
38-39
Olry R, Motomiya K. 1997: Paolo Mascagni, Ernest
Alexandra Lauth and Marie Philibert Constant Sappey
on the dissection and injection of the lymphatics. J Int
Soc Plastination 12(2): 4-7
Ostrom K. Fixation of tissue for plastination: general
principles. J Int Soc Plastination 1(1): 3-11
Plastination hologram
(https://www.gettyimages.com.au/detail/news-
photo/three-dimensional-hologram-of-a-plastinated-
couple-during-news-photo/98631343 (accessed 15/1/19)
Suriyaprapadilok L, Withyachumnarnkul B. 1997:
Plastination of stained sections of the human brain:
comparison between different staining methods. J Int
Soc Plastination 12(1): 27-32
Visible Human Project
https://infocus.nlm.nih.gov/2014/12/31/the-visible-
human-project-at-20/ (accessed 12/1/19)
Weiglein AH. 1993: Plastinated Brain Specimens in the
Anatomical Curriculum at Graz University. J Int Soc
Plastination 7: 3-7
The Journal of Plastination 30(2):15-18 (2018)
TECHNICAL REPORT
Establishing for the First Time the Use of the Standard S10 Technique for Plastination in The Sudan
MOHAMED AMA1
AHMED AA2
ADAM ASIA I2
ALI AM1
AND TAHA AAM2
1 Department of Anatomy,
College of Veterinary
Medicine, King Faisal
University, Al-Ahsa 31982,
Kingdom of Saudi Arabia1.
2-Department of Anatomy,
Faculty of Veterinary
Medicine, University of
Khartoum. PO Box: 32,
Postal code: 13314
Shambat, Khartoum North,
Sudan.
ABSTRACT:
Plastination is method for long-term preservation of biological tissue, to produce dry,
durable, convenient and natural looking specimens that are useful as a unique teaching
aid for anatomy, pathology, radiology and surgery. The present study describes, for the
first time, the plastination laboratory in the Faculty of Veterinary Medicine, University of
Khartoum, Sudan. The standard Biodur ®S10 plastination technique was carried out in
formalin-fixed specimens of goat and donkey. They were first dehydrated in acetone.
Forced impregnation was then carried out using a vacuum chamber, and lastly, the
specimens were hardened in a gas curing chamber. Plastinated specimens were long-
lasting, and can be an important adjunct to traditional methods of teaching; they are
also excellent museum specimens.
KEY WORDS: acetone; formaldehyde; laboratory; museum; plastination; S10; Sudan * Correspondence to: Dr. Mohamed, A.M.A., Department of Anatomy, College of Veterinary Medicine, King Faisal University, Al-Ahsa 31982, Kingdom of Saudi Arabia. Mobile: +966530588021, email: [email protected]
Introduction
Plastination is the method of long-term preservation of
biological tissues with excellent surface details and high
durability. It was developed by Dr. Gunther von Hagens
in 1978 at the Heidelberg University in Germany (von
Hagens, 1979). Although it is difficult to prepare a well-
plastinated specimen, it is the most promising method to
preserve specimens as an alternative to formalin
preservation (Dawson, 1990).
In recent years, plastination has revolutionized the way
in which gross anatomy can be presented to students
(Latorre et al., 2007). Therefore, many Departments of
Anatomy in medical colleges throughout the world
started to establish plastination techniques in their own
laboratories (Briggs et al., 1997; Asadi, 1998; Reina-de
la Torre et al., 2004; Ali and Al-Thnaian, 2007; Suganthy
et al., 2012 and Sawad and Al-Asadi, 2014)
The aim of this study was to initiate and establish S10
plastination, in the plastination laboratory in the
Department of Anatomy, Faculty of Veterinary Medicine,
University of Khartoum, Sudan.
Materials and Methods
The standard Biodur® silicone S10 technique for
preservation of specimens in the laboratory of
plastination at the Department of Anatomy, Faculty of
Veterinary Medicine, University of Khartoum, Sudan,
was established in 2017. The laboratory was designed
according to the Plastination Technical Leaflets of
Heidelberg (Von Hagens, 1986). Financial resources
were obtained from the University of Khartoum, to
TECH
NIC
AL R
EPO
RT
16 – Mohamed, et al.
Figure 1: Plastination laboratory set-up, showing:
freezers (a) Bennert manometer (b) and separator (c).
Figure 2: Plastination laboratory set-up, showing gas
curing unit (a); stainless steel drums (b) and conveyor
pump (c).
Figure 3: Plastination laboratory set-up, showing
dehydration and impregnation containers.
promote new technologies in teaching anatomy in
faculties of Medicine and Veterinary Medicine in Sudan.
After the plastination lab had been set up, all specimens
for plastination were obtained from the Department of
Anatomy as follows: kidneys, hearts, spleen and liver,
from goats that had been infused with 10% formalin; and
heart and whole stomach from old donkey specimens
that had been fixed with 10% formalin for over a year.
For preparing specimens for plastination, the standard
silicone (S 10) method (von Hagens, 1979; 1986; von
Hagens et al., 1987) was used. The basic steps for
plastination technique are: specimen preparation,
dehydration & degreasing, impregnation, and curing.
Specimens were prepared, fixed with 10% formalin at
room temperature for 2 days, and then refrigerated at 4
°C for 24 hours. They were then dehydrated in cold
acetone (–25 °C) with three weekly changes to minimize
tissue shrinkage. After cold dehydration, the specimens
and acetone bath were brought to room temperature for
two days, for lipid removal. Forced impregnation was
then carried out, by placing the specimen into the
silicone polymer/catalyst S10/S3 mixture (100:1)
(Biodur®, Germany) in the vacuum chamber at -20 °C,
and gradually reducing the pressure. The final vacuum
ranged between 2 and 15 mmHg, and impregnation time
was four weeks.
Specimens were removed from the silicone bath, and
kept on a strainer at room temperature for 24 hours.
Specimens were then positioned in a gas curing
chamber containing S6 (Biodur®, Germany) in a small
glass container, at room temperature, for three weeks.
After the gas curing step, the specimens were ready to
use.
Results
The laboratory in the Department of the Anatomy had
been organized in order to host the plastination
production process. The total area of the laboratory is 52
m2 (3 rooms of different sizes), with large windows, and
extraction fans for providing adequate ventilation. The
freezers for dehydration are located in a separate room
(Fig. 1[a]) with a Bennert manometer (Fig. 1[b]) and
separator for oil and solvents (Fig. 1[c]); the freezer’s
compressors are located in another, adjacent, room of
the laboratory. The gas curing unit (Fig. 2[a]) is located
in the third room, with other necessary materials of the
plastination process: stainless steel drums (Fig. 2[b]),
conveyor pump (Fig.2[c]); and wire baskets, and
impregnation containers (Fig. 3).
Plastinated specimens of the goat: longitudinal section of
kidney (Fig. 4), liver (Fig. 5), cranio-lateral view for heart
(Fig. 6), longitudinal section for heart (Fig. 7), and spleen
(Fig. 8) were found to be quite similar to their natural
appearance, and kept their previous morphological
features, with minimal shrinkage. They were odorless,
durable, non-hazardous, easy-to-handle, formalin-free,
and life-like, and could be handled without a need for
personal protective equipment. In contrast, the
plastinated specimens of the donkey, longitudinal
section of the heart (Fig. 9) and whole stomach (Fig. 10),
were dark brown in color.
First Time S10 Plastination Technique in The Sudan - 17
Figure 4
Figure 5
Figure 6
Figure 7
Figure 8
Figure 9
Figure 10 Figures 4 – 10 showing various S10 plastinated
specimens of goat: longitudinal section of kidney (4);
liver (5); cranio-lateral view of heart (6); longitudinal
section of heart (7); spleen (8); and donkey:
longitudinal section of heart (9) and whole stomach
(10).
18 – Mohamed, et al.
Discussion
The Faculty of Veterinary Medicine at the University of
Khartoum represents the first research laboratory in the
Sudan where plastination has begun to improve the
teaching in practical anatomy, and to reduce the
exposure to toxic fumes for teachers, technical staff and
students.
The preservation of anatomical specimens has been a
long-standing goal of anatomists, pathologists and other
medical educators (Baptista et al., 1989). In recent
decades, plastinated specimens are near ideal, and are
excellent for teaching gross anatomy and neuroanatomy
(where routine specimens are delicate and scarce).
Their anatomical structure is well preserved, and
appears like a fresh specimen (Henry, 2004).
The plastination laboratory at the Faculty of Veterinary
Medicine, University of Khartoum, has produced good
quality plastinated specimens for the first time in Sudan.
There were a limited number of unsuccessful
plastinates, where the specimens changed color to dark
brown, which was likely due to the old formalin-fixed
specimens, as reported by Miklošová and Mikloš (2004).
The good quality plastinated specimens are used as
anatomical specimens for education and for study, as
predicted by Dibal et al, (2018). It is also intended to use
the laboratory to train technicians in the techniques of
plastination, and to encourage higher degree students in
the area. Financial support for this laboratory was
obtained from the University of Khartoum, Sudan, to
improve the capabilities of the laboratories, and to aid
new technologies in teaching anatomy. The general
belief that production of plastinates is expensive needs
revision. Thus, future research should target the
development of fast and cost-effective techniques of
plastination.
References
Ali AM, Al-Thnaian TA. 2007: Preservation of ruminant
and equine anatomical specimens by silicone
plastination. Sci J King Faisal University (Basic and
Applied Sciences) 8:111- 119.
Asadi MH. 1998: Plastination of sturgeons with the S10
technique in Iran: the first trials. J Int Soc Plastination
13:15-16.
Baptista CAC, Skie M, Yeasting RA, Ebraheim N,
Jackson WT. 1989: Plastination of wrist: potential uses
in education and clinical medicine. J Int Soc Plastination
3:18-21.
Briggs CA, Robbins SG, Kaegi WH. 1997: Development
of an anatomical technologies laboratory. J Int Soc
Plastination 12:8-11.
Dibal NI, Garba SH, Jacks TW. 2018: Plastinates:
possible tool for medical education in the near future:
mini review. Res Dev Med Educ 7:3-7.
Dawson TP, James RS, Williams GT. 1990: How do we
teach pathology? Silicone plastinated pathology
specimens and their teaching potential. J Path 162:265-
272.
Henry RW. 2004: Polyester plastination techniques,
specific troubles and problems. Murcia, Spain, 12th
International Conference on Plastination.
Latorre RM, García-Sanz MP, Moreno M, Hernández F,
Gil F, López O, Ayala MD, Ramírez G, Vázquez JM,
Arencibia A, Henry RW. 2007: How useful is plastination
in learning anatomy? J Vet Med Educ 34:172-176.
Miklošová M, Mikloš V. 2004: Plastination with silicone
method S 10 – Monitoring and analysis cause of failure.
Biomed Papers 148: 237–238.
Reina-de la Torre F, Rodríguez-Baeza A, Doménech-
Mateu JM. 2004: Setting up a plastination laboratory at
the Faculty of Medicine of the Autonomous University of
Barcelona. Eur J Anat 8: 1-6.
Sawad AA, Al-Asadi FS. 2014: Establishing a
plastination laboratory at the College of Veterinary
Medicine, University of Basra, Iraq. J Plast 26:30- 33.
Suganthy J, Deepak Vinod Francis. 2012: Plastination
using standard S10 technique - our experience in
Christian Medical College, Vellore. J Anat Soc India 61:
44-47.
von Hagens G. 1979: Impregnation of soft biological
specimens with thermosetting resins and elastomers.
Anat Rec 194: 247- 255.
von Hagens G. 1986: Heidelberg plastination folder.
Collection of all technical leaflets for plastination. 2nd
Edn. Heidelberg, Anatomische Institut, Universität
Heidelberg.
von Hagens G. Tiedemann K, Kriz W. 1987: The current
potential of plastination. Anat Embryol 175:411-421.
The Journal of Plastination 30(2):19 (2018)
Report of the 19th International Conference on Plastination.
Dalian, China, July 19 -July 22, 2018
By Sui, Hong-Jin
The 19th international conference of plastination was held in Dalian Medical University, Dalian, People’s
Republic of China (PRC). The conference commenced after the hands-on plastination training workshop held
at the Hoffen Biotechnique company. The workshop was attended by 34 people from nine countries. The
workshop was on the S10 plastination technique, and P45 sheet plastination. The participants testified that
P45 sheet plastination is relatively easy and produces good results.
Forty participants from fifteen countries attended the conference which was graced with posters and oral
presentations on plastination and research in plastination. At the oral presentations, new developments in
plastination on teaching, research and popular science were discussed. The Editor-in-Chief invited every
member of the International Society for Plastination to submit at least one paper a year in the Society’s
journal, The Journal of Plastination. The International Society for Plastination’s biennial Business Meeting was
not left out, it was held on Saturday, 21st July.
The participants will have a life-long memory of the Welcome party which was held in the world’s first
museum to display only plastinated collections, the ‘Mystery of Life Museum’. The participants dined in the
museum and had the opportunity to have a look at what the museum has to offer.
The participants visited the Department of Anatomy, Dalian Medical University, and where acquainted with
the Department’s teaching and student activities. They toured Dalian Hoffen, the world’s biggest and most
advanced plastination laboratory. There was also a tour around Lushunkou, which is a small historic town
where the University is situated. The Gala Night was another very memorable social activity, where the
participants enjoyed traditional Chinese opera, and tasted various Dalian sea foods, thereby having a feel of
the Chinese culture.
The Journal of Plastination 30(2):20 (2018)
Abstracts Presented at the 19th International Conference of the ISP
Dalian, China
18-22nd July 2018
3-D VISUALISATION OF THE RETROBULBAR ORBITAL SEPTA USING BIODUR E12® AND BIOVIS-3D SOFTWARE
ADDS PJ, CHEUNG A
Institute of Medical and Biomedical Education (Anatomy), St Georges, University of London, London, UK
Introduction: The retrobulbar fat body in the orbit is supported by a network of collagenous septa that support the
ocular adnexa, and aid in coordinating precise eye movements. The exact anatomical nature of these septa and their
contribution in ocular motility are not completely understood, but they are thought to act as pulleys for the extra-
ocular muscles. This has implications for orbital pathology and surgery.
Objectives: The aim of this study was to create 3D models of the retrobulbar fat septa in the human orbit, using epoxy
resin and 3D serial reconstruction software, in order to compare the morphology of the septa in left and right orbits, and
between individuals.
Materials and Methods: Four formalin-fixed human orbits were dissected, decalcified, dehydrated in acetone at -20° C,
and impregnated in Biodur® E12 epoxy resin. Serial sections (0.3 mm) were cut with a slow-speed diamond saw, stained
with Gomori’s trichrome for elastin and collagen, and photographed with a digital SLR camera. BioVis3D software was
then used to create 3D models of the fat septa.
Results: This project generated four x 3D reconstructions of the connective-tissue fat septa, which could be rotated on
all axes. The individual reconstructed structures could be isolated and manipulated. Using 3D reconstructions and serial
histological sections, common characteristics and variations of connective tissue septa among different orbits were
described.
Conclusions: Orbits from the same individual were noted to share a similar arrangement and areas of condensation of
fat septa. Although sharing broadly similar morphology, the orbital septa of different individuals displayed variations in
thickness, density and fine arrangement. The results reported here could serve as the groundwork for defining the
normal anatomy of the septa, and for investigations into the clinical and surgical implications of the variations between
individuals.
This study received no outside funding.
The Journal of Plastination 30(2):21 (2018)
ESTABLISHING A PLASTINATION LABORATORY MAY REDUCE RUNNING COSTS FOR THE VETERINARY ANATOMY
SECTION AT CVAS, JHANG, PAKISTAN
ANSARI AR
Section of Anatomy and Histology, Department of Basic Sciences, College of Veterinary and Animal Sciences (CVAS)
Jhang, University of Veterinary and Animal Sciences (UVAS), Lahore, Pakistan.
Introduction: Plastination is a laboratory preservation technique applied to specimens so that they can be used as
models in the study of anatomy, for both undergraduate education and research purpose. There are several limitations,
including financial constraints, in replacing traditional veterinary anatomy preservation and dissection with the
plastination technique in developing countries, because establishing such high-tech laboratories, equipped with costly
infrastructure, needs a lot of capital investment.
Materials and Methods: Animals can be purchased from the local market for preserving through the standard silicone
plastination technique, by following von Hagens’ previously described method.
Results: This technique may reduce the exposure to various harmful gases and toxic fumes, and provides fixed, non-
perishable, long-lasting veterinary anatomy specimens. After plastination, the preserved organs and animal bodies can
be used for undergraduate veterinary anatomy teaching for several years at the College of Veterinary and Animal
Sciences (CVAS), Jhang, Pakistan.
Conclusions: Hence, provision of equipment for plastination in the Anatomy Lab, CVAS, Jhang, will act as a double-edged
sword, because, on the one hand, it will protect the environment and health of veterinary anatomy professionals as well
as veterinary students, and on other hand, it will reduce the running costs and budget for the purchase of healthy and
expensive animals and toxic chemicals essentially needed for the dissection and embalming of equine, bovine, canine,
and avian species on a yearly basis.
Grant support: This work will be supported by Institutional Strengthening and Upgrade of Labs and Libraries number
HEC/ACAD/ISULL/2017/1323; the application is under processing.
The Journal of Plastination 30(2):22 (2018)
PRE-IMPREGNATION RATE OF FAT REMOVAL BY ACETONE DURING PLASTINATION
DEZSE KE, BAPTISTA CAC
University of Toledo, College of Medicine and Life Sciences, Department of Medical Education,3000 Arlington Avenue,
Toledo, Ohio,43614-1721 USA
Introduction: It is well known in the plastination community that the dehydrating and defatting phases are not
determined by quantitative measurements. Samples are soaked in acetone for varying intervals of time while using the
scientist’s discretion to determine when and how often the acetone bath should be changed. This makes the process
susceptible to human error and an imperfectly dehydrated/defatted sample. To minimize this inevitable error,
measurements were taken to ascertain the rate of penetration for acetone on a given sample, which can be
corroborated by histological images. Additionally, the overall effectiveness of the acetone was measured by collecting
the total amount of fat released after a set time interval.
Materials and Methods: Four x 7.6 cm (three inch) cubes of fat were collected from the lower back of one cadaver, and
their weight recorded. The lower back was chosen because of its low concentration of vessels and nerves. Samples
were soaked in cold (-25o C) acetone, and then room temperature (25o C) acetone for 21 days each. Acetone was
changed at 7-day intervals, and the “dirty”/used acetone was collected and total fat content was determined using a
rotary evaporator, and recorded. After three cold changes of acetone, the samples were deemed to be dehydrated
(>99% purity of acetone, the “dehydration phase”). The samples were removed from the cold temperature into room
temperature for another 21 day soak (a “defatting phase”) with three x 7-day interval changes. The dirty acetone was
saved, and fat content determined using a rotary evaporator, and recorded. Total fat extraction was measured from the
acetone/fat-mix remaining in the evaporator (approximately 50 ml). This mix was placed in a fume hood to allow
complete evaporation of the acetone from the fat. Twenty-four hours later, the remaining fat was weighed and total fat
extraction was recorded.
Results: The initial weight of each sample was as follows: Sample 1: 107.14 g; Sample 2: 74.19 g; Sample 3: 105.47 g;
Sample 4: 83.68 g. Figure 1 illustrates the results of 42 days of acetone soaking. In order to compare the samples, the
weight of fat extracted was divided by the initial weight of the sample to give a percentage (range 0.38% to 38.46%,
dependent on time). The rate of change between each time interval was calculated by using the “% of fat extracted” as
data points. These K-values were then used to compile a line of best fit representing all four samples’ fat extraction rate
over time (Table 1).
Figure 1
Table 1
The Journal of Plastination 30(2):23 (2018)
Discussion/Conclusion: Based on these data, several points can be concluded. Dehydration was complete after three
weeks and samples’ density was over 99%. This indicates that the samples are dehydrated, and defatting should be
started. This 21-day period is interesting because when you compare it to the defatting phase it disproves the initial
assumption that the defatting phase constitutes the majority of the fat loss. In other words, the dehydrating phase sees
more defatting than the actual defatting phase. This is supported by the fact that it takes seven less days to reach <10%
fat extracted. Additionally, it is concluded that the presence of density readings should be accompanied with each step
to ensure that dehydration and defatting is fully complete. Finally, we find that the plastination community should
adjust their procedures according to these findings in order to optimize aesthetic results and effectively save time. The
degree to which dehydration and defatting should be lengthened and shortened respectively, however, is still elusive
due to our samples not being representative of every potential plastinated specimen. Furthermore, we can safely say
that our goal for quantifying a “successful extraction of fat” was achieved and supported by density readings with the
change in slope of fat extraction. It is true that fat extraction can be proven quantitatively.
By collecting measurements of fat extraction during various times in the plastination process, this study provides
evidence of acetone’s efficiency in penetration of tissue samples. This offers incredible value to the plastination
community, because of the ability to point out where time and money can be saved, while also guaranteeing optimal
aesthetic quality of the tissues. This project also has the opportunity for extensive follow-up studies. Different
dehydrating/defatting agents and procedural techniques can be compared to the ones demonstrated in our study to
determine which is more effective. Samples can be taken from different locations on the body and from various organ
systems to show how different tissue compositions affect fat extraction. Studying changes in temperature during
different time intervals could also reveal further information on fat extraction. Finally, histological imaging should be
incorporated into this type of study. By coupling imaging with our study’s extraction data, we can observe cellular
changes as the fat concentration within the cell changes. There are a plethora of possible combinations to be explored,
and what we learn gives us the opportunity to perfect the plastination process and expand the plastination society’s
knowledge as a whole.
The Journal of Plastination 30(2):24 (2018)
SILICONE-INJECTED CADAVERIC HEADS FOR NEUROSURGICAL DISSECTION
DHINGRA R1, MOCHAN S1, SOUBAM P2, MISHRA S2, LALWANI S3, SURI A2, MAHPATRA AK2, ROY TS1
1Department of Anatomy, 2Department of Neurosurgery, 3Department of Forensic Medicine and Toxicology, All India
Institute of Medical Sciences, New Delhi, India
Introduction: Surgical simulation using cadaveric human heads is one of the most valid strategies, and is still considered
to be the gold standard for ex-vivo simulation among all the models available for neurosurgical training. The educational
value of these cadaveric heads for neurosurgical dissections can be enhanced by the injection of colored dyes in the
intracranial vascular tree. The knowledge of vascular anatomy, being integral to the all the neurosurgical procedures,
gets more clearly defined in these injected specimens. The technique of silicone injection in the cerebral vasculature of
human heads has been reported in the literature. However, the technique and materials for the injection of blood
vessels of cadaveric human heads in Indian conditions have not been standardized.
Materials and Methods: Four freshly-donated human cadavers were used. The technique of dye injection was
standardized in goat heads, and subsequently translated into humans. In the human cadavers, the carotid arteries and
internal jugular veins were dissected out from the carotid sheath, whereas the exposure of vertebral arteries required
deeper dissection of the neck. The arteries were then irrigated with warm normal saline till the returning fluid was clear
and free of blood. This was followed by perfusion with the embalming solution. The vessels were then clamped for 30
minutes to one hour. The silicone (red and blue) dyes were mixed with the catalyst, polymerization time was
standardized for the individual dyes and then injected into the common carotid arteries, vertebral arteries and the
internal jugular veins bilaterally. The injection was stopped once it was ensured that the entire vascular tree was filled
with the respective dyes. The cadavers were then decapitated after one week, and then immersed in fixation solution
consisting of 10% formalin and 20% ethanol, for 30 days.
Results: A small craniotomy was performed around the vertex of the cranium. The condition of the brain and quality of
the injection in the meningeal and cerebral vessels was observed. All the injected cerebral vessels, including external
and internal carotid arteries and their branches, along with the vertebro-basilar system, were filled with red silicone dye.
A well-opacified blue silicone dye was seen in the cerebral veins and dural venous sinuses. A favorable consistency of the
brain, without features of putrefaction, was observed. The brains were devoid of any disagreeable odor and could be
dissected for prolonged periods.
Conclusion: Self-curing silicone dyes injected in the cerebral vessels enhanced the educational potential of the human
cadavers, and served as a useful tool for the understanding of neurosurgical anatomical features, and learning various
surgical approaches.
The Journal of Plastination 30(2):25 (2018)
STUDENT-TO-STUDENT TOURS AND THE MUSEUM OF PLASTINATES: ENGAGING THE NEXT GENERATION OF
HEALTHCARE PROFESSIONALS AND THE PUBLIC
GANGISETTY AS, SHULKA V, LEE D, BAPTISTA CAC
University of Toledo, College of Medicine and Life Sciences, Department of Medical Education,3000 Arlington Avenue,
Toledo, Ohio,43614-1721 USA
Introduction: The Student-to-Student (S2S) program was launched in March 1986 by happenstance after three medical
students showed a plastinated heart to a group of primary school children. Through this outreach program the medical
students would bring wet specimens into community schools for the educational benefit of elementary and secondary
students. The medical students agreed that this experience not only benefited the community schools, but also helped
medical students to learn speaking skills and gain confidence in meeting the public. The S2S program was very successful
but the excessive number of requests from the community schools made the coordination and availability of the medical
students impractical. The solution was to bring the school children to meet the medical students on the medical college
campus. Until 2013, medical students provided a presentation to the school groups in the gross anatomy lab, followed
by a demonstration of select plastinated specimens. With the creation of the Interactive Museum of Anatomy and
Pathology in 2013, the medical students were provided with a high-quality venue for the delivery of more structured
tours. The Plastination Museum was created to provide valuable educational resources not only for the University of
Toledo students in healthcare and related disciplines, but also for the public at large. It was also created with the
purpose of educating students in the Toledo school districts and vicinity. The Museum is located in the College of
Medicine, Health Science Campus, at the University of Toledo. The Museum provides a dynamic study and teaching
space. The Museum was named after Dr. Liberato DiDio, former chairman of the Department of Anatomy, and Dr. Peter
Goldblatt, former chairman of the Department of Pathology. The establishment of the Museum and its use will be
discussed.
Materials and Methods: The “construction” of the Plastination Museum started 20 years ago when several plastinated
specimens were created as a resource of didactic material to advance the teaching mission of the department of
anatomy. Monies received by the plastination laboratory’s specimen preparation services (for other institutions)
provided funding for the project. The original project budget rationale involved the following: relocating the plastination
lab to free up space for the museum, renovating the old space with painting, installation of an acoustic ceiling and tile
flooring, and the purchase of new wooden/glass display cabinets. The museum was constructed in three months and
now houses approximately 300 specimens comprising the anatomical and pathological collections of the College of
Medicine. Each cabinet was divided according to function (digestion, breathing, circulation, filtration, control, support,
development, and comparative). Each cabinet has an android tablet containing explanations of each specimen, thereby
providing a self-guided tour.
Results: The space allocated to the Museum, even though small, has been used appropriately by medical students and
other healthcare students and professionals. Thousands of high school students from the Northwest Ohio and southern
Michigan area have toured the Museum through the S2S Program. The S2S educational outreach program is organized
by 1st and 2nd year medical students, who coordinate the Museum tours. Since 2013, the number of community
students touring the Museum each year has increased as follows: 1,291 (2013-2014), 1,375 (2014-2015), 1,707 (2015-
2016), 1,981 (2016-2017) and 2,507 (2017-2018).
Conclusion: The Plastination Museum was created with a limited budget but has proven to be an excellent method for
housing the anatomical and pathological collections of the College of Medicine in a single location. The Museum has also
proven to be an enormous asset to educate community students and the general public on normal human anatomy and
diseases.
The Journal of Plastination 30(2):26 (2018)
NEW PERSPECTIVES IN PRACTICAL LESSONS USING PLASTINATED PARASITES IN VETERINARY DEGREE
1GONZÁLVEZ M, 1ORTIZ J, 1RUIZ DE YBÁÑEZ R, 2LÓPEZ-ALBORS O, 2LATORRE R
1Department of Animal Health (Parasitology and Parasitic Diseases); 2Department of Anatomy and Comparative
Pathological Anatomy; Regional Campus of International Excellence “Campus Mare Nostrum”, University of Murcia,
Murcia, Spain.
Introduction: Plastinated specimens have been used as an education tool in different subjects, mainly related with
anatomy. However, few references exist about the plastination of parasites, most of them about the necessity of
changes in conventional plastination protocols.
Objectives: The goal of this study was to evaluate the use of plastinated parasites as an innovative teaching and learning
tool in parasitology practicals.
Materials and Methods: Seven different species of plastinated macroparasites were used: arthropods (Oestrus ovis);
nematodes (Parascaris equorum, Ascaris suum, Macracanthorynchus hirudinaceus) and platyhelminthes (Fasciola
hepatica, Dicrocoelium dendriticum, Taenia sp.). A set of 89 students were involved, all of them assessed for previous
background on the subject before any contact with the parasites. The experimental group of students used plastinated
specimens in the practicals, whereas the control group used conventional wet specimens (preserved in ethanol and
formaldehyde). After each practical session both groups were assessed with the same evaluation and also had the
opportunity to score their level of satisfaction with the material (plastinated or not).
Results: The scores relating to knowledge and satisfaction after practical sessions did not show statistically significant
differences between both groups (p>0.05). Most of the students in the experimental group highlighted the facility of
handling the plastinated specimens.
Conclusion: Plastination can be used to replace the traditional wet parasites in the practical sessions of Veterinary
Parasitology without a decrease in the students’ performing and evaluation scores. As has been already demonstrated in
other subjects such as Anatomy, this is an effective way of avoiding the traditional use of wet, irritant, toxic and even
carcinogenic chemicals in Parasitology.
The Journal of Plastination 30(2):27 (2018)
ACCESSORY LIGAMENT OF THE DEEP DIGITAL FLEXOR TENDON IN THE HORSE FORELIMB. A FLUORESCENCE STUDY
WITH E12 PLASTINATED SECTIONS
GUILABERT R, LÓPEZ ALBORS O, JORDAN J, LATORRE R
Department of Anatomy and Comparative Pathology, Campus International Mare Nostrum, University of Murcia, Murcia,
Spain.
Introduction: The accessory ligament of the deep digital flexor tendon (AL-DDFT) is important for the correct function of
the horse forelimb. Recently, a lateral fibrous lamina, binding the AL-DDFT to the superficial digital flexor tendon (SDFT)
has been described by ultrasonography and MRI (FL-AL-DDFT). The main goal of this study was to describe its
topographic anatomy by E12 plastinated serial cross-sections. The low refractive index of the epoxy resin E12, with its
minimal shrinkage during polymerization, makes it the method of choice to study different tissues, in different planes of
sectioning, from macroscopic to microscopic levels. The absence of manipulation and decalcification ensures that the
topography of anatomical structures is unaffected. The removal of fat tissue allows connective tissue, blood vessels, and
nerves to be identified quite clearly, without suffering any manipulation.
Materials and Methods: Ten forelimbs from two adults and four yearlings were used. Dissection techniques and epoxy
(E12) transparent plastinated serial cross-sections were used to study in detail the topographical relationship between
the AL-DDFT with the SDFT. The thickness of the sections was 2 mm. Plastinated sections were scanned with 1200 dpi
resolution. Sections were studied under a Leica stereoscopic microscope. Plastinated collagen tissue had endogenous
autofluorescence (488-nm excitation). Differentiation among fibers was based on their anatomical distribution and
florescent intensity. A Nikon confocal scanning microscope was used to observe the plastinated sections. Thickness
frame of the optical sections used was 15 – 20 μm and 10X.
Results: At microscopic level the FL-AL-DDFT appears as a fibrous band in close association with the synovial membrane
of the common digital flexor vaina synovialis (CDFVS). The fibrous connective tissue includes some capillaries and
melanin pigment, and is likely to be covered by synovial epithelium at inner and outer sides. Thus, all along its proximo-
distal projection, the FL-AL-DDFT may establish a lateral compartment in CDFVS which has not been described in detail
so far. Further studies at higher (microscopic) magnification are required.
Conclusion: In addition to a mechanical bond between the AL-DDFT and the SDFT, the FL-AL-DDFT may be relevant for
the synovial compartment between the DDFT and SDFT, which must be considered in all the pathologies affecting the
function of the flexor component of the distal thoracic limb. Further studies are required, to further characterize the
anatomical features of the FL-AL-DDFT depending on the age, breed, or different types of lameness.
The Journal of Plastination 30(2):28 (2018)
INFLUENCE OF SILICONE VISCOSITY IN TISSUE SHRINKAGE DURING THE PLASTINATION IMPREGNATION PHASE
1JUVENATO LS, 1MONTEIRO YF, 2,3BITTENCOURT APSV, 4BAPTISTA CAC, 1,2BITTENCOURT AS 1Department of Morphology, Federal University of Espirito Santo, Brazil, 2Biochemistry and Pharmacology Graduation
Program, Federal University of Espirito Santo, Brazil, 3Department of Physiology Science, Federal University of Espirito
Santo, Brazil, 4College of Medicine and Life Sciences, University of Toledo, Ohio, USA
Introduction: Impregnation is the most important step in the plastination process, and is decisive for good specimen
quality. The viscosity of the polymer is an important variable in the level of shrinkage observed during the impregnation
process. Because shrinkage is one of the disadvantages of the technique, several researches seek ways to circumvent or
reduce this retraction. The objective of this work was to test the influence of the viscosity of three different silicones on
the shrinkage during the forced impregnation process at different temperatures.
Materials and Methods: For this experiment, 24 bovine kidneys were used, which were previously fixed in 10% formalin
for one month, and were then organized into 2 groups: 12 for plastination at room temperature (RT) (25° C) and 12 for
plastination at low temperature (-15° C). The silicones used were: Poliplast 1 (P1) and Poliplast 10 (P10) (Polisil Silicones
Ltd.) and S10 (Biodur). Statistical analysis was performed using one way ANOVA followed by Duncan's post-hoc test or
by two way ANOVA. The level of significance was considered p <0.05.
Results: Comparing the three silicones of different viscosities, P10 showed the greatest shrinkage when used in the
impregnation, with a mean of 39% at room temperature and 42.5% at low temperature. This is due to the fact that it has
the approximate viscosity of 1250 mPas at 20° C (value supplied by the manufacturer), that is, the highest viscosity
among the compared silicones (2.5X more viscous than S10). It is known that the higher the viscosity, the slower the
speed at which the fluid moves, thus presenting a greater difficulty of penetrating the tissues. In the impregnation with
P10, the shrinkage was much more evident at low temperature. Silicone P1 presented the best results, since the mean
shrinkage was 4.75% at room temperature, and 15.5% at low temperature.
Conclusion: Polisil® P1 silicone appeared to be a good alternative to Biodur® S10 silicone, since it has a viscosity of at
least 4X at room temperature, and 3X at low temperature, making it a viable substitution silicone of reference, since its
cost of commercialization is more accessible in Brazil.
Grant Support: CNPq (458328/2013-8; 440729/2017-3); FAPES (5537479411); UFES-PROEXT and CAPES.
The Journal of Plastination 30(2):29 (2018)
PRINCIPALS OF THE E12 TECHNIQUE
LATORRE R
Veterinary Anatomy and Comparative Pathology, University of Murcia, Spain.
The E12 technique was designed to preserve transparent body sections. Some technical aspects will now be emphasized
regarding the E12 protocol.
Sectioning of specimens:
Fresh or fixed specimens can be used. Vascular injections should be with epoxy instead of latex or silicone. The specimen
should be frozen at the lowest possible temperature, at least -70 or -80 °C before, to obtain the transparent sections.
Sections of 1.5-3 mm thickness are made with a band saw, and it is recommended to use dry ice or liquid nitrogen. The
use of -40 °C acetone during the cleaning of the sawdust prevents sections from thawing.
Dehydration by freeze substitution and defeating:
Dehydration of sections is carried out with cold acetone. Moreover, it is necessary to remove fatty tissue to get the
highest transparency. This is done with several baths in acetone or methylene chloride at room temperature.
Forced impregnation:
The impregnation solution employed consists of epoxy E12, plus the hardener E1. The dehydrated sections are
immersed in the impregnation mixture and forced vacuum is applied at room temperature for 6-12 h. Impregnation is
complete when the pressures reached are below 5 mmHg.
Polymerization:
Polymerization must be done immediately after finishing the impregnation, to prevent polymerization in the
impregnation chamber. Impregnated sections are introduced into flat glass chambers surrounded by E12+E1. The
polymerization agent is the temperature: 40 °C for 4-6 days, combined with the hardener E1. There are alternatives to
the flat glass chambers like the sandwich method, which saves time and is easier.
The Journal of Plastination 30(2):30 (2018)
RESULTS OF A RECENT SURVEY ABOUT ACTIVE PLASTINATION LABS
LATORRE R1, BAPTISTA C2, HENRY R3, TUNALY S4, STARCHIK D5, SUI H-J6, LÓPEZ ALBORS O1
1 Department of Anatomy and Comparative Pathology, Veterinary Faculty, University of Murcia, Spain, 2 Department of
Medical Education, College of Medicine, University of Toledo, Toledo, Ohio, USA, 3College of Veterinary Medicine, Lincoln
Memorial University, Seymour, Tennessee, USA, 4 Department of Anatomy, Faculty of Medicine, TOBB University of
Economics and Technology, Ankara, Turkey, 5International Morphological Centre, Saint Petersburg, Russia, 6 Department
of Anatomy, Dalian Medical University, Dalian, China
This survey was addressed to the plastination community registered in the database of the ISP, and the historical list of
participants in the workshops held in Murcia (Spain). The survey was open for a month (May 2018), submitted to 592
people (academics and technical staff), and answered by 62. Results showed that most participants were academics
(61%), males (61.3%) and currently active in plastination techniques (74.2%). Around 66 % were members of the ISP, and
at least 50% of them had been involved with plastination for a minimum of 5 years. However, 40 % of the participants
never attended ISP conferences, or just once (20%). Plastination is mainly carried out in anatomy labs (36.8% human,
23.2% vets), but also in other venues or with other materials: museums (15.8 %), zoology labs (8.4%), fungi, flowers, etc.
There is still a great interest in plastination workshops, especially those focused on advanced topics such as vascular
injection, plastination of hollow and flexible organs, brains, clinical applications, and the research potential of
plastination. Other suggested topics were: different methods of sectioning and casting in sheet plastination, brain
reconstruction with P40 slices, staining of ultrathin epoxy slices, typical mistakes in plastination, and plastination for
educational purposes. In conclusion, there is a need to define a strategy aimed at improving the coupling of the active
plastinators with the ISP; plastination is quite well consolidated in non-anatomical fields; and there is an important
potential for workshops focused on advanced topics in plastination.
The Journal of Plastination 30(2):31 (2018)
A WORKFLOW UTILIZING PLASTINATIONS FOR VISUALIZING AND ANNOTATING X-REALITY ANATOMICAL MODELS
WITH HEAD MOUNTED AND CONSOLE BASED TECHNOLOGIES
LOZANOFF S, THOMPSON J, HONG TM, LOZANOFF BK, LABRASH S
Department of Anatomy, Biochemistry, and Physiology, University of Hawaii, John A. Burns School of Medicine,
Honolulu, HI, USA
Objectives: Plastinated anatomical specimens are very useful as educational instruments. However, utilization in self-
directed learning activities remains problematic since plastinations cannot be easily annotated. Augmented reality (AR)
may provide a novel method to enable visualization and annotation of plastinated specimens. The purpose of this study
was to develop an annotation system within an AR environment facilitating the use of labeled plastinated specimens.
Materials and Methods: A plastinated heart was imaged using a photogrammetry system. The digital model was
polished using Z-brush software and then imported and processed using Vuforia software within the Unity3D engine.
The 3D digital model of the plastination was converted into target points that recognized and tracked the heart in real
time. Annotations could be added directly onto the tracked object within the Unity environment. In conjunction with a
smartphone or head-mounted display (HMD) such as the Microsoft Hololens, plastinations could be supplemented with
annotations or additional contextual information within a text box display. The annotated virtual model can also be
used for X-Reality applications as well as 3D printing.
Results: Results showed tight correspondence between the heart and associated annotations. The student could
physically handle the plastinations while simultaneously visualizing the annotations and supplemental text box
information. Qualitative comments among students indicated that they could utilize the models independently or in
small groups, facilitating independent learning.
Conclusion: While such workflows were originally meant for CAD models and gaming, our application of
photogrammetric methodologies enable real world plastinations to be overlaid with virtual assets and information. This
AR tool in conjunction with plastinated resources may also be useful in clinical training sessions or museum displays.
Work is being directed at developing quantitative tools to assess the educational usefulness of AR annotated
plastinations.
The Journal of Plastination 30(2):32 (2018)
INFLUENCE OF THE TEMPERATURE ON THE VISCOSITY OF DIFFERENT TYPES OF SILICONE
1MONTEIRO YF, 1JUVENATO LS, 2,3BITTENCOURT APSV, 2SIQUEIRA BMM, 2MONTEIRO FC, 4BAPTISTA CAC, 1,2BITTENCOURT
AS
1Department of Morphology, Universidade Federal do Espírito Santo, Brazil, 2Biochemistry and Pharmacology
Graduation Program, Federal University of Espirito Santo, Brazil, 3Department of Physiology Science, Universidade
Federal do Espírito Santo, Brazil, 4College of Medicine and Life Sciences, University of Toledo, Ohio, USA
Introduction: The term silicone, or polysiloxane, describes mixed polymers of organic and inorganic materials, whose
crude formula is [R2SiO]n, where ‘R’ are organic groups such as methyl, ethyl and phenyl. The main silicone used in the
plastination process is polydimethylsiloxane (PDMS), referring to linear polymers, where the organic radical is methyl.
One of the main external factors influencing the viscosity of a silicone is the temperature. The objective of this work
was to test the influence of temperature on the viscosity of three silicones of different molecular weights (Biodur® S10,
Polisil® P10 and P1), commonly used in the plastination technique.
Materials and Methods: For this study, the RheolabQC model rotational rheometer was used to measure the dynamic
viscosities of the chosen polymers at the following temperatures: -5, 0, 5, 10, 15, 20, 25, 30, and 35° C. From the 9
measurements of viscosities obtained from each sample, a viscosity vs. temperature graph was constructed. The
equation of the dynamic viscosity curve of each polymer was analyzed.
Results: Poliplast® 1 silicone had a much lower viscosity compared to other silicones (about 80 mPa.s at 25° C and 550
mPa.s at -25 ° C). Poliplast® 10 silicone presented the highest viscosity of the polymers analyzed (approximately 1180
mPa.s at 25° C and 3730 mPa.s at -25° C). The Biodur® S10 silicone showed an intermediate viscosity (about 410 mPa.s
at 25° C and 1500 mPa.s at -25° C). The different viscosities found in the tested silicones are determined by the degree
of polymerization. The larger the silicone chain (P10> S10> P1), the more intermolecular bonds are made with adjacent
molecules and thus the less fluidity of the chain.
Conclusion: We conclude that Polisil® P1 silicone presented the best physico-chemical characteristics of the tested
silicones for plastination, because it has high fluidity and low viscosity. It is noteworthy that the viscosity of Polisil® P1 in
cold impregnation temperature (-15° C) is still lower than the viscosity of the Biodur® S10 (control) at room
temperature (20-25° C). We also conclude that knowledge of the intrinsic and extrinsic physicochemical characteristics
of the silicone, and its dynamic viscosity is helpful in choosing the ideal silicone for use in the cold or room temperature
plastination techniques.
Grant Support: CNPq (458328/2013-8; 440729/2017-3); FAPES (5537479411); UFES-PROEXT and CAPES.
The Journal of Plastination 30(2):33 (2018)
ENHANCING EFFECTIVE AND INNOVATIVE LEARNING IN ANIMAL ANATOMY DAF200
1OLIVIER W, 2VAN MARLE-KὅSTER, E
1Department of Anatomy and Physiology, Faculty of Veterinary Science, Onderstepoort 2Department of Animal & Wildlife Sciences, University of Pretoria, Gauteng, South Africa
Animal science consists of three major disciplines, namely: animal physiology, animal breeding, and genetics and animal
nutrition. Animal anatomy and physiology (DAF200) is presented as a year module where practical sessions make up an
important part of presentation and learning, as students need to observe and experience hands-on the texture,
conformation, and anatomical organization, including splanchnology and topography of the animal body, for an
improved understanding. Furthermore, reproduction physiology is presented on a third-year level, and students need to
handle the reproductive organs before techniques such as artificial insemination can be practised on live animals.
Approximately 120 and 75 students annually need to be taught in the anatomy and reproduction physiology modules,
respectively.
In the past, dissected material was preserved in formalin, posing health hazards for both lecturer and students. This has
been replaced with dissection of fresh cadavers for the past decade, but this practice has become costly, and has animal
welfare implications, which can be regarded by some as unnecessary slaughtering and a waste of used carcasses. The
use of fresh material may also pose other potential health risks to students and lecturers, such as Rift Valley fever or
brucellosis.
In 2017, the Department of Animal & Wildlife Sciences decided to decrease the number of practical sessions on
slaughtered sheep, due to costs and concerns with animal welfare and health risks. This has implications for effective
learning outcomes, which led to the investigation of alternative methods to enhance future teaching and learning.
A project is underway where the different organs will be plastinated to be used during practical sessions. To enhance
teaching and learning, plastinated organ specimens and even complete cadavers have become commonplace in human
and veterinary faculties. In this project the various models will be developed, for example for the complete digestive
system and reproductive system. An Anatomy Model Teaching Room is envisaged, representing the anatomy sections
required for the modules presented by the department of Animal and Wildlife Science.
The use of plastinated specimens holds a number of advantages, including more effective hands-on teaching, fewer
health risks for students and lecturers, more cost-effectiveness in the long-term. It will provide a sustainable alternative
for practical teaching of anatomy and physiology in the Animal Sciences.
The Journal of Plastination 30(2):34 (2018)
COMPARISON OF BACTERIA ISOLATED IN THE ANATOMICAL LABORATORY AND OTHER BIOMEDICAL LABORATORIES
IN THE MEDICAL FACULTY OF MUHAMMADIYAH UNIVERSITY OF PURWOKERTO, INDONESIA
1PUTRA RAN, 1PUTRI PM, 2FEBRIYANTI RW, 3PANGESTIKA TR
1Anatomy Department, Universitas Muhammadiyah Purwokerto, Indonesia, 2Microbiology Department, Universitas
Muhammadiyah Purwokerto, Indonesia, 3Undergraduate student of the Medical Faculty, Universitas Muhammadiyah
Purwokerto, Indonesia
Background: Infectious diseases are still in the top 10 causes of death in the world. The laboratory is a specific
environment for the development of infectious bacteria. The anatomical laboratory, as a cadaver preparation area, plays
an important role in the development of bacteria that can cause laboratory-acquired infection. Comparisons of bacterial
isolates detected in the anatomical laboratory, and other biomedical laboratories, have not hitherto been widely known.
Objectives: To compare the bacterial isolates from the anatomical laboratory and other biomedical laboratories, at the
Medical Faculty of Muhammadiyah University of Purwokerto, Indonesia.
Materials and Methods: The sampling technique used the settle plate method for environmental monitoring. Samples
were taken with 5-point surface swabs from the anatomical laboratory and other biomedical laboratories (Histology,
Microbiology and Clinical Pathology Laboratories) at the Medical Faculty, Muhammadiyah University of Purwokerto.
Samples were grown on blood agar, nutrient agar and MacConkey agar, and then subjected to Gram staining and
biochemical tests. Bacterial isolate data were analyzed descriptively.
Results: The total number of samples for this study was 20 samples. The bacteria identified were Staphylococcus aureus,
coagulase-negative staphylococci, Pseudomonas sp., Enterobacter sp., Vibrio vulnificus, Aeromonas hydrophila, E. coli,
Enterobacter intermedius, Klebsiella pneumonia, Serratia fonticola and Streptococcus sp. The Microbiology Laboratory
had the highest contamination (63 colonies), while the Anatomical Laboratory was in third rank (17 colonies). The
bacterial isolates found varied per laboratory. Vibrio vulnificus was the dominant bacterium in the Anatomical
Laboratory (41.6% of 17 colonies), unlike the Histology Laboratory (Enterobacter intermedius), the Microbiology
Laboratory (Pseudomonas sp.) and the Clinical Pathology Laboratory (Enterobacter intermedius).
Conclusion: Different bacterial isolates were found in each laboratory. The Anatomical Laboratory was dominated by
Vibrio vulnificus, in contrast to the other three biomedical laboratories.
The Journal of Plastination 30(2):35 (2018)
COMPARISON OF BACTERIA NUMBER BETWEEN THE ANATOMICAL LABORATORY AND OTHER BIOMEDICAL
LABORATORIES IN THE MEDICAL FACULTY OF MUHAMMADIYAH UNIVERSITY OF PURWOKERTO, BEFORE AND AFTER
DISINFECTION WITH 70% ALCOHOL
1PUTRA RAN, 1PUTRI PM, 2FEBRIYANTI RW, 3PANGESTIKA TR
1Anatomy Department, Universitas Muhammadiyah Purwokerto, Indonesia, 2Microbiology Department, Universitas
Muhammadiyah Purwokerto, Indonesia, 3Undergraduate Student of Medical Faculty, Universitas Muhammadiyah
Purwokerto, Indonesia
Background: The laboratory is a specific environment for the growth of pathogenic and non-pathogenic bacteria. The
anatomical laboratory, as a cadaver preparation area for medical students’ practicals, can be a route for bacterial
transmission. Disinfection is a useful process to reduce bacterial contamination, using chemicals such as 70% alcohol, to
prevent bacterial transmission.
Objectives: To compare the number of bacteria between the anatomical laboratory and other biomedical laboratories,
of the Medical Faculty of Muhammadiyah University of Purwokerto, Indonesia, before and after disinfection with 70%
alcohol.
Materials and Methods: Duplicate samples were taken from the anatomical laboratory and other biomedical
laboratories (Histology, Microbiology, and Clinical Pathology Laboratories) in the Medical Faculty of Muhammadiyah
University of Purwokerto. The first and second samples were taken from the surface swab of each sampling point before
and after disinfection. Samples were grown on blood agar, nutrient agar and McConkey agar, and then subjected to
Gram staining and biochemical tests. Data of bacterial number differences before and after disinfection were analyzed
using a paired t-test, while data of bacterial number comparison between each laboratory was analyzed by using an
independent t-test and Mann Whitney test.
Results: The average number of bacteria before disinfection was higher than after disinfection. The difference between
the two groups is very significant (p = 0.010, p <0.05). The Clinical Pathology Laboratory had the highest bacterial
contamination, and the Histology Laboratory had the lowest bacterial contamination. There was no bacterial number
difference between the Anatomical Laboratory and the other biomedical laboratories (p> 0.05).
Conclusion: The disinfection procedure is useful in reducing bacterial contamination. There was no significant bacterial
number difference between the Anatomical Laboratory and other biomedical laboratories. The number of bacteria in
the Anatomy Laboratory was not higher than the Clinical Pathology Laboratory and the Microbiology Laboratory.
The Journal of Plastination 30(2):36 (2018)
COMPARISON OF FIXATIVE SOLUTIONS AND THEIR INFLUENCE ON PLASTINATION PROTOCOLS IN HEPATIC,
MUSCULAR AND ARTERIAL TISSUE
1RAMOS ML, 1DE PAULA TAR, 2SARRIAS L, 2ALBORS OL, 2LATORRE RM
1 Department of Veterinary Medicine, Federal University of Viçosa, Brazil, 2 Department of Anatomy and Comparative
Pathology, University of Murcia, Spain.
Introduction: Plastination allows the use of specimens for histology purposes, but there are no references regarding
specific results of different tissues. The objective of this study is to investigate if plastinated tissue samples from artery,
liver and skeletal muscle can be used for routine histology, and how the fixation and plastination protocol used can
influence the results.
Materials and Methods: Nine tissue samples from each organ (artery, liver and skeletal muscle) were fixed with three
different solutions. Group I: 10% formalin; Group II: 2.5% formalin and Group III: Cambridge solution (an alcohol-based
fluid containing phenol). Two samples from each group were plastinated with Biodur S10 technique, half of them cured,
with the standard protocol, and the other half uncured (without curing). Sodium methoxide was used for the
deplastination protocol. After deplastination, samples were paraffin-embedded for histology. Microtome sections were
stained with H&E. The last sample from each group was a control sample, these were not plastinated, and processed
following the classical histology protocol. During evaluation of results, each section was assigned an overall score based
on the histological quality of the cellular components of the tissue. Sections were scored from 1 to 3 (1, good; 2,
satisfactory/useful; 3, poor).
Results: Satisfactory sections were obtained from all tissues. The control sample from group I (formaldehyde 10%)
resulted in consistently good quality of the tissue histology. Control samples from groups II and III resulted in
consistently useful quality sections. In Group I, fixed with formaldehyde 10%, artery and skeletal muscle produced useful
slides, and liver produced poor slides. In Group II, fixed with formaldehyde 10%, artery produced good slides, liver and
skeletal muscle produced poor slides. For Groups I and II, fixed with Cambridge solution, artery produced poor slide
quality, liver and skeletal muscle produced useful slides.
Conclusion: The tissues have different behaviors with different fixatives and plastinated protocols.
The Journal of Plastination 30(2):37 (2018)
HISTOLOGICAL EVALUATION OF SILICONE PLASTINATED SAMPLES PROCESSED UNDER DIFFERENT CONDITIONS.
1RAMOS ML, 1DE PAULA TAR, 2SARRIAS L, 2ALBORS OL, 2LATORRE RM
1 Department of Veterinary Medicine, Federal University of Viçosa, Brazil, 2 Department of Anatomy and Comparative
Pathology, University of Murcia, Spain.
Introduction: Plastinated specimens can be used for light microscopy studies by means of deplastination techniques to
obtain histological slides. The results of these deplastination techniques are susceptible of several improvements. One of
them would be to minimize the negative effect of the fixation. On the other hand, previous authors have described a
better preservation of the histological structure when the curing step from plastination protocol is avoided. Therefore,
this study aims to assess the effect of several different fixation solutions, as well as the use of curing during plastination,
on the histological structure of silicone-plastinated samples.
Materials and Methods: Twelve samples from each of the following nine organs: artery, esophagus, liver, small
intestine, large intestine, skeletal muscle, nerve, pancreas, and trachea, were used for this study; in total 108 samples.
Samples from each organ were divided into three groups (four samples each) depending of the fixative solution used.
Group I: samples fixed with formaldehyde 10%; Group II: samples fixed with 2.5% formaldehyde, and Group III: samples
fixed with Cambridge embalming solution. After fixation, tissue samples were plastinated using the S 10 Biodur® silicone
technique, however, each group was divided into two subgroups, depending on whether samples were cured or not,
with two samples per organ in each subgroup. After the plastination process, tissues were deplastinated using sodium
methoxide. After deplastination, specimens were processed for paraffin embedding. Sections were stained with routine
H & E. The histology preservation was evaluated using a scale of 1 to 3, being: “1” the absence of morphological
changes, good morphology; "2" minor morphological alteration; "3" intense morphological change.
Results: Tissues from groups I and III, fixed with 10% formaldehyde solution or with Cambridge solution and curing
protocol, demonstrated the best histological results under light microscopy (P˂0.05). Similar results (P˂0.05) were
observed in the slides obtained from uncured specimens of these two groups. Tissues from group II, fixed with 2.5%
formaldehyde, cured or uncured, produced histological slides of level 3, with intense morphological changes, compared
with groups I and III (P˂0.05).
Conclusion: Plastinated specimens fixed with either 10% formaldehyde or Cambridge solution have a good histological
preservation after deplastination. The curing step during the plastination procedure does not affect the histological
results.
Grant support: Coordenação de Aperfeiçoamento de Pessoal de Nível Superior – CAPES - Brasil.
The Journal of Plastination 30(2):38 (2018)
NEAR FIELD COMMUNICATION (NFC) DEVICES AND PLASTINATION: AN INTEGRATED TUTORIAL SYSTEM TOOL FOR
SELF-DIRECTED LEARNING
ROSOLOWSKI BL, TENBRINK P, BAPTISTA CAC
University of Toledo, College of Medicine and Life Sciences, Department of Medical Education, 3000 Arlington Avenue,
Toledo, Ohio, 43614-1721 USA
Introduction: The purpose of integrating Near Field Communication (“NFC”) with plastination is to provide students with
a self-directed learning tool that offers intelligible flexibility for the study of anatomy. The objective of this learning tool
is to offer students an interactive learning environment outside of the usual academic setting. This self-directed learning
tool allows students to point at different parts of the plastinated anatomical specimen, which are tagged with smart
chips, directing students to a web-page that presents a description of the specific anatomical muscle, nerve, or artery to
which the NFC wand or smartphone is directed.
Materials and Methods: An application was designed to integrate the NFC reader, NFC tags, smartphones, Android
tablet devices, and plastinates. The NFC is a set of standards for radio communication between tablets, smartphones
and similar devices that allows communication with each other by proximity or touch. We used a vWand
(SistelNetworks) in order to provide NFC connectivity for the Android tablet using Bluetooth connection. This wand
allowed flexibility and uniformity within the system. Fifteen structures were identified in silicone- plastinated brain
slices. Each anatomical description was entered in HTML format and stored as a webpage in the server. An NFC tag was
attached to each structure and the URL web address of each structure was written into the tag using a vWand Pro
Android app. When the tag was read by the vWand, an application launched the web browser containing a link to the
webpage with the anatomical description of the structure. Similar techniques can be used with a smartphone by
downloading an NFC Reader application. The smartphone will use its NFC reader to detect the NFC smart-chip tags on
the plastinate. The smartphone will then open the URL containing the desired information within the NFC Reader
application. The integration of the smartphone will allow for flexibility and convenience.
Results: The prototype consisted of 15 NFC tags implanted in multiple brain slices. Each tag when opened corresponded
to a URL containing the description of the structure.
Conclusions: The NFC device is a platform for self-directed learning that integrates plastinates with digital technology,
providing flexibility for the study of anatomy and pathology outside the usual academic settings. In addition, it provides
an interactive environment, structure, and guidance to the student, and a powerful educational tool to promote
meaningful learning through the integration of words, sounds and visuals.
The Journal of Plastination 30(2):39 (2018)
PLASTINATION OF BOVINE EYES AT ROOM TEMPERATURE
SAVOINI EH
Department of Biology, Yavapai College, Arizona, USA
Introduction: Bovine eyes are used for dissection in a wide range of anatomy courses. Use of plastinated bovine eyes
will have long-term cost and time benefits for the institution and instructor.
Materials and Methods:
Acquisition& Preservation
Bovine eyes (10) were obtained from slaughter within 1-4 hours from the time of death. The optic nerve and ocular
muscles were exposed via dissection. Eyes were placed in an aqueous solution (40% ethanol and 10% formalin
concentrate) for 14 days, and rinsed in a circulating water bath for 2 days.
Dehydration
An 18 gauge needle was inserted through the sclera into the posterior chamber approximately 1 cm posterior to the
edge of the cornea. A second 18 gauge needle, attached to a 10 ml syringe, was inserted into the posterior chamber on
the opposite side. Some vitreous humor (2-5 ml) was removed and discarded. Acetone (>98%) was injected into the
posterior chamber via the syringe, positioned so that the acetone entered the eye from the bottom, with the first
needle, at the top, as a vent. Acetone injection ceased when it began to spill out of the top needle. Acetone-filled eyes,
with both needles inserted, were placed in a bucket of acetone (>98%) for 10 days, after which, each eye was
reinjected/flushed with 10 ml of acetone (>98%) through the bottom needle, with old acetone escaping via the top
needle. Eyes were placed into another bucket of acetone (>98%) for 10 days, repeating for a total of four cycles.
Impregnation
A mixture of silicone polymer (North Carolina Silicones, NCS-X) and 3% catalyst (North Carolina Silicones, NCS-III) was
injected into the posterior chamber of each eye as acetone had been previously. Eyes were then submerged in the
polymer-catalyst mixture and placed into a vacuum chamber, at room temperature. Pressure was decreased from
atmospheric to 20 mmHg, over 5 days.
Dissection & Curing
Eyes were removed from the chamber and allowed to drain (2-7 days). The lateral surface of the posterior chamber was
cut with a scalpel. The now opaque and gelatinous vitreous humor was delicately removed, leaving the retina intact and
lens in place. Eyes were placed with the opening downward on an absorbent cloth, to allow the posterior chamber to
drain (2 days), then placed inside an enclosure with vaporized NCS-VI (North Carolina Silicones) for 12 hours. Eyes were
turned over and exposed to NCS-VI for another 12 hours, then removed.
Results: Dissection after silicone polymer impregnation of bovine eyes provided the best means to maintain placement
of the retina and lens.
Conclusions: Plastinated bovine eyes are a useful tool in any anatomy course, due to the same general gross anatomy
features as human eyes, but are large enough to easily visualize structural features. They are commonly available fresh
from slaughter, and produce excellent plastinated specimens.
The Journal of Plastination 30(2):40 (2018)
PLASTINATION OF LARGE MAMMAL KNEES
SAVOINI EH
Department of Biology, Yavapai College, Arizona, USA
Introduction: Plastinated knees that retain the ability to flex and extend are a significant improvement over rigid
cadaver knees or plastic models.
Methods:
Acquisition& Preparation
Elk knees were obtained from a slaughterhouse. Connective tissue, remaining muscle, and most of the periosteum was
removed to expose the ligaments of the knee. Tubing was placed between the medial and lateral collateral ligaments
and their respective medial and lateral condylar surface of the femur. The tubing was to prevent the ligaments from
shortening during the dehydration process.
Dehydration
After dissection, knees were placed in the first acetone bath (>98%) for 10 days, followed by three more changes of
acetone (>98%) baths, each 10 days in duration.
Impregnation
Knees were submerged within in a mixture of silicone polymer (North Carolina Silicones, NCS-X) and 3% catalyst (North
Carolina Silicones, NCS-III) and placed in a vacuum chamber at room temperature. Pressure was reduced incrementally
from atmospheric pressure to 20 mmHg over 5 days.
Curing& Flexing
Knees were removed from the polymer-catalyst mixture. Gripping the femur and tibia, knees were made to flex and
extend several times. Knees were flexed and extended daily for 5-7 days and then every 2-3 days over two weeks. After
3 months, knees were cured by exposure to the cross-linker (North Carolina Silicones, NCS-VI) overnight inside a sealed
chamber.
Results: Flexion and extension abilities of large mammal knees were achieved by placing the fresh tissue directly into
acetone without prior fixation with formalin, moving the knee daily after impregnation, and delaying the curing process
by several months.
Conclusions: Large mammal knees are ideal for human anatomy joint lessons because of their large menisci and thick
ligaments. The size of the elk knee is larger than human knees, and does not have a fibula. The health and integrity of
the ligaments are superior to most of the knees from aged human donors.
The Journal of Plastination 30(2):41 (2018)
SMALL-SCALE, INEXPENSIVEACETONE DISTILLATION AND PURIFICATION
SAVOINI EH, SMOLENYAK P
School of Engineering and Science, Yavapai College, Arizona, USA
Introduction: In the initial stages of a plastination laboratory set-up, purchase of a solvent recycler for acetone may be
as cost prohibitive as purchasing large quantities of acetone. A distillation apparatus assembled from typical laboratory
equipment and standard organic ground glass jointware provides a cost effective way to recover pure acetone.
Methods:
Vacuum Filtration
Dirty, used acetone from specimen baths was placed in a bucket in a freezer (<-15° C) overnight. The cold, dirty acetone
was vacuum filtered to remove solidified lipid and specimen debris.
Fractional Distillation
The vacuum-filtered acetone was placed in the distillation apparatus. The fractionating column was fashioned from a 30
cm Leibig condenser loosely packed with wire from a stainless steel kitchen scouring pad. The vertical condenser
connected via a vacuum adapter minimized the hood space required for the apparatus. Fitting the heating flask with a
Claisen adaptor provided easy access for the addition of acetone with minimal disassembly. The heating mantle
accommodated a 2000 mL round-bottom flask providing sufficient volume for processing on a convenient scale that
required minimal attention. Condenser cooling was provided by circulating ice water with an inexpensive aquarium
pump.
Purification
Distilled acetone was placed into 4 L flasks with 1 L of 3 Å molecular sieves, and left for 12-24 hours.
Results: The three-step process from water-fat-debris-containing acetone to >99% pure acetone took a total of 2 days
for 6-8 L. Fractional distillation produced approximately 1 L per hour of acetone (96-98% purity) and >90% recovery of
available acetone. After exposure to the molecular sieves, acetone achieved >99% purity.
Conclusions: This method produces 6-8 L of pure acetone (>99%) per day, to accommodate the smaller volumes, cost-
constraints, and space limitations of a small-scale plastination lab.
The Journal of Plastination 30(2):42 (2018)
SOFT PLASTINATION METHOD FOR PREPARATION OF FLEXIBLE TEACHING MODELS
SAWAD AA
College of Veterinary Medicine, University of Basrah, Basrah, Iraq
At the college of veterinary medicine, University of Basrah, the ambition was to produce low-cost educational
anatomical specimens, using local materials that are easy to use and inexpensive, that are useful for dissection, and
reduce the formalin exposure hazard. In order to confront these problems, a new soft plastination method was modified
to obtain flexible, clean, curable, odorless, portable and non-toxic specimens that can be kept for long durations without
deterioration. The advantages of this process are that it can be managed at room temperature by local inexpensive
materials, and it can be accomplished with little experience in co-operation with the silicone plastination method.
Students found the use of soft plastination specimens in the dissection room to be helpful in anatomical education, and
it is a good and enjoyable method that improved understanding of anatomy, biology, embryology, and their related
sciences.
The Journal of Plastination 30(2):43 (2018)
COMPARING EPOXIES BIODUR® E12 AND E12A FOR SHEET PLASTINATION
SCHILL VK
BIODUR® Products GmbH, Im Bosseldorn 17, 69126 Heidelberg, Germany
In this study, epoxy resins BIODUR® E12 and BIODUR® E12A were compared with regard to their use in sheet plastination.
While E12 is a well-established epoxy resin, E12A has been developed recently in order to obtain an epoxy especially
suited for the flat chamber method.
Slices of approx. 3 mm thickness were plastinated using BIODUR® E12 or E12A in combination with hardener E1. The
plastinated slices were cast either in a flat chamber or between sheets of polyester foil for the drain (sandwich) method.
Results illustrate that the mixture of E12A, plasticiser AE21, and hardener E1 shows a viscosity of less than 1000 mPa*s
during a time span of at least 30 hours at room temperature. E12A therefore, is perfectly suited for the flat chamber
method. On the other hand, E12 with its higher viscosity, is better suited for the sandwich method, because of its low
tendency to allow for air entering the interstice between the slice and the polyester foil. Visual inspection showed that
both E12 and E12A, yielded transparent plastinated slices of comparable, excellent quality.
The Journal of Plastination 30(2):44 (2018)
COLORING MUSCLE TISSUE FOR PLASTINATION
1SIQUEIRA BMM, 2 MONTEIRO YF, 2 JUVENATO LS, 3 BITTENCOURT APSV, 4 BAPTISTA CAC, 1,2BITTENCOURT AS
1Biochemistry and Pharmacology Graduation Program, Federal University of Espírito Santo, Vitória, Brazil, 2Department
of Morfology, Federal University of Espírito Santo, Vitória, Brazil, 3Department of Physiology Science, Federal University
of Espírito Santo, Vitória, Brazil, 4College of Medicine and Life Sciences, University of Toledo, Ohio, USA
Introduction: The visual appearance of the plastinated specimens is a very important factor to obtain an ideal product,
which conserves the original appearance of the specimen. The skeletal muscle tissue comprises approximately 40% of
the total body, being always very much in evidence in anatomical preparations. Therefore, the objective of this work was
to test the staining and selectivity of different histological dyes (acidic and basic) by muscle tissue for use in the
plastination technique.
Materials and Methods: Skeletal muscle tissue from carcasses of rats destined for disposal was used for staining. Each
test dye was placed in 10% buffered formaldehyde solution, totaling a final volume of 300 mL of dye solution. The
following dyes and their respective amounts were employed: ferrous fuchsin solution (7.5 mL), floxin B (0.0051 grams),
safranin (0.0060 grams), Masson's trichrome solution (7.5 mL), and control (no dyes) for staining during the fixation step
(30 days). After that, the tissues underwent the plastination process. Microscopy of stained tissues (prior to plastination)
was also performed to evaluate the selectivity and adherence of dyes in tissues.
Results: All dyes had affinity for muscle tissue and did not stain underlying tissues. However, the affinity of Masson's
trichrome was also observed in epidermis with microscopy analysis. It was possible to distinguish more easily the stained
muscle tissue from the other tissues. The dyes that presented the best results and selectivity were those of acidic
character (Masson's trichrome and floxin B), since they did not undergo the metachromasia phenomenon (color change
of basic dyes in biological tissues). Among these, the most promising was the Masson’s trichrome, since it more closely
approached the actual color of muscle tissue in vivo.
Conclusion: It was verified that the coloration of the skeletal muscle tissue of anatomical pieces helps in the
differentiation of tissues, such as the conjunctiva, adipose and epidermis. Masson's trichrome allowed coloration closer
to the real thing, showing it to be the most suitable for the technique.
Grant Support: CNPq (458328/2013-8; 440729/2017-3); FAPES (5537479411); UFES-PROEXT and CAPES.
The Journal of Plastination 30(2):45 (2018)
ROOM TEMPERATURE SILICONE PLASTINATION
STARCHIK D
International Morphological Centre, Saint-Petersburg, Russian Federation
Introduction: There are two silicone plastination techniques all over the world. The classical method (S10) was
introduced by Gunther von Hagens in 1977. It uses a reaction silicone impregnation mixture, and needs a freezer to keep
it cold for slowing down polymerization. The second technique was proposed by Daniel Corcoran & Dow Corning
Corporation in 1998, and it uses a non-reaction mixture silicone mix, and carries out impregnation at room temperature
(RT). Cold and RT silicone plastination techniques have differences in polymer components in the impregnation mixture,
and also in the sequence of their combination in the curing stage. Because the RT technique is less popular than the
cold-temperature method, it is advisable to know how to realize RT plastination stages, and what features those
plastinated specimens have.
Materials and Methods: We plastinated a variety of organs, regions, and whole body specimens with the RT technique
using standard procedures (dissection, dehydration, defatting, impregnation, & curing) and evaluated the advantages
and shortcomings of this method. Criteria used for evaluation included shrinkage, duration of impregnation and curing,
quality of plastinated specimens, need for extra equipment and its maintenance, as well as other cost considerations.
Cylindrical core samples of parenchymatous organs were used to efficiently evaluate shrinkage and plastination
duration. Core cylinder volume was evaluated at the end of each stage of the process by fluid displacement.
Results: The first three steps for the RT procedure are identical to the cold technique; the difference is in the
impregnation and curing steps only. The RT impregnation mixture consists of 93% silicone and 5% cross-linker. That is
not a reaction mixture, and so there is no need to keep it in a freezer. Molecular weight is about 500, and 12-second
dynamic viscosity. The average shrinkage calculated for the tissue cores plastinated by the RT technique (16.2 ± 1.49 %)
was 1.5 times less than by the cold method (p < 0.05). The total duration of the impregnation and curing stages of
samples for the RT plastination was found to be 1.54 times shorter than that of the S10 technique. The silicone
impregnation-mix for the RT technique, because of its low viscosity, drains very easily from impregnated hair, fur, and
feather specimens, which is a large time saver. Hollow organs and body part specimens plastinated by the RT procedure
were less flexible, more fragile and harder after curing than those made with the cold technique. There is no need for an
additional freezer for the impregnation vacuum chamber, or a special chamber equipped with a fan, an aquarium pump
and desiccant for the RT technique.
Conclusion: The specimens plastinated with the RT technique were less flexible and elastic, but this process allows
production of good quality specimens with minimal shrinkage, and in a shorter period of time. This method is preferable
for whole brain, parenchymatous organs, parts of the body, fetus, fur/hair/feather-covered specimens, reptiles & fish, as
well as for long time formalin-fixed specimens, for archaeological and fossil objects. A room temperature plastination
laboratory is more economical to set up than the cold-temperature system.
The Journal of Plastination 30(2):46 (2018)
E12 PLASTINATION FOR RESEARCH INTO STENTED CORONARY ARTERIES
1STARCHIK D, 2SORA MC, 3SHISHKEVICH A, 1ANDREEV Y
1Department of Human Morphology, North-western State Medical University, Saint Petersburg, Russia, 2Department of Anatomy and Molecular Medicine, Sigmund Freud University, Vienna, Austria, 3The First
Department and Clinic of Surgery, Military Medical Academy, Saint Petersburg, Russia
Introduction: Stenting of coronary arteries is the most promising method of myocardial revascularization for ischemic
heart disease. At the same time, detailed visualization of the metal stent inside the coronary artery, and study of the
relationship of its elements to the arterial wall in histological specimens, is not possible. Use of a special technique of
epoxy plastination for stented coronary arteries contributes to new treatment methods for one of the most common
cardiovascular pathologies.
Materials and Methods: A portion of the heart wall with an implanted stent was dehydrated in cold acetone at -25° C.
After completely replacing the water, the preparation was immersed in mixture of low-viscosity E12 epoxy resin and
hardener in a ratio of 20: 1, and impregnated in a vacuum chamber at 30° C, decreasing pressure gradually until release
of acetone bubbles ceased. The heart wall was then taken out of the resin, and left for 7 to 10 days at a temperature of
40° C until it was completely cured. Tissue was removed around the stented artery by grinding, or the specimen was cut
with a diamond saw in 0.5 – 1 mm sections. The sections were placed in polymethylmethacrylate flat chambers, and
filled up with a composition of E12 epoxy resin and hardener in the ratio of 10: 1. After hardening of the resin, the
transparent specimens of stented arteries were scanned at 600 or 1200 dpi, or were microscopically examined up to 20X
magnification.
Results: Plastinated E12 slices provided good anatomical details down to the microscopic level. The proposed method is
fine for detecting changes in the geometry and morphology of stented lesions of coronary arteries. It permits good
visualisation of the contact of the stent with the inner vessel wall and atherosclerotic plaques, and it easily provides
measurements and statistical analysis. In comparison with radiography, this method makes possible evaluation of the
topography of the metal implants in arterial bifurcations, and it demonstrates the degree of deformation of the
atherosclerotic plaque after stenting. Hard transparent sections with whole stents retain transparency and can be cut
transversely for more detailed examination.
Conclusion: Adequate impregnation of the heart wall is achieved by using low-viscosity epoxy resin, and heating the
impregnating composition, which increases the fluidity of the resin, and enabled impregnation of specimens with
thickness up to 25 mm. This method can be applied for clinical research for development of new methods of stenting of
coronary arteries, and for the study of other organs with implanted metal constructions and devices.
The Journal of Plastination 30(2):47 (2018)
Minutes
19th Biennial Business Meeting of The International Society for Plastination
Held in Dalian, China, July 21, 2018
1. The President, Professor Rafael Latorre opened the meeting. The membership was informed of the death of Lance
Nash, an active member of the ISP for many years. The President paid tribute to Dr. Nash, and a minute of silence was
observed.
2. The minutes of the previous AGM of the ISP (June 30th, 2016, Toledo, USA) were presented and approved.
3. President’s report
The President Rafael Latorre reported that grant applications to attend this meeting had been received from students 1
from Africa, and 2 from Murcia, however the applicants had no budget to purchase tickets. In view of this, the President
requested approval for financial help for transport to meetings for students.
Dr. Carlos Baptista reminded the meeting that this was the source of membership for the future, but it raised the issue
of where to get funds, as conferences operate on a very tight budget. Should sponsorship from companies be explored?
The President: companies present (Biodur and Hoffen) were asked and they agreed to support one transportation
student grant for the next ICP.
The President: all attendees at the workshops in Murcia automatically become members for 2 years. There is a budget
from Murcia to sponsor 1 student to the ICP. This was to be decided by committee, the applicant must be a member of
the ISP, they must present at the meeting, and they must submit a paper to the Journal.
Prof Sui (H-JS): it should be limited to students working in plastination. H-JS proposed that the host of the ICP could
sponsor 1 student for registration and accommodation. This proposal for the 2020 meeting was approved.
The President stressed that it was important to update email addresses for the website/Groupspace. It was agreed to
share email queries with all members.
4. Treasurer’s report
Balance Transferred from Joshua Lopez (2016) $23,309.30 Income (ISP Membership is $75. Total received after fees is $72.52) Membership Dues (July-December 2016) $725.20 (10 Membership) Membership Dues (2017) $432.88 (6 Membership) Membership Dues (January-July 1, 2018) $507.64 (23 Membership) Total Membership Dues $1,813.00
The Journal of Plastination 30(2):48 (2018)
Expenses Printing & Production Journal ($2,761.85) Scholarship paid to students attending the Interim S Africa Paypal Charges (2016:; 2017:$17.12;2018) Bank Charges ($143.53) Bank Charges relates to Chase Bank only Total Expenses ($2,905.38) Current Chase Account Balance $21,294.75 PayPal Account: Payments received: $2,400.00 Fees $97.66 Current PayPal Account Balance $2,302.34 Chase Account: Fees Current Chase Account Balance Total Accounts $22,838.75
The Treasurer Carlos Baptista reported that there was a problem that the ISP was failing to retain new members from
the workshops after 2 years.
The new website hosting and domain name had been paid for in advance for 3 years.
The hard copy of the Journal cost 600 USD/issue for printing 40 copies. This was a considerable outlay for the ISP. There
was discussion on whether the Journal should continue to be printed or issued as an electronic version only. The
possibility of a two-tier membership (with or without the hard copy) was discussed. It was also suggested that an option
to order a hard copy through a print-on-demand service should be included on the website.
A proposal to discontinue the hard copy of the Journal of Plastination was carried.
Dr. Octavio Lopez (OL) offered to help with marketing/support, for example communication with members, particularly
targeting non-active members.
The possibility of automatic renewals was discussed. Renewals would automatically continue until cancelled.
Carlos Baptista: Groupspace has a link with PayPal, could generate an automatic reminder. Carlos Baptista agreed to
look into automatic payments via PayPal
5. Editor’s report
The Editor, Philip Adds (PA), reported that applications were in the process of being submitted to Scopus and Web of
Science. Following feedback from the NLM on our application to Medline, the Journal now carries a statement of ethics.
PA intends to prepare a new submission to Medline in the near future.
The Journal of Plastination 30(2):49 (2018)
There were 6 papers currently undergoing review/revision, and 2 more had been submitted that were waiting to be sent
out for peer review. Because English is not the first language of many of our contributors it is frequently necessary for
manuscripts to undergo quite heavy English-language editing before they can be sent out for peer review. This takes up
quite a lot of the Editor’s time, and therefore, unfortunately, delays the peer review process. Because of the low number
of submissions, PA is reluctant to reject papers outright, and some need a lot of work to get them up to an acceptable
standard for publication.
PA reminded the meeting that it had been the intention of the Editorial Board to republish the ‘Cook Book’ of standard
plastination procedures (last published in 2007). This was agreed several years ago, but despite repeated requests to the
authors of the original articles for updated versions, only 1 has so far been submitted for publication. PA wondered if
perhaps it was time to revisit this project, and aim to publish the new, updated cookbook in 2019.
RL proposed that the Journal should carry an Obituary of Lance Nash. Approved unanimously. RL to ask Dr. Ming Zhang
to write it.
The use of a professional English-language editing service was discussed. It was generally felt that it would be too
expensive, and unlikely to have the necessary expertise in the techniques of plastination.
Matthew Tinney kindly offered to help with copy editing.
7. New business
i. There had been no volunteers to host the 13th Interim Meeting
Carlos Baptista suggested emailing the membership, if it was felt that an interim meeting was still necessary.
Dr. Dmitry Starchik suggested that the unsuccessful candidate to host the ICP might wish to host the Interim Meeting.
ii. 20th ICP (2020)
Two bids were received: Temuco (Chile) in July, and Honolulu (Hawaii) in March, 2020. The two bids were put to a vote.
The majority favored Temuco.
Carlos Baptista offered to contact Scott Lozanoff with a view to either hosting the 13th Interim Meeting, or submitting a
new application to host the 2022 ICP.
iii. 21st ICP (2022)
There was discussion on two possible venues for the 21st ICP: Hawaii and Denmark. Further discussion with the
potential hosts was needed.
The Journal of Plastination 30(2):50 (2018)
8. Report of the Elections.
Carlos Baptista gave a verbal report on the outcome of the recent ISP elections.
President: Rafael Latorre; Vice-president: Dmitry Starchik; Secretary: Nicolas Ottone; Treasurer: Carlos Baptista; and
Councillors Kees De Jong, Hong-Jin Sui, Onyemaech Okpara Azu, and Telma Masuko.
RL: thank Anthony Weinhaus for his support with this election process. Special thanks to Selcuk Tunali for his tenure as
Secretary since 2016. Thanks to Bob Henry, Ming Zhang and Athelson Bittencourt for serving as councilors since 2016.
He greatly appreciates the time they have dedicated to the ISP.
The meeting closed at 15.30
The Journal of Plastination 30(2):51 (2018)
The Plastination Journal 30 (2):52 (2018)
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: [email protected]. If you experience any problems or need further information, please contact Philip J. Adds, [email protected].
Authors must have an e-mail address at which they may be reached.
Necessary Files for Submission Include:
Cover letter
Manuscript (including references and 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 ([email protected]) 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. HOLLADAY1*, B. L. BLAYLOCK2 and B. J. SMITH1 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: [email protected]
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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 Plastination Journal 30 (2):54 (2018)
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
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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