The Kagoshima University Museum / The Research Institute for Humanity and Nature

Fish Collection Building and Procedures Manual
English Edition

Hiroyuki Motomura and Satoshi Ishikawa(translated by Akira Takagi and Yuka Ogata)

The Kagoshima University Museum / The Research Institute for Humanity and Nature
1

Fish Collection Building and Procedures ManualFish Collection Building and Procedures Manual. English Edition
edited by Hiroyuki Motomura and Satoshi Ishikawa
translated by Akira Takagi and Yuka Ogata
The Kagoshima University Museum / The Research Institute for Humanity and Nature
27 February 2013This book is published as parts of the following two projects:
Laboratory of Fish Systematics
The Kagoshima University Museum, Japan“Kagoshima Fish Diversity” Project
Project leader: Hiroyuki Motomura
Inter-University Research Institute Corporation,
National Institutes for the Humanities, JapanThe Research Institute for Humanity and Nature“Coastal Area Capability Enhancement in Southeast Asia” Project
Project leader: Satoshi Ishikawa

Copy Right © 2013 by the Kagoshima University Museum, JapanAll rights reserved. No part of this publication may be reproduced or transmitted in any form or by anymeans without prior written permission from the publisher. Copyrights of the photographs are held by thephotographers.
For bibliographic purposes this book should be cited as follows:
Hiroyuki Motomura and Satoshi Ishikawa (eds.). 2013. Fish collection building and procedures manual.
English edition. The Kagoshima University Museum, Kagoshima and the Research Institute for Humanity
and Nature, Kyoto. 70 pp.
ISBN 978-4-905464-01-3
Desktop Publishing by Hiroyuki Motomura (The Kagoshima University Museum)2

The Kagoshima University Museum / The Research Institute for Humanity and NaturePreface

The recent destruction and deterioration of the environment has led to aworldwide decline in biodiversity. To
conserve the current biodiversity, there
is an urgent need to identify and catalog
species. Therefore, natural history museums and biological research institutes,
including universities, must keep primarysources, including voucher specimens
and photographs, of organisms to record
the current biodiversity and pass them on
to posterity as humanity's common properties.
This book describes curatorial procedures for fish specimens and fish collections as practised by the Kagoshima
University Museum. The curatorial andcataloging methods and procedures described here, while perhaps not ideal
for large museums such as the Natural
History Museum, London, or National
Museum of Nature and Science, Tsukuba,
are suitable for local small-sized museums or universities with lower budgets.
This book also provides details of various
equipment for photography, preservationand storage, including specifications andnames of manufacturers. It also mentions
methods for the creation of specimen
databases and procedures for specimenloans.

The book was originally publishedin 2009 as a manual written in Japanese[Motomura, H. (ed.) 2009. Fish Collection Building and Procedures Manual.
The Kagoshima University Museum,Kagoshima]. The English edition of themanual is herein published as a part ofthe editors’ research projects, Kagoshima
Fish Diversity and Coastal Area Capability Enhancement in Southeast Asia projects, and supported by the Kagoshima
University Museum and the ResearchInstitute for Humanity and Nature.
I am grateful to M. Aizawa, H. Endo,
Y. Iwatsuki, S. Kimura, K. Matsuura,and H. Senou for sharing their extensive knowledge of curatorial proceduresfor fish specimens, M. McGrouther, D.
Catania, J. Maclaine, A. Bentley, K. Murphy, D. Nelson, M. Sabaj, K. Swagel, E.
Holm, G. Dally, Y.-C. Liao, L. Kelvin,
and G. Yearsley for providing informationon their fish collections, photographs ofcollection labels and jars, and commentson the book, and volunteers and studentsof the Kagoshima University Museum fortheir assistance.
Hiroyuki Motomura
The Kagoshima University Museum

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Fish Collection Building and Procedures Manual

Preface

Underway since 2012, “The coastal
area capability enhancement in Southeast
Asia” project, directed by the ResearchInstitute for Humanity and Nature (RIHN)
system, will continue through 2017. Its
purpose is to realize “area capability” andto generate a new approach toward ruraldevelopment evaluation based on theharmonization between ecosystem healthconservation and improvement of local
people’s quality of life. Because rural
people’s lives are invested in the capital
involving goods and services that ecosystems provide, we believe that strengthening the link between natural capital andlocal people is key for the sustainable development of rural areas. Thus far, however, the benefits of natural capital andservices have not been fully recognized.
As part of the RIHN project, we conduct
a detailed field survey of environmental,
biological, social, and economic aspects
through a holistic joint approach to grasp
the actual situation regarding ecosystem
health, the livelihoods of local people,
and the connection between the two.
The original Fish Collection Building and Procedures Manual was published by Kagoshima University Museum
and edited by Prof. Motomura. We arepleased that the publication of this English edition marks one of RIHN’s most4

memorable achievements. We are certainthat it will contribute to the understanding of biodiversity and the variety ofnatural capital through the establishmentof a sophisticated fish collection at museums and research institutes. Though
Southeast Asian coastal fauna and flora
hold high biodiversity, taxonomic studies
and food web analyses, including population studies, have not been thoroughly
investigated. We hope that this manual
will promote biodiversity studies in theSoutheast Asian coastal area and provide
people with a unique opportunity to improve their research skills.
The RIHN project is based on the joint
research efforts of Southeast Asian Fisheries Development Center (SEAFDEC),
Faculty of Fisheries of Kasetsart University, the University of the Philippines
Visayas (UPV), and Japanese researchers
who are members of the RIHN project.
Aklan State University and Eastern Marine Fisheries Research and Development
Center of Department Fishery, Thailand,
are active participants as well. Throughthis collaboration, we share the same vision for future biodiversity studies.
Satoshi Ishikawa
The Research Institute for Humanity and Nature

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Editors
Hiroyuki Motomura, PhD
Professor, the Kagoshima University Museum
1-21-30 Korimoto, Kagoshima 890-0065, Japan
Ph: +81 99 285 8111; fax: +81 99 285 7267
E-mail: motomura@kaum.kagoshima-u.ac.jp
URL: http://www.museum.kagoshima-u.ac.jp/staff/motomura/motomura.html
Satoshi Ishikawa, PhD
Associate Professor, Research Institute for Humanity and Nature
457-4 Motoyama, Kamigamo, Kita-ku, Kyoto 603-8047, Japan
Ph: +81 75 707 2320; fax: +81 75 707 2507
E-mail: oounagi@chikyu.ac.jp
URL: http://www.chikyu.ac.jp/index_e.html

Translators
Akira Takagi, PhD
Senior Project Researcher, Research Institute for Humanity and Nature
457-4 Motoyama, Kamigamo, Kita-ku, Kyoto 603-8047, Japan
Ph: +81 75 707 2317; fax: +81 75 707 2507
E-mail: akirapt@chikyu.ac.jp
Yuka Ogata, PhD
Project Researcher, Graduate School of Frontier Science, the University of Tokyo
5-1-5 Kashiwanoha, Kashiwa, Chiba 227-8564, Japan
Ph: +81 3 5841 5018; fax: +81 3 5841 5018
E-mail: aogata@mail.ecc.u-tokyo.ac.jp

5Fish Collection Building and Procedures Manual

Authors
Kaoru Kuriiwa
Research Assistant, Department of Zoology, the National Museum of Nature and Science
4-1-1 Amakubo, Tsukuba, Ibaraki 305-0005, Japan
Mizuki Matsunuma
PhD student, the United Graduate School of Agricultural Sciences, Kagoshima University
1-21-24 Korimoto, Kagoshima 890-0065, Japan
Masatoshi Meguro
PhD student, the United Graduate School of Agricultural Sciences, Kagoshima University
1-21-24 Korimoto, Kagoshima 890-0065, Japan
Hiroyuki Motomura
See Editors
Gota Ogihara
PhD student, the United Graduate School of Agricultural Sciences, Kagoshima University
1-21-24 Korimoto, Kagoshima 890-0065, Japan
Tomohiro Yoshida
PhD student, the United Graduate School of Agricultural Sciences, Kagoshima University
1-21-24 Korimoto, Kagoshima 890-0065, Japan

6

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Contents

Step 1

Transport ............................................................................................... H. Motomura

9

Step 2Freezing ........................................................................... M. Meguro and H. Motomura 11

Step 3

Defrosting ................................................................................. M. Meguro and T. Yoshida 13

Step 4

Rinsing ........................................................................................ H. Motomura and M. Meguro 14

Step 5Assigning tags to specimens ................................................................................... H. Motomura 15

Step 6

Obtaining tissue samples for DNA analysis ................................. H. Motomura and K. Kuriiwa 17

Step 7Pre-fixation .............................................................................. M. Matsunuma and H. Motomura 19

Step 8

Photography ............................................................................................................. H. Motomura 29

Information 1

Details about specimen photography .......................................................................... K. Kuriiwa 35

Information 2

Specimen photography in field ................................................................................. H. Motomura 46

Information 3

Side-view photography ............................................................................................. H. Motomura 47

Step 9

Tagging .................................................................................... M. Matsunuma and H. Motomura 49

Step 10

Measurement ................................................................................................................ G. Ogihara 53

Step 11

Identification ................................................................................... G. Ogihara and H. Motomura 54

Step 12Fixation .................................................................................................................... H. Motomura 55

Step 13

Replacement of formalin with alcohol ..................................................................... H. Motomura 57

Step 14Storage ..................................................................................................................... H. Motomura 58

Information 4

Specimen data labels ................................................................................................ H. Motomura 60

Step 15

Creation of database ................................................................................................. H. Motomura 63

Step 16

Image processing ..................................................................................................... H. Motomura 64

Step 17

Loan ......................................................................................................................... H. Motomura 67

7

Fish Collection Building and Procedures ManualCollecting fishes by team of the Kagoshima University Museum

8The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 1

Transport
Hiroyuki Motomura

After fish samples are obtained, theyshould be transported in an appropriate
way to the laboratory at the university
or institute for preparing specimens forreference collection. Therefore, a suitabletransportation method should be selected,
depending on the conditions. In this section, methods of sample transportation
are discussed in detail.
Live fish samples are the best for
preparing beautiful specimens. To keep
fishes alive, they should be carried insmall, water-filled plastic bags, taking
care that they are not hurt in any way. For
example, if gobies and scorpionfishes arecarried in the same plastic bag, the latter may prey on the former; even if thegobies survive predation by the scorpionfishes, their body surface and fin mem-branes will be frayed and torn.
Although live-fish transportation is
the most appropriate way to prepare good
specimens, this method exposes the fishes
to the risk of being damaged. If live fishsamples do not have access to abundantair, the fishes will suffocate and die.
When suffocated, most fishes spread open
their mouths and gill covers wide. In thiscondition, accurate measurements likethose of standard length of the fishes cannot be obtained, since the open mouths
and gill covers rigidify because of rigor
mortis and it is difficult to restore them tothe normal positions. Data for calculating
accurate standard length are the most important in academic researches. Furthermore, if fishes die because of lack of oxygen in room-temperature water (including
seawater), their body coloration whenfresh cannot be recorded, since the bodycolor fades rapidly and significantly.
Although live-fish transportation isvery valuable for preparing beautiful
specimens, it should be noted that transportation of some fishes alive, especiallyfreshwater fishes, is legally prohibited.
These laws differ among countries, andall laws in a country should be respected
while collecting and transporting fishsamples.
After successful transportation of live
fish samples to the laboratory, the fishes

Some fishes carried in a plastic bag.

9

Fish Collection Building and Procedures Manual

Largemouth bass (Micropterus salmoides), listed under theInvasive Alien Species Act of Japan.

Fresh fishes carried in ice-cold water.

Bluegill (Lepomis macrochirus), listed under the Invasive
Alien Species Act of Japan.

should be sacrificed by immersion in icecold water or anesthetized.
If sample treatment cannot be initiatedimmediately after collection (live fishes
from sampling sites or dead fishes fromplaces like fish markets), the fishes shouldbe transported in a cold box filled with
ice-cold water (seawater for marine fishes
and freshwater for freshwater fishes). It
should be noted that if fishes are put on
ice directly, the color of the body part
touching the ice changes. Moreover, icecold water is crucial for preserving the
freshness of the samples before treatment.
Thus, it is better to temporarily keep fishsamples in ice-cold water for at least several minutes before initiating treatment.
Freezing is a better method of preserving fish samples than keeping in ice-coldwater, if the samples cannot be treated
for a long time and/or they need to be

10

Fish samples temporarily kept in ice-cold water to preserve
their freshness.

transported over long distances. However, fishes begin decaying, especiallythe internal organs, on long-term chilledstorage. Fish samples should be stored ina freezer if they cannot be treated within
1–2 days after collection. Methods offreezing fish samples are discussed in
STEP 2.

1. Freezing of samples (for preservation andstorage)→ Step 2

2. Treatment of samples (for specimen preparation)→ Step 4

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 2Freezing
Masatoshi Meguro and Hiroyuki Motomura

It is very important to obtain fresh fishsamples for preparing beautiful specimens with a long shelf life, but specimens may not be prepared from fresh
fish samples at any time. Freezing is the
only way to keep fish samples fresh for along time before treating them to prepare
specimens. However, freezing is not suitable for all fishes such as those of Gobiidae and Blenniidae, which have very
weak fin membranes, and of Clupeidae,
whose scales readily exfoliate on freezing. It should also be carefully considered
whether or not to choose freezing for interim storage.
We use “National NR-FC28FG” deep
freezers built by Panasonic Corporation
for freeze preservation in our museum,the Kagoshima University Museum.
Fish samples are stored at –20°C in NRFC28FG deep freezers; ideally, samples
should be stored at –80°C.
Preservation temperature is an important factor to be considered to prevent

clouding of the eye lens. Regular hometype refrigerators can also preserve fishsamples, although their humidity level ishigher than that of deep freezers. However, regular home-type refrigerators can-Deep freezer for preservation of fish samples.

Freezer-burned fish specimen. It is difficult to spread each fin, since the specimen gets dehydrated and becomes hard.

11Fish Collection Building and Procedures Manual

Specimen freeze preserved in seawater. Tip of the caudal fin
is bent, and hence, it is not a well-preserved specimen.

Specimen freeze preserved in seawater. It is a well-preserved
specimen. This specimen had been preserved since 2007. It
could well be treated in 2009.

not prevent clouding of the eye lens, andtherefore, they are not suitable for preserving fishes, especially those of Labridae.
If fish samples are left in a freezer fora long time, the fish bodies become rigid(not frozen) and are not restored to thenormal state after defrosting. This condition is called “freezer burn.” Freezer burnoccurs when the tissues are damaged bydehydration and oxidation, because of air
reaching them. Freezer burn causes irreversible denaturation of samples. Freezerburned samples cannot become normal
again even if immersed in sufficientamount of water. The fins cannot spread
well, which makes it difficult to record
fin color and pedicel length correctly.
It is very difficult to prevent freezer
burn completely; however, the use of
appropriate water type can reduce therisk of symptom development. Seawater should be used for marine fishes andfreshwater for freshwater fishes duringfreeze preservation of fish samples.
A note indicating the sampling dateshould be included with the sample fish
before freezing. It is very important that
all the data are recorded before the information slips from the mind. All available
information about the samples should
be written down in detail, including
the names of the people who collected

the samples, the places of origin of thesamples, the collected data, the sampling
methods, and the depth at which the samples were collected.
The data should be written with a regular or a mechanical pencil on waterproof
paper. Ballpoint pen and ordinary paper
are not suitable for recording the data to
be included with the freezing samples.
This is because the information will belost on thawing the samples, since the paper will tear and the ink will get washedaway. Moreover, important fish samples
will be rendered worthless without therelevant background information. Thus,
data recording is very important.
Nowadays, all specimens used forexperiments, even those for molecularbiological research, registered in research
institutes, including museums and universities. Sometimes, fish samples are
stored in a freezer for later DNA analysis.
However, prolonged freezing of samples
not only deters preparation of beautiful
specimens but also causes freezer burn,which inhibits species identification. It ispreferable to first obtain some tissues for
DNA analysis and then immediately proceed to sample preservation.

12

Defrosting of samples→ Step 3

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 3

Defrosting
Masatoshi Meguro and Tomohiro Yoshida

Defrosting is the first step in preparing
specimens from fish samples frozen fora long time. It does not involve simply
thawing at any time but should be initiated at the right time.
Defrosting should be initiated considering the room temperature and the massof the ice supplied with the samples. If
thawing is started too early, the fishes
will begin decaying, especially the internal organs. On the other hand, if thawing
is delayed, sufficient ice will not melt. In

this situation, however, the remaining unmelted ice should not be pulled from thefish bodies, since it will break the scales
and surface skin.
In addition, scrupulous attention isrequired while thawing a few frozen samples together, to avoid mixing up of datanotes.
Rinsing of samples→ Step 4

Defrosting under running water.

13

Fish Collection Building and Procedures Manual

Step 4

Rinsing
Hiroyuki Motomura and Masatoshi Meguro

Defrosted fish samples should be
rinsed to remove grime and mucous
membranes. Rinsing requires scrupulous
attention, since scales and fin membranes
may easily break after long-term storage
in the freezer. Usually, the body surface
is gently rubbed with fingertips to remove
extraneous material. If necessary, a softbristled paintbrush can be used. Some
fishes have a mucous membrane on theirbody surface. This membrane becomescloudy if formalin is applied directly(without rinsing well) for fin spreading.
Furthermore, if the fishes are not rinsed
enough, their body coloration cannot berecorded well. Therefore, mucous membranes should be completely removed.
However, it is slightly difficult to ensure
this, since the mucous membrane is colorless and transparent.

Assigning tags

→ Step 5

An imperfect catfish specimen, with mucous membrane not removed completely (KAUM–I. 3508, 451.8 mm standard length).
The membrane is removed only around the nape, and the original body color can be observed.

A good catfish specimen, with mucous membrane removed well (KAUM–I. 4806, 72.9 mm standard length).

14

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 5 Assigning tags to specimens
Hiroyuki Motomura
After rinsing, each cleaned fish is assigned an individual number for individual identification of the specimen. Whena few samples are treated together, eachsample needs to be individually identified; they should not be thrown into disarray, so that there is no confusion arisingin subsequent processes.
All specimens should be assigned
individual numbers, especially for obtaining tissue samples for DNA analysis
and for taking photographs. However,the number tag should not be attached to
the fish body before taking photographs.
This is because the tag will need to be
removed while photographing and reattached to the fish body after the specimen
is photographed.
As an individual number is assigned
to a specimen, a specimen list should be
simultaneously prepared using the sampling-date note as reference.
A list prepared using a regular or amechanical pencil is maintained almost
permanently. The specimen list should
include the scientific name of the species.

However, quick species identification ofsome fishes is quite difficult. In this case,the scientific name need not be written at
this stage. If a long time is taken for identification, it will not be possible to record
body coloration, since the body color
fades with every moment. Identification
can be performed taking enough time after the specimen is photographed.

Specimen tags used in our museum. Each numbered tag is
cut and used for an individual specimen.

Left: numbering machine (D51, Lion Office Products Co.
Ltd.). Right: inkpad.

■Preparation of a number tag
The number tag is made of cloth. Calico is the best cloth for the tag, since it
absorbs inks well, is not a stretchy fabric,
and does not tear easily. Serial numbers
are printed on white calico by using a
numbering machine with pigmented ink.
The brevity code of the inventory location of the specimens is also printed in
front of each individual number. A brevity

15Fish Collection Building and Procedures Manual

Specimen tag. Serial numbers and museum abbreviation
code are printed on calico (before applying collodion).
Below: Collodion bottle.

code is an establishedcode for the nameof an international
research institution.
For example, the abbreviation code of the
Kagoshima University
Museum-Ichthyology
is KAUM–I. The
printed informationon the calico tag iscompletely dried byair seasoning for atleast 1 day, followed by a few more days.
Thereafter, the calico tag is coated witha formulated concentrated collodion solution to make it waterproof. The wet
calico tag is then hung outside to dry, likewashed clothes, secured by a clothespin.
If collodion is applied before completely drying the ink, the print will be
smudged and become unreadable. A tag
with an unreadable or missing number will obviously have to be remade.
When a number tag is remade, the same
numbering machine is not used. This isbecause the numbering machine will automatically print the next serial number.
Thus, the use of the same numberingmachine for reprinting a number tag iscertainly wrong, since it will cause either
overlapping or omission of numbers. A
16

Tapewriter (DM1585-B, Orient Enterprise Co. Ltd.) andnumber-tag tape (9 mm width).

Its use is convenient. Cut the calico tag 30–50 numbers androlled and preserved.

tapewriter is a useful tool for remaking aparticular number tag.
After the calico tag is coated withcollodion, it becomes hard and waterproof
like a plastic board. It can be easily cut
using a pair of scissors and the ink too
does not get washed in water. However,collodion application does not confirm
alcohol resistance.

1. Obtaining tissue samples for DNA analysis→ Step 6

2. Pre-fixation (no tissue samples taken for
DNA analysis)→ Step 7

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 6 Obtaining tissue samples for DNA analysis
Hiroyuki Motomura and Kaoru Kuriiwa
In current research, DNA analysis isperformed intensively by many scientists.
Many museums collect and store tissue
samples for DNA analysis, besides whole
specimens.
In this section, methods of collection
and storage of tissue samples in our museum are discussed.
Tissue samples should be collected
after assigning the number tags (STEP 5)
and before applying formalin (STEP 7) tothe specimens, so that each tissue sample
is assigned the same individual number as
the specimen. Moreover, formalin consti-

tutes a limiting factor for DNA analysis.
In taxonomic study of fishes, scientists
examine the left side of the fish body.
Therefore, muscle tissue samples should
be collected from the right side of thebody, so that the important characters fortaxonomy such as the lateral line and pectoral fins are preserved. Enough genomic
DNA can be extracted from at least a
5-mm2 muscle tissue section. However,
if possible, it is better to obtain a 1-cm2
muscle tissue section, as reserve. In addition, the muscle tissues samples should
not contain other material such as scurf,
fat, and blood. This is because the scurf
will produce a smear on the tissue sam-Fish (right side of body) with tissues excised for samples.
White part indicates the point of excision.
Tissue sample and data label (specimen number and species
name) in a screw-cap sampling glass bottle.

Screw-cap sampling glass bottles used for tissue samples.

Data (specimen number, species name, and sampling site)
written on a screw-cap.

17

Fish Collection Building and Procedures Manualples when they are immersed in ethanol,and the fat and blood cells will hinder
DNA purification.
It is difficult to collect muscle tissues
from some small fish species. In this case,the right pelvic (abdominal) fin is collected as an alternative for muscle tissue.
Fishes have a pair each of pectoral finsand pelvic fins. Pelvic fins have relatively
low mutations as compared with pectoral
fins and are not used as an alpha-leveltaxonomy. Thus, they can be excised.
The obtained muscle or fin tissue
samples are put into screw-cap sampling
glass bottles filled with 99.5% ethanol.
A 20-cc screw-cap sampling glass bottle is well suited for a 1-cm2 muscle tissue
section. A large tissue sample should not
be put into a small bottle to prevent penetration of ethanol into the whole tissue. A
small note indicating the individual number and species name should be included
with each tissue sample. The note should
be written with a regular or a mechanicalpencil on waterproof paper.
For better storage of the tissue samples, ethanol should be changed regularly,
as it becomes hazy because of water oozing out from the tissues.
In our museum, the screw-cap sampling glass bottles containing the tissue samples are stored in boxes. This
arrangement makes it easy to locate aparticular sample in order to write thesame individual number as the specimen
and the species name on the cap. For thispurpose, an ethanol-proof pen should be
used, since the ink of a normal felt pen
will get washed away by ethanol, erasing
all information. The tissue sample boxes
are stored in the deep freezer for subsequent analysis.
Previously, the liver was used to
extract enough DNA from fresh fishsamples, since it contains the largest
amount of DNA. However, it contains alarge amount of fat, as well as sugar andprotein. Therefore, nowadays, a liver tissue sample is not suitable for conducting
DNA analysis.

Tissue samples in screw-cap sampling glass bottles ina box. The box is placed in deep freezer after closing
the lid.

Pre-fixation

18→ Step 7

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 7Pre-fixation
Mizuki Matsunuma and Hiroyuki Motomura

Fish specimens are usually made forscientific research. Measurement andcounting of traits are quite convenientwhen the fish body is mounted straight
and all the fins are spread well. It is difficult to obtain the real, discrete value ofan imperfectly fixed specimen.
Some fishes have significantly longbases of the dorsal and/or anal fins, suchas those of Muraenesocidae (pike congers) and Ophichthidae (snake eels).
Thus, it is really difficult to count the
number of fin rays if the fins are fixed ina folded condition. Perfectly fixed speci-

An imperfect specimen of Scorpaenodes evides. The fish
is fixed with the mouth and gill cover open, and therefore,
body length cannot be measured correctly.

An ideal specimen of Scorpaenodes evides (KAUM–I. 4371,
48.2 mm standard length).

mens obviously have a higher academic
value than imperfectly fixed ones.
■Equipment
1. Pins
Pins are the most important tools for
spreading fish fins. Normally, insect pins
are used for this purpose. Shiga Insect
Pins are the best choice for spreading
fins. They are particularly useful as they
are available in many sizes. We usually
use No. 00 to No. 6 pins and micropins.
The thickness of the pin increases with
its number; for example, the thickness increases from 0.3 mm to 0.65 mm for No.
00 to No. 6 pins. The most appropriate
pin should be selected according to thefish size and density of its fin membrane.
A micropin is thinner than a No. 00 pin.
It is used for very small fishes with very
weak fin membranes, such as those of
Gobiidae and Tripterygiidae. Since micropins are very thin and small, it is preferable that they are handled with forceps.
Sometimes, bamboo skewers are used
to spread the fins of large fishes such as
scombrids. In our museum, we use a longand thick pin to spread the fins of a largescaled fish. We use an E979 needle supplied by Australian Entomological Supplies, which has a 70-mm stainless-steelsolid head and a thickness of 1.37 mm.
The sizes of a micropin and different
Shiga Insect Pins are provided below.
19Fish Collection Building and Procedures Manual

Shiga Insect Pin No. 5 and outer packaging.

Shiga Insect Pin No. 00 to No. 6 and long needle made byAustralian Entomological Supplies.

•Micropin (stainless steel)
0.18-mm thick, 17.5-mm long
•Shiga Insect Pin (stainless steel)
No. 00: 0.30-mm thick, about 40-mm long
No. 0: 0.35-mm thick, about 40-mm long
No. 1: 0.40-mm thick, about 40-mm long
No. 2: 0.45-mm thick, about 40-mm long
No. 3: 0.50-mm thick, about 40-mm long
No. 4: 0.55-mm thick, about 40-mm long
No. 5: 0.60-mm thick, about 40-mm long
No. 6: 0.65-mm thick, about 40-mm long
2. Plastic foam boards and food trays
Plastic foam boards and trays areused as the base for pinning of spreadfins. Boards and trays of different sizes
should be arranged to suit different sizes

20

Fin spreading in formalin solution in a plastic food tray.

of fishes. The ideal thickness of a board
is at least 3 cm. A small fish can be fixedin formalin solution in a food tray. One
should collect a diverse range of flatbottomed food trays on a regular basis for
fixation. Sometimes, when a tray is used
several times, formalin leaks from thetiny holes created by the pins. This canbe prevented by overlapping trays of thesame size and/or placing the trays withformalin on hard plastic trays during fixation.
3. Soft sponge boards
Thin and weak pins like micropins
bend easily and sometimes break thefin membrane when pulled out from theplastic foam boards and/or food trays.
Boards made of a soft material, e.g.,sponge boards, reduce the risk of damage
to the specimen and pins. A black spongeboard is better than a white one, since it
facilitates examination of the transparent
fin membrane. The sponge board is fixed
on a flat-bottomed ceramic casserole or atupperware.
4. PaintbrushA paintbrush is used to apply formalin on the fins to spread them. A brushThe Kagoshima University Museum / The Research Institute for Humanity and Nature

Black soft sponge board fixed on a plastic storage case.

Concentrated formalin solution in small bottles, used for fin
spreading.

Paintbrushes with different sizes.

with moderate softness and good waterretention quality is the best choice. A
hard-bristled brush will abrade the bodysurface and/or even rip some scales.
Therefore, the softness of bristles should
be checked while purchasing a brushfrom the stationery. Brushes with tips ofdifferent diameters, around 3–10 mm,should be arranged to suit different sizesof fishes.
5. Formalin
A formulated concentrated formalinsolution is used for spreading the fins to
prepare a specimen. It is convenient touse formalin packaged in small bottles,
which can be hermetically sealed, before starting fin spreading in a numberof specimens. Formalin has serious toxic
consequences, and therefore, the air
should be cleared of its fumes after using
it indoors. A 10% dilute solution of formalin is used to spread the fins of a small
fish in a food tray.

Sprays to prevent dehydration of fishes during sample
treatment.

6. Atomist spray
Atomist spray is used to prevent damage due to dehydration. It can be purchased from garden supply shops.
In addition to the above-describedequipment and reagent, we use commonly used laboratory instruments, including
forceps and trays, for fin spreading.

21

Fish Collection Building and Procedures Manual■Preparation for fin spreading
Before starting fin spreading, thefishes are rinsed in water to remove themucilaginous solution and dirt adheringto the body surface (STEP 4).
Some fishes like those of Mullidae
are rinsed gently, because their scales
can easily exfoliate. On the other hand,sharks and rays and members of Channidae (snakeheads) have a hard bodysurface and can therefore be rinsed vigorously. Mucilaginous solution adhering to
the body surface should particularly beremoved taking extra care, since it often
remains at the gill pores and around themouth and fin bases.
For effective removal of the mucilaginous solution, a dishwashing sponge andhousehold detergent may be used. The
bodies of some defrosted fishes continue
to rigidify after death. They should besoftened gently at the time of rinsing tofacilitate their fixation.
After rinsing the fish body, the moisture should be wiped off the body surface
by using a paper towel. If formalin is applied on wet body surface, it will spreadon the plate and/or tray, causing discomfort to us by its odor.
When there are many fishes lined up
for fin fixation, they are considered in theorder of precedence. Fishes like herrings,
anchovies, and small gobies decay easily and therefore should be treated before
tough fishes like those of Scorpaenidae
and Holocentridae (North Pacific squirrelfish). Fishes that are to be treated later
should be stored in ice-cold water in an
ice chest to prevent spoilage.
■Fin spreading
There are numerous types of fishes in
the world, having different body shapes.
Thus, a different approach for fin spreading needs to be adopted for different
22

fishes. A general method of fin spreadingis provided below, following which specific methods for different fish shapes are
discussed.
1. Positioning of the fish
In a conventional fish specimen, thefish body is laid with the left side facing up for fin spreading. The fish is laid
in a coolite tray in such a position thatthe body axis is horizontal, with the left
side of the body facing up and the rightside facing down. An exception includes
anglerfishes and flatfishes, which are laidwith the dorsal surface facing up. Afterthe fish is laid straight in the tray, slightly
thick insect pins should be inserted at the
4 corners around the fish. If the mouthis open, it should be closed by pinningunder the jaw. In the case of fishes with aslender and round body shape, like herrings and mackerels, the head often inclines downward. To keep the body axis
straight, the fish head should be held ina slightly raised position by using a pin
and/or placing a small piece of coolite
under the head. Fishes with long bodies, like sea snakes and pike congers, areusually laid in an S or inverted C shape.
Moreover, the tails of fishes like seahorses are laid in their normal round shape.
2. Fin spreading
As much as possible, thin pins should
be used during fin spreading to prevent
damage to the fin membrane. The pins
should be inserted near the fin base along
the fin rays and not in the center of thefin. A good specimen photograph cannot
be obtained if there are distinct holes in
the fins.
After fixing the body axis, the caudal
fin (tail) should be pinned at both theends and spread naturally. The order of
spreading the different types of fins is im-

The Kagoshima University Museum / The Research Institute for Humanity and Nature

1. Fixed body axis.

2. Spread and fixed caudal fin.

3. Spread and fixed dorsal fin ray from the rear.

4. Spread and fixed anal fin.

5. Spread and fixed dorsal spiny ray.

6. Spread and fixed pelvic fin.

Process of fin spreading for specimen of Seriola dumerili.

Dragonets (Callionymidae). Above: before fin spreading.
Below: after fin spreading.

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Fish Collection Building and Procedures Manual

Fin spreading.

Spread fins just before applying formalin solution.

portant. The body does not move forward
if the caudal fin is spread and fixed before
the dorsal and anal fins. Small fishes, inparticular, tend to float in formalin because of the buoyant force. Therefore, thecaudal fin should be spread and fixed firstto prevent the floating and moving of the
fish body.
After the caudal fin, the dorsal andanal fins are spread. The fins will be wrinkle free and beautifully spread if they are
spread from the rear fin rays toward thefront. Next, the pelvic fin on the left sideis spread completely and fixed naturally.
The pelvic fin on the right side is along
the body line with pins and completely
hidden. Finally, the left pectoral fin canbe spread naturally by using the fingers.
Some fishes have free soft rays on the
pectoral fins. These soft rays, found infishes of the order Scorpaeniformes, including Scorpaenidae and Triglidae, arenot spread. On the other hand, the softrays are an important taxonomic character for threadfins, and therefore, they are
spread for these fishes.
Goatfishes and catfishes have barbels
on their chin. All the barbels are spreadand barbel angles are adjusted such thatthe barbels do not overlap when photographs are taken.

3. Application of formalin
A concentrated formalin solution isapplied around the fins after spreading all
the fins. In the case of large fishes, whichhave thick and tough fin membranes, formalin is applied on the entire fin. On theother hand, it should be very carefully applied in the case of tiny fishes and/or fishes having thin and weak fin membranes.
If it is over applied, fin membranes will
break because of shrinking after fixation.
Thus, it should only be applied at thebases of the fins. In addition, it is appliedaround the mouth to keep it closed.
Formalin fixation normally takes 5–10
min for small fish specimens, e.g., smaller than 10 cm. Large fish specimens canbe fixed in approximately 15 min.
However, it is difficult to fix a large
fish, which has thick muscle tissue (especially, the pelvic fin and/or jaw), by
simply applying a layer of formalin. To
properly fix each body part, the target
parts should be covered with a formalinsoaked paper towel.
To prevent the fish body from drying
after application of formalin, it should becovered with a paper towel and sprayedwith plenty of water.
Some fishes have elongated pelvic-finrays, such as threadfin bream (Nemipter-

24

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Application of formalin solution.

Fish body covered with a wet paper towel, sprayed withwater to prevent drying of the fish.
It is difficult to spread the pelvic fin of a large fish.
Therefore, the fin is covered with a formalin-soaked paper
towel.

us), which may come out from under thewetted paper towel. In this case, a longpin should be inserted around the fin tip.
Extreme care is required to pull out thepins after fixation. The fins may be fixed
successfully, but irreversible damage
may be caused if the pins are pulled out
roughly. Furthermore, the pins should be
carefully pulled out such that they do notcreate wide holes in the fins. After formalin fixation, the next treatment should beinitiated before the eyes become clouded
and body color changes.

4. Rinsing
After formalin fixation, the fish bodyis re-washed and prepared for obtaining photographs. To prevent any damage
before the photographs are taken, thefish specimen is stored in ice-cold water.
However, the specimen should not be
stored in ice-cold water for a long time.
■Special cases of fin spreading
For fin spreading, some fishes essentially require the most suitable treatment
according to their morphological characters.
1. Small fishes [e.g., Gobiidae (gobies),
Tripterygiidae (triplefins), and Adrianichthyidae (ricefishes)]
Small fishes and larval fishes havevery weak fin membranes, which may
25Fish Collection Building and Procedures Manual

Fin spreading in formalin solution in a plastic food tray.

Right pectoral fin (inner side) of Chelidonichthys spinosus.

Fin spreading on a black soft sponge board fixed on a plastic
storage case.

easily break on dehydration. For small
fishes, fin spreading should be performedin 10% diluted formalin solution in aplastic food tray or a plastic case bedded with a soft sponge mat. The fish
body (especially, the bodies of gobies)
tends to move by the buoyant force in theformalin solution. Thus, the fish mouthshould be held using a relatively thick
pin to prevent the body from moving or
floating. After all the fins are spread andfixed, the pin holding the mouth in placeis removed and the mouth is pinned using
new pins. For small fishes, the thinnest
possible pins should be used and thesepins should be handled with forceps.

26

Right pectoral fin (inner side) of Inimicus japonicus.

2. Fishes with ornamental pectoral fins[e.g., Triglidae (sea robins), Exocoetidae(flying fishes), and Synanceiidae (stonefishes)]
In the case of fishes in which pectoral fin is a taxonomic character, the rightpectoral fin should be removed, spread,and photographed. This makes it easy toidentify the species for taxonomic purpose.

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Fin spreading in Pisodonophis zophistius.

3. Fishes with long bases of the dorsal
and anal fins [e.g., Hydrophiidae (seasnakes), Muraenesocidae (pike congers),
and Trichiuridae (hairtails)]
In the case of fishes with long bodies
and long bases of the dorsal and anal fins,
it is easy to spread the fins by individually spreading the fin rays from the rear

Fixed specimen of Gymnothorax kidako.

toward the front. While spreading thefins, dorsal fin and anal fin rays are fixedalternately on the 2 sides, not on one side
after the other.

Fin spreading in Pterois lunulata in formalin solution in a styrene foam box. It is a good way of fin spreading for fishes with skinflaps and long fin rays.

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Fish Collection Building and Procedures Manual

Fin spreading in Pterois volitans. Fin membranes and skin flaps spread beautifully.

4. Flatfishes [e.g., Paralichthyidae (flounders) and Soleidae (soles)]In the case of flatfishes (Heterosomata), spreading of the anterior parts ofthe dorsal and anal fins is often forgotten.
The number of fin rays is an importanttaxonomic character for flatfishes. Thus,
the fins should not be fixed in a folded
condition. Before fixing the fins, the baseand tip of each fin should be checked
carefully. Since it is difficult to spreadthe fin rays on the front perfectly, forceps
should be used to manipulate the fin.
5. Fishes with long fin rays and/or skinflaps [e.g., Pteroinae (lionfishes) and species of the carangid genus Alectis]
Members of Scorpaenidae (especially,
lionfishes) have skin flaps and long finrays, such as larval stages of threadfishes.
Their fins are best spread in 10% diluted
formalin solution in a plastic case.
28

Sharks and rays do not require fin spreading.

6. Sharks and rays
Members of the shark and ray families
do not require fin spreading.

Photography→ Step 8

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 8

Photograpy
Hiroyuki Motomura

Photography of specimens needs to be
performed very carefully. Fish specimens
are fixed in formalin (STEP 12) and preserved in alcohol. However, in the courseof the fixation and preservation procedures, most of the original body color ofa fish, except black, fades away. Therefore, we have only one chance to record
the body color of a specimen.
In this section, the photography procedure is explained, along with the introduction of the equipments used in the
Kagoshima University Museum. Refer
to Information 1 for the basic technicalinformation about specimen photography
and cameras.
■Close-up copy standFish specimens have body width.
Hence, for good specimen photography,
the depth of field should be increased (seeInformation 1 for details). This requires

Close-up copy stand, light, and glass aquarium for medium
specimens.

stopping down the camera lens. However,stopping down the lens slows the shutter
speed. Therefore, it is necessary to use aclose-up copy stand for specimen photography.
■LightingFish specimens are photographed indoors, and the lens requires to be stopped
down. Therefore, lighting is a very important element of good specimen photography. In our museum, a photo-reflector lamp was used until 2007. Thereafter,
it was replaced with a fluorescent lamp
because of high heat generation (making photography in summer difficult)and short life (requiring to be turned on
and off frequently) of the photo-reflector
lamp. The following fluorescent lamps
are used for specimen photography in our
museum.

Close-up copy stand, light, and glass aquarium for small
specimens.

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Fish Collection Building and Procedures Manual

For medium to large specimens: Copy light
Company: LPL Co., Ltd.
Model: FL-217 L18527
Japanese Article Number (JAN) code:
4988115, 185309
For small specimens: Web dot studio light
Company: LPL Co., Ltd.
Model: WL-230 L18552
JAN code: 4988115, 185521■Glass tank (aquarium)
After pre-fixation (STEP 7), the photography procedure should be initiatedimmediately. The specimen should be
re-washed according to STEP 4 and immersed in a glass tank filled with water.
In our museum, 3 different sizes of tanks
are used, depending on the sizes of fish
specimens.
Small tank: 20 cm (length) × 15 cm (width) × 5 cm (height)
Medium tank: 30 cm (length) × 20 cm (width) × 10 cm (height)
Large tank: 60 cm (length) × 30 cm (width) × 20 cm (height)
The tanks are custom made, butthey are relatively inexpensive (large:
about ¥10000; medium and small: about¥5000). At the time of ordering a glass
tank, it is important to ask the manufacturer to use transparent silicone for gluing

Glass aquarium should be placed a little above the whiteboard on the close-up copy stand. Here, square wooden
blocks are used as legs for the aquarium.

30

each glass panel. If normal white silicone
is used, photography lighting will produce shadows, marring the clarity of thephotographs.
An acrylic tank is lighter and moreconvenient than a glass tank. However,its surface easily develops scratches(which will be visible in the photographs). Therefore, an acrylic tank is
not recommended. On the other hand,a glass tank is heavy and breakable. In
our museum, a glass tank is used in thelaboratory, and an acrylic tank is used in
the field. This is because a glass tank is
so fragile that it can break by slight carelessness. If the tank breaks, it will not be
possible to record the body color of thespecimen in the fresh state, even when arare specimen is obtained. To mitigate therisk, we arrange for 2 glass tanks each of
the large, medium, and small sizes in our
museum.
After the aquarium is filled withwater, it is placed on a close-up copy
stand. Prior to this, a clean white board(matte type) is placed on the close-up
copy stand, following which 4 blocks areplaced on the board as legs for the aquarium. The white board will serve as a whitebackground for the photographs (details
explained later). If the aquarium is directly placed on the white board, a shadowof the specimen will appear on the board.
Therefore, the aquarium should be about
3–10 cm above the white board.
The swimbladder is not evolved in themembers of Scorpaenoidei and abyssal
fishes, and therefore, they can easily andstably sink to the bottom of the aquarium.
On the other hand, fishes such as thoseof Perciformes live in shallow water,
and therefore, it is often difficult to sinkthem to the bottom of the aquarium. In
such a case, the right side of the abdomenshould be punctured with a needle to let

The Kagoshima University Museum / The Research Institute for Humanity and NatureSpecimen is stably lying on the bottom of the water-filled aquarium. Above: view from the back. Below: view from the front.

A slight tilt in the fish position can be adjusted using a rubber piece (e.g., an eraser piece).

the air out of the swimbladder, and thefish should then be re-immersed in water.
If the fish still floats, the right side of theabdomen should be incised to let the airout of the abdominal cavity. Small fishes
may not sink even after incising the abdomen, or even if they sink, they may
lie tilted. In such a case, a thick needle

should be inserted in the abdomen or the
pectoral area to stabilize the body by theweight of the needle. However, it should
be ensured that the needle does not overlap the pectoral and pelvic fins. It will not
be possible to process the image (STEP
16) later if the needle is visible throughthe fin membrane.
31Fish Collection Building and Procedures Manual

Specimen is not stable. The body is leaning backward. Further, the photograph is not taken from the right angle. This isan example of a poor photograph.

Specimen is stabilized using a needle. Ensure that the needle
does not overlap the pelvic fin.

■Photography technique
Refer to Information 1 for the general
specimen photography technique andcamera characteristics. Here, the actual
photography technique is explained, taking a digital single-lens reflex (SLR)
camera (Pentax K100D Super) as an example.
We use 2 kinds of lenses: generalzoom lens (DA 18-55mm F3.5-5.6 AL)
and micro lens (D FA 50mm F2.8 Macro). With one camera body and 2 different lenses as one set, we always prepare 3

sets, so that the situation can be managed
when multiple groups of students andvolunteers want to go to the field at thesame time.
The photography technique involves
the following steps.
1. Fix the camera on the pole of theclose-up copy stand. Ensure that thecamera is not tilted and has a memory
card. Replace the lens with the micro lens
while photographing small fishes. Con-

Specimen (Upeneus tragula) ready for photography (light not turned on). Remote release is hanging behind the pole of the closeup copy stand.

32

The Kagoshima University Museum / The Research Institute for Humanity and Nature

The first photograph should be taken with the tag.

nect the alternate current (AC) adapter
to the external power port of the camera.
If a battery is used, the photographs will
not be clear when the battery reaches exhaustion, and further, the camera settings
may not be saved due to battery exhaustion. Therefore, we recommend the useof external power for the camera. The
AC adapter is compatible with 240 V, andtherefore, it can directly be used outside
Japan.
2. Set the aperture priority mode (Av)to stop down the lens to focus the fish
body. Configure the aperture value settingwith the mode dial (the extent of stopping
down the lens depends on the size andthickness of the fish body; see Information 1 for details.)
3. Turn on the light and adjust thewhite balance. If the light in the photograph is the same as that in the previous
photographs, it need not be re-adjusted(the setting does not change even when
the camera is switched off). A flash lamp
may not be used.
4. Push the “Function (Fn)” buttonon the side of the monitor, and use theAuto Bracket function (this setting needs
to be configured every time). The Auto
Bracket function allows 3 successive
shots by clicking the shutter button only
once. When this function is used for the

Photography with a black background. A black board isplaced between the white board and the aquarium base. In
order to avoid reflection of roof barre and lamp in the glassof the aquarium base, it is necessary to hold a large blackboard above the camera.

first time, go to “Settings” in “Menu” andset the increment level of the exposure of
the 3 serial shots as 0.5 steps (this setting
does not change even when the camera isswitched off).
5. Configure Remote Release on thecamera. Since the aperture priority modeis used, the shutter speed becomes slow,
and the photograph can be blurred because of a little camera shake when the
shutter button is pushed. Further, since 3
successive shots are taken using the Auto
Bracket function, the shutter button needs
to be kept pressed. Therefore, it is necessary to configure Remote Release.
6. Adjust the camera lens and pole ofthe close-up copy stand to bring the fishon the bottom of the aquarium inside theviewfinder. In the case of most cameras,
the view from the viewfinder and the actual photograph do not match completely—a point that should be consideredwhile photographing the specimen. The
viewfinder field of view in Pentax K100D
Super is 96%, which means 4% of theactual photograph (the outer edge) is notobserved from the viewfinder.
7. Take the first shot such that it includes both the fish body and the specimen tag (STEP 5). Next, take 3 serial
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Fish Collection Building and Procedures Manual

shots, using the Auto Bracket function.
When a white background is used, set theinitial Auto Bracket level as 0 and thentake the photographs (the increment levelshould be set as 0.5 steps in point 4, sothat the photographs are taken at 3 exposure levels: 0, +0.5, and +1.0).
8. After 3 photographs are taken withthe white background, place a blackboard (or black velvet fabric) between
the aquarium base and the white board,and take 3 successive shots with theblack background. Each specimen should
be photographed with both white andblack backgrounds. This is because whiteparts (e.g., fins) will not be clearly visible against a white background, whereas
black parts (e.g., black spots on the finmembrane) will not be visible against a
black background.
If the specimen is photographed witha black background, with the initial Auto
Bracket level set as 0, the photographmay be too bright and overexposed.
Therefore, set the initial level as –1.5, andthen take the photographs (the specimen
will be photographed at the 3 exposurelevels of –1.5, –1.0, and –0.5). Note that
this setting depends on the model of thecamera and the brightness of the light.
When a black background is used, aroof lamp and/or patterns of roof may bereflected in the glass of the aquarium baseand on some part of the fish body. Somemeasure should be adopted to avoid thereflection. In our museum, we hold alarge black board above the camera fixed
on the close-up copy stand to avoid thereflection.
The above steps describe the actual
photography technique. With regard tosaving the photographs in the memory
card, 7 photographs for each specimen
should be saved as follows: one photo34

graph of the specimen with the tag (witha white background), 3 specimen photographs (taken in 3 steps of exposure compensation) with a white background, and3 specimen photographs (taken in 3 steps
of exposure compensation) with a black
background. These photographs are obviously digital images and can be modified
on the computer, but the overexposedparts cannot be modified because they do
not contain any graphic information. The
body color of fishes like those of Carangidae and Trichiuridae is silver gray,which is easily reflected. Therefore, overexposure should particularly be avoided
in their case. For these types of fishes, it
is safe to take photographs by using the 3steps of exposure compensation.
Very large specimens (e.g., sharks andhairtails), which cannot fit on the close-up
copy stand, can be placed on the floor andphotographed. When we photographed a
2-m-long largehead hairtail, we placed iton the floor and clicked the photographfrom the second-floor balcony. Extreme
caution should be exercised while photographing specimens outdoors, since cats
may target the specimens.

1. Details about specimen photography→ Information 1

2. Photography in field→ Information 2

3. Tagging→ Step 9

4. Image processing→ Step 16

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Information 1 Details about specimen photography
Kaoru Kuriiwa■Choice of camera and lens
1. Compact or SLR camera
The most commonly used cameras canbe divided into 2 main types: (1) compact
digital camera, which has an integrated
lens and (2) SLR camera, which has a replaceable lens. The question is which of
the 2 types is suitable for specimen photography.
The basis of selecting a compact camera will obviously be convenience. A
compact camera is small, light, and canbe used anywhere. No complicated setup
is required, and most people can easily
handle it. Basically, it captures a deepfocus shot (everything from the objects inthe background to those in the foregroundare evenly focused), without adjusting theaperture or shutter speed. On the other
hand, the lens of an SLR camera needs
to be replaced depending on the purpose;
furthermore, the body is large and heavy
as compared with that of a compact camera. Exposure needs to be set according
to the aperture and shutter speed, and thedepth of field (the depth and thickness
that come into focus) should be carefully
selected. Thus, it requires some knowledge and skill to take a good photograph.
The performance of both types of cameras is improving with time, but there still
exist significant differences in 2 basic
parts, irrespective of the extent of sophistication of the machines. The difference isin the size of the light-receiving element(sensor) and that of the aperture of thelens.
A camera receives optical information
through the sensor, and there is considerable difference in the sensor size between
compact and SLR cameras. Suppose there
are 2 cameras—a compact camera and an
SLR camera—with the same resolution,i.e., ten megapixels (10 MP). In a 10-MP
camera, the optical information received
by the sensor is delimited into 1 × 106
dots, and an image is produced. The sensor of a commonly used compact camera
is 5.7 mm × 4.3 mm (1/2.5-inch-typecharge-coupled device [CCD] sensor),whereas that of a general SLR camera isapproximately 24 mm × 16 mm (advanced
photo system [APS] sensor). As evident,
there is a difference of more than 15
times. In fact, despite both being 10-MP
cameras, there is a significant differencein the optical information of each pixel.
Thus, an SLR camera has more opticalinformation than a compact camera, allowing it to duplicate the object faithfullyand describe it in more detail. Imagine
sharing a table-sized cake and a 20-cmwide cake, each with 10 people. It isneedless to say a piece of which cake will
be larger (consider cake size as sensor
size, number of people as pixel value, andquantity of the cake as optical information). Furthermore, there is a differencein the bore diameter of the lens between
compact and SLR cameras, and an SLR
camera, which has a larger bore diameter,
has higher resolution.
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Fish Collection Building and Procedures Manual

Figure 1. Comparison of the light-receiving element (sensor)
of compact and SLR cameras.

Specimen photography for academic
purpose should be taken under better
photographic equipments to take a highresolution image. Therefore, it is preferable to use an SLR camera.
2. APS or full-frame camera
There are actually 3 main sensor sizesfor digital SLR cameras (Figure 1). One
is full-frame sensor, the size of which isthe same as that of the sensor of a 35-mm
film camera (full-frame camera). Anotheris APS sensor, the size of which is thesame as that of the sensor of an APS camera. The third type is called FourThirds
sensor, which is mainly adopted by
OLYMPUS. Full-frame sensors have so
far been adopted only by a few companies (e.g., Nikon, Canon, and SONY) in
their top models, and APS sensors are
adopted by other companies in their SLR
cameras. Here, we focus on full-frameand APS cameras.
Full-frame and APS cameras havedifferent field of view (field angle) evenwhen lenses with the same focal length
are used (Figure 2). This is due to the difference in sensor size. For example, whena lens with 50-mm focal length is used,
the field angle of a full-frame camera be36

Figure 2. Difference in the effective image field angles offull-frame and APS cameras.

comes 50 mm, but that of an APS camera
becomes approximately 75 mm, which is
1.5 times smaller (effective image field
angle). In general, field angle is expressedusing that of a 35-mm film camera (fullframe camera) as a reference. That is
why the field angle of an APS camera
lens is often indicated as “35-mm camera
conversion: __ mm.” In other words, if
a 50-mm focal length lens is used in an
APS camera, the field angle corresponds
to that of a 35-mm film camera with anapproximately 75-mm focal length lens.
However, the field angle is not actually
reduced by 1.5 times, but the visible 50mm field angle of a full-frame camera is
reduced to make it approximately 75 mm
for an APS camera.
Whether the photographer chooses afull-frame or an APS camera depends on
his preference based on the characteristics of both. The advantages of a fullframe camera include high sensitivity
because of its large sensor, broad range of
tones, and wide field angle. However, it
is expensive, large in size, and may produce limb darkening. On the other hand,the advantages of an APS camera include
telephotography, ability to achieve highresolution in the center of an image because of reduction of field angle, small

The Kagoshima University Museum / The Research Institute for Humanity and Naturesize, light weight, and cost effectiveness.
Its disadvantage is that the focal length ofthe lens is not the same as the field angle,
but is a 35-mm format equivalent (for example, a 50-mm focal length lens cannot
be used for 50-mm field angle). Further,to get the same aperture value from thesame effective image field angle, the aperture diaphragm in a full-frame camera
needs to be adjusted to narrow down theaperture by about one level. In short, it
should be noted that there is a differencein the depth of field at the same aperturelevel (f-number, described later with Figure 8).
3. Zoom or micro lensThere are 2 kinds of lenses: zoom
lens and fixed-focal-length lens. The difference lies in that the focal length of
the former can be changed, whereas thatof the latter is fixed. Zoom lens is very
convenient because it does not require
the position of the camera to be changed
to change its focal length. However, it
should be noted that its release f-number(maximum aperture, described later with
Figure 8) is generally darker and its resolution is lower than that of fixed-focallength lens. Dark release f-number is disadvantageous for specimen photography
and often requires narrowing down of theaperture. Since resolution is importantin specimen photography, low resolution of zoom lens is also a disadvantage.
In addition, zoom lens can best reduceaberration around the center of the focal
area; the aberration increases at the edgesof the focal area (barrel distortion occurs
on the wide-angle side, and pincushion
distortion occurs on the telescopic side).
Thus, fixed-focal-length lens, called micro lens, appears to be the best choice for
specimen photography.

The characteristics of micro lens are
as follows: (1) high resolution in close-up
photography, (2) high magnification (canphotograph with life-size or increased
magnification), and (3) minimum aberrations (especially distortion). In termsof resolution of a close-up photograph,micro lens shows much better performance than zoom lens. There are microlenses with zoom function (zoom microlenses) and zoom lenses with micro function, but they are quite inferior to fixedfocal-length micro lenses. In addition, themaximum magnification of micro lens is
1:1 (life-size magnification) or 1:2, whilethat of zoom lens with micro function is
only approximately 1:4.
Many companies sell various kindsof micro lenses such as standard (focal
length, approximately 50 mm), medium
telephoto (focal length, 100 mm), andtelephoto (focal length, 150 mm or more).
In specimen photography, the suitablefocal length of lens differs depending on
the size of the specimens. This is because
overhead photography is conducted usinga tripod and camera base. Furthermore,shooting distance is an important factor.
The distance between the sensor and thesubject is called shooting distance (Figure 3). A lens with shorter focal length
has a longer minimum shooting distance,and vice versa. When a large specimen is
photographed using a lens with long focal
length, the field angle lacks width (short

Figure 3. Shooting distance and working distance.

37

Fish Collection Building and Procedures Manual

Figure 4. Difference in the effective image field angles offull-frame and APS cameras. Nikon D700, AF-S Micro Nikkor 60mm f/2.8G (aperture: f/16).

Figure 5. Difference in the effective image field angles offull-frame and APS cameras. Nikon D60, AF-S Micro Nikkor 60mm f/2.8G (aperture: f/8).

shooting distance), and the image does
not fit in frame. This should be borne inmind that the actual field angle (effective
image field angle) of an APS camera isconverted into 35-mm format. The distance from the front of the lens to the object is called working distance (Figure 3).
The description of lens performance by acompany mentions the minimum shooting distance, but not working distance.
Note that even though the minimum
shooting distance is the same, working
distance can be different among lenses
with different body length. In addition, towhat extent a lens can enlarge the imageof an object is expressed as the maximum
magnification, and this is not related to
focal length. When lenses with the same
maximum magnification and different focal lengths are compared, the difference
is found to be only in the shooting distance for maximum magnification.
As mentioned above, the effective
image field angle of an APS camera becomes equal to that of 90-mm medium
telephoto macro lens in 35-mm format,
even though standard macro lens is used.
When a specimen measuring more than
40–50 cm is photographed using 90-mm
medium telephoto macro lens, the image does not fit in frame because of shortshooting distance (Figure 2). Figures 4
and 5 show how difference in field angle
due to difference in sensor size affects
the actual photograph. The photographs
are taken with a full-frame camera and an
APS camera, with both fixed in positionand having the same focal length lens (60mm) (standard length of the specimen:
26 cm). Figure 4 shows the photographtaken with a full-frame camera, and theeffective image field angle is 90 mm in
35-mm format. To photograph the same
area as in Figure 4 by using an APS camera, a lens with 40-mm shooting distance(35-mm format equivalent: 60 mm) needs
to be used or the camera needs to be positioned comparatively high. If a large
specimen is taken, an APS camera will
not photograph it as a full-frame camera
because of narrow field angle. If an evenlarger specimen is taken, macro lens will
not allow its image to fit in frame, regardless of which camera is used. In thiscase, zoom lens should be used. As statedabove, aberrations of zoom lens increaseat the edges of the focal area. Therefore,
with zoom lens, the photograph should betaken being as close to the center of the
focal area as possible.

38

The Kagoshima University Museum / The Research Institute for Humanity and Nature■Shooting procedure and settings
1. Storage formats of digital images
Photographs taken with a digital camera are stored in RAW, TIFF, and JPG formats. RAW format, as the name suggests,
is a raw image data file. It is “developed”
into image files in TIFF and JPG formats.
The word “developed” implies that a
RAW file is “converted” into a TIFF or
JPG file on the computer. A TIFF image
does not deflate during development;
thus, the image quality is good, although
the file size increases. On the other hand,a JPG image is compressed during development; thus, the file size decreases but
the image quality is low. As explained,
each format has its own characteristics.
The question is which is the best format
for storing specimen photographs.
As stated above, a RAW file contains
raw image data, and the image is simply
a digitized output signal from the sensor. The data are not adjusted by the image processor engine of the camera, andhence, the image can be retouched without deteriorating its quality. On the other
hand, TIFF and JPG files contain processed image data, and hence, the image
quality deteriorates with every retouch.
Allegorically, raw ingredients collected
for cooking correspond to a RAW image,and cooked food corresponds to TIFF and
JPG images. The concept is the same thatcooked food cannot really be re-cooked.
Many characteristics of a RAW image,such as exposure level, white balance,noise, and color tone, can be changed;
thus, RAW format is undoubtedly more
flexible than TIFF and JPG formats. If
retouch is presupposed, the image should
definitely be saved in RAW format. If
a specimen photograph is to be used ina research paper, the background often
needs to be processed and the photographsize also needs to be changed. For thispurpose, the image should be saved in
RAW format. On the other hand, data in
RAW format are very voluminous and require extra effort for conversion into JPG
format. Therefore, when many specimen
photographs are taken, the images may besaved in JPG format, and only when it isdecided beforehand that a particular image has to be used in research papers, it
may be saved in RAW format. In terms oftint of specimen photographs, white balance is very important. This point needs
to be considered while saving an image
in JPG format.
2. Camera installation and lighting
There are mainly 2 ways of placing a
specimen for photography: (1) the specimen can be placed in a transparent glasstank filled with water (Figure 6) and (2)the specimen can be placed on anti-reflex
glass and photographed on land (Figure7). Fishes with many skin flaps, such as
those of Scorpaenoidei and Lophiidae,
should be photographed in water in order
to correctly visualize the dimensions of
the skin flaps. A very large specimen thatcannot fit in the aquarium should be photographed by the latter method.
When an immersed specimen is photographed, lighting plays a very important role. Shine flashlights from eitherside of the specimen, taking care that
there is no reflection or shadow on thefish body. It often works well if a tracing paper is fixed in front of the light tosoften it. Measure the exposure with the
flashlights on, and photograph the specimen with only the flashlights on and theroom lamp off. Suitable backgroundcolor—whether black or white—should
be decided depending on the skin andscale color of the fish, although the best
way is to photograph the specimen withboth backgrounds. Furthermore, if there39

Fish Collection Building and Procedures Manual

Figure 6. Photography technique for a specimen placed inwater.

Figure 7. Photography technique for a specimen placed onland.

is air inside the abdominal cavity, thespecimen may float on the water surface
or heel over in the aquarium. In this case,puncture the right side of the abdomenwith a surgical knife to remove the air. Be
careful not to damage the organs, so thatthe water remains clean. If an organ is accidentally damaged, wash with runningwater. A glass aquarium is better than anacrylic aquarium, because it has resistance against scratches and tarnish.
For photography on land, the specimen should be placed outdoors in a shaded area. Even in a shaded area, sometimes
a different shade of the specimen may berecorded. This should be taken care of.
Pay attention to white balance too, andconfigure the settings such that, as muchas possible, the real color of the specimen
is recorded. Mist the fish frequently withwater to prevent dehydration. However,wipe the body surface lightly while photographing in order to avoid refection due
to the water adhering to the body surface.

down, and this decreases shutter speed(described later with Figure 8). Slow
shutter speed increases camera shake,and therefore, photographs should not
be taken with a handheld camera. Nowadays, many companies equip cameras
with a function of hand-shake correction.
However, this function can only “alleviate” hand shake, and there is a limit ofthe degree of correction. In specimen
photography, slow shutter speed exceeds
the limit of the hand-shake correction
function. Further, hand-shake correction
should be turned off while using a tripod,
because it will cause another shake if
kept turned on. Release is used to avoid
shake when the shutter button is pushed.
If release is not available, a self-timer (setat approximately 2 s) or a remote control
can be used.
An SLR camera is equipped with anin-built mirror. At the moment when the
shutter button is pushed, the mirror flips
up and light is received by the sensor
behind the mirror. The flipping up of themirror generates a slight shake (mirror
shock). This shake cannot be avoided byusing a tripod, camera base, release, selftimer, or even the hand-shake correction
function (this is a different kind of shake).
To avoid mirror shock, raise the mirror
before pushing the shutter button. This iscalled mirror lock-up photography and is

3. Minimization of shaking
Shaking should be minimized while
photographing. For this purpose, a tripod or camera base and remote releaseshould be used. If possible, use a camera
level vial to position the camera horizontally. It has earlier been stated that thelens aperture often needs to be narrowed
40

The Kagoshima University Museum / The Research Institute for Humanity and Naturea very good technique for specimen photography. However, all cameras do nothave the feature of mirror lock-up photography. Incidentally, Nikon provides
the exposure-delay mode (not available
in lower models). With this mode activated, the mirror flips up when the shutter
button is pushed, and then after approximately 0.4 s, the shutter is released. This,
like mirror lock-up photography, is amethod to control mirror shock.
■Photography
1. Exposure
Exposure depends on aperture andshutter speed. The aperture value, whichcorrelates with the depth of field, is very
important. Therefore, exposure is decided using manual setting or aperturepriority autosetting. In manual setting,the photographer decides the exposure
by setting the aperture value and shutter
speed. In aperture priority autosetting, thephotographer decides the aperture value,
and then, the camera automatically decides shutter speed that suits the aperture.
Please decide the correct exposure, referring to the histogram.
Figure 8 shows a correlation chart ofaperture, depth of field, light entering theaperture, and shutter speed. The aperture
value (f-number) starts from 1 and increases in multiples of 1 and √2 (1.4) insequence (that is, a sequence of multiples
of 1 and 1.4 in alternation). As the aperture is narrowed down by one level, theamount of light entering it decreases 2
times. Accordingly, for the same exposure
after narrowing down the aperture by one
level, shutter speed should be decreased
to half (shutter speed should be expressed
as multiples of 2).
Overexposure (too much light) causesclipping, and underexposure (too lesslight) . Be careful that clipping particu-

larly tends to occur with a digital camera.
If an image is saved in RAW format,
the clipped or suppressed part can be retouched, and information can later be retrieved (this technique cannot be adopted
with an image saved in JPG format).
However, if an image is completely whiteor black, the color in each part is saturated, and in this case, no information canbe retrieved even from RAW format. Theview through the viewfinder differs witheach camera model and manufacturer.
Therefore, a few photographs should betaken at each exposure level from lightto dark. It is also useful to use the Auto
Bracket function, with which a camera
automatically takes a few photographs
at each exposure level (this function iscalled the automatic exposure bracketing[AEB] function in Canon).
2. Aperture and depth of field
Fishes have body width. Hence, theyshould be photographed by narrowing
down the aperture to increase the depth offield (Figure 8). In a general portrait, thefocus is on the eyes; similarly, in a fish
specimen photograph, the focus should
be on the eyes of the fish. If the depth offield is not enough and each fin is not in
focus, the specimen photograph cannot
be used for academic purpose. Therefore,
the aperture generally needs to be narrowed down. Note that if the aperture isnarrowed down too much, resolution becomes low because of diffraction of light.

Figure 8. Correlation chart of aperture, depth of field, lightentering the aperture, and shutter speed.

41

Fish Collection Building and Procedures Manual

Figure 9. Difference in depth of field depending on f-number. Standard length of the specimen, 5.6 cm; total length of the specimen, 7.1 cm; shooting distance, approximately 30 cm. Nikon D3, AF-S Micro Nikkor 105mm f/2.8G.

This is called narrow-aperture defocus.
When the aperture is narrowed down, thepath of light entering the lens becomesnarrow. If the aperture is narrowed down
too much, the light travels around theback of the aperture diaphragm, and because of interference of this light, resolution becomes low. Diffraction occurs independently of the camera, whether film
or digital, but the effect is more notablein the case of a digital camera. This is because the smaller the sensor is, the more42

it can be affected by diffraction. An APS
camera is particularly affected severely.
Degradation of resolution due to diffraction can be interpreted as degradation ofcontrast, to be precise. In other words,
a photograph in “white - black - white”
changes to one in “white - gray - white.”
Depth of field depends not only on aperture but also on shooting distance. Long
shooting distance increases the depth offield, whereas short shooting distance decreases it. That is why the depth of field

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Figure 10. Difference in depth of field depending on f-number. Shooting conditions and equipment are the same as indicated for
Figure 9.

becomes extremely shallow in close-up
photography, in which macro lens is used.
The resolution peak of the lens itself isgenerally at f/5.6–f/8; however, although
the fish eyes or head are in focus aroundthat f-number, the caudal fin and soft raysof the anal fin sometimes appear blurred.
In this condition, the aperture may need
to be narrowed down to as much as f/32.
However, this decreases the lens resolution, and at the same time, degradation ofresolution also occurs because of diffrac-tion due to an excessively narrowed aperture. Check the focus by scaling up the
viewfinder, and look for the appropriate
f-number and depth of field. The depthof field also depends on focal length. The
shorter the focal length is, the deeper thefield becomes (pantoscope), and the longer the focal length is, the shallower thefield becomes (telescope).
3. Examples
Let us take a look at examples to un43

Fish Collection Building and Procedures Manual

Figure 11. Difference in depth of field depending on f-number and shooting distance. Standard length of the specimen, 2.2 cm;
total length of the specimen, 2.7 cm; shooting distance, approximately 20 or 35 cm. Nikon D700, AF-S Micro Nikkor 60mm
f/2.8G.

derstand the correct f-number and depthof field for photographing a specimen.
Figures 9–11 show photographs takenwith a full-frame camera. The f-numberof a full-frame camera is about one level
lower than that of an APS camera, which
has a different sensor size, to generate
the same aperture values from the same
effective image field angle. Thus, in thefollowing explanation, f/8 of a full-frame
camera corresponds to f/5.6 of an APS
44camera.
Specimen photographs taken at f/4–
f/40 (APS: f/2.8–f/36) are shown in Figures 9 and 10. The standard length of thespecimen is 5.6 cm (total length, 7.1 cm),and in all the photographs, the focus is on
the eyes. This is almost the limit of closeup photography (shooting distance, approximately 30 cm). The left side shows
the complete image; the middle shows
the image at the same f-number, with

The Kagoshima University Museum / The Research Institute for Humanity and Nature

the cephalic part cropped; and the rightside shows the image with the caudal fin
cropped as well. The photograph obviously lacks the depth of field at f/4–f/8(APS: f/2.8–f/5.6). The camera is beginning to focus on the caudal fin at f/11
(APS: f/8), and focuses it enough at f/22
(APS: f/16). In the cephalic part, on theother hand, degradation of resolution due
to diffraction becomes noticeable at f/22
(APS: f/16). It is particularly notable at
f/40 (APS: f/36). The effects of the depthof field and diffraction are only noticeable in the original image in large size,
not when the image size is scaled down
as shown here. Nevertheless, even in the
reduced image, the difference in the depthof field is quite obvious. Recent digital
camera models may be able to control theeffect of diffraction well. Figure 11 shows
the photographs taken with the lens at
two distances and f-numbers from a small
specimen (shooting distances, 20 and 35
cm; f-numbers, f/8 and f/11). When the
distance from the object was increased,large depth of field was possible evenwith small f-number.
Small fishes, e.g., a few centimeterlong ones, should be photographed froma very close distance, so that they occupy
the full frame. Therefore, the optimal fnumber tends to be large. On the other

hand, a large fish needs to be photographed from a long distance, and hence,
the aperture need not be narrowed down
too much to obtain the optimal depth offield. Figure 12 shows the photograph ofa specimen with less body depth (standard
length, 18.6 cm; total length, 22.7 cm),taken with a full-frame camera (shooting
distance, approximately 56 cm). The aperture value is f/11 (APS: f/8). Figure 13
shows the photograph of a specimen withgreater body depth (standard length, 13.9
cm; total length, 17.1 cm), taken with an
APS camera (shooting distance, approximately 68 cm). The aperture value is f/16
(full-frame: f/22).
The examples mentioned above indicate that the optimal f-number for fishes
longer than a few centimeters is mostly
acceptable up to f/22 of a full-frame camera and about f/16 of an APS camera (microscope photography is better for fishes
smaller than a few centimeters, not SLR
photography). Body depth differs depending on species. Therefore, every time aphotograph is taken, the view through the
viewfinder should be checked to ascertain
the optimal f-number. With practice, one
can roughly estimate the standard value
from the specimen size, body shape (bodydepth), and shooting distance before
pushing the shutter button.

Figure 12. Specimen without much depth. Standard length,18.6 cm; total length, 22.7 cm; shooting distance, approximately 56 cm. Nikon D700, AF-S Micro Nikkor 60mm
f/2.8G (aperture: f/11).

Figure 13. Specimen with good body depth. Standard length,13.9 cm; total length, 17.1 cm; shooting distance, approximately 68 cm. Nikon D60, AF-S Micro Nikkor 60mm f/2.8G(aperture: f/16).

45

Fish Collection Building and Procedures Manual

Information 2 Specimen photography in field
Hiroyuki Motomura

To photograph a specimen in field,
whether in the native country or abroad,facilities of local research organizations
such as local museums and universities
can be used. If they have the equipments
for specimen photography, such as closeup copy stand and appropriate lighting
system, the specimen can be photographed at the organization as explainedin STEP 8. However, if there is no suchorganization, specimen photography may
have to be conducted elsewhere such asat the hotel of stay. In this case, a small

foldable close-up copy stand and a small
light need to be carried along to the field.
Photography equipments can then be set
up using the desks and chairs in the hotel
room.
Sometimes, even though facilities of
local research organizations can be used,
the organizations do not have perfect photography equipments. If the stay is short,photography can be conducted at the hotel, as indicated above. However, if thestay is supposed to be long, as in our caseat the Kagoshima University Museum, adifferent approach should be adopted. We
brought some acrylic boards cut beforehand, and framed them in the field. Acryl
is convenient since it is light, splinterless, and easy to frame. In addition, we
brought a dismantled close-up copy standand purchased 2 stand lights found in thelocal area. There was no need to arrange
for an electric transformer for them.

Photography using a desk in hotel room.

Photography using the chairs and a table in hotel room.

46

Assembling of acrylic aquarium. Use vinyl tape to instantly
join the acrylic boards, and apply glue with the cylinder. You
can use it from the next day.

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Information 3 Side-view photography
Hiroyuki Motomura

Overhead specimen photography is
explained in STEP 8 and Information 2.
A specimen can also be photographedfrom the side, and the technique for sideview photography will be explained inthis section. As shown in the figure, thefish specimen is stabilized by placing between a glass board (or acrylic board) andfront side of the aquarium and then photographed from the side. The glass board
should be fixed using clips, like clothes-pins, so that it can easily be adjusted tochange its angle and position dependingon the thickness of the fish.
Dirt and incrustations from the fish
settle at the bottom of the aquarium.
Thus, this photography technique hassome benefits such as low frequency ofwater replacement as compared with thatin overhead photography and stabilization of the fish body without removal of
air from the bladder and/or abdominal

Specimen photography room at the Museum Support Center, Smithsonian Institution National Museum of Natural History. Sideview photography of the type specimen in progress. Camera is fixed on a tripod, and flash lamp is used for lighting. The photographed image is scanned directly into the computer.

47

Fish Collection Building and Procedures Manual

Glass aquarium seen from the front. The black board at theback of the aquarium is used for a photograph with black
background.

Glass aquarium seen from the side. Fish is stabilized byplacing between a glass board and front side of the aquarium.

Acrylic aquarium with white background.

48

Acrylic aquarium with black background.

Acrylic aquarium with black background.

cavity. However, the drawbacks are that
it is difficult to stabilize very small fishes
in the right position and that the glass
board appears in the photograph since
the fish body is placed between the glass
board and the front side of the aquarium.
The latter is especially troublesome in thecase of a fish with dermal flap on the surface of pectoral fins, because the dermal
flap is pressed against the surface of theglass board.
This photography method is adopted
by the Smithsonian Institution National
Museum of Natural History, the Kagoshima University Museum, and the Fisheries
Research Laboratory at Mie University.

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 9

Tagging
Mizuki Matsunuma and Hiroyuki Motomura

Management of specimens varies depending on the organization possessing
them. Specimens can be stored individually in bottles, with the corresponding
data note included in each bottle, or specimens can be stored in bottles groupedby species/families, with register number
tags. The latter method is adopted in our
museum; specimens are tagged and stored
in bottles, which are grouped together byHome-sewing, thick thread (white) by Yokota Co., Ltd.,
Japan. Considering its strength and price, this is the mostsuitable thread for tagging.

General surgical needle, 1/2 circle and spring eyed (#1–#10),
by Natsume Seisakusho Co., Ltd., Japan. The spring eye(needle hole) and curve are perfect for tagging.

species. In this section, tagging of specimens is explained.
After photographs are taken (STEP
8), the specimens should immediately
be assigned tags with register numbers
printed on them. Refer to STEP 5 for thematerials required and method used toprepare specimen tags.
In general, a specimen tag should bethreaded through the mouth from a rightgill slit, trying as much as possible not
to damage the fish body. In our museum,“Daruma” home-sewing, thick thread(white) by Yokota Co., Ltd., Japan, isused. Other types of textile threads canalso be used, provided they are strong
enough for the purpose. For threading,a general surgical needle, 1/2 circle
and spring eyed (#1–#10), by Natsume
Seisakusho Co., Ltd., Japan, is used. The
thread is easily drawn through the needle
eye, as it simply needs to slide down thegroove in the needle (from spring eye);

Typical method of tagging. The threaded needle is passed
through the right gill slit to the mouth.

49Fish Collection Building and Procedures Manual

moreover, the half arc makes it easier topass the needle through the gill slits.
Prepare the needle, thread, andspecimen tag. Open a right gill slit with
fingers, and pass a needle threaded witha tag through the gill slit to the mouth.
Finally, cut the thread at a reasonable
length and tie a knot to ensure that the
tag does not come off. Sometimes, it isdifficult to pass the needle through thegill slits to the mouth depending on thetypes of fishes. Different methods fordifferent types of fishes are explained
below.
■Large fishes/fishes with a long
distance between gill slits and mouth
When it is difficult to pass a needle
through the gill slits to the mouthbecause of large size of the fish, the tag
can be attached through the lower jaw.
For tagging in this case, the soft tissue
lining around the teeth should be pierced.

Tagging through the lower jaw. The soft tissue between dentaries should be pierced.

50

For tagging fishes with small gill slits and a long distance
between the gill slits and the mouth, such as those of Tetraodontidae, the gill slit area can be pierced.

Specimen in a Ziploc bag for preservation. Tags should be
inside the bag, not attached to the fish.

Specimens preserved in screw-cap vials with tags.

The Kagoshima University Museum / The Research Institute for Humanity and Nature

For tagging sharks and rays, the base of the right pelvic fin
should be pierced.

A sampling-site tag is prepared in the same manner as anumber tag in the Kagoshima University Museum.
For tagging members of Syngnathidae, tie the thread at thesulcus of arthromeres.

Alternatively, for tagging members of
Tetraodontidae and Lophiidae, whichhave small gill slits and a long distance
between the gill slits and the mouth, thegill slit area can be pierced.
■Small fishes
When the body of the fish is too small
and there is a risk of damage to the lower
jaw while trying to pass a needle throughthe gill slits to the mouth, the tag can be
enclosed with the fish body in plastic
food-storage bags (e.g., Ziploc, which is abrand of polyethylene bags with zippers).
For the large range of sizes, “Unipac”
by Seisannipponsha Ltd. (Seinichi) is
suitable. Furthermore, specimens ofhigh academic value and fragile larvae/
fingerlings should be preserved in storage

Rubber stamp for sampling-site tag.

bags. In addition, screw-cap vials and test
tubes can be used for storage.
■Members of Chondrichthyes
Since sharks and rays are generally
large and it is difficult to tag the mouth
through the gill slits, the base of theright abdominal fin should be pierced fortagging.
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Fish Collection Building and Procedures Manual

Specimen with a number tag
and a sampling-site tag. The
latter is useful to know sampling locality of the specimen.

■Elongated fishes (such as those ofSyngnathidae)
Fishes with extremely small gillslits and mouth, such as those ofSyngnathidae, need to be tagged on thecauda, with a knot tied in the thread. The
risk of losing tags can be reduced by tying
the thread at the sulcus of arthromeres.
The unique approach adopted by our
museum, that is, the use of sampling-sitetags, besides number tags, can be used.
Local museums (or local universities)
play an important role by providing
information about the local environment.
It would not be worthwhile to collectfishes from the Atlantic Ocean for
Kagoshima University. It is important
to collect local fishes from Kagoshima
and manage them. Thus, the samplingsite becomes confined to an extent,
and sampling-site tags can be prepared
beforehand.
At central and large museums (such as
National Museum of Nature and Science,
Tokyo) and Western museums, specimens
in bottles are managed by including labels inside the bottles (STEP 5), and thus,
52

information about the sampling site of thespecimen is instantly available. However,in our museum, the bottles are sorted by
the species of the specimens, in whichcase it will not be possible to know thesampling site of the specimen without
noting the specimen number and searching the database. Therefore, we prepare
sampling-site tags prior to our main surveys and attach them to the specimens
together with number tags. This method
is very useful because the sampling siteof the specimen can be instantly known.
A sampling-site tag (rubber stamped) canbe purchased for about ¥105. It is delivered within 24 h when ordered online,
and thus, sudden requirement at the field
can be managed.

Measurement→ Step 10

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 10

Measurement
Gota Ogihara

After photography (STEP 8) and tagging (STEP 9), measure the size of thespecimen with a caliper. Use millimeters(mm) as the measuring unit, and record
up to one decimal place.
Measurement methods used for fishes
vary depending on the taxonomic groups.
The projective method, which measures
the three-dimensional figure as flat figure, is used for sharks; the total length(distance between the tip of the rostrum
and the end of the caudal fin) is also measured for sharks. In case of rays, the tipof the tail is often removed (especially inmarkets), which does not allow accurate
measurement of the total length. In thiscase, the disk width (the longest distance
between the right and left pectoral fins) isalso measured. For teleosts, the standard
length (the shortest distance between thelower jaw and the end of the hypural) is
measured. The end of the hypural can beascertained by flexing the caudal fin to
the left side and observing the line on thepeduncle. In fishes with the lower jaw
longer than the upper jaw, such as those

Caliper used in the Kagoshima University Museum. Above:
VC-30, M-type standard caliper by Mitutoyo, Japan. Below:
CD-SC, series 500 ABS digimatic solar caliper by Mitutoyo,
Japan.

of Synanceiidae, Serranidae, and Hemiramphidae, measurement should be takenfrom the top of the upper jaw. In fisheswith scutes, such as those of Carangidae,
the end of the hypural cannot be easily
ascertained; therefore, fork length (theshortest distance between the top of theupper jaw and the most indented part ofthe caudal fin) is alternatively measured.
Measurement data are eventually input
and registered into a computer database.
Therefore, it is very useful in many ways
to measure and process specimens simultaneously. For instance, if it is known
how big a particular specimen is, it caneasily be identified from the numerous
individual specimens in bottles with theregistered size information. Further, if
a request is received to investigate thespecimen collection, the specimens will
not require to be physically rechecked,
but database search will provide the answers. Furthermore, by checking bodylength and sampling date in the database,valuable information such as growth
speed and appearance time of a particularspecies can be deduced at one glance. For
large specimens, it is particularly better tomeasure the size while processing them,since such specimens cannot be removed
from and placed in bottles frequently.

Identification→ Step 11

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Fish Collection Building and Procedures Manual

Step 11

Identification
Gota Ogihara and Hiroyuki Motomura

Identification involves determinationof the name (scientific name) in biological classification. In this section, the identification method used in our museum
is discussed. If the specimen could be
identified anytime between STEP 5 (Assigning tags to specimens) and STEP 10(Measurement), the species name should
be noted in a notebook. However, if thespecimen cannot be easily identified at an
early time point, identification should bepostponed so as to preserve the freshnessof the specimen for better photography.
Pepole who is not quite familiar withfishes, should first identify the taxonomic
group of the specimen (family, genus,etc.). We call this method “matchingidentification,” for which pictorial bookswith colored pictures are used. The specimen is compared with the pictures in thebook, searching for similar image (figure)and color (pattern). This method does
not require any professional knowledge.
Following books are useful for matchingidentification of fishes in Kagoshima.
The fishes of the Japanese Archipelago (Tokai University Press)
Sea fishes of Japan (Yama-kei Publishers Co., Ltd.)
Fishes of the Kuroshio Current, Japan (Osaka Aquarium Kaiyukan)
A photographic guide to the gobioid fishes of Japan (Heibonsha)
Once the family (or genus) is roughly
identified by matching identification,
search for the species on the page of thefamily (or genus) by using “Fishes of54

Japan with pictorial keys to the species(Tokai University Press)”. This bookshows a graphic representation of classification characters (characters used to distinguish a taxonomic group) and is easy
to use, but this requires some knowledgeof fish taxonomy.
If the species cannot be identified despite using the above book, refer to scientific papers (often written in English) on
each taxonomic group. This may be difficult for an inexperienced volunteer. Whenthe species cannot be identified even byreferring to scientific papers, a researcher
who specializes in the taxonomic groupto which the fish belongs should be requested to identify the specimen. If eventhe specialist cannot identify the specimen, there is a possibility that the fish has
not been taxonomically described (i.e., itwill be a new species).
Although external morphology ismainly used for identification, there aresome taxonomic groups in which external
morphology is not as clear as in others(such as, Muraenidae and Sciaenidae). In
this case, check the difference in the number of vertebrae by examining an X-ray
image and in internal morphology (suchas otolith and swimbladder) by studying
the anatomy of the fish.

Fixation→ Step 12

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 12Fixation
Hiroyuki Motomura

After identification (STEP 11), thespecimen should be immediately fixed informaldehyde solution (formalin). The
stores usually sell approximately 37%
formalin, and we call it “formulated formalin.” For fixing fish specimens, decuple-diluted formulated formalin is used.
This is called “10% formalin,” but the actual formaldehyde content of this solution
is 3.7%.
For fixation, the specimens should becompletely soaked with 10% formalin.
For medium specimens, incise the rightside of the abdomen to facilitate absorption of formalin. For large specimens,
additionally incise the right side of thedorsum. At many research organizations, formalin is injected into a body
cavity by using a syringe. However, in
our museum, surgical knife and syringe
are not used because of the presence ofmany general museum volunteers. Surgical knife and syringe are not preferable
themselves, and there is a risk of formalin
spill if excessive pressure is exerted on
the specimen. Formalin is assigned to thecategory of poisonous and deleterious
substances (specific chemical substances,second category) by Occupational Safety
and Health regulation on disability prevention from specific chemical substances and therefore needs to be handled withextreme care.
Small specimens can be soaked directly, but to avoid damage, they should

not be soaked together with medium
specimens in the same container. In our
museum, we fix very large fishes suchas giant mottled eel and oarfishes, which
need to be fixed straight, in a large tankplaced on the roof. For medium and small
specimens, we use large plastic containers and Tupperware, respectively.
Fish body consists mostly of water.
Therefore, the longer the specimens stayin a formalin fixation container, the more
the formalin gets diluted. Formulated
formalin needs to be added regularly, depending on the size and number of newly
added specimens. If too many specimens
are added without adding formalin, they
will not be fixed and begin to decay. Pres-

Formulated formalin: 37% formalin solution. The product islabeled as “third type,” but it is currently designated as “second type” by Occupational Safety and Health regulation.

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Formalin fixationof medium to large
specimens.

ence of bubbles on the surface of formalin solution, even only a few, is an initial
sign of contamination. Once the specimen
begins to decay, formalin contamination
cannot be inhibited even by adding highly
concentrated formalin. As a strategy, thedecaying specimen can be fixed in 98%
ethanol; this will fix the specimen well

Formalin fixation of small specimens.

A tank for fixation of a large specimen. It is placed on theroof and locked.

56

and the problem of contamination will
be resolved. However, ethanol has a very
bad odor, and it cannot be reused. Therefore, it is important to monitor the densityof formalin in order to avoid contamination.
The period of soaking a specimen informalin (fixation period) varies depending on research organizations. Mostly, it
is 7 days to 3 weeks; in our museum, it
is approximately 10 days. In 7 days, thespecimen is not fixed well, and it will
dehydrate on alcohol treatment (STEP
13). Dehydration will produce many
wrinkles on the body surface and makethe specimen quite hard (especially fishesof Carangidae). In our museum, weregularly prepare a large number of new
specimens, and we do not have enoughnumber of (and space to place) formalin
fixation containers to soak specimens forweeks. Therefore, 10 days is the best duration for both complete fixation and fast
rotation.

Replacement of formalin with alcohol→Step 13

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 13 Replacement of formalin with alcohol
Hiroyuki Motomura
After fixation, replace formalin withalcohol for long-term storage of specimens. There are several reasons to replaceformalin with alcohol, a couple of which
are that 1) formalin produces formic acidon oxidation, which demineralizes hardtissues such as fish bones and 2) formalincauses nose irritation and is very poisonous, which makes prolonged examinationof specimens difficult. However, alcohol
is more expensive than formalin, andmany research organizations actually preserve large specimens, which need a large
amount of alcohol, in formalin solution(without replacing with alcohol). Whena specimen is to be retained in formalin,
add sodium hydrogen carbonate (bakingsoda) or hexamethylenetetramine in formalin to neutralize formic acid, and thenuse it to avoid degradation of bony tissue.

If a formalin-fixed specimen is directly placed in alcohol, it can simply be assumed that the same amount of formalinas the volume of the fish will percolate
into alcohol, which will later create problems while examination of the specimen.
Therefore, the specimen should be first
immersed in real water to exude formalinfrom it. Thereafter, it should be placed in“substitutive alcohol” for several days;
used alcohol can be used as substitutive
alcohol. At this stage, formalin and realwater are completely removed from thespecimen and replaced with alcohol. Alcohol used in our museum is “general alcohol; ethyl alcohol, 99 vol%; synthesis,
unmodified; 18L TNS.”
Storage→ Step 14

As the first step of replacing formalin with alcohol, the formalinfixed specimen is soaked in realwater for 1 day to exude formalin from it.

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Step 14Storage
Hiroyuki Motomura

After replacement of formalin withalcohol, the specimen should be stored
in a specimen bottle filled with alcohol.
In foreign museums, glass jars are often
used as specimen bottles, but in Japan,
glass jars of reasonable size and price are
not readily available. Therefore, in many
Japanese museums, specimen bottles
made of polyvinyl chloride are used. In
our museum, we choose the bottle size
depending on the specimen size. We use
a 3-L wide-mouthed T-type bottle, which
is useful for fishes with large body width,such as blue-striped angelfish, although
air circulation is more in this bottle.
Sometimes, the lid can slip while carrying the bottles; therefore, they should be
carefully handled. Following is a list of
specimen bottles used in our museum.

AS ONE Corporation
Wide-mouthed T-type bottle with rubber seal (clear vinyl chloride)
0.3 L (width, 75 mm; height, 92 mm)
0.5 L (width, 90 mm; height, 118 mm)
1 L (width, 97 mm; height, 167 mm)
2 L (width, 112 mm; height, 255 mm)
3 L (width, 134 mm; height, 263 mm)
Wide-mouthed bottle with inner lid(clear vinyl chloride)
0.1 L (width, 48 mm; height, 82 mm)
0.25 L (width, 61 mm; height, 119 mm)
Wide-mouthed BB-type bottle without
stopcock
20 L
Plastic container 75 L
Sanko Co., Ltd.
Plastic drum (open type)
25 L (width, 296 mm; height, 522 mm)
30 L (width, 312 mm; height, 502 mm)
120 L (width, 490 mm; height, 792 mm)
210 L (width, 580 mm; height, 974 mm)
Power drum (open type)
60 L (width, 400 mm; height, 618 mm)

Specimen bottles by AS ONE Corporation. From the left:
0.25-L, 1-L, and 2-L bottles.

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Specimen bottles are labeled withfamily number and species name and arranged according to the number. Specimen preparation and arrangement aredone by volunteer staff members. Therefore, we use standard Japanese names

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Storage containers for medium to large specimens.
Upper row: 20-L wide-mouthed BB-type bottle without
stopper cock. Lower row: 25-L plastic drum. Label thefront side with white vinyl tape, and write the family
number and species name with alcohol-resistant
Sharpie Fine Point pen (sold in Europe and the UnitedStates).

Specimen bottle labeled with family number and speciesname (2-L wide-mouthed T-type bottle).

Thermoelectric printer (Name Land; CASIO) for printing labels for specimen bottles. We use black ink for normal specimens and red ink for type specimens. Further, we use 12mm-wide tape for 0.5- to 3-L specimen bottles and 9-mmwide tape for 0.25-L specimen bottles.

Specimen fixed in 99% ethanol (for DNA analysis). Label isprinted using a thermoelectric printer, which prevents deterioration or loss of print.

instead of scientific names as speciesnames. Labels are printed using “NameLand” by CASIO, which is a type ofthermoelectric printer. A thermoelectric
printer provides excellent resistance to alcohol (see Information 4), which preventsloss of label print because of alcohol.

Shelf for storage of specimen bottles.

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Information 4

Specimen data labels
Hiroyuki Motomura

Because of the storage capacity of thespecimen room in the Kagoshima University Museum, we allocate one number tag
to one immersion specimen, and store/
control it in a specimen bottle that issorted by species. Other museums, however, including many foreign museumsof natural history and the National Science Museum in Japan, store one lot of
specimens in one specimen bottle. In thiscase, various information regarding thespecimen is written on the label (specimenlabel), which is placed together with thespecimen in the bottle. In this section, I

will explain the pertinent details regarding specimen labels.
Because immersion specimens are
stored in alcohol, which is highly volatile, the label and ink that are use must be
sure to take this into account. Moreover,since specimens are intended to be passed
down from generation to generation as
assets shared by all human beings, theymust be preserved properly not just on ascale of years or decades, but on a scaleof centuries.
Presently, there are four types of printers that are commercially available forthe printing of labels.
■Ink jet/ Bubble jet printer (In this technology, the ink is sprayed onto the paper)
Normal ink will blur if it is soaked
in alcohol, but a specifically alcoholresistant ink has recently been introduced
to the market. In the Natural History
Museum, London, for example, specimen

Specimen label used in the Smithsonian National Museumof Natural History, USA.

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Specimen label printed by an ink jet printer using alcoholresistant ink, in the Natural History Museum, London (photoby J. Maclaine).

The Kagoshima University Museum / The Research Institute for Humanity and Naturelabels are printed using an ink jet printer.
However, it is an open question whether
today’s alcohol-resistant ink will remain
legible into the long-term future.
■Laser printer (This kind of printerprojects the picture via a laser onto a
photoconductive drum. The drum thenprints the picture onto paper in the formof toner, via direct contact and heat)
Since the script or image is not printed
under pressure, the ink will come off
if the paper is soaked in alcohol. In theAustralian Museum, Sydney, and theMuseum and Art Gallery of the Northern
Territory, Darwin, printed specimen labels are heated in an oven and soaked inalcohol after the ink has been completely
released onto the paper. Acrylic-coatedpaper, as we will explain later, is not suitable because it will not accept the ink.

Specimen label printed with laser printer and general ink
used in the Academy of Natural Sciences, Philadelphia. Thislabel is acrylic-coated.

Specimen label printed with laser printer and general ink.
The print has become illegible (photo by D. Catania).

In the Field Museum of Natural History, Chicago, they print labels using alcohol-resistant ink and use specimen labels
without subjecting them to heat treatment.
At the University of Michigan Museumof Zoology, Ann Arbor, and the Academy of Natural Sciences, Philadelphia,
on the other hand, they print specimenlabels using a general ink toner, acryliccoat the labels using spray (Krylon Matte
Finish), and then apply them. Acrylic
coating, however, can be dissolved by fatthat is exuded from specimens (especially
Chondrichthyes), and thus is not practicalfor such applications.
In the Academia Sinica, Biodiversity
Research Center, Taipei, specimen labels
are printed using normal ink, and are thenlaminated so they can be soaked in alcohol. The problem is that the laminate will
eventually come unstuck. This method is
also not recommended due to the cost andthe labor involved.
■Dot impact printer (This kind of
printer applies pressure to a ribbon that issaturated with ink, and thus prints words
and images using thousands of tiny dots)
In this case, the printed label does notblur and come unstuck, but one cannot
expect clear printing because this is a
low-resolution technology. A dot impact
printer was traditionally used in the Cali-

Specimen label printed with dot impact printer and alcoholresistant ink. The print is too light and hard to read (photo by
D. Catania).

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Thermoelectric printer: Datamax I Class Thermal Printer(photo by D. Catania).

Specimen label printed by Datamax I Class Thermal Printer(photo by D. Catania).

moelectric printers are predominantly
used for the production of specimen labels.

Datamax I Class Thermal Printer. In the photo, the black rollis the ink ribbon and the white roll is acrylic-coated paper(Photo by D. Catania).

fornia Academy of Sciences, San Francisco, but the alcohol-resistant ink ribbons
it uses are no longer available. Therefore,
this method is practical only in museums
that still have a considerable stock ofalcohol-resistant ink ribbons.
■Thermoelectric printer (This kindof printer transcribes words to paper byheating an ink ribbon on film)
This printing technology is characterized by its resistance to blurring and coming unstuck, and is thus the best printerfor the printing of specimen labels. It has
the disadvantage, though, of the overall
cost of the ink ribbons it uses, which areexpensive. In American museums, ther62

As mentioned above, various printing
methods are used in different museums to
produce specimen labels. There are drawbacks and advantages involved in the
choice of a given product, due to the rapid
development of printers and ink, and thediscontinuation of existing products. Thatsaid, the safest (since the print will endure for the long-term) and clearest (since
the print can easily be read) method at
the moment is printing on acrylic-coatedpaper (a thick, shiny, water-resistant paper) using a thermoelectric printer. The
Datamax series of thermoelectric printers,
made by Alpha System, in the U.S., isthe most widely used printer in American
museums.
Three different kinds of ink ribbon
can be used in a thermoelectric printer:waxed, plastic, and a combination wax/
plastic ribbon. The plastic ribbon is forlight printing, and the waxed ribbon cancome unstuck, so the combination ribbon
is regarded as being best for the printingof specimen labels.

The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 15 Creation of database
Hiroyuki Motomura
tion system saves the trouble of inputting
Input data immediately after cominformation.
pleting a series of specimen preparation
In order to manage the data safely forprocedures. In our museum, we use Filea long period, we print thousands of copMaker Pro 9 as the database software andies of the input data and bind them for
set up the database to enable people tostorage.
access it, input data into it, and search itfrom any computer on the campus.
In our museum, there is a dictionary file in which fish name (Japanesename and Latin name) and species name(standard Japanese name and scientificname) are linked. If the standard Japanese name of a species is input, the Japanese and Latin names of the family andthe scientific name of the species willautomatically be generated. For autoScreen shot of list-style display. If the standard Japanesename is input in the second row from the top, family namemated input system, the correct standard
and species name will automatically be generated in the firstJapanese name should be input, and not
row and third row, respectively. This figure shows selectionof habitat environment from the pop-up menu.
the scientific name, which is mistakable.
The settings cancompletely prevent
input error. We use
pop-up menus forselecting samplingsite (Pacific Ocean,
Sea of Japan, etc.),
habitat environment(saltwater, freshwater, brackish water, etc.), sampling
method (use of fill
net, set net, angling,etc.), and the optionof “with or without
shot of chart-style display. Information can be input/searched in the database in 3
picture.” This selec- Screen
styles, namely, list style, chart style, and form style, with FileMaker Pro 9.
63Fish Collection Building and Procedures Manual

Step 16 Image processing
Hiroyuki Motomura
In this section, image processing
method is explained. The following steps
explain the actual procedure employedin our museum. We use the Adobe Photoshop 7.0 software, but basically any
version may be used for this method. The

following procedure may appear cumbersome, but once one gets used to it, an image can be processed in about 3 min.
Steps 4 and 5 can be omitted for animage with a white background.

1. Unprocessed original image.

2. Fine tune the image. Select “Image” → “Compensate
Color Tone” → “Level Compensate.” A dialog box will appear. Adjust the triangle and click “OK.”

3. Rotate the image such that the specimen is horizontal. Select “Image” → “Rotate Canvas” → “Input Angle.” A dialog
box will appear. Type the angle value and click “OK.”

4. Roughly paint the background black. Select “pencil tool”
and increase the diameter of the brush. The area close to thefish need not be painted.

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The Kagoshima University Museum / The Research Institute for Humanity and Nature

5. If needed, zoom in and paint the background black.
Roughly select the borderline of the image and backgroundby using “rectangle select tool” and click on the selectedarea with “fill paint tool.” If this automatically paints the
specimen black, reduce “Latitude” and click the selectedarea again.

6. After painting the background black, crop the excess
background.

7. The image after cropping the excess background.

8. Before saving the image, adjust resolution and image size.
Select “Image” → “Image Resolution.” A dialog box willappear. In the case of this image, the resolution, width, andpixel value are 72 pixels/inch, 804.33 mm, and 8.64 MP, respectively. In the dialog box, change the resolution from 72
to 300 pixels/inch.

9. Pixel value can be increased by increasing resolution. To
decrease the pixel value to as small as it was before changing resolution, decrease the width (image quality cannot be
improved beyond that of the original image even by increasing the pixel value). In this image, by changing the widthfrom 804.33 mm to 190 mm, the pixel value was increased
to 8.37 MP, which was almost the same as it was beforechanging the width (8.64 MP). Image quality at 190-mm
width and 300 pixels/inch resolution is acceptable for almost
any type of printing.

10. Finally, select “File” → “Save as.” Set the image quality
to 12, and confirm that the compression is set as “Highest(Low Compression Rate).” Select the “Baseline (Standard)”
option.

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Fish Collection Building and Procedures Manual

Unprocessed original
image.

Processed image withblack background.

Processed image withwhite background.

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The Kagoshima University Museum / The Research Institute for Humanity and Nature

Step 17

Loan
Hiroyuki Motomura

Stored and managed specimens are
not simply maintained in the collection
but utilized for various research purposessuch as for biodiversity, taxonomic, andecological research. Some researchers visit organizations and examine
specimens, but most of the others borrow
specimens to examine them at their organization. In our museum, there were 52
cases of loan requests (681 specimens)
only in the fiscal year 2008 (which does
not include a huge number of requests foron-campus use).
In this section, mailing of specimens
is discussed. Wrap each specimen carefully in a gauze dampened with 20%
alcohol. Transportation of highly concentrated alcohol is prohibited by regulations
such as Special Provision A58 in theInternational Air Transport Association(IATA) Dangerous Goods Regulations;
therefore, alcohol is diluted to 20%. Pack

the wrapped specimen in 3 folds of thickplastic bag. Use sealer to seal the package, and try to enclose as less air as possible while sealing. Ziploc or any otherthin vinyl should not be used, because
alcohol can leak even on a small change
in atmospheric pressure.

FV-801 sealer by Hakko Corporation, used for sealing specimens in vinyl.

Packed specimen is placed in a cardboard box filled withcushioning materials.

Specimen is wrapped in a gauze dampened with alcohol andpacked in vinyl.

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Fish Collection Building and Procedures Manual

Place the hermetically sealed specimen
in a cardboard box filled with cushioning
materials. Ensure that the cardboard box
is fairly strong.
Before shipping the box abroad, attacha clear Ziploc bag on the top or side of
the box and enclose an invoice (specificlist of the content) in the bag. This helps
to confirm at the Customs that the content is not a Convention on International
Trade in Endangered Species of Wild
Fauna and Flora (CITES)-listed speciesor (tax-free) product. There is a column
for the price of the content on an international package, and an object that costs
more than US $20 is subject to taxation.
NVC (which means “no commercial
value”) or a price of US $20 or lower canbe written in the price column.

Even when the invoice is included, thecontent is sometimes checked at the Customs. In this case, there is a possibility
that the vinyl package is opened. Therefore, a written warning should be enclosed with the box. The following points
should be mentioned: (1) the content is aformalin-fixed specimen for research purpose and has no trouble with quarantine;(2) the species is not listed in CITES; (3)the specimen is preserved in 20% alcohol; and most importantly (4) the package
should be immediately resealed.
Two same invoices should be mailed
with the package while shipping. One ofthem should remain with the borrower ofthe specimen, whereas the other should
be signed by the borrower and returned to
the lending organization as receipt. The
borrower usually writes the result of reidentification on the copy of the invoice.
For general specimens, the loaning period
is normally for 1 year.

Loan invoice for specimen. Information such as
specimen data and loan period is listed.

Important memorandum to be enclosed with thespecimen in the cardboard box.

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The Kagoshima University Museum / The Research Institute for Humanity and Nature

Fish photographs, taken with methods described in this manual, were
published in the following books. These books can be downloaded as PDFs from:http://www.museum.kagoshima-u.ac.jp/staff/motomura/dl_en.html

Motomura, H. and K. Matsuura (eds.). 2010
(Mar.). Fishes of Yaku-shima Island – A
World Heritage Island in the Osumi Group,
Kagoshima Prefecture, Southern Japan.
National Museum of Nature and Science,
Tokyo. viii + 264 pp., 704 figs.

Matsunuma, M., H. Motomura, K. Matsuura, N.
A. M. Shazili and M. A. Ambak (eds.). 2011
(Nov.). Fishes of Terengganu – East Coastof Malay Peninsula, Malaysia. National
Museum of Nature and Science, Tokyo,
Universiti Malaysia Terengganu, Terengganu, and Kagoshima University Museum,Kagoshima. viii + 251 pp., 678 figs.

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Fish Collection Building and Procedures Manual

Front page left images: Fishes of Kagoshima Bay, Japan
Front page central image: Devil Scorpionfish, Scorpaenopsis diabolus, from Tanegashima, Kagoshima, Japan
Back page right images: Fishes of Yakushima, Kagoshima, Japan70