. 

NOAA Technical Memorandum NMFS 

PROCEDURES FOR SAMPLING DOLPHINS: "I 

. A 'HANDBOOK FOR SHIPBOARD OBSERVERS 

Albert C. Myrick, Jr. 

JULY 1986 

NOAA-TM-NMFS-SWFC-62 

U.S. DEPARTMENT OF COMMERCE 

National Oceanic and Atmospheric Administration 

National Marine Fisheries Service 

Southwest Fisheries Center 
NOAA Technical Memorandum NMFS 

The National Oceanic and Atmospheric Administration (NOAA), organized in 

1970, has evolved into an agency which establishes national policies and 

manages and conserves our oceanic, coastal, and atmospheric resources. An 

organizational element within NOAA, the Office of Fisheries is responsible for 

fisheries policy and the direction of the National Marine Fisheries Service 

(NMFS). 

In addition to its formal publications, the NMFS uses the NOAA Technical 

Memorandum series to issue informal scientific and technical publications when 

complete formal review and editorial processing are not appropriate or feasible. 

Documents within this series, however, reflect sound professional work and may 

be referenced in the formal scientific and technical literature. 
NOAA Technical Memorandum NMFS 

This TM series IS used for documentation and timely communication of preliminary results. irnterirn reporis, or 

special purpose informallon. and have not recetved complete formal renew. editortal control, ci delailed edibng 

JULY 1986 

PROCEDURES FOR SAMPLING DOLPHINS: 

A HANDBOOK FOR SHIPBOARD OBSERVERS 

Albert C. Myrick, Jr. 

Southwest Fisheries Center 

National Marine Fisheries Service, NOAA 

La Jolla, California 92038 

NOAA-TM-NMFS-SWFC-62 

U.S. DEPARTMENT OF COMMERCE 

Malcolm Baldrige. Secretary 

National Oceanic and Atmospheric Administration 

Anthony J. Calio, Administrator 

National Marine Fisheries Service 

William G. Gordon. Assistant Administrator for Fisheries 
TABLE OF CONTENTS 

Page 

INTRODUCTION ............................................... 1 

SAMPLING EQUIPMENT AND USE ................................. 2 

Measuring Devices ..................................... 10 

Labels. Cable Ties. and Labeling ...................... 14 

Gloves ................................................ 14 

Containers. Preservatives. and Specimen Handling ...... 15 

Porpoise Life History Form ............................ 2 

Pruning Shears ........................................ 17 

Knives ................................................ 17 

COLLECTION EFFORT AND PROCESSING DECISIONS ................. 18 

PARTIAL-SAMPLE SELECTION ................................... 19 

COLLECTING CARCASSES FOR SHORE-BASED PROCESSING ............ 19 

SPECIMEN PROCESSING ON BOARD ............................... 20 

GETTING READY ............................................ 21 

NON-DOLPHIN ITEMS ........................................ 21 

EXTERNAL ANATOMY ITEMS ................................... 21 

Coloration/Sketch/Fin Trace/Photos .................... 21 

Total Length .......................................... 24 

Sex ................................................... 25 

Specimen Number ....................................... 25 

Advance Preparation of Labels ......................... 25 

Lactating/Milk/Mammary Gland .......................... 31 

Teeth ................................................. 31 

Head .................................................. 34 

INTERNAL ANATOMY ITEMS ................................... 36 

Organization ........................................... 36 

Mid-ventral Incision ................................... 38 

Short Incision ........................................ 38 

Extended Incision and Thoracic Flap ................... 38 

Sampling from the Abdominal Cavity ..................... 40 

Urinary Bladder ....................................... 40 

Female Reproductive Tract ............................. 43 

Major Organs of the Thoracic Cavity .................... 53 

Liver/Diaphragm/Lungs ................................. 55 

Heart/Dorsal Aorta .................................... 55 

Parasites .............................................. 55 

Male: Testis and Epididymis ........................... 47 

Stomach ............................................... 51 

i 
FINISHING ................................................ 57 

ACKNOWLEDGMENTS ............................................ 57 

FURTHER READING ............................................ 58 

INDEX ...................................................... 59 

ii 
LIST OF TABLES 

Table Page 

1 Labeling, packaging, and storage requirements for 

some dolphin organ and tissue samples ............. 30 

iii 
Figure 

1 

2 

3 

4 

5 

6 

7 

8 

9 

10 

LIST OF FIGURES 

Page 

Porpoise Life History Form.................... 3 

A. Measuring sticks and moveable caliper jaws positioned for calibrating calipers. Arrow at A indicates 

where the measurement should be read. Arrows from B 

indicate position of setscrews for locking or adjusting 

the jaws. B. Taking a dolphin's body length measurement with measuring stick. One caliper jaw is in the 

tail notch and the other one is at the tip of the upper 

jaw ...........................................I1 

Carcass of dolphin in sketch position (photo taken by 

J. Greene) .................................... 22 

Sketch and fin-trace requirements. A. Outlining the 

pigmentation pattern on a Porpoise Life History Form 

to fulfill the sketch requirement for spinner dolphins. 

B. Using the blank side of the form to trace the fin of 

a spinner dolphin ............................. 23 

Ventral surface of female dolphin showing genital slit, 

anal slit, and paired mammary slits. The genital and anal 

slits are almost continuous in female dolphins (photo 

courtesy of W. Perrin) ........................ 26 

Ventral surface of male dolphin showing umbilical scar, 

genital slit, and anal slit. The genital and anal 

slits are well separated in male dolphins (photo 

courtesy of W. Perrin) ........................ 27 

Use of blunt end of pencil to determine the sex of a 

dolphin. If probe can be pushed backward when inserted 

into genital opening, the specimen is male. If it can 

be pushed forward, the specimen is female. Photo shows 

a male dolphin ................................ 28 

Regional Observer Life History Requirements 

chart.........................................29 

Two-step check of female dolphin for lactation. 

A. Each mammary is pinched several times. B. If no 

milk shows, each mammary is cut transversely to a depth 

of about two cm and pinched again ............. 32 

Collecting a jaw sample of teeth. A. A slit is made 

along the inside of the left jawbone. B. One blade 

of the shears is inserted into the slit and the jaw is 

severed transversely. C. The jaw is sheared again about 

eight to ten middle-teeth away from the first truncation. .............................................. 33 

iv 
11 Collecting the head of a dolphin. A. The initial 

incision is lengthened and deepened as the beak is 

pulled down toward the belly. B. The occipital 

condyles (arrow) pull away from the atlas vertebra; 

then the neck is girdled and the head completely 

severed .......................................35 

12 Line drawing showing organization of a female dolphin 

internal anatomy in ventral view (length protracted) 

..............................................37 

13 Short mid-ventral incision of female dolphin for 

collecting posterior abdominal organ samples. 

A. Incision extends from behind the umbilical scar to 

the anal slit. B. Incision is deepened. C. Incision 

completed. Internal abdominal organs can be seen 

(arrow) .......................................39 

14 Extended mid-ventral incision for collecting organ 

samples from thoracic and abdominal cavities. 

A. Incision is made anteriorly from the anal slit to 

the sternum. B. Ribs are cut from their cartilaginous 

attachments to the sternum. C. The left side is 

reflected to expose organs of the thoracic and 

abdominal cavities (white and gray knobs at upper 

margin of cavity are knife handles). Especially note 

the double sheet of peritoneum in the genital region. 

One sheet supports the urinary bladder; the other (more 

dorsal) supports the uterus ................... 41 

15 Position of the urinary bladder. A. Finger at right 

points to bladder, the most ventral abdominal organ. 

B. Bladder lifted to show sheet of supporting 

peritoneum ....................................42 

16 Freeing the left uterine horn and labeling it in place. 

A. The left ovary (arrow) and the end of the left horn' 

are held for protection as the peritoneal tissue is cut 

away from them. B. Cutting is halted when the body of 

the uterus is reached. A cable-tied label is then 

cinched securely to the left horn, half-way between the 

ovary and the uterine body .................... 44 

17 Handling large fetuses. 

A. A nearly full-term whitebelly spinner dolphin fetus 

in the left uterine horn. The left ovary is in the 

observer's left hand. B. For fetuses larger than 25 

cm, the uterus is slit and the fetus removed, sexed, 

measured, and discarded, but the uterus and ovaries are 

collected. .................................... 46 

V 
18 Paired organs of the abdominal cavity in male dolphins. 

A. Testes with epididymides. The epididymis is longer 

than the testis on either end (arrows). B. Kidneys and 

adrenal glands. C. Comparison of position, size, and 

shape of kidneys and testes of mature male dolphin. 

Testes are suspended from the lateral walls by 

peritoneum; kidneys are technically excluded from the 

body cavity by peritoneum .................... 48 

19 Collecting the right testis and epididymis. 

A. The testis is pulled away from wall, making taut the 

suspensory peritoneum for easier cutting. 

B. Peritoneum is cut along the outside of the 

epididymis and the posterior end of the organ complex 

is severed well behind its actual terminus. 

C. Testes longer than 15 cm are slit lengthwise down to 

their centers to allow the fixative to penetrate the 

entire sample ................................. 50 

20 Defining the limits of the stomach. 

A. The connecting tissues of liver, diaphragm, and 

peritoneum are cut away to fully expose the esophagus. 

B. The tissues covering the three stomach compartments 

are removed. C. The pyloric sphincter (arrow) and the 

duodenal ampulla (D.A.), at the anterior end of the 

small intestine, are found by searching along the 

posterior end of the stomach ................. 52 

21 Removal of the stomach. 

A and B. The stomach is tied off at the esophagus and 

at the small intestine below the duodenal ampulla. The 

esophagus is severed above its tie and the small 

intestine below its tie. The sample is lifted up and 

out of the cavity as the remaining connecting tissues 

are cut away from it. 

C. The stomach is examined to ensure that it is intact. 

..............................................54 

22 Major organs of the thoracic cavity. 

A. The liver and lungs may be confused. They are 

similar in shape and size and sometimes in color. Lungs 

are spongy to the touch; liver is more compact. 

B. The heart lies ventromedial to the lungs, encased in 

a sac of peritoneum. The largest chamber of the heart 

is the left ventricle. It is a potential source of 

aerated blood if blood is to be sampled. 

C. The dorsal aorta lies along the dorsal wall of the 

thoracic cavity above the lungs and esophagus. This 

artery - is a - potential source for blood samples. .............................................. 56 

vi 
PROCEDURES FOR SAMPLING DOLPHINS: 

A HANDBOOK FOR SHIPBOAD OBSERVERS 

Albert C. Myrick, Jr. 

Southwest Fisheries Center 

National Marine Fisheries Service, NOAA 

La Jolla, California 92038 

INTRODUCTION 

Every year, since the late 1950's when purse-seine fishing 

on dolphins to catch yellowfin tuna was first introduced, there 

has been an incidental kill of dolphins by purse seiners in the 

eastern tropical Pacific Ocean (ETP). As part of an effort to 

evaluate the biological consequences of this inadvertent "take" 

on the affected dolphin populations, biological technicians 

(observers) of the National Marine Fisheries Service (NMFS) and 

the Inter-American Tropical Tuna Commission (IATTC) accompany the 

seiner fishermen on selected cruises to collect biological 

samples and data from dolphins that are killed. Upon completion 

of each cruise, the observer delivers his samples to the NMFS 

laboratory of the Southwest Fisheries Center (SWFC) in La Jolla, 

California, for study. These samples and data are important to 

the scientific monitoring effort because they represent all of 

the life history data that NMFS scientists have to work with in 

assessing the status of these wild dolphin populations. 

Because the life history samples are studied together, it is 

vital that all samples be comparable. This requires that they be 

collected with as much uniformity as possible by all observers. 

Divergent methodologies increase the risk of sampling error. For 

example, if each observer chose his own method or equipment for 

measuring body length, a considerable variance in the data could 

result. Square measurements of a straight body axis might 

provide a true approximation of maximum length, while tape 

measurements following the curvature of the body would produce 

highly erroneous values. Conceivably, length measurements of the 

same adult taken by different methods could differ by many 

centimeters depending on the method and the equipment used. 

Because one of the criteria for distinguishing between certain 

ETP dolphin stocks is average adult length, the substantial error 

introduced by various methodologies in collecting length data 

could render the length criterion useless for stock 

discrimination. Thus, for body length measurements and for many 

other aspects of data collecting and specimen sampling, standards 

are needed to minimize procedural drift by observers. 
2 

This handbook was written to provide procedural guidelines 

for shipboard observers in collecting dolphin organ and tissue 

samples and associated data. Its focus is on current and 

potential sampling needs of the SWFC and IATTC. The handbook 

contains a description of the sampling equipment and its proper 

use and care. It covers a number of sampling situations and 

explains the priorities that must be considered in deciding what 

sort of collection to make. Elements of some organ systems are 

described and illustrated in this handbook, but it is not 

intended as a complete atlas of dolphin anatomy. (I have 

appended a short reading list for those readers interested in 

more thorough treatments of mammalian and cetacean anatomy.) 

Finally, included in the handbook is a suggested collecting 

routine that minimizes specimen processing time without 

sacrificing sample integrity. Although the handbook was prepared 

withthe observer trainee in mind, others should find parts of it 

useful also. It is indexed for ready referral to key-word topics. 

SAMPLING EQUIPMENT AND USE 

The sampling equipment described in this handbook is the 

equipment that is regularly issued to observers who accompany 

tuna purse seiners. 

Porpoise Life History Form 

A Porpoise Life History Form (the form) is used to report 

biological and other field data for each dolphin carcass 

processed (Fig. 1). The form is your most important sampling 

tool. Use it to remind yourself of what you have and have not 

done in sampling a specimen. 

The importance of the data contained on a completed form 

cannot be overemphasized. Used in concert with other reporting 

documents, such as the Marine Mammal Set Log and the Marine 

Mammal Sighting Record, the form provides researchers with 

information not only about the stock, sex, body length, and 

reproductive condition of the sampled specimen, but also about 

the other dolphins associated with it, and where, when, and under 

what conditions they were together. When finally completed at the 

lab, the form virtually becomes the biological specimen that the 

researchers analyze. For that reason, it is imperative that all 

observers complete the form in the same way, with a strict 

understanding of what each item calls for and how to respond to 

its requirements. 

1. Guidelines for Completing the Form 

The form consists of two sections: the in-field section (the 

upper half) and the in-lab section (lower half). You should 

complete the in-field half of the form with a soft-lead pencil to 

record the field data for each specimen processed. Each item of 

the in-field section is followed by a line with non-coded and 
3 

IIllln IIIIIn 

33 

IN2 I I I I I I 

LAB 12 13 20 26 27 TOTAL WEIGHT (gm) L GONAD w/epi (gm) L GONAD W/O epi (gm) 

PORPOISE NOM FORM 88 129 

NMFS 11 82 

CR.# AJ 

I 4 

Ill IIn- 34 40 

R GONAD w/epi (gm) 

IIIlln IIII In 41 47 48 Ln(rnrn) ISG I E 53 54 56 57 

R GONAD w/o epr (gm) ' RIGHTTESTIS ~ EL GLG'S CL 

11111 I I 

58 I 60 I 62 64 66 

CL DIAMS CA(L) CA(R) (mm) 

CARD 

In IIIIIIIII 3 I 11111llll11 I, 

68 FO~L. 71 1 (73 2 175 3 177 4 (79 5 12 13 6 15 1 (17 2 I 19 3 121 4 123 5 125 6 27Tota, 

DIAM. (mm) C.A. IN LEFT OVARY C.A. : IN RIGHT OVARY CORP 

(CA+CLl 

Figure 1. Porpoise Life History Form 

Im 111i11 

"TUBULE 32 33 39 

DIAM (mrn) PREG, FETUS WEIGHT (gm) 
4 

coded divisions (see Fig. 1). Only non-coded responses should 

be made in the field. All coding and completion of the in-lab 

section of the form is done by technicians at the La Jolla 

Laboratory. To ensure that other persons will have no difficulty 

in reading your records, you should print all data and all names 

and words (except compass directions and dates) in unabbreviated 

form. For example, print and spell out llStenella ---_--__ __-______ attenuata1I 

instead of using llS. attenuataV1; spell out llfemalell rather than 

using llfll or the biological symbol for female. 

If you make an entry error, do not erase; line through the 

error with a single line and a d d t h e c o r r e c t d a t a b e s i d e or above 

it. Records of errors are often useful to data editors and 

researchers in rectifying cases of mixed-up specimens or data. 

You should respond to all items in the in-field section of 

the form appropriate to the specimen that you are processing. 

This means, for instance, that if the specimen is a male, 

responses are required for all non-sex dependent items, such as 

llSPECIES/STOCK,ll 'IHEAD,I1 and IITOTAL LENGTH (cm.) ,I1 and for the 

male-dependent item llTESTIS,ll but female-dependent items, such as 

"OVARIES & UTERUS,l1 "LACTATING ?,I1 or llFETUS,ll should be left 

blank. 

The only situation in which you would not respond to all 

appropriate items on the form is when you collect the whole 

carcass. If you collect the carcass because there are more 

carcasses than you can measure and physically sample within the 

time limits set by the skipper, fill in only lines one, three, 

and four of the form, plus the item llCARCASS.ll If, however, you 

collect a carcass because of some special request or standing 

requirement and you have the time, you should measure total 

length, take a jaw sample containing teeth, and fill in the top 

five lines of the form, plus the items llTEETH1l and llCARCASS.ll 

(The tooth sample is taken in the latter case to ensure that the 

specimen will be represented by at least a tooth sample if the 

carcass becomes lost after the contents of the ship's wells are 

unloaded in port.) 

2. Explanation of the Data Items: Definitions and Responses 

The in-field items are explained below in the order that they 

appear on the form. Item titles are listed exactly as they appear 

on the form, from left to right, top to bottom, except for my 

notes enclosed in brackets (see Fig. 1). 

CR.# [cruise number] 

This item asks for the unique cruise number that identifies 

your cruise. A cruise number is assigned sequentially to each 

observer-accompanied cruise by the NMFS San Diego Field Office. 

The cruise number is usually transmitted to the observer by radio 

shortly after the seiner leaves home port because of the 

uncertainty of departure times and the need to preserve the 

chronology of cruise numbers. 
5 

SPECIMEN # [specimen number] 

The specimen number consists of a unique three-letter group, 

followed by a four-digit number. The three-letter group is 

assigned to you permanently by NMFS when your observer training 

is completed. You will assign the four-digit number to a 

specimen, but the number is not a valid specimen number without 

your three-letter prefix. If you are assigned letters ABC and on 

your first cruise you process your first carcass, you will assign 

it specimen number ABC0001. If you process a second specimen on 

your first cruise, you will assign it specimen number ABC0002, 

and so on. On your next cruise in which there is mortality, the 

first specimen that you process will receive ABC and the next 

four-digit number following the number that you assigned last on 

your first cruise. In this way every processed specimen is 

represented by a unique field number, and because letter groups 

are different for each observer, no two specimens should have the 

same specimen number. 

You should assign a specimen number to a specimen after you 

have processed it and have filled in at least the first five 

lines of the form for the specimen. You should also assign a 

specimen number to a carcass that you have collected whole and 

tail-tagged, i.e. secured a cable tie bearing specimen 

identification to the base of the tail (tail-stock) just in front 

of the flukes. In this case, you must also copy onto the upper 

left corner of the form any other identification number on the 

carcass's tail-tag. 

SPECIES/STOCK 

For this item, you should print the full scientific name 

(genus and trivial) and stock name according to the established 

field criteria. If you are uncertain about the stock 

identification, the scientific name should be followed by the 

word Wnidentifiedl' and you should try to take photographs of the 

specimen. Because species and stock identifications are based 

primarily on observations of the school composition, your entry 

for this item should be one of the names that you have provided 

on other reporting documents for the same set on the school. 

SEX 

Print out in full either llmalell orllfemalell in response to 

the implicit question: llWhich sex is this specimen?" Be sure 

that you understand the procedure for determining the sex of a 

specimen. It is described below under "EXTERNAL ANATOMY ITEMS." 
6 

DATE (YR/MO/DAY) 

This item refers to the year, month, and day in which the 

specimen was killed. You should indicate the year with four 

digits, the abbreviated name of the month, and the day of the 

month indicated with two digits. For example, for a specimen 

killed on 2 January 1987, your entry should read: 111987/Jan/02.11 

SET # [set number] 

Each time the net is !!let go," i.e. the skiff with net 

connected is launched from the stern of the seiner, a set is 

considered to have occurred. On every cruise, each set, 

regardless of whether it is a "marine mammal set," is numbered 

sequentially beginning with the number 001. If, for instance, 

your first set was a school-fish set, your second set was a 

marine mammal set that involved no mortality, and your third set 

resulted in 12 specimens for processing, the form for each 

specimen processed should have 1100311 entered for this item. 

POSITION (LAT/LONG) 

For this item, furnish the coordinates, in degrees and 

minutes, of the set in which the specimen being processed was 

taken. For example, if the coordinates of the marine mammal set 

were 14 degrees 30 minutes north, 120 degrees 15 minutes west, 

you would enter 11140 301N/1200 151W11 next to this item. 

QUAD [quadrant] 

This item asks you for the quadrant of the hemisphere in 

which the set was made. For the eastern tropical Pacific, the 

correct response for sets made above the equator would be (IN/W,If 

and for sets below the equator it would be "S/W." 

TOTAL LENGTH (cm.) 

Basic specimen data include a total body length of the 

carcass, rounded to the nearest whole centimeter. For example, if 

the specimen were 138.4 cm long, you would enter 11138;11 if it 

were 159.5 cm long, you would enter 11160.11 See I1Measuring 

Devices1! for instructions on making the body length measurement. 
7 

(SPOTTED ONLY) 

COLORATION 

Among the dolphin stocks involved in the yellowfin tuna 

fishery, only spotted dolphins undergo several distinct changes 

in pigmentation (color phases) as they mature. The stages are 

classified as: 1. neonate (dark cape, white ventrally and 

laterally, with total body length under 100 cm), 2. two-tone 

(similar to neonate, but over 100 cm and grayer ventrally than 

the neonate),3. speckled (spotted ventrally and laterally), 4. 

mottled (ventral and lateral spots touching and merging, dorsal 

spotting), and 5. fused (ventral and lateral fields gray 

monotone, spots dorsally). Use one of these classifications in 

the blank. Complete this item only if you are processing a 

spotted dolphin, _____-___-_______ Stenella attenuata. Leave this item blank for 

any other species. 

LACTATING? 

This is a sex-dependent item and you should complete this 

blank with a "yes" or a %o1I only if the sex of the specimen 

is female. The item asks: "When you checked the mammaries, was 

the female lactating?" The two-step test for lactating is 

described below under EXTERNAL ANATOMY ITEMS. To leave this item 

blank when the specimen is a female means that you did not check 

for lactating. (So, donlt unless you didn't!) 

FETUS: SEX 

Complete this blank by printing the sex of the pregnant 

female's fetus only if the fetus is 25 cm or more in length and 

you discard it. In other cases, no response is required. Fetuses 

less than 25 cm long are not sexed or measured in the field, but 

are collected with the reproductive tract of the pregnant female 

(see note at bottom of in-field section of form, Fig. 1). 

[FETUS] LENGTH (cm.) 

In this blank, you should enter the total body length 

measurement of the pregnant female's fetus, to the nearest whole 

centimeter, only if the fetus is 25 cm or more in length and you 

discard it. In other cases, no response is required. 

COLLECTED? : TEETH 

You should collect eight toten teeth fromthe middle ofthe 

tooth row of the left mandible (i.e. the left lower jaw). Answer 

"yes" if you collect the teeth; otherwise answer lcno.ll A jaw 

sample containing teeth should be collected separately when the 

head is collected or when a carcass is collected because of 

special or standing instructions. 
8 

[collected?: 3 TESTIS 

A response to this item is required only if the specimen 

that you are processing is male and you do not collect the 

carcass. If the specimen is male, the proper response is llyesll if 

you collect the right testis or llnoll if you do not. 

[collected?:] OVARIES & UTERUS 

This item requires a I1yes1l or llnoll response only if the sex 

of the specimen is female and the carcass is not collected. A 

Ilyesll response indicates that you collected the entire 

reproductive tract. 

[collected?:] FETUS 

This item asks: "Did you collect a fetus?ll You should 

respond only if the adult specimen is female and after you 

inspect the uterus from the outside to try to ascertain the 

presence or absence of a fetus. A llyesll to this item indicates 

that you observed (through the uterine wall) or felt what you 

interpreted to be a fetus, that it was less than 25 cm long, and 

that you collected it along with the reproductive tract. A I1nog1 

response indicates that you did not collect a fetus although it 

does not necessarily mean that no fetus was present. (Current 

requirements instruct that all fetuses 25 cm or more in length 

should be sexed, measured, and discarded.) A third response to 

this question may be a question mark. A question mark indicates 

that you checked, but that you were not able to confirm the 

presence or absence of a fetus. Caution: Do not open the uterus 

to look for a fetus unless you are certain that it is 25 cm or 

more in length. A small embryo may be lost through the opening 

that you make. An unpunctured uterus ensures that its contents 

are collected intact. 

[collected:?] STOMACH 

This item requires a I1yesl1 or llnoll answer to the question: 

"Did you collect the stomach from this carcass?11 Do not leave 

this item blank unless you collect the carcass. 

[collected:?] HEAD 

If the carcass is not collected, this item requires a llyesll 

or llnoll answer to the question: "Did you collect the head?I1 

Leave the item blank only if you collect the carcass. If you 

collect the head, you should collect a jaw sample separately. 
9 

[collected:?] CARCASS 

If you collect the carcass, you should answer this item with 

a llyes;ll otherwise answer with a llno.lt Do not leave this item 

blank. If you collect the carcass for purposes other than to save 

time, you should collect a jaw sample separately. 

[collected?: 3 MAMMARY GLAND 

If the specimen is female, this item should have either a 

llyesll or a llnotl response depending on whether you collect a 

tissue sample from the mammaries. No response is required if the 

specimen is male, or if you collect the carcass. 

[collected?: 3 MILK 

If the specimen is notalactating female, or if youcollect 

the carcass, you need not respond to this item. Otherwise, a 

Ityes1l or %otl answer to the question: "Did you take a milk sample 

from the mammaries?11 is necessary. 

[collected?:] PARASITES 

This item should not be left blank unless you collect the 

carcass. You should provide a I1yesO or IInoII answer to the 

question: @!Did you collect any parasite from any part of the 

carcass?lI 

[collected?: 3 BLOOD 

This item asks: "Did you take a blood sample?I1 You should 

respond with either a I1yes1I or a llno.ll Do not leave this item 

blank unless the carcass is collected and blood is not collected 

from it separately. 

PHOTOS? ROLL # FRAME(S) # 

These three items are concerned with the question: "Did you 

take photographs of this specimen and, if so, how many exposures 

were taken and what was the number used to identify the roll of 

film?t1 The item tlPHOTOS1l must be answered with a I1yesl1 or a 

llno.ll If your response to PHOTOS is I1no,l1 do not respond to the 

following two items. If it is llyes,ll then you should furnish the 

roll identification and the numbers of frames exposed. No 

response to these items should be made if you collect the 

carcass. 
10 

COLOR PATTERN [sketch] & DORSAL FIN [trace]: 

To decrease processing time per specimen and maximize 

sample volume, sketches are not required for spotted dolphins and 

only the outline of the ventral hump, dorsal fin, and color 

pattern is required for spinner dolphins. If you are working up 

a spinner dolphin, your instructions may also require you to 

trace the dorsal fin on the back of the form (see EXTERNAL 

ANATOMY ITEMS for procedure). 

If you have plenty of time, you may use the sketch outline 

to illustrate features of interest, such as scarring, unusual 

pigmentation patterns, or points of injury. Although not a 

required part of processing, the use of the sketch in this way 

frequently has proven useful to researchers in the past. If you 

collect the carcass, do not respond to this item. 

BASIS FOR STOCK ID: 

This item asks you for the criteria that you used to 

determine the stock to which the carcass being processed belongs. 

The criteria used should emphasize the general appearance of the 

adults in the school from which the carcass came. In cases where 

it is obvious that a school consists of animals of a single 

stock, identification is usually straightforward. However, in 

cases where it appears that more than one form of the same 

species is represented, you should include in your evaluations 

such criteria as homogeneity or heterogeneity of behavior between 

groups in the encircled school. 

In any case, the carcass that you are processing should 

belong to one of the stocks that you identified from observations 

of the school. You should make clear whether you have identified 

the carcass strictly according to your observations of the 

school, without considering the characteristics exhibited by the 

carcass. If you collect the carcass and do not identify the 

specimen, leave this space blank. 

Measuring Devices 

1. Sticks 

Two, two-meter measuring sticks and one pair of moveable 

caliper jaws (Fig. 2) may be included in your equipment package 

for measuring total body lengths of carcasses and for providing 

scale in photographs of single carcasses. 

Care 

The sticks are constructed of wood and vulnerable to 

breakage and warpage. You should use special care in handling and 
11 

-BB 

Figure 2. A. Measuring sticks and moveable caliper jaws 

positioned for calibrating calipers. Arrow at A indicates where 

the measurement should be read. Arrows from B indicate position 

of setscrews for locking or adjusting the jaws. B. Taking a 

dolphin's body length measurement with a measuring stick. One 

caliper jaw is in the tail notch and the other one is at the tip 

of the upper jaw. 
12 

storing them. To minimize warpage, keep them dry and lubricated 

with s i . l i c o n e s p r a y (included in your equipment package). And, 

whether at the workup site or the storage area, lay (or store) 

them flat when they are not in use. 

Metric Measurement 

The sticks are marked in inches on one side and in 

centimeters on the reverse side. Measurements must be made in 

centimeters, so it is important to cultivate the habit of 

consciously checking which side of the stick you are reading. On 

the metric side of most sticks, the numbers go up to 100 cm and 

begin over again at one. Thus, the number at 110 cm is only lglO.gl 

Failure to note that the first 100 centimeters has been traversed 

during carcass measurements could lead to records of remarkably 

short-bodied animals. 

Calibrating the Caliper Jaws Between Sets 

Both caliper jaws are moveable and the one nearest the zero 

end of the stick that remains stationary during measurements, 

(the "stationary" jaw) may come out of calibration because of a 

loose setscrew, a sudden jolt in transit between decks, or during 

storage, simply through vibrations from the boat. To ensure 

that measurements with the stick are accurate, you should 

recalibrate the stationary jaw at the workup site before 

processing specimens from a set. 

Begin calibration by placing the stick without jaws (the 

first stick) on the deck, metric side up. Place the stick to be 

calibrated (the second stick) parallel to the first stick so that 

the zero ends of both sticks are in the same direction and both 

caliper jaws of the second stick extend at a right angle across 

the metric scale of the first stick (Fig. 2A). While keeping the 

two sticks parallel, slide the non-stationary caliper jaw (i.e. 

the jaw farthest from the zero end of the stick) to the 20-cm 

mark of the first stick and to the 10-cm mark of the second 

stick. The first stick should now extend 10 cm in the zero-end 

direction beyond the second stick. Tighten the setscrew of the 

non-stationary jaw. Now, move the stationary jaw to the 10-cm 

mark of the first stick and tighten its setscrew. Calibration of 

the second stick should be complete and the stick ready for use. 

As a test, loosen the setscrew on the non-stationary jaw, 

slide the jaw along the stick to its 35-cm mark, and tighten the 

setscrew. Place the two jaws across the metric scale of the first 

stick. If the stationary jaw of the second stick is calibrated 

correctly, the distance showing between the two jaws on the first 

stick should be 35 cm. 
13 

Taking Body-length Measurements 

The total body length measurement may be taken with the 

carcass resting slightly on its right side on the deck. The long 

axis of the body should be straight. The lower yaw usually 

extends farther forward than the tip of the upper jaw, but 

despite this common llunderbite,ll standard total body length 

measurement is made from the tip of the upper jaw to the caudal 

notch of the flukes at midline. 

To measure the carcass with the stick, place the stick above 

the midline of the long axis of the body, parallel to the deck, 

with the zero-end pointing posteriorly. Loosen the setscrew of 

the non-stationary jaw, place the stationary jaw into the notch 

between the tail flukes, and slide the non-stationary jaw along 

the stick to the anterior tip of the upper jaw of the specimen 

(Fig. 2B). After checking to make sure that the stick is parallel 

to the deck and that the caliper jaws are not closed toward each 

other so tightly that the body is being bent, tighten the 

setscrew of the non-stationary jaw and remove the stick from the 

carcass. Read the measurement from the metric side of the stick, 

round off to the nearest whole centimeter, and record the number 

next to TOTAL LENGTH (cm.) on the form. 

When you are measuring a carcass close to or more than two 

meters in length, which is not uncommon, begin by following the 

same procedure as described above. The non-stationary jaw will 

not reach to the tip of the upper jaw, so move it to a prominent 

landmark, such as the anterior margin of the melon. Be sure that 

the measurement is as square as possible; then tighten the 

setscrew, turn the stick to the metric side, take the reading 

without rounding off, and write the number down. Now, move the 

stationary caliper jaw to the point where your first measurement 

ended and slide the non-stationary jaw out to a point even with 

the tip of the upper jaw of the specimen. Tighten the setscrew, 

take the reading without rounding off, and add the first reading 

to the second. Now, round off to the nearest whole centimeter and 

enter this number on the TOTAL LENGTH (cm.) blank. 

2. Other Devices 

You may be issued equipment other than measuring sticks 

with which to take length measurements. Flexible tapes with 

various accessories that permit square measurements have been 

used. They have several advantages over sticks: 1) they are 

compact and easily transported, 2) they are less vulnerable to 

damage, and 3) they do not require recalibrating between sets. 

Their main disadvantage is that they are flexible and may 

consistently sag a bit during measuring. This requires the 

measurerer to take special care that the tape is taut and 

parallel to the deck. Because of the problem of tape sag, it is 

best to use a rigid measuring stick whenever possible. 
14 

Labels, Cable Ties, and Labeling 

Among your supplies will be several hundred labels (tags), 

approximately 6 x 12 cm, with metal eyelets for accommodating 

ties which are used to attach the labels to samples. The labels 

are specially constructed to resist tearing, deterioration, and 

dissolution. Each may already have a string in the eyelet, but 

because string adds knot-tying to the problem of attaching a 

label, rib-lock cable ties, also found in your equipment package, 

are now used. If you remove all of the strings before you reach 

the fishing grounds, you will have saved time by not having to 

unstring labels during specimen workups. 

Cable ties will be in two lengths: - 15 and 60 cm, the 

shorter for attaching labels to organ and tissue samples, the 

longer (pre-embossed in packs of 25 or 50) for tail-tagging 

carcasses. To attach a cable-tie label to a specimen, thread the 

tie through the eyelet of the label, encircle the organ sample 

with the tie, then pass the end of the tie through the tie 

sleeve and cinch it down tightly on the sample to lock it. To 

ensure that the correct side of the tie is against the sleeve, 

give a few tugs on the tie to try to back it out. If it backs 

out, thread the tie back through the sleeve using the opposite 

side of the tie against the sleeve to lock it down. Test again 

for locking. 

Print the specimen number on the label in soft-lead pencil. 

No other data should appear on the label and no writing implement 

other than a soft-lead pencil should be used. Finally, all of the 

labels needed for collecting samples from a specimen may be made 

up at the same time, but never make up labels for more than one 

specimen at a time. Prenumbering labels greatly increases the 

chance of data mix-ups. 

Gloves 

Your equipment includes one pair of heavy cotton gloves, 

several pairs of Playtex-type rubber gloves, and up to a halfdozen pairs of surgeons disposable latex gloves (optional). 

Always wear gloves to protect you hands from minor injuries while 

you are handling carcasses and to prevent contact with 

potentially harmful microorganisms or fixative. The cotton gloves 

are intended for use in moving carcasses. The rubber gloves are 

for use during specimen processing and handling of fixative or 

fixed samples. If the rubber gloves are used for carrying or 

dragging carcasses, they will puncture or tear. You will have no 

protection for your hands during specimen workups if all of the 

rubber gloves have been damaged through misuse, because cotton 

gloves are porous and too cumbersome for removing most anatomical 

samples. 
15 

Containers, Preservative, and Specimen Handling 

1. Buckets and Bags 

You will be taking aboard up to three plastic five-gallon 

buckets with watertight lids, small and large plastic bags and 

burlap bags. Buckets are for: 1) receiving samples being 

collected on the working deck, 2) formalin-fixation of samples, 

and 3) specimen storage and boat-to-lab transport of 

consolidated preserved samples. Plastic and burlap bags are for 

packaging specimens that require frozen storage. 

2. Fixative: Preparation and Cautions 

A one-gallon plastic bottle containing 100% formalin (a 

saturated solution of 37% formaldehyde) is part of your regular 

issue. A 10% formalin solution is needed for optimal fixation of 

tissues (a process that may take about one week depending on 

tissue thickness and permeability). Concentrations greater than 

about 13 or 14 % usually result in formalin ffburnff (actually a 

bleaching/hardening process), so be sure to follow the recipe. 

To prepare the fixative, mix one part (100%) formalin to 

nine parts seawater from the deck hose. (For the sake of 

maintaining congenial relations with the crew, it is best not to 

use the boat's limited freshwater supply for making fixative or 

for cleaning tools or specimens.) Prepare enough 10%-strength 

solution to fill one bucket about half full. Keep this bucket of 

fixative in the storage area for receipt and complete fixation of 

the freshly collected samples. (One of the best places for 

storage of specimens, data forms, and collecting equipment is 

near the bow on the wet deck because it is out of the way of 

heavy traffic, and despite its name, the wet deck is usually 

dry- 1 

Avoid skin contact with the fixative or prolonged inhalation 

of its fumes. Besides being an irritant to skin, eyes, and nasal 

membranes, formaldehyde is generally considered to be a potential 

occupational carcinogen that is known to cause certain kinds of 

cancer in laboratory animals. Remember to wear rubber gloves 

when preparing fixative, or when transferring fixative or fixed 

specimens from one container to another. Handle the fixative only 

in a well-ventilated area. If you can smell the formaldehyde, you 

may be inhaling enough to do you harm. 

3. Handling and Preservation of Samples: Recommended Routine 

You will be preserving samples in two ways: frozen storage 

and fixed-wet storage. Carcasses, heads, and stomachs should be 

frozen after being collected: carcasses and heads, because they 

are too large for buckets and they cannot be fixed on board, and 

stomachs, because formalin is acidic and will etch or dissolve 

fish otoliths and squid statocysts that are contained in the 
16 

stomachs. Frozen specimens smaller than heads are normally 

stored in the cook's food freezer. You must obtain permission 

from the ship's cook to use freezer space for storage of samples. 

Once permission is obtained, try to make any samples stored there 

as presentable as possible. Carcasses and heads are stored in the 

wells (where the tuna are stored) with the permission of the 

skipper or chief engineer. The heads are generally tied to the 

coamings of the wells. To remind yourself where you have stored 

your frozen samples, keep a running inventory of them in your 

green book. 

When you collect a head, put a label in the mouth and cinch 

the jaws tightly shut with a cable tie or strong cord (a length 

of seine-net twine will do). Make a slit through the skin and 

muscle near the base ofthe head and pass a cable tie throughthe 

slit. Attach a label to the tie and cinch it securely. Place the 

head in a large plastic bag and tie it shut with twine to prevent 

leakage of blood. Finally, place the package into a burlap bag to 

protect it and use a cable tie (with an outside label to make it 

identifiable) to close the bag. 

Before removing the stomach from a carcass, seal off each 

end of the stomach with a 15-cm cable tie (one with a label) to 

avoid spilling its contents. After removing the stomach, place it 

in a small plastic bag and close the bag with a cable tie and 

outside identifying label. 

Jaw samples with teeth and testis or uterus and ovaries 

samples are preserved in formalin. Each of these samples should 

be labeled once with a cable-tied label and placed in your 

sampling bucket without additional packaging. 

If a kill set includes carcasses from which stomachs, heads, 

or other samples needing frozen storage are required, take your 

sampling equipment, a supply of bags, and an empty bucket 

(sampling bucket) up to the working deck with you. (If no frozenstored samples are to be collected, do not take bags.) Fill the 

sampling bucket half full of seawater from the deck hose and use 

the sampling bucket to store tooth and gonad samples until 

specimen processing is completed. To keep your forms clean and 

to minimize slippage of hand tools during sampling, use seawater 

fromthedeckhosetowashyourhands inbetween workups anddata 

recording. 

After you have processed the specimens from a set, take the 

samples packaged for freezing to the freezer or to the well. Then 

take your sampling bucket down to the specimen storage area and 

transfer the samples to the fixation bucket. Keep the lid tight 

on the fixation bucket to prevent evaporation of fixative and 

leakage of its fumes. You should always keep enough fixative in 

the bucket to cover all specimens and the specimens should remain 

for at least a week in the fixative. After fixation is complete, 

transfer the specimens to the storage bucket to make the fixation 

bucket available for new samples. The storage bucket should be 

lined with doubled plastic bags with only a little preservative 
17 

to keep specimens moist. Intermittent monitoring and maintenance 

of the fluid level in the storage bucket will ensure that the 

samples do not dry out. If the take becomes high and sets with 

large kills become frequent during a cruise, it may be necessary 

to add another fixation bucket to your system. You may substitute 

a large plastic bag for your sampling bucket and use the sampling 

bucket for fixation. 

At the end of the cruise, close up'the double bags of 

consolidated specimens in the storage buckets with cable ties and 

close the lids tightly. Cable-tie a label onto the handle of 

each bucket with your last name and the cruise number printed on 

it in soft-lead pencil. As an extra precaution against 

misplacement of the buckets, use one of the waterproof markers, 

included in your equipment package, to print your last name and 

the cruise number on the lid and on the side of each storage 

bucket. 

You will find it convenient during a cruise to keep all 

(separately packaged) stomachs together in the freezer in a large 

plastic or burlap bag. As you collect more stomach samples, just 

add them to your stomach bag. At the end of the cruise, cabletie a label to it with your last name and cruise number printed 

in waterproof marker. Make sure that there are outside labels 

with the same information for any heads that you have collected. 

Pruning Shears 

One of your collecting tools will be a pair of pruning 

shears for use in collecting jaw samples containing teeth. The 

blades of the shears are subject to corrosion from salt water and 

the edges tend to dull from shearing bone and connective tissue. 

Dull and rusted shears slow the jaw sampling procedure and 

prolong the overall processing time. After processing specimens, 

clean the blades of organic materials, dry the blades, and spray 

them with silicon. If they become heavily rusted, ask permission 

of the chief engineer or others in charge to use the wire wheel 

on the bench grinder to burnish the blades. (Use eye protection 

while using the bench grinder.) The shears may be sharpened with 

the whetstone, included in your pack, or with the ship's bench 

grinder or any flat metal file (be sure to ask permission to use 

the grinder or a file). 

Knives 

Two boning knives are your standard-issue dissection tools. 

The knives should be kept sharp if they are to be used 

effectively. This means that practice with the whetstone 

(included) and oil is important. A crew member may sometimes 

offer to show the beginner the proper way to set a knife edge. 

Take him up on it! A dull-edged knife can be extremely dangerous 

because too much uncontrolled pressure must be applied to cut 

with it. In slicing a soft tomato with a dull paring knife, it is 
18 

easier to cut fingers than to cut tomato. In processing 

specimens, if you apply pressure to a dull-edged knife against a 

carcass, and the carcass, your hand, or your knife slips, you 

maybe seriously cut. A knife woundmay require the services of a 

physician, 

Of course, even when they are sharp, knives should be used 

with care. During processing, you should always embed them in the 

heavy musculature of the carcass when they are not being used. 

However, do not forget to remove the knives from the "knife 

holder11 before it is discarded or you may run out of knives long 

before you run out of carcasses. Do not hold the knife while 

trying to adjust the position of a carcass. A sudden slip may 

cause an injury. Hold the knife only when you intend to cut with 

it. When you cut, you should always keep the point and sharp 

edge of the blade turned away from yourself. 

which are usually in short supply on tuna seiners. 

COLLECTION EFFORT AND PROCESSING DECISIONS 

Processing or collecting all specimens brought on board 

after each set should be high on your list of priorities. Based 

on recent estimated kill rates, one hundred percent of the take 

must be sampled to obtain an adequate annual sample for 

monitoring the affected dolphin stocks. To support this effort, 

NMFS has offered to reimburse U.S. boats at tuna prices for well 

space to store dolphin carcasses that cannot be processed by the 

on-board observer. 

For sets in which fewer than 15 or so animals are taken, 

complete workup on all carcasses is usually not a problem as long 

as there is no impending set. Processing of so few carcasses 

should require two to three hours of work. With larger kills, 

however, certain factors such as time of day, fishing success, 

and weather must be considered in deciding how many specimens you 

have time to work up and how many you should tag and send to the 

wells for processing on shore after the cruise. You must decide 

this while the chute to the wells is still set up. If, after the 

chute is dismantled, you have retained more carcasses on deck 

than can be worked up before a new set is initiated, you will 

have no choice but to dump some overboard to clear the deck. On 

the other hand, if you hedge your bet by always retaining on deck 

an exceedingly small number of carcasses to work up and by 

sending all of the other specimens into the wells unprocessed, 

the costs of well-space rental for unprocessed specimens could 

rise prohibitively. 

If the large kill set is made near the end of the day, it 

would be reasonable to expect no additional sets for the day and 

you can usually be certain about this by observing the crew's 

non-searching activity or by asking the skipper. In that case, 

you would retain on deck a maximum number of specimens that you 

could process (up to 30) and send the others down the chute. 
19 

If, after a kill set, the skipper gives you a time limit for 

clearing away specimens from the deck (for whatever reason), you 

should try to collect or process as many specimens as possible. 

If the chute is still assembled, you might opt to tail-tag and 

send all of the carcasses that you have time for down the chute. 

But, if the chute is dismantled, you should work up as many 

specimens as time permits. 

PARTIAL-SAMPLE SELECTION 

In cases where only part of the specimens brought aboard 

from a set can be collected or sampled, carcasses should be 

selected for processing arbitrarily, as they come onto the deck, 

or as the crew has stacked them for you. The purpose of choosing 

specimens randomly is to try to minimize any age or sex bias in 

the sample caused, for example, by selecting only the smallest, 

or the largest, etc. Analyses of samples at the laboratory will 

include an attempt to reconstruct the size and age distributions 

as well as the sex ratios of schools that were set upon. By using 

hundreds of school samples in their analyses, researchers can 

obtain an age and size distribution and sex ratio of the sampled 

population. With this information and other age and reproductive 

data, predictions canbe madeaboutthe futuregrowth rate ofthe 

population from which the samples came. 

If, in collecting specimens representing only part of the 

kill, you choose mainly larger animals, a bias toward adults 

would exist in your sample. Because males usually grow larger 

than females, the bias might also be toward males. Such samples 

not accompanied by a description of the sampling procedure used 

by the observer might mislead scientists conducting the analyses 

into concluding that the schools were made up chiefly of adult 

males, with few juveniles. 

COLLECTING CARCASSES FOR SHORE-BASED PROCESSING 

The large (-60 cm) cable ties are used for rapidly tagging 

carcasses that go into the wells for processing ashore. They are 

designed primarily for situations in which there are more 

carcasses than you have time to process. You should put aside 

the specimens you have time to process and then send the 

remainder tothe wells. The large cableties come inpacks of 25 

or 50 and each tie has a pre-embossed letter/number code, 

numbered sequentially. Before passing the first carcass down the 

chute, detach the first cable tie from the pack and secure it, 

without a label, around the tail stock. Tail-tag the next 

specimen using the next cable tie of the pack, and so on. Then 

process your on-deck specimens. 
20 

After all ofthe carcasses from a setareeitherprocessedor 

tail-tagged and in the wells, prepare a separate Porpoise Life 

History Form for every cable-tie number used. Record a cable-tie 

number in the upper left-hand corner of each form and then enter 

the set number, the date, position, enter llyesll for CARCASS, and 

assign and enter a specimen number (three-letter, four-number 

code). No other data should be entered. The remaining blanks on 

the form are filled in when the tail-tagged carcass goes through 

the complete workup at the laboratory. For purposes of good 

bookkeeping and to help keep track of carcasses from each set, 

enter the beginning and ending cable-tie numbers used, by set, in 

your greenbook. Also, to facilitate recovery of carcasses when 

the wells are unloaded, you should include beside each entry, 

thenumber ofthe well that contain sthe carcasses from the set. 

SPECIMEN PROCESSING ON BOARD 

It is a good idea to establish a uniform routine for onboard processing of specimens early in your tour. If all 

specimens are aligned the same way and all are sampled and 

repositioned always in the same sequence, fewer errors and 

omissions will be made, more specimens can be processed, and 

workup time per specimen can be appreciably shortened. 

The order in which the following procedures are described 

need not be the order of the routine you adopt. Try out several 

systems and decide what order is most convenient and efficient 

for you. Then keep to that order. Periodically, you should 

review the sample needs listed in the llRegional Observer Life 

History Requirements" chart, as well as the procedures described 

in this handbook. By keeping your system current, you can ensure 

that your methodology does not drift from the requirements to the 

extent that the collection of important data is systematically 

omitted. After you have processed and discarded 123 females and 

you suddenly realize that you forgot to determine whether they 

were lactating, it is impossible to retrieve the information. 

I recommend a processing order that progresses through the 

items on the form by operational blocks: 

1) non-dolphin items (data that can be answered 

immediately without a specimen), 

2) external anatomy items (data that can be collected 

without opening the body cavities), and 

3) internal anatomy items (data that must be gathered 

from the cavities). 

This or a similar order should be adopted because it is simpler, 

smoother, and faster than following the order of items as they 

appear on the form. It allows you to respond to operationally 

related requirements all at once, without having to change 

sampling modes repeatedly. 
21 

GETTING READY 

When you go out on deck with your equipment to work up 

specimens after a kill set, lineup as many carcasses as youhave 

time to do side-by-side and position them slightly on their right 

sides, body axes straight, heads pointed to your left as you view 

their left sides (Fig. 3). You should note that this relative 

position (sketch position) is similar to the sketch outline on 

the form (Fig. 1). (Remember that if for some reason you can 

sample or collect only some of the carcasses, select them 

arbitrarily.) After aligning your specimens, immediately prepare 

yourequipment for work up of the first specimen in line. This 

includes calibrating your stick on site, filling your bucket with 

seawater, and inventorying your equipment. 

NON-DOLPHIN ITEMS 

Take a sharpened soft-lead pencil and enter all of the "nondolphin" data on a new form: 

1. cruise number 

2. date 

3. set 

4. position 

5. quad. 

EXTERNAL ANATOMY ITEMS 

Then, examine the first specimen. With the carcass in sketch 

position you are ready to respond to the first half of the 

external anatomy items block: 

6. species/stock 

7. basis for stock ID 

8. (if spotted dolphin) coloration 

9. (if spinner dolphin) color pattern ...( sketch) 

10. photos? 

11. total length 

Coloration/Sketch/Fin Trace/Photos 

If you have identified the specimen as a spotted dolphin, 

(Stenella attenuata), you should now respond to the ltCOLORATION1f 

item by entering the color phase of the specimen. If the 

specimen is a spinner dolphin (Stenella longirostris), you should 

indicate, by lines on the dolphin outline, the shapes of the 

ventral hump, dorsal fin, and basic pigmentation pattern (Fig. 

4A). 
22 

. 

h 

a, 

c 

a, 

a, 

LI 

CJ 

b 

. 

c 

a, 

Y 

Q 

4J 

0 

4J 

0 

c 

a 

tn 

0 

a 

c 

a 

d 

0 

a 

u-l 

0 

. 

m 

a, 

LI 
23 

Figure 4. Sketch and fin-trace requirements. A. Outlining the 

pigmentation pattern on a Porpoise Life History Form to fulfill 

the sketch requirement for spinner dolphins. B. Using the blank 

side of the form to trace the fin of a spinner dolphin. 
24 

When you are working up a spinner dolphin and requirements 

include a fin trace, turn the form over to the blank side, place 

that side behind the dorsal fin, and trace the outline of the fin 

on the paper with your pencil (Fig. 4B). Include a bit of the 

margin of the body outline in front and behind the fin so the 

angle of the fin on the body is evident. If there is not room 

enough to include the whole fin outline, try to position the 

paper diagonally. If results are still poor, use backs of two 

forms placed together (if you do this, make sure that the same 

specimen number is on both forms). To distinguish anterior from 

posterior, draw an arrow pointing forward from the anterior 

margin. Fin traces are used to measure variation in dorsal fin 

angle among spinner dolphin populations. 

If you have some doubt as to the stock to which the specimen 

belongs, or if the life history requirements so indicate, you 

should take photographs and enter the photo and film data in the 

appropriate blanks. Before photographing a specimen, place some 

sort of scale with it (a measuring stick works well). Finally, 

be sure that only one specimen is photographed on deck at a time 

and that the photographing has been approved of in advance by the 

skipper. 

Total Length 

Check that the body axis is straight. Now, take the total 

length measurement. (If you are using a measuring stick, do not 

forget to read from the metric side and from the inside of the 

caliper jaw slide (Fig. 2A)). Round off to the nearest whole 

centimeter and enter the number. 

By rolling the carcass fully onto its right side to expose 

the venter, you are ready to respond to the rest of the block of 

external anatomy items: 

12. 

13. 

14. 

15. 

16. 

17. 

18. 

19. 

sex 

specimen number 

(if female) lactating 

(if lactating) milk 

(if female) mammary gland 

teeth 

head 

carcass 

Sex 
25 

Sex 

Examine the body along the mid-ventral line, posterior to the 

level of the dorsal fin. Three criteria should be used together 

to determine sex from the external anatomy. If the specimen is a 

female, the genital and anal slits--the two openings on the midline of the venter--should be almost continuous (Fig. 5). In 

males, there is a considerable hiatus between the genital slit 

and the more posterior anal slit (Fig. 6). A female has a 

mammary slit on either side of the genital slit, but in males 

mammary slits seldom develop. When they do occur in males, they 

are usually found on either side of the mid-ventral line between 

the genital and anal slits. Finally, to ensure correct sex 

determination, insert a small blunt-ended probe, such as the 

eraser end of a pencil, eraser first, into the genital opening of 

a specimen. Try to push the pencil gently forward, then backward, 

at a slight upward angle. You should find that if the pencil goes 

easily forward, you cannot push it backward. If the pencil goes 

forward it enters the vagina which is angled forward and slightly 

upward and the specimen is female. In males, the pencil can be 

moved backward into the chamber that houses the penis, but not 

forward (Fig. 7). 

Specimen Number 

Now that you have recorded all of the minimum data necessary 

for assigning a specimen number, you should print your threeletter prefix followed by the next succeeding four-digit number 

beside this item on the form. This should be the same as the 

number on the labels that you are about to make up in advance for 

the specimen. 

Advance Preparation of Labels 

Having determined the species and stock of the specimen, you 

are ready to check the I1Regional Observer Porpoise Life History 

RequirementsI1 to find what samples are to be collected (Fig. 8). 

Knowing this will permit you to save time by preparing labels 

(printing specimen number) for all of the samples from the 

specimen inone operation. But donot number labels formore than 

one specimen at a time. Prepare labels for the first specimen, 

work it up completely, then prepare labels for the next specimen, 

and so on. 

For standard workups (in which just the jaw, with teeth, and 

the gonads are sampled) only two labels are needed. If you 

collect the stomach, you need two more labels. If the head is 

collected, you should make up three labels for it. (But 

remember, if the head is taken, you still must collect a jaw 

sample separately.) Thus, for a specimen from which the jaw 

sample, reproductive tract, stomach, and head are collected, you 

should prepare seven labels (see Table 1). 
26 

Figure 5. Ventral surface of female dolphin showing genital slit, 

anal slit, and paired mammary slits. The genital and anal slits 

are almost continuous in female dolphins (photo courtesy of W. 

Perrin) . 
27 

Umbilical scar 

Genital 

Ana 
28 
29 

PLH-3 / 85 -1 3 

REGIONAL OBSERVER LIFE HISTORY REQUIREMENTS 

OFFSHORE COASTAL EASTERN AND COSTA STREAKERS DELPHINUS ALL 

SPOTTER SPOTTER WHITEB ELLY RICAN OTHER 

SPINNER SPINNER 

TOTAL 

LENGTH X X X X X X X 

COLOR 

PATTERN 5 5 5 5 

DORSAL FIN 

TRACE X X X X 

PHOTOS X 

~~ 

X X 

TEETH X X X X X X X 

HEAD X X 1 1 X 

REPRODUCT ION 

DATA X X X X X X X 

REPRODUCTIVE 

TRACT 2 X 2 X X X X 

STOMACH X X 3 3 X 

LN ER 4 4 

X=COLLECT/RECORD 

1-COLLECP HEADS OF ADULTS FROM BELCW EQUATOR OR OUTSIDE CYRA 

2=COLLECT TESTIS/OVARIES AND UTERUS IF 150CM OR LONGER. DISCARD FETUSES NOT LEFT IN UTERUS (AFTER 

3=COLLECT STOMACH FROM ALL AGES FROM BELCW EQUATOR OR OUTSIDE CYRA 

4=COLLECT 200GMS AND FREEZE AS SOON AFTER DEATH AS POSSIBLE 

SrRECORD COLOR PATTERN OF SPOTTERS1 SKETCH COLOR PATTERN OF SPINNERS 

PEASURING AND SEXING) 

Figure 8. Regional Observer Life History Requirements Chart 
30 

Table 1. Labeling, packaging, and storage requirements for some 

dolphin organ and tissue samples. 

No. of 

Sample labels 

Needed 

Testis or 

Reproductive 

Tract 1 

Jaw sample 

with Teeth 1 

Head 3 

Stomach 2 

Label Type Type Placement Packaging Storage 

organ none fixed wet 

(bucket) 

jaw sample none fixed wet 

(bucket) 

a. mouth frozen 

b. back of head plastic bag (wells) 

c. outside bag inside burlap 

bag 

a. organ plastic bag frozen 

b. outside bag (freezer) 
31 

Lactating/Milk/Mammary Gland 

1. Sampling Procedure (test for lactating) 

The test to determine whether a female is lactating is in 

two steps. Locate the mammary slits on each side of the genital 

slit. Grasp each mammary, one at a time, between the thumb and 

forefinger and pinch firmly several times (Fig. 9A). If the 

pinch test fails to produce positive results, slice each mammary 

through the center to a depth of about two cm (Fig. 9B) and pinch 

again several times. 

If no whitish fluid is exuded, enter IlnolI on the life 

history form in response to lactating. If fluid is produced by 

the test, inspect the incised area for cestode parasites. 

Cestodes sometimes encyst in the genital areas, encasing 

themselves in a membraneous sac filled with a yellowish-white 

fluid that can be mistaken for milk. If you find no cestodes or 

other similar parasites and you suspect the fluid to be milk, 

enter lfyestl for lactating. A blank means that you did not check 

for lactating, so do not leave this item blank unless you did not 

perform the test. 

2. Utility 

The presence of milk indicates either that the female is in 

the late stages of pregnancy or that she has been nursing a calf. 

If a non-pregnant lactating female is taken, the dependent calf 

probably would not have survived even if it hadnot been taken in 

the set. This information is important to biological researchers 

because it can be used in estimates of calf mortality. 

Presently, there are no requirements for collecting milk and 

mammary-gland tissue. Procedural instructions will be issued if 

such sampling becomes necessary. 

Teeth 

1. Sampling Procedure 

A sample from the middle of the left lower jaw containing 

between eight and ten teeth should be collected except when you 

collect the carcass in lieu of on-board processing. When the 

carcass is on its right side, the left mandible is fully exposed 

for examining and sampling. Open the mouth slightly and inspect 

the left tooth row. Locate the middle of the dental series. Make 

a 15-cm-long slice from below along the inside of the middle of 

the left mandible, so that the point of your knife protrudes into 

the mouth medial to the tooth row (Fig. 10A). Insert one of the 

blades of the pruning shears into the slit; then sever the jaw 

transversely four o r f i v e t e e t h in front of the middle tooth of 

the dental row (Fig. 10B). With another snip of the shears, 

sever the jaw behind the first truncation so that the jaw sample 
32 

A 

Figure 9. Two-step check of female dolphin for lactation. A. Each 

mammary is pinched several times. B. If no milk shows, each 

mammary is cut transversely to a depth of about two cm and 

pinched again. 
33 

A 

B 

C 

Figure 10. Collecting a jaw sample of teeth. A. A slit is made 

along the inside of the left jawbone. B. One blade of the shears 

is inserted into the slit and the jaw is severed transversely. C. 

The jaw is sheared again about eight to ten middle-teeth away 

from first truncation. 
34 

contains the required complement of undamaged teeth (Fig. 1OC). 

The sample is now easily removed by using your knife to cut away 

remaining soft tissue connections. Label the jaw section once, 

record utyestl next to TEETH on the form, and place the sample into 

your sampling bucket. 

2. utility 

After they reach the laboratory, the teeth are treated 

chemically, cut into thin sections, and read with a microscope to 

estimate the specimen's age. Ages from hundreds--even thousands-- 

of specimens are analyzed together to try to determine whether 

the age structure or age-specific reproductive rates are changing 

within a given population. 

The jaw sample is taken from the middle series of the 

mandible because that is where the clearest tooth layers form. 

The teeth are sampled from the left mandible for uniformity. 

Eight to ten teeth are needed because sometimes tooth 

preparations do not turn out well on initial tries and additional 

teeth must be used. A reserve supply is also desirable so that 

teeth may be available for future studies. 

Head 

1. Sampling Procedure 

It is usually easiest to remove the head with the carcass 

resting on its right side if you follow the routine recommended. 

(First, collect and label a jaw sample.) Locate the blowhole on 

top of the head with your hand. Move your hand backwards to feel 

for the hard braincase beneath the skin and notice that the bony 

roof slopes downward at an abrupt angle posteriorly. The 

incision to separate the head from the vertebral column should be 

made just slightly (-3 cm) behind the sloped region. Make a 

small cut there with your knife blade. Make a small cut with 

your knife about 2.5 cm in front of the leading edge of the 

insertion of the left pectoral fin. 

Sever the head from the body by connecting the two small 

cuts and deepening and lengthening this incision above and on 

both sides of the neck. As the incision progresses, pull the 

beak downward toward the specimen's belly (Fig. 11A). This 

expandsthedorsal opening ofthe incisionandallowsyouto feel 

for the juncture between the occipital condyles of the skull and 

the first (atlas) vertebra. Continue to cut as you pull the beak 

down and as the skull and the vertebral column separate, tension 

will be suddenly released (usually accompanied by a popping 

noise). Peek in, and if you have made a proper incision, you 

will be able to see the separation between the convex condyles 

and the concave articular surfaces of the atlas vertebra from the 

top of the incision (Fig. 11B). 
35 

A 

B 

Figure 11. Collecting the head of a dolphin. A. The initial 

incision is lengthened and deepened as the beak is pulled down 

toward the belly. B. The occipital condyles (arrow) pull away 

from the atlas vertebra; then the neck is girdled and the head 

completely severed. 
36 

Extend and deepen the incision to girdle the neck using 

short knife strokes. Continue cutting until the head is 

completely severed. Exercise care to ensure that no damage is 

done to the condyles. 

The final task is labeling, packaging, and storing the 

sample (see page 16 for description). Enter next to the 

item HEAD as you complete the collection. 

2. Utility 

The head is collected because measurements of and 

relationships between cranial bones are among the most useful 

characters in diagnosing different races and subspecies. At the 

laboratory, the head is flensed, and the skull is cleaned and 

studied. Morphometric and meristic data gathered from the 

specimen are added to the appropriate data base for later stock 

discrimination and distribution studies. 

INTERNAL ANATOMY ITEMS 

Organization 

The organ systems of delphinids correspond generally in 

position (but not necessarily in proportions or morphology) to 

those of most other placental mammals, i.e. eutherians (Fig. 12). 

The body cavity is partitioned transversely by a muscular 

diaphragm into a thoracic portion containing a four-chambered 

heart, trachea and paired lungs, the esophagus, and various major 

blood vessels including the dorsal aorta: and an abdominal 

portion containing a liver, a stomach, a spleen, pancreas, 

intestines, reproductive tract, and urinary bladder. 

Retroperitoneally, the abdominal cavity contains paired kidneys 

and paired adrenal glands. 

Some of the features that distinguish delphinid internal 

anatomy from that of most other eutherians, however, include: 1) 

the numerous stomach compartments (non-ruminant eutherians 

typically have one), 2) the small number of lobes of the liver, 

3) the lack of a gall bladder, 4) the peculiar grape-bunch-like 

(botryoidal) morphology of the paired kidneys, 5) the extensive 

system of blood vessels ("retia mirabilia") in the thoracic 

cavity and elsewhere, 6) the remarkably long intestines, and 7) 

in males, the internal testes. Despite these general 

differences, however, dolphin anatomy is very much mammalian and 

the identification and collection of internal anatomical samples 

from dolphins is easily learned, especially if you have had prior 

experience with mammalian or comparative vertebrate anatomy. 
37 

ANTERIOR 

Esophagus Trachea 

-Heart 

Liver t 

Forestomach 

Spleen 

Pancreas 

- Kidney 

\* 

POSTERIOR 

- Intestine 

. Dorsal Peritoneal Sheet 

' Ventral Peritoneal Sheet 

' Urinary Bladder 

Anus 

Figure 12. Line drawing showing organization of internal anatomy 

of a female dolphin in ventral view (length protracted). 
38 

Mid-ventral Incision 

By now, all non-dolphin and external-anatomy items have been 

addressed and you have reached the point where you must collect 

the remaining data from inside the body cavities. For this, the 

carcass should be rolled high up on its right side, or as close 

to an upside-down position as possible. Observations and handling 

are best accomplished by straddling the specimen and bracing it 

with your legs. This allows both hands to be free for examining 

and cutting. 

(the breastbone which 

the ribs are attached to in front) and the umbilical scar 

situated on the midline about halfway between the head and the 

flukes. The umbilical scar usually is visible as an oblong, 

discolored depression two to four cm in length, one to two cm in 

width, and a few mm deep (Fig. 6). Run your hand along the 

midline and you should be able to feel this hard knot of scar 

tissue surrounded by relatively soft skin and muscle. After 

having located the umbilical scar, you should note the position 

of the genital/anal slit(s) (Figs. 5, 6, and 7). 

Inspect the venter for the sternum 

Short Incision 

If your instructions require you to sample only the gonads, 

you may not have to cut as far forward as the umbilical scar. 

(Your own short-cut method for quick-sampling of gonads may be 

developed for such cases.) For the short version of the 

incision, straddle the carcass and face the tail flukes. Make a 

shallow, straight-line cut from a point about 15 cm behind the 

umbilical scar to the posterior margin of the anal slit (Fig. 

13A). Once the length of the incision has been established, 

deepen it, but be careful not to damage the underlying organs 

that you intend to sample. Use the thumb or index finger of your 

freehand tohold the incision open (follow with your handbehind 

the knife) so that you can track the progress of the incision. 

Your knife will first pass through the thin layer of skin, 

then the white layer of blubber, and then the red abdominal 

muscle (Fig. 13B). Finally, it will pass through the external 

lining of the abdominal cavity (peritoneum) and you should be 

able to see the abdominal organs covered by the thin transparent 

peritoneal tissue ( i n t e r n a l l i n i n g ) , which can be cleared away 

with the fingers when necessary (Fig. 13C). 

Extended Incision and Thoracic Flap 

If samples of the stomach, blood, or other more anterior 

organs or tissues are needed, more working space inside the body 

cavity will be required than if you sample only the reproductive 

tracts. Both ends of the stomach must be found and tied off and 

you may have to locate and remove blood (probably) from the 
39 

A 

C 

Figure 13. Short mid-ventral incision of female dolphin for 

collecting posterior abdominal organ samples. A. Incision extends 

from behind the umbilical scar to the anal slit. B. Incision is 

deepened. C. Incision completed. Internal abdominal organs can 

be seen (arrow). 
40 

dorsal aorta situated dorsal to the lungs. You will know 

immediately that the mid-ventral incision has to be extended 

anteriorly into the thoracic cavity. You will save time if you 

make the extended a n t e r i o r c u t at the same time that you make the 

initial incision. 

Feel along the midline in front of the umbilical scar for 

the xiphoid process (posterior process) of the sternum. Make a 

straight-line cut from the posterior margin of the anal slit, 

through the umbilical scar, to the xiphoid process. Deepen the 

incision as in the short-incision procedure and stop (Fig. 14A). 

Roll the carcass back onto its right side. After feeling 

for the junctures of the ribs with the sternum, extend the 

incision anteriorly from the xiphoid process by cutting forward 

through the cartilagenous rib attachments (at the base of the 

ribs) between the ribs and the left side of the sternum. To form 

a flap of ribs that can be reflected to expose organs in the 

thoracic cavity and the anterior portion of the abdominal cavity, 

cut upward on the left side between the ribs immediately behind 

the left scapula, i.e. s h o u l d e r b l a d e (Fig. 14B). 

Reflect the flap that you have just made and you are ready 

to collect samples from any part of the thoracic or abdominal 

cavities (Fig. 14C). As you raise the flap, notice the prominent 

transverse sheet of muscle, the diaphragm, that at once forms the 

floor of the thoracic cavity and the ceiling of the abdominal 

cavity. The diaphragm must be cut away from the left wall to 

fully raise the flap anteriorly, and one side of the sheet of 

peritoneum attached to the urinary bladder must be severed to 

raise the flap posteriorly. 

Sampling from the Abdominal Cavity 

Urinary Bladder 

If the mid-ventral incision you have made is well-centered, 

the most ventrally positioned organ that you should see in the 

abdominal cavity is the urinary bladder. It is a heavily 

muscular, cigar-shaped organ, held in place (mid-ventrally and 

close to the body wall, immediately anterior to the genital 

opening) by a strong sheet of peritoneum that is attached to the 

wall of the body cavity on either side of the bladder (Figs. 12, 

14C, 15A and B). 

The bladder is a useful point of reference for you. If the 

carcass is female, the entire reproductive tract that you will be 

examining, labeling, and collecting will be located in a second 

sheet of peritoneal tissue immediately above (dorsal to) the 

sheet supporting the bladder (Figs. 12, 14C). During your 

initial workups, you may want to remove the bladder for a better 

view of the female reproductive tract, but there are no sampling 

requirements for bladders, so do not collect them. If the 
41 

A 

B 

C 

I 

Diaphragm (cut) 

Liver 

Kidney 

Intestine 

-eft Uterine 

Left Ovary 

I Uterus 

Horn 

#ladder 

Peritoneal SheetsFigure 14. Extended mid-ventral incision for collection of organ 

samples from thoracic and abdominal cavities. A. Incision is 

made anteriorly from the anal slit to the sternum. B. Ribs are 

cut from their cartilaginous attachments to the sternum. C. The 

left side is reflected to expose organs of the thoracic cavities 

(white and gray knobs at upper margin of cavity are knife 

handles). Especially note the double sheet of peritoneum in the 

genital region. One sheet supports the urinary bladder; the 

other (more dorsal) supports the uterus. 
42 

A 

Figure 15. Position of the urinary bladder. 

points to bladder, the most ventral abdominal organ. B. Bladder 

lifted to show sheet of supporting peritoneum. 

A. Finger at right 
43 

specimen is male, you will be collecting only t h e r i g h t testis 

and epididymis located immediately dorsolateral to the bladder. 

Because the bladder's position does not usually interfere with 

the sampling of male reproductive organs, there is little need to 

remove the bladder in males. 

Female Reproductive Tract 

1. Description 

You can best view the female reproductive tract and its 

sheet of peritoneal tissue (the broad ligaments) that holds the 

tract in place by excising the urinary bladder and its peritoneal 

support. The female reproductive tract includes the uterus 

(oriented along midline), a right and left uterine horn branched 

anterolaterally from the body of the uterus, and a small, beanto egg-shaped, reddish-brown ovary delicately attached to the 

fallopian tube at the anterior end of each horn via the broad 

ligament (Fig. 12). The uterus opens posteriorly into the 

vagina, which is separated from the uterus by the cervix. The 

cervix (literally, "neck") is actually the posterior constriction 

of the uterus that, together with a number of circular folds in 

the lining of the vaginal canal, among other things, may function 

in sealing the uterus. 

2. Sampling procedure 

The standard procedure is to tag the left horn of the uterus 

and collect the entire reproductive tract. If carcasses are 

always sampled in the same position, you will always know which 

horn is the left one. Although the left ovary usually produces 

more ova and is larger than the right ovary in sexually mature 

females, exceptions are common and it may be impossible to 

distinguish left horn from right horn if one of the horns is not 

identified and labeled before removal. To avoid possible 

confusion, you should free and label the left horn before freeing 

the right one. 

The physical connection between ovary and horn is tenuous 

and must be protected during sampling. While you cut, keep the 

left ovary and the anterior end of the left horn in your free 

hand.Pullthem slightlyaway fromtheleftwall. This makes the 

peritoneal tissue taut, enabling you to sever it more easily from 

the reproductive tissues. Draw the knife along the outside edge 

of the left ovary and horn and continue to cut the peritoneum 

away as you proceed down the left horn toward the uterus (Fig. 

16A). Because the peritoneum covers the ventral surface of the 

uterus and uterine horns, it is easier to distinguish between 

peritoneum and horn if you i n t e r m i t t e n t l y e v e r t the peritoneal 

sheet and examine its dorsal surface. 
44 

B 

Figure 16. Freeing the left uterine horn and labeling it in 

place. A. The left ovary (arrow) and the end of the left horn 

are held for protection as the peritoneal tissue is cut away from 

them. B. Cutting is halted when the body of the uterus is 

reached. A cable-tied label is then cinched securely to the left 

horn, half-way between the ovary and the uterine body. 
45 

After the left horn is freed, attach a cable-tied label to 

it halfway between the ovary and the horn's juncture with the 

uterus and cinch it tightly to ensure that it does not slip on 

the horn (Fig. 16B). Free the right horn and all of the uterus, 

except for its connection with the vagina, in the same manner as 

you did the left. 

In completing removal of the sample, use the cervical 

constriction as a natural seal to ensure that anything contained 

in the uterus is not lost. Locate the cervix by moving your 

finger up along the vaginal canal until you reach a firm, 

transverse, muscular barrier; then cut the reproductive tract off 

below it. Sever any connecting tissues and remove the sample. 

Now, place the reproductive tract on top of the carcass to be 

sure that all required parts are intact and that one horn (the 

left) is labeled. 

3. Utility 

Ovaries of dolphins contain data relating to their 

reproductive biology. Each time a female ovulates, a scar, 

'tcorpus albicans," is eventually formed on the ovary and it is 

generally thought that in dolphins corpora albicantia never 

disappear. A count of these scars provides an estimate of the 

number of opportunities a female may have had to become pregnant. 

Age estimates of females with only one scar, or only one freshly 

erupted Graafian follicle, furnish data about the average age 

that females of a certain population become sexually mature and 

how the age atsexua1maturationmaychange.Youareaskedtotag 

the left horn so that technicians at the laboratory can tell left 

from right sides when they count ovulation scars on the ovaries 

and tabulate pregnancies for the right or left horn. 

4. Pregnancies 

Inspect the reproductive tract and feel for any firm mass 

inside the uterus or horns that may indicate a fetus. If a fetus 

less than 25 cm is found, leave it in the uterus and print "yes" 

for FETUS (print lmnoll if not: enter a question mark if you are 

not sure). Finally, tag the left horn, remove the reproductive 

tract, place it into your sampling bucket, and print "yesl' beside 

UTERUS AND OVARIES. 

It is usually obvious when a female is carrying a large 

fetus. Even before the ventral cut is completed, the fetus-filled 

uterus may begin to slide out of the body cavity through the 

incision (Fig. 17A). Without puncturing the uterus, make an 

approximate length measurement of the fetus through the wall of 

the uterus to see if it is less than 25 cm long. If its length 

is equal to or more than 25 cm, slit theuterinewall, remove the 

fetus, and sex and measure it (Fig. 17B). Before you discard 

it, record FETUS SEX and LENGTH data on the life history form and 
46 

A 

B 

Figure 17. Handling large fetuses. 

A. A nearly full-term whitebelly spinner dolphin fetus in the 

left uterine horn. The left ovary is in the observer's left 

hand. B. For fetuses larger than 25 cm, the uterus is slit and 

the fetus removed, sexed, measured, and discarded, but the uterus 

and ovaries are collected. 
47 

enter llnotl beside FETUS [COLLECTED?]. Then, tag the left uterine 

horn, and collect the reproductive tract. 

One of the most important pieces of life history data 

collected for assessing dolphin populations is the percentage of 

mature females that are pregnant in a mortality sample. 

Laboratory researchers estimate this by adding all of the I1yest1 

responses in the FETUS [COLLECTED?] blank to all of the nonblank responses to FETUS:SEX and then dividing this sum by the 

total number of females in the sample that have at least one 

ovarian scar. Inferences about calving rates and trends in 

population growth for wild dolphins can be made from data on the 

percent of pregnant females sampled in the kill. 

Male: Testis and Epididymis 

1. Description 

The testes are paired, sperm-producing organs, situated 

slightly posterior and lateral to the kidneys. They are 

surrounded by peritoneum from which they hang from the 

dorsolateral walls of the posterior part of the abdominal cavity, 

in somewhat the same position as the ovaries in the females. The 

dolphin testes are sausage shaped and range in length from a few 

centimeters in calves, to well over 15 centimeters in some mature 

adults. 

The epididymis is a long, narrow, vermiform organ, with a 

sperm-storage function, that lies immediately dorsal to and in 

close association with the testis (Fig. 18A). The two organs are 

connected by a thin casing of peritoneum, which can be discerned 

by holding them up to a light and gently pulling them away from 

each other. The epididymis is longer on either end than the 

testis, and this sometimes creates a collecting problem with 

adult specimens because of the difficulty in defining the full 

extent of the epididymis posteriorly where it is continuous with 

the vas deferens. 

The requirements for sampling male reproductive organs are 

to collect the RIGHT TESTIS WITH EPIDIDYMIS ATTACHED. The right 

side is sampled simply for the sake of uniformity. If you 

develop a sampling routine that includes placing all carcasses in 

the same position at the same stage of sampling, you will always 

know which organs are on t h e r i g h t side without having to stop 

and think about it. 

If you move the intestines out of the way, along the back 

(dorsal) wall you should immediately see the paired kidneys that 

faintly resemble two bunches of grapes wrapped in transparent 

plastic. (Just anterodorsal to each kidney, is a small, firm, 

lobular adrenal gland--a potential candidate for future sampling 

(Fig. 18B).) If the carcass is that of an adult, the testes 

could be as large as the kidneys. The testes lie in almost the 
48 

A 

B 

Kidney 

Testis 

C 

Figure 18. Paired organs of the abdominal cavity of male 

dolphins. A. Testes with epididymides. The epididymis is longer 

than the testis on either end (arrows). B. Kidneys and adrenal 

glands. C. Comparison of position, size, and shape of kidneys and 

testes of mature male dolphin. Testes are suspended from the 

lateral walls by peritoneum; kidneys are technically excluded 

from the body cavity by peritoneum. 
49 

same regions and are about the same shape as the kidneys, but 

testes have a smoothly textured surface as compared to the 

kidneys' lumpy or botryoidal surface (Fig. 18C). 

2. Sampling Procedure 

Pull the testis and epididymis away from the body wall with 

one hand. Beginning at the anterior end of the organ complex, use 

short knife strokes along the dorsal margin of the epididymis to 

free the two organs from their peritoneal connection with the 

body wall (Fig. 19A). When you reach the posterior end of the 

testis, you must determine the limits of the epididymis so that 

part of the posterior portion will not be cut off and left with 

the vas deferens. The epididymis and vas deferens are continuous 

organs and they are nearly indistinguishable from each other. 

Often, the transition between epididymis and vas deferens can be 

discerned by an abrupt thickening on the end of the otherwise 

tapering epididymis. This enlargement marks the anterior extent 

of the vas deferens and you may safely sever the epididymis from 

the vas deferens at this point. Whenever you are in doubt about 

the exact limits of the epididymis, collect part of the vas 

deferens too (Fig. 19B). Lab technicians can always discard the 

excess, but they can never regenerate what is missing. 

Sever the testis and epididymis from all connecting tissue 

and remove them fromthebodycavity. Place the sample on top of 

the carcass and inspect it. If the testis is longer than six 

inches (-15 cm), it must be sliced to its center along its entire 

longitudinal extent so that the fixative will reach all parts of 

the organ (Fig. 19C). Large testes not prepared in this way will 

rot inside. Now, lift the sample by the middle of the epididymis 

and make a small slit with your knife just below, through the 

middle of the peritoneal connection between the epididymis and 

testis. Be careful not to puncture either organ. Thread a cabletie, with a specimen-numbered label, through the slit and secure 

it tightly to the epididymis. Finally, place the sample in your 

sampling bucket and print llyeslt next to the TESTIS item on the 

form. 

3. utility 

In the laboratory, the testis is measured and weighed with 

and without the epididymis. Histological sections are made from 

each organ to determine the presence and abundance of spermatids 

and spermatozoa. The data are used in studies of male 

reproductive maturity and seasonality. 
50 

B 

C 

Figure 19. Collecting the right testis and epididymis. 

A. The testis is pulled away from wall, making taut the 

suspensory peritoneum for easier cutting. 

B. Peritoneum is cut along the outside of the epididymis and the 

posterior end of the organ complex is severed well behind its 

actual terminus. 

C. Testes longer than 15 cm are slit lengthwise down to their 

centers to allow the fixative to penetrate the entire sample. 
51 

Stomach 

1. Description 

The stomach is located immediately under the diaphragm/liver 

complex in the anterior part of the abdominal cavity. As in other 

mammals, the delphinid stomach communicates with the pharynx 

anteriorly through the esophagus. The posterior part is in close 

contact with the spleen and pancreas and connects through the 

pyloric sphincter with the anterior end of the small intestine 

(i.e. the duodenum). Unlike that of other mammals, the delphinid 

stomach consists of three distinct compartments: 1) a highly 

muscular, non-secretory forestomach, 2) an enzyme-secreting midor main stomach, and 3) a thin-walled, non-muscular pyloric 

stomach (Fig. 12). 

2. Sampling procedure 

Because the stomach is collected for its contents, you 

should take every precaution to ensure that it is sealed and 

removed intact. Remember to store stomachs frozen because the 

stomach may contain calcareous fish otoliths and squid 

statocysts, important to prey studies, and formalin is acidic. 

If you have no instructions to collect the stomach, print l1nof1 

next to the item STOMACH on the life history form and go on to 

the next requirement. If your workup includes stomach collection, 

follow the steps below. 

Give yourself enough room to work. Open the anterior part of 

the abdominal cavity sufficiently to follow the course of the 

digestive tract from the small intestine to the point at which 

the esophagus penetrates the diaphragm. If you have trouble 

locating the esophagus, open the posterior part of the thoracic 

cavity and cut through the diaphragm and liver to reach and 

expose the esophagus. 

Define the limits of the stomach. Beginning at the 

esophagus, tease and cut away connecting tissues of liver, 

diaphragm, and peritoneum until you have cleared enough of the 

esophagus to close your free hand completely around the hind 10 

cm of it (Fig. 20A). Carefully clear tissues from the three 

stomach compartments. The tissues and organs surrounding the 

pyloric stomach usually cause the most trouble. The walls of the 

pyloric stomach are very thin and flaccid and they are closely 

associated with the spleen (a compact, semi-spherical, darkly 

colored gland) and, especially, with the rather amorphous and 

flabby pancreas (Fig. 20B). The problem is further complicated by 

the system of twists and turns typically exhibited by the pyloric 

stomach and the duodenal (upper) portion of the small intestine. 

Pull gently on the pyloric stomach and clear away connecting 

tissue (mostly mesenteries, ducts, and blood vessels) with only 

your hands at this point. Follow the tract posteriorly until you 

reach the pyloric sphincter, a muscular constriction that 
52 

A 

B 

C 

Figure 20. Defining the limits of the stomach. 

A. The connecting tissues of liver, diaphragm, and peritoneum are 

cut away to fully expose the esophagus (arrow). 

B. The tissues covering the three stomach compartments are 

removed. 

C. The pyloric sphincter (arrow) and duodenal ampulla (D.A.) at 

the anterior end of the small intestine are found by searching 

along the posterior end of the stomach. 
53 

physically and f u n c t i o n a l l y s e p a r a t e s the stomach from the the 

small intestine (Fig. 20C). The sphincter is identified by an 

abrupt change from flabbiness to firmness and wideness to a 

marked narrowness as you feel backwards along the pyloric stomach. 

Just posterior to the sphincter, the tract is wide for a short 

distance. This wide region of the small intestine is the duodenal 

ampulla. Posterior to the duodenal ampulla the intestine is (and 

remains) firm and narrow. 

the sphincter or the duodenal ampulla. If you are unsure about 

theposterior limits ofthe stomachwhen it is timetoremove it, 

tie off and sever the connection far down on the (more obvious) 

small intestine (Fig. 21A). 

Sometimes it is not easy to identify 

Close off both ends of the stomach. Because the esophagus 

and small intestine are difficult to seal off completely, it is 

important that you cinch down the cable ties as tightly as 

possible. Lock one around the esophagus about four or five cm 

above the stomach. Lockthe other aroundthe duodenum five cm or 

more below the pyloric sphincter. 

You now are ready to begin the final cutting and clearing 

process to remove the sample. Sever the esophagus above the tie 

and hold the sample in one hand and pull it up (or roll it out) 

toward you as you tease away the connecting tissue (Fig. 21B). Cut off the spleen and discard it. You may risk puncturing the 

pyloric stomach if you try to remove much of the pancreas, so cut 

around the pancreas and collect it with the sample. Sever the 

connection with the small intestine below the tie and remove the 

sample from the body cavity (Fig. 21C). Finally, attach a cabletiedlabelto any part ofthe sample, place the sample ina small 

plastic bag, cable-tie a label tightly around the outside to 

close the bag, and put it with the samples destined for the 

freezer. After you have done so, print a 88yes88 beside the item 

STOMACH on the form. 

3. Utility 

Stomachs are collected in order to study what, when, and how 

much the animals ate. A stomach chamber may contain freshly 

captured prey, such as whole fish or squids, that can be readily 

identified and measured, or all chambers may be empty except for 

undigested hard parts from which prey size and species may be 

inferred. Data from analyses of large samples of stomachs can be 

used to address questions about resource use, species 

competition, prey-type and prey-size preference, and energy 

requirements of the sampled dolphins. 

Major Organs of the Thoracic Cavity 

The following discussion is included for general reference 

in case you are asked to collect samples from the thoracic cavity. 
54 

B 

Figure 21. Removal of the stomach. 

A and B. The stomach is tied off at the esophagus and at the 

small intestine below the duodenal ampulla. The esophagus is 

severed above its tie and the small intestine below its tie. The 

sample is lifted up and out of the cavity as the remaining 

C. The stomach is examined to ensure that it is intact. 

connecting tissues are cut away from it. 
55 

Liver/Diaphragm/Lungs 

The diaphragm forms the floor of the thoracic cavity, but it 

is in close association with the liver lying just inside of the 

abdominal cavity and with the posterior margins of the lungs that 

occupy the pleural portion of the thoracic cavity. When the 

diaphragm is cut back and cleared away, it is possible to mistake 

lung tissue for liver tissue, especially if you are in too great 

a hurry to follow the lobes medially to determine their origins. 

Inspect both tissues and note how closely they can resemble each 

other in gross shape and (sometimes) in color (Fig. 22A). Compare 

them for texture and resiliency by squeezing lung and liver 

lobes with your hand. Notice that the liver tissue is much more 

compact and muscle-like, while the tissue of the lung has more 

of a spongy consistency. 

Heart/Dorsal Aorta 

The heart lies ventromedially to (between) the lungs and is 

encased in the pericardium (or cardiac sac). Reach between the 

lungs and feel for this softball-sized, muscular organ. To 

expose the heart you must cut a hole in the tough sac and then 

open it up with your hands (Fig. 22B). Pull the heart out from 

between the lungs to inspect it. The largest of the four 

chambers of the heart is the left ventricle (Fig. 22B). If you 

are instructed to collect blood from the heart, you will likely 

take the sample from that chamber. 

Aerated blood is pumped from the left ventricle t o a l l parts 

of the body via the aortic arch. Blood is sent dorsally and 

posteriorly via the dorsal aorta which lies along the dorsal wall 

of the thoracic and abdominal cavities (Fig. 22C). Open the 

thoracic cavity wide with your hands and pull the lungs ventrally 

to expose this large artery. You may be asked to collect blood 

from the d o r s a l a o r t a if such samples are needed in the future. 

Precise details for blood sampling, handling, and preservation 

will accompany any blood collecting requests. Nevertheless, 

do not forget to answer item BLOOD with a vtyestv or a vvnovv on the 

form. 

Parasites 

I recommend that you respond to this final "internal 

anatomyvv item last (if it is still necessary), because parasites 

maybe a s s o c i a t e d w i t h any organor area ofthe carcass andthus 

should be addressed when examining or sampling the applicable 

body part(s). There are no instructions for the sampling of 

parasites at the time of this writing, but you should remember to 

respond to this item with a vtyesvv or a vtnotv answer when 

processing a specimen. Special sampling requests will be 

accompanied by instructions when specific parasite samples are 

needed. 
56 

-- 

Figure 22. 

A. The liver and lungs may be confused. They are similar in 

shape and size and sometimes in color. Lungs are spongy to the 

touch: liver is more compact. 

B. The heart lies ventromedial to the lungs, encased in a sac of 

peritoneum. The largest chamber of the heart is the left 

ventricle. It is a potential source of aerated blood if blood is 

to be sampled. 

C. The dorsal aorta lies along the dorsal wall of the thoracic 

cavity above the lungs and esophagus. This artery is a potential 

source for blood samples. 

Major organs of the thoracic cavity. 
57 

FINISHING 

When you finish sampling the specimen, check your work. 

Make certain that you followed the requirements for the proper 

species, stock, and area. Check that you have responded to all 

of the appropriate data items on the form that you have just 

completed. Inventory your samples, and. ..remember to take the 

knife out of the knife holder before you discard it. 

I hope that this handbook has helped to make your job a 

little easier and more relevant for you. You have a difficult 

job that not everyone can do. But it is an important job, 

indispensible to the research effort. Without the care and 

concern exercised by observers in making the collections, very 

few biological studies could proceed. You should t a k e p r i d e in 

that. 

ACKNOWLEDGEMENTS 

For their helpful comments and recommendations on the 

manuscript, I thank A. Jackson, R. Miller, P. Akin, J. 

Gilpatrick, A. Dizon, A. Hohn, H. Bernard, J. Barlow, D. Holts, 

R. Holt, W. Perryman, W. Perrin, N. Mendes, B. Norberg, J. Young, 

M. Scott, M. Hall, and L. Rodriquez. I am especially grateful to 

B. Norberg who offered many suggestions and who helped in working 

out some of the details in the processing routines. K. Raymond 

prepared the figures. J. Michalski provided editorial assistance. 
58 

FURTHER READING 

Fraser, F.C. 1952. Handbook of R. H. Burne's cetacean 

dissections. British Museum (Nat. Hist.), London. 70 pp. 

Green, R.F. 1972. Observations of the anatomy of some cetaceans 

and pinnipeds. Pp. 247-297. In: S. Ridgway (ed.), Mammals of 

the sea. C.C. Thomas, Ill. 812 pp. 

Harrison, B.M. 1970. Dissection of the cat (and comparisons with 

man). C.V. Mosby, St. Louis. 198 pp. 

Walker, W.F. Jr. 1965. Vertebrate dissection. W.B. Saunders, 

Phila. 374 pp. 

Winchell, Jane M. 1982. A field manual for small cetacean 

dissection. 67 pp. (unpublished, copyright May, 1982 by 

College of the Atlantic, Bar Harbor, Maine 04609) 
59 

INDEX 

A 

Abdominal cavity 36, 37 

sampling 40-53 

Adrenal glands 47, 48 

Age distributions 19 

Aortic arch 55 

Atlas vertebra (head collection) 34, 35 

B 

Bags 

Basis for stock I D 

Bench grinder 

uses 15 

explained 10 

eye protection 17 

permission to use 17 

sharpening shears 17 

minimizing 19 

explained 9 

description 43 

fixation 16 

sampling 16 

storage 16 

Bias in selecting carcasses 

Blood 

Broad ligaments 

Bucket 

end-of-cruise preparations 17 

fixative in 16-17 

labeling 17 

preparing 16-17 

Buckets 

issued 15 

uses 15 

C 

Cable-tie number 

entered on form 20 

keeping track by set 20 

for attaching labels 14 

lengths 14 

locking 14 

pre-embossed for tail-tagging 19-20 

uses 14 

Cable ties 14 
60 

Carcass 

explained 9 

used as knife holder 18 

when chute assembled 18-20 

rapid 19-20 

dumping 18 

keeping track by set 20 

keeping track by well 20 

tail-tagging 18-20 

when to collect 4 

cutting behind 45 

description 43 

locating 45 

seals uterus 43 

Checking your work 57 

Chute 

carcass collecting 18-20 

Coamings (of wells) 

head storage 16 

Collection effort 18 

Coloration 

Carcass collecting 

Carcasses 

Cervix 

explained 6-7 

external anatomy item 21 

explained 10 

Color pattern and dorsal fin 

Containers 15 

Corpus albicans 

Cruise number 

sexual maturity 45 

explained 4 

non-dolphin items 21 

D 

Data item 

Date 

when not to respond 4 

explained 6 

non-dolphin item 21 

divides body cavity 36, 55 

description 55, 56 

potential blood-sampling source 55 

tying off 53, 54 

defining stomach 52-54 

Diaphragm 55 

Dorsal aorta 

Duodenum 

Duodenal ampulla 
61 

E 

Epididymis 

defining posterior end 47, 49 

description 47-48 

internal anatomy item (with testis) 43 

correcting 4 

exposing 53 

tying off 53, 54 

External anatomy items 

definition 20 

discussed 21, 23, 25, 26, 27-36 

Errors, entry 

Esophagus 53 

F 

Fallopian tube 

Fetus 

description 43 

explained 8 

measuring 45-47 

sexing 45-47 

when to collect 45 

when to discard 45, 46 

Fetus length 

explained 7 

Fetus sex 

explained 7 

Fixation of specimens 15-16 

fixation time 16 

fixative level 16 

cautions 15 

preparation 15 

Fixative 

Formaldehyde 15 

Formalin 

acidic 15 

burn 15 

damage to stomach contents 15 

permission to use 16 

specimens stored in 16 

Frozen samples 

carcasses 15 

heads 15 

inventory 16 

stomachs 15 

Frozen storage 

wells 16 

Freezer, cook's 
62 

G 

Gloves 14 

cotton, uses 14 

rubber, uses 14 

surgeons, uses 14 

used during fixative preparation 15 

keeping track of carcasses by set 20 

Greenbook 

H 

Head 

bending beak bellywards 34-35 

end-of-cruise preparation 17 

explained 8 

external anatomy item 24 

jaw sampled first 34 

labeling 16 

landmarks for collecting 34 

packaging 16 

sampling procedure 34-36 

use in biological studies 36 

description 55, 56 

description 43 

labeled in place 43-44 

Heart 

Horn, uterine 

I 

Internal anatomy 

compared to other mammals 36 

organization 36-37 

Internal anatomy items 36-56 

J 

Jaw sample (teeth) 

labeling 16 

preservation of 16 

sampling procedure 31, 33-34 

use in biological studies 34 
63 

K 

Kidneys 

compared to testes 47-49 

description 47-49 

holder 17 

safety 18 

sharpening 17 

hazards 17-18 

Knife 

knives 

L 

Labeling 

carcass (tail-tagging) 19 

epididymis (with right tes-is) 

head 16, 30 

head bag 16, 30 

left uterine horn 44, 45 

stomach 16, 30, 53 

advance preparation 14, 25, 30 

attached with cable ties 14 

Labels 14 

6, 30, 49 

printed with soft-lead pencil 14 

required 25, 30 

unstringing 14 

explained 7 

external anatomy item 24 

implication 31 

parasite fluid mistaken for 31 

test for 31, 32 

how many specimens to process 18-20 

processing 18-19 

compared with lung 55, 56 

described 55, 56 

compared with liver 55, 56 

described 55, 56 

Lactating 

Lactation 

Large kills 

Liver 

Lung 
64 

M 

Mammary gland 

explained 9 

external anatomy item 24, 31 

females 25, 26 

males 25 

Mammary slits 

Measuring devices 10-13 

Measuring sticks 10-13 

calibrating caliper jaws 11-12 

on site 21 

care of 10 

measuring with 11, 13 

metric measurement 12 

advantages, disadvantages 13 

rounding off 13 

extended (for complete sampling) 38, 40 

short (for quick-sampling of gonads) 38, 39 

landmarks for 38 

explained 9 

external anatomy item 24, 31 

Measuring tapes 13 

Measurements 

Mid-ventral incision 

Milk 

N 

Non-dolphin items 

definition of 20 

listed 21 

0 

Occipital condyles (head collection) 34, 35, 36 

Ovary 

Ovaries and uterus 

description 43 

collecting 43, 44 

explained 8 

P 

Pancreas 51, 53, 54 

description 51 

leave with stomach 53 
65 

Parasites 

explained 9 

fluid mistaken for milk in lactation test 31 

item completed at any time 

minimizing bias in 19 

for printing specimen number 14 

for sex determination 25, 28 

to complete forms 2 

55 

Partial-sample selection 

Pencil 

Pericardium 55 

Peritoneal sheets 

bladder suspension 40, 41, 42 

description 40, 41, 42, 43 

uterus suspension 41, 43 

Peritoneum (lining the cavity) 38 

Permission 

take photos 24 

use bench grinder 17 

use cookc's freezer 16 

use wells 16 

Pharynx 51 

Photos 

permission to take 24 

scale with 24 

Photos (Rolls, Frames) 

explained 9 

external antomy item 21, 24 

Porpoise Life History Form 2-10 

data items expained 4-10 

guidelines for completing 2, 4 

Position 

explained 6 

non-dolphin item 21 

Positioning carcasses 

first (sketch position) 21 

second (right side) 24 

third (upside-down) 38 

uniformly 21, 43, 47 

Pregnancies 45-47 

detection of 45, 46 

Probe (pencil) 

use in sex determination 25, 28 

Processing 

decisions 18-19 

Processing specimens 

order 20 

routine 20-57 

time limits 18-19 

against cuts (see knife safety) 

eye 17 

skin (see gloves) 

Protection 
66 

Pruning shears 

care of 17 

sharpening 17 

use 17, 31, 33, 34 

posterior limits 51-53, 54 

Pyloric sphincter 

Q 

Quad 

explained 6 

non-dolphin item 21 

R 

Random sampling (partial-sample selection) 19, 21 

Reproductive tract, female 

description 37, 47-49 

labeling 44, 45 

sampling procedure 43-45 

use in biological sudies 45 

description 47-49 

labeling 49 

sampling procedure 49-50 

use in biological studies 49 

labeling 25, 30 

packaging 16-17, 30 

sampling 25, 29 

storage 16-17, 30 

sample preservation 15-17 

specimen handling 15-17 

Reproductive tract, male 

Requirements 

Routine 

S 

Sample preservation 15 

Sampling equipment 

description 2-17 

use 2-18 

Sampling error (increased by divergent methodologies) 1 

Seawater 

for sampling bucket 16 

for washing hands 16 

preparing fixative 15 

Selection of specimens (in sampling part of kill) 19, 21 
67 

Set number 

explained 6 

non-dolphin item 21 

explained 5 

external anatomy item 24, 25 

direction of probe 25, 28 

distance between ventral openings 25, 26, 27 

mammary slits 25, 26 

Sex 

Sex-determination criteria 

Sex ratios 19 

Size distributions 19 

Sketch/fin trace 

explained 10 

external anatomy item 21-24 

Sketch position 21, 22 

Species/stock 

explained 5 

external anatomy item 21 . 

explained 5 

external anatomy item 24, 25 

printed on labels 14 

when to assign 5 

description 51 

landmark for mid-ventral incision 38, 40 

external anatomy item 21 

defining limits 51-53 

description 37, 51, 52, 54 

explained 8 

fore- 51, 54 

labeling 16, 53 

mid- 51, 54 

packaging 53 

pyloric 51, 53, 54 

sampling procedure 51-54 

tying off with cable ties 16, 53, 54 

storage 16, 51, 53 

labeling 16-17 

packaging 16 

storage 17 

end-of-cruise preparation of 17 

separate packaging of 17 

use in biological studies 51, 53 

Specimen number 

Spleen 51, 52, 53 

Sternum 

Stock ID (basis for) 

Stomach 

trouble in collecting 51, 53 

Stomach bag 

Stomach samples 

Storage area, recommended 15 
68 

T 

Tail-tagging carcasses 14 

Teeth 

explained 7 

external anatomy item 24 

in jaw samples 16 

sampling procedure 31, 32, 33-34 

use in biological studies 34 

compared to kidneys 47-49 

description 47-48 

explained 8 

internal anatomy item 43 

labeling (epididymis) 16 

large specimens sliced open 49, 50 

preservation of 16 

definition 36, 37 

organs of 53, 55, 56 

for processing 18-19 

explained 6 

external anatomy item 21, 24 

definition 11, 13 

measuring 11, 13 

for packaging heads 16 

Testis 

Thoracic cavity 

Time limits 

Total length 

Twine, seine-net 

U 

Umbilical scar (as landmark for mid-ventral incision) 38, 39 

Urinary bladder 

description 40 

reference point for internal organs 40, 43 

Uterus (and ovaries) 

description 43 

labeling 16 

preserving 16 

Vagina 

Vas deferens 

Ventricle, left 

description 43 

continuous with epididymis 47-48 

potential source of blood sample 55 
69 

W 

Wells 

Well-space 

Wet deck 

Whetstone 

permisssion to use 16 

rental for carcass frozen storage 18 

good storage area 15 

sharpening knives 17 

sharpening shears 17 

X 

Xiphoid process (of sternum) 

mid-ventral incision landmark 40 

Y