NOAA Technical Report NMFS 1

Synopsis of Biological Data

on the Blue Crab,

Callinectes sapidus Rathbun

Mark R. Millikin and Austin B. Williams

March 1984

FAO Fisheries Synopsis No. 138

u.s. DEPARTMENT OF COMMERCE

Malcolm Baldrige, Secretary

National Oceanic and Atmospheric Administration

John V. Byrne, Administrator

National Marine Fisheries Service

William G. Gordon, Assistant Administrator for Fisheries
The National Marine Fisheries Service (NMFS) does not approve, recommend or endorse any proprietary product or proprietary material

mentioned in this publication. No reference shall be made to NMFS, or

to this publication furnished by NMFS, in any advertising or sales promotion which would indicate or imply that NMFS approves, recommends

or endorses any proprietary product or proprietary material mentioned

herein, or which has as its purpose an intent to cause directly or indirectly

the advertised product to be used or purchased because of this NMFS

publication.
CONTENTS

Introduction .

Identity . .

1. 1 Nomenclature .

I. 11 Valid name .

1.12 Objective synonymy .

1.2 Taxonomy .

1.21 Affinities .

1.22 Taxonomic status .

I. 23 Subspecies .

1.24 Standard common names, vernacular names .

1.3 Morphology .

1. 31 External morphology .

1.32 Cytomorpho1ogy*

1.33 Protein specificity .

1.34 Aging .

1.35 Endoskeleton .

1.36 Hemolymph (blood) .

1.37 Internal anatomy .

2 Distribution .

2. 1 Total area .

2.2 Differential distribution .

2.21 Larvae and juveniles .

2.22 Adults .

2.3 Determinants of distribution changes*

2.4 Hybridization *

3 Bionomics and life history .

3.1 Reproduction .

3.11 Sexuality .

3. 12 Maturity .

3.13 Mating .

3.14 Fertilization .

3.15 Gonads .

3.16 Spawning '" .

3.17 Eggs .

3.2 Preadult phase .

3.21 Embryonic phase .

3.22 Larval phase .

3.23 Juvenile phase .

3.3 Adult phase .

3.31 Longevity .

3.32 Hardiness .

3.33 Competitors .

3.34 Predators. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . .

3.35 Diseases, parasites, and commensals , .

3.4 Nutrition and growth .

341 Feeding .

3.42 Food . .

3.43 Growth and morphometry .

3.44 Metabolism .

3.45 Molting. . . . . . . . . . . . . . . . . . . .. .. . .

3.5 Behavior. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . .

3.51 Migrations and local movements. . . . . . . . . . .. . .

3.52 Schooling .. .

3.53 Responses to stimuli. . . . . . . . . . . . . . . . . . .. . .

4 Population . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . .. . .

4. I Structure...... . .

4.11 Sex ratios .

4.12 Age composition*

4.13 Size composition .

111

I

1

I

1

I

I

I

4

4

5

5

5

5

5

5

5

6

6

6

7

7

8

8

8

8

8

8

9

9

9

9

9

9

10

15

16

16

16

16

17

17

20

20

2i

21

23

23

24

24

25

25

25

25

25

25
4.2 Abundance and density (of population) 25

4.21 Average abundance *

4.22 Changes in abundance .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

4.23 Average density*

4.24 Changes in density. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26

4.3 Natality and recruitment 26

4.31 Reproduction rates*

4.32 Factors affecting reproduction *

4.33 Recruitment 26

4.4 Mortality and morbidity 26

4.41 Mortality rates*

4.42 Factors affecting mortality rates 26

4.43 Factors affecting morbidity 26

5 Exploitation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26

5.1 Crabbing equipment 26

5. I I Gear 26

5.12 Boats. . . . . . . . . . . . . .. . . . . .. . . . . . . . .. . . .. . . .. . . . . . . . . .. . . . . . . . . .. . . .. .. . . . . . . .. . .. . . .. . 27

5.2 Blue crab areas. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

5.21 General geographic distribution. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

5.22 Geographical ranges. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

5.23 Depth ranges. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

5.24 Conditions of the grounds 27

5.3 Crabbing seasons. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

5.4 Crabbing operations and results 27

5.41 Effort and intensity. . . . . . . . . . . . .. . . . . .. . . .. .. . . . . . . .. . . . . . .. . . . . . .. . .. . .. . . . . .. . . . .. . .. . 27

5.42 Selectivity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27

5.43 Catches..... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

6 Protection and management . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

6.1 Regulatory measures 30

6.11 Limitation or reduction of total catch 30

6.12 Protection of portions of population 30

6.2 Control or alteration of physical features of the environment*

6.3 Control or alteration of chemical features of the environment*

6.4 Control or alteration of the biological features of the environment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

6.41 Control of aquatic vegetation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

6.5 Artificial stocking 30

7 Soft crab shedding operations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

7. 1 Use of cultured crabs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

7.2 Procurement and maintenance of stock. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

7.3 Genetic selection of stocks *

7.4 Spawning.. .. .. .. . . .. .. .. . . 31

7.5 Shedding operation management . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31

7.6 Foods; feeding. . . . . . . . . . . . . . . . . . , . . . . . . . .. 31

7.7 Diseases . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31

Acknowledgments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32

Literature CIted 32

*No information available.

iv
Synopsis of Biological Data on the Blue Crab,

Callinectes sapidus Rathbun 1

MARK R. MILLIKIN2 and AUSTIN B. WILLIAMS 3

ABSTRACT

This synopsis reviews taxonomy, morphology, distribution, life history, commercial hard and soft shell

crab fisheries, physiology, diseases, ecology, laboratory culture methodology, and inlluences of environmental

pollutants on the blue crab, Callinectes sapidus. Over 300 selected, published reports up to and including 1982

are covered.

INTRODUCTION

The importance of the commercial and recreational fisheries of

the blue crab, Callinectes sapidus Rathbun, along the Atlantic and

Gulf of Mexico coasts of the United States is reflected in the large

amount of basic and applied research conducted with this species,

both past and present. Life history and the fishery for the blue crab

have been described for Delaware (Hall 1976), Chesapeake Bay

(Hay 1905; Churchill 1921; Pearson 1948; Van Engel 1958), North

Carolina (Pearson 1951), South Carolina (Eldridge and Waltz

1977), Georgia (Palmer 1974), St. Johns River, Fla. (Tagatz

1968a, b). the Atlantic coast states in general (Sholar 1982), gulf

coast of Florida (Oesterling 1976), Alabama (Tatum 1979), Mississippi (Perry 1975), Louisiana (Jaworski 1972, 1982; Adkins

1982; Roberts and Thompson 1982), and Texas (More 1969). A

partial bibliography by Cronin et al. (1957) and an annotated bibliography by Tagatz and Hall (1971) on the bl ue crab have served

researchers, fishery managers, and commercial crabbers. In view

of the continuing contributions to the knowledge of life processes

of this species, a synopsis of biological data of C. sapidus is

presented.

1 IDENTITY

1. I Nomenclature

1.11 Valid name

Callinectes sapidus Rathbun 1896.

l. 12 Objecti ve synonymy

It is not necessary to cite here all references to descriptions or

figures attributable to C. sapidus. The synonymy below is a

modified version of that given by Williams (1974) and does not

distinguish objective and subjective synonyms.

I Contribution No. 83-23C from the Southeast Fisheries Center, Charleston

Laboratory.

'Southeast Fisheries Center Charleston Laboratory, National Marine Fisheries

Service, NOAA, P.O. Box 12607, Charleston, SC 29412.

'Northeast Fisheries Center Systematics Laboratory, National Marine Fisheries

Service, NOAA, National Museum of Natural History, Washington, DC 20560.

Portunus hastatus Bosc 1802 (eastern North America; not Cancer

(= Portunus) hastatus Linnaeus 1767).

Lupa hastata Say 1817 (eastern North America; not L. hastatus

Desmarest 1823 = Cancer hastatus Linnaeus 1767).

Portunus diacantha Latreille 1825 (variety; North America,

Antilles, Brazil, etc., types not extant; restricted to Philadelphia,

Pa., by Holthuis (1962), and name suppressed by International

Commission on Zoological Nomenclature (1964) ).

Lupa dicantha Gould 1841 (variant spelling of P. diacantha

Latreille 1825; also DeKay 1844; Holmes 1858).

Callinectes diacanthus Stimpson 1860 (western Atlantic).

Callinectes hastatus Ordway 1863 (western Atlantic; also A. Milne

Edwards 1879; Rathbun 1884; Young 1900).

Neptunus (Callinectes) diacanthus Ortmann 1894 (Florida; Haiti;

Brazil).

Callinectes sapidus Rathbun 1896 (eastern North America and

Caribbean).

Callineetes sapidus acutidens Rathbun 1896 (Brazil).

Callinectes africanus A. Milne Edwards and Bouvier 1900 (= a

figure of C. sapidus).

I.2 Taxonomy

1.21 Affinities

Suprageneric

Phylum Arthropoda

Class Crustacea

Subclass Malacostraca

Order Decapoda

Suborder Pleocyemata

Infraorder Brachyura

Superfamily Portunoidea

Family Portunidae

Subfamily Portuninae

Generic

Callinectes Stimpson, 1860

The generic concept is that of Stimpson (1860), emended by

Rathbun (1896), and Williams (1974), who gave the following
description: Portunid crabs lacking an internal spine on carpus of

chelipeds. Abdomen of males broad proximally, narrow distally,

roughly T-shaped; first segment broad, almost hidden; second

segment broad, slightly overlapping coxae of fifth pereopods at

each side; third-fifth segments fused and tapering sinuously from

broad third to distally narrow fifth; sixth segment elongate and

narrow; telson ovate with acute tip. Abdomen of females exhibiting

two forms: Immature females with abdomen triangular from fourth

segment to tip of telson, segments fused; mature females with

abdomen broadly ovate (excluding telson), segments freely articulated; first segment almost hidden; second and third segments

slightly overlapping coxae of fifth pereopods at each side; fifth and

sixth segments with greatest sagittal length; sixth segment narrowing distally in irregular broad arc to articulate with triangular telson.

Abdomen and telson of both sexes reaching anteriorly beyond

suture between thoracic sternites IV and V.

The type-species is Callinectes sapidus Rathbun 1896, by designation of the International Commission on Zoological l"omenclature (1964:336).

Stimpson (1860) created the genus Callinectes to contain

portunids in which the males have a T-shaped abdomen and the

merus of the outer maxillipeds is short, sharply prominent, and

curved outward at its external angle. Today only Stimpson's abdominal character helps to distinguish Callinectes from similar

portunid genera, the obvious affinities leading both Stephenson and

Campbell (1959) and Stephenson (1962) to challenge the validity of

Callinectes, although Stephenson et al. (1968) agreed to its

distinctness after numerical analysis of 57 characters in 41 species

of portunids in 4 genera. Strengthening this position is the lack of

an internal distal spine on the carpus of the chelipeds of both the

megalopa and all crab stages in Callinectes, whereas this spine is

always present, occasionally prominent, in all Portunus species.

Specific

The following species diagnosis for adult specimens of Callinectes sapidus (Fig. I) is quoted from Williams (1974:779):

"Carapace bearing two broad either obtuse or acuminate,

triangular frontal teeth with mesial slopes (incorporating a pair of

rudimentary submesial teeth) longer than lateral slopes [Fig. I].

~etagastric area with posterior width approximately 1.2 times

length, anterior width about 2 times length. Anterolateral margins

slightly arched; anterolateral teeth exclusive of outer orbital and

lataral spine obtuse to acuminate and directed outward more than

forward. Much of surface smooth, with scattered granules, but

granules concentrated locally on mesobranchial, posterior slope of

p

..........•.......

I

1

I

,I

I

~_-:':'C"''''''''-''''-=::: .. ~:C~~;=~'~~~~~~1~:f1;~~~~~~~~~;~;?==>-_::::""'L-S---?/ ____~-~::..-~_____ C,A

I

I

I

I

I

I

Figure I.-Matu.·e male Callinectes sapidus from North Carolina in dors"t view. Measured features indicated by numbered lines: 1, carapace length; 2, carapace width to

base oflateral spines; 3, carapace width including lateral spines; dimensions of metagastric area: a, anterior width, b, length, c, posterior width. Other features include: F,

frontal teet:,; 0, outer orbital tooth; AL, anterolateral teeth; LS, lateral spine; PL, posterolateral margin; Ep, epibranchialline; ES, epistomial spine; MB, mesobranchial

area; CA, cardiac area; BL, branchial lobe; cheliped: M, merus; C, carpus; P, propodus; D, dactyl (figure reproduced from Williams 1974).

2
cardiac, and anterior portion of mesogastric area; a tendency to

crowding of granules into transverse ridge at summit of cardiac and

mesobranchial area in some individuals. Sculpturing of surface

varying individually from low to raised relief. Lateral spines varying from rather stout, blunt, and forward trending to slender,

elongate, and slightly backward trending. Epibranchialline nearly

straight over branchial region, otherwise sinuously curved.

"Propodus and carpus of chelipeds with moderate finely granulate ridges, width of chelae similar, propodal finger of major hand

occasionally with lower margin decurved proximally.

"Male abdomen and telson reaching about midlength of thoracic

sternite IV; telson lanceolate, much longer than broad; sixth

segment of abdomen broadened distally. Mature female abdomen

and telson reaching about midlength of thoracic sternite IV; telson

with inflated sides almost equilaterally triangular, fifth and sixth

abdominal segments equal in length [Fig. 2]. First gonopods of

male very long, reaching beyond suture between thoracic sternites

1V and V but not exceeding telson; sinuously curved and overlapping proximally, diverging distally, twisting mesioventrally on axis

lateral to abdominal locking tubercle and recurving to termination

near midline; armed distally with row of large and small retrogressive spinules following ventral and lateral borders with twist of axis;

tip membranous, tlared portion suggesting an elongate quadri lateral

in outline. Gonopores of female paraboloid in outline with apex on

iong axis directed anteromesad, aperture of each sloping from

surface on mesial side under irregularly rounded and linearly

wrinkled anterior border superior to bulbous posterolateral border."

Size: A male from Chesapeake Bay measuring 9 in (227 mm)

in carapace width has been reported (Williams in press). A female

measuring 204 mm across the carapace, including lateral spines,

with a length of75 mm was reported by Williams (1974). He stated

(p. 779) that "mature size of females varies considerably, the

smallest examined having a carapace length of 21 [mm], width at

base of lateral spines 41 [mm] , including lateral spines 55 [mm] ."

Color: (From Williams 1974:780) "Grayish, bluish, or brownish green of varying shades and tints dorsally on carapace and

chelipeds; spines may have reddish tints. tubercles at articulations

of legs orange, and legs varyingly blue and white with traces of red

or brownish green. Males with propodi of chelae blue on inner and

outer surfaces ... and tipped with red. Mature females with orange

fingers on chelae tipped with purple. Underparts off-white with tints

of yellow and pink."

According to Williams (1974:780), "Color variations other than

those associated with sexual dimorphism and molt cycle are known.

Albinos or partial albinos are in museum collections and have been

reported both in systematic literature and elsewhere." Other colors

reported are: An adult male lacking dorsal green coloration and

bright blue and scarlet markings on the legs, the upper surface of the

carapace being "robins egg blue" and the appendages paler than

usual, but abdomen and underparts with normal color; a bilateral

gray and brown colored specimen (summarized by Williams 1974).

Type-specimens: No longer extant.

Type-locality: Restricted to "east coast of United States"

(Williams 1965).

For nomenclatural purposes, Holthuis (1962) selected a lectotype

from Latreille's (1825) original material from Philadelphia, Pa. It is

almost certain that Latreille's material no longer exists, though it is

equally probable that his specimens were from North America. By

that designation, the "typical" sapidus is associated with a

population belonging to the eastern United States and particularly

to the Middle Atlantic States.

Subspecies: Will iams (1974:780) pointed out that "there are

morph<)logical variations in this species having far greater systematic interest than si7e and color." Study of many specimens from

throughout the range of the species bears out the conclusion of

Chace and Hobbs (1969) that extreme variants "are so different

from each other that they could easily be interpreted as distinct

species," but there is "no point of demarcation" -morphological,

geographic, bathymetric-between the "typical" rather bluntspined form predominating along the east coast of the United States

and the acute-spined form, named C. sapidus acutidens by Rathbun, predominating from Florida southward.

According to Williams (1974:780), "Rathbun (1896) characterized the 'acutidens' form (paraphrasing) as being wider than the

'typical' with all prominences more strongly marked, areolations

separated by deeper depressions, granules more raised, gastric

ridges stronger and more sinuous, a transverse granulate ridge on

each cardiac lobe, frontal teeth narrower and more acute and bearing two small intervening teeth, anterolateral teeth broad at base and

narrowing abruptly to long acuminate tips with margins granulate,

Figure 2. -Composite ventral view of thoracic steroites (Roman

numerals), abdomen (Arabic numerals), and telson (T) in situ. a,

mature male; b, mature female; c, immature female (figure reproduced from Williams 1974).

3

III

5-'-3

J_ -- .11

h

c
lateral spines longer than in 'typical' specimens of equal size, and

ridges of chelipeds quite prominent and strongly granulate.

"I thought for a time that a species distributed through approximately 85° of latitude from North Temperate through Tropic to

South Temperate Zones might reflect responses to temperature in

spination or other characters. 'typical' structure being prevalent in

the temperate zones and sharp spination in the tropics, the differences thereby justifying nomenclatural recognition. There is weak

but inconsistent evidence for this pattern. Though 'acutidens'

individuals are uncommon outside the tropics, intermediates occur

everywhere to some degree, and some 'typical' individuals occur in

the tropics. Genetic pooling or environmental response reflected in

morphology seems poorly structured.

"I consider the whole C. sapidus complex to be a single species

which has diverged into ill defined populations in certain portions

of its range. The 'acutidens' form predominates over most of the

latitudinal range. but there are variations. Among these are 'typical'

features that reach their most pronounced expression in the population along the east coast of the United States. Taxonomic thinking of

biologists has been clouded by the fact that the form originally

described was the North American variant which became the

standard against which all comparisons were made.

"Callinectes sapidus is the member of the genus which has most

successfully invaded the Temperate Zone, and in this respect it may

bc that speciation into forms associated with temperature regimes

is progressing, but the process is not yet complete enough that

morphological separation is distinct."

A detailed key for identification of the various species of Callinectes was presented by Williams (1974). Diagnostic characters of

importance for adults include granulation of the carapace, shape

and size of frontal and anterolateral teeth, and shape of the male

gonopods. Distinguishing characters of various Callinectes sp.

occurring in the western central Atlantic Oat. 35°N to 5°N: long.

50 0 W to 95°W) were described by Williams (1978).

Keys to species occurring in eastern and southern United States

Callinectes sapidus is the only species in the genus having 2

broad based, triangular teeth located on the front between the inner

orbital teeth. The 7 species of Callinectes whose ranges may

overlap in parts of the eastern and southern United States can be

identified with the aid of the following keys adapted from Williams

(1974). Also, color and morphology of these species were discussed

by Norse and Fox-Norse (1982).

Key to species based on carapace (excluding juveniles)

lao Front with 2 prominent, broad based, triangular teeth

between inner orbitals; each with or without rudimentary

submesial tooth on mesial slope C. sapidus

Ib. Front with 4 teeth between inner orbitals or 2 prominent

teeth separated by a space often bearing pair of rudimentary submesial teeth 2

2a. Submesial pair of frontal teeth well developed and more

than half as long as lateral pair (measuring from base of

lat:::ral notch between teeth) C. bocourti

2b. Frontal teeth decidedly unequal in size, submesial pair

no more than half as long as lateral pair (measuring from

base of lateral notch between teeth) 3

3a. Carapace very smoothly granulated, lines of granules

visible but barely perceptible to touch (except epibranchial line variably prominent) C. similis

4

3b. Carapace not so smooth, scattered granules and lines

of granules quite evident to sight and touch 4

4a. Carapace coarsely granulated; anterolateral teeth (exclusive of outer orbital and lateral spine) lacking shoulders

and swept forward 5

4b. Carapace finely granulated; anterolateral teeth 2-6 (exclusive of outer orbital, 7-8 and lateral spine) with shoulders,

not swept forward 6

5a. Anterolateral teeth well separated, all except first 3 and

lateral spine with anterior margins concave; chelipeds

with ridges finely granulated C. larvatus

5b. Anterolateral teeth adjacent, stout, anterior margins not

noticeably concave, fifth tooth often largest; chelipeds

with ridges coarsely granulated C. exasperatus

6a. Submesial pair of frontal teeth absent or vestigial

...................................... C. ornatus

6b. Submesial pair of frontal teeth never vestigial, but no

more than half length of lateral pair C. danae

Key to mature or nearly mature males

based primarily on first pleopods

Ia Tips of first pleopods fall ing well short of suture between

thoracic sternite VI and mesially expanded sternite VII . 2

lb. First pleopuds reaching to, almost to, or beyond suture

between thoracic sternite VI and mesially expanded

sternite VII 4

2a. First pleopods well separated from each other. never

touching or crossed: tips not lanceolate 3

2b. First pleopods overlapping each other, often crossed;

tips lanceolate C. ornatus

:~a. First pleopods slender distally, nearly straight, tips bent

slightly mesad C. simi/is

3b. First pleopods fairly stout distally, angled toward midline

then abruptly bent forward in short slender terminal

extension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. larvatus

4a. Tips of first pleopods curved abruptly mesad

................................... C. exasperatus

4b. Tips of first pleopods not curved abruptly mesad 5

5a. First pleopods with tips never extending beyond abdominallocking tubercle on thoracic sternite V, slender distal

part almost straight, minutely spined. tips almost always

bent ventrolaterally C. danae

5b. First pleopods with tips extending beyond abdominal

locking tubercle on thoracic sternite V, slender part

definitely curved or sinuous, variously spined, never

bent ventrolaterally at tip 6

6a. Front with 2 prominent, broad based, triangular teeth

between inner orbitals; each with or without rudimentary

submesial tooth on mesial slope C. sapidus

6b. Front with 4 teeth between inner orbitals reaching nearly

common level C. bocourti

1.22 Taxonomic status

This is a morphospecies exhibiting some variation.

1.23 Subspecies

None. See section 1.21 for discussion on the "forms" of Callinectes sapidus.
1.24 Standard common names, vernacular names

Blue crab

I.3 Morphology

1.31 External morphology

Detailed accounts of external morphology of the blue crab are

presented by Pyle and Cronin (1950) and in section 1.21 of this

synopsis.

1.33 Protein specificity

Polymorphism of several enzymes of the blue crab have been

determined. Polymorphic enzymes for both Chesapeake Bay and

Chincoteague Bay blue crab populations include: Gonad acid

phosphatase, glutamate oxaloacetate transaminase-I, phosphoglucose isomerase, hydroxybutyrate dehydrogenase, malic enzyme, 6-phosphoglucomate dehydrogenase, phosphoglucomutase,

esterase, peroxidase, a-glycerophosphate dehydrogenase, and

glyceraldehyde-3-phosphate dehydrogenase (Cole and Morgan

1978a). Only the first three enzymes provided reliable zymograms

in the gonad, muscle, and green gland tissues, respectively.

Monomorphic enzymes for both populations included acid phosphatase of the green gland, adenylate kinase, creatine kinase,

esterase (from gonad and gill), hexokinase-I and -2, lactate

dehydrogenase- I and -2, leucine amino peptidase-I and -2, malic

dehydrogenase, sorbitol dehydrogenase, and xanthine dehydrogenase. Comparison of muscle, eye, and serum protein from adult

male crabs from the Choptank River, Md. (8 to 12%0), Backbay,

Va. (17 to 20Yoo), and Chincoteague Bay (35:100) with discontinuous

polyacrylamide electrophoresis yielded no differences in muscle

protein and low polymorphism in eye and serum proteins (Cole and

Morgan 1978b). In adult male crabs, lactate dehydrogenase

occurred as one band of activity and 6-phosphogluconate dehydrogenase occurred as two bands of activity following electrophoretic

separation from each of seven tissues (muscle, heart, gill, vas

deferens, midgut gland, testes, and hypodermis) (Dendinger 1980).

Conversely, glucose-6-phosphate dehydrogenase occurred as four

bands of activity with different bands in different tissues.

Genetic similarity in the Chincoteague and Chesapeake Bay

blue crab populations is suggested from similar percentage polymorphism and similar gene frequencies of the polymorphic enzymes, acid phosphatase, glutamate oxaloacetate transaminase-I,

and phosphoglucoisomerase-I. Therefore, Cole and Morgan

(1978a) suggested that genetic uniformity is most Iikely maintained

by larval intermixing offshore and that recruitment to estuarine

populations of blue crabs is accomplished by immigration of

megalopae.

1.34 Aging

Aging of blue crabs is difficult because of the lack of annuli

formation on hard body parts. Variation in age to maturity is

covered in section 3.12.

1.35 Endoskeleton

A briefdescription ofthe endoskeleton was presented by Pyle and

Cronin (1950).

5

1.36 Hemolymph (blood)

Osmoregulation and ionic regulation

Except for the larval stages of development, blue crabs are

euryhaline and hyperosmoregulate in the majority ofthe salinities in

which they occur. Blue crabs sometimes occur in freshwater

habitats that have access to estuarine waters, at least seasonally

(Mangum and Amende 1972). Blue crabs have been reported in

freshwater in Louisiana (Gunter 1938), Florida (Odum 1953),

Virginia (Mangum and Amende 1972), and Delaware (Ettinger and

Blye 1981). Freezing-point depression determinations of blood

samples have shown that all adult crabs sampled were hyperosmotic

to salinities of 10,20, and 30100 (Tan and Van Engel 1966). Males

had more ability than females to regulate blood sodium concentrations, especially at a salinity of IQ%., , thereby suggesting a physiological adaptation which allows adult male crabs to inhabit low

salinities (Tan and Van Engel 1966). Ovigerous females do not

regulate as well as nonovigerous, mature females or adult males at

salinities of 1.7 or 17Yoo (Tagatz 1971). This is consistent with the

fact that ovigerous females usually occur in salinities higher than

17Yoo . Temperatures between 10° and 30°C did not significantly

affect the osmoregulatory ability of various stages or sex of blue

crabs. According to Leffler (I975a), juvenile blue crabs (30-50

mm carapace width) reared in 25°C, hyperosmoregulate in media

of < 700 mOsm/l (25100 salinity) and conform osmotically at

higher salinities. Regarding sodium ions, juvenile blue crabs

hyperosmoregulate in environments of up to 720 mOsm/1 (26100

salinity), whereas adult blue crabs hyperosmoregulate below 28Y""

(Colvocoresses ct al. 1974). Engel (1977) examined the acclimation abilities of the blue crab and C. similis to osmoregulate in

salinities of 5, 20, and 35%". In agreement with distribution in

natural waters, hemolymph and muscle concentrations of Na + ,

CI-, and K + indicated that adult C. sapidus is better adapted for

ionic regulation than adult C. similis at lower salinities (e.g., 5:100).

Callinectes .nmilis had significantly lower hemolymph Na+ and

Cl- concentrations than C. sapidus at 5%" . Accl imation at the two

higher salinities (20 or 35%,,) resulted in similar hemolymph Na+

and Cl- concentrations for both species. Engel and Eggert (1974)

showed that ovigerous and nonovigerous adult female blue crabs

have lower respiration rates than adult male blue crabs at a salinity

of 5Yoo during summer.

The physiological mechanism allowing blue crabs to hyperosmoregulate was discussed by Mangum and Towle (1977) based

upon several experimental studies. Gilles (1973) reported increased

14C02 production as a result of amino acid catabolism, following

120 min exposure of isolated axons of blue crabs to salines consisting of one of two salinities containing one of several uniformly

labelled (U_14 C) amino acids (arginine, alanine, leucine, glutamate,

aspartate, or serine). Oxygen consumption also increased in the

diluted medium (850 mOsm/l) compared with seawater (1,100

mOsm/l). Weiland and Mangum (1975) reported that decreased

salt concentrations of the blood in blue crabs are accompanied by

increases in blood pH. Thereafter, Mangum et al. (1976) demonstrated that male blue crabs (100 to 235 g) produced higher amounts

of ammonia (and subsequent higher pH due to combination of NH 3

+ H+) when acclimated to salinities of 28 or 5:100 compared with

crabs acclimated to 35%". This is probably the result of increased

amino acid catabolism which occurs when blue crabs are located in

salinities lower than that of their blood (Gerard and Gilles 1972;

Gilles and Gerard 1974). An enzyme believed to be involved in the

"salt pump" of the blue crab, Na + + K+-dependent ATPase is

responsive to increased Na+ uptake in blue crabs migrating
upstream (Mangum and Towle 1977). Specific actIVIty of this

enzyme in gill microsomes from crabs at a salinity of 5roo was nearly

twice the value for crabs acclimated to a salinity of 34roo . Mangum

and Towle (1977) suggested that an increase in blood NH4+ might

increase "extrusion" of Na+ from the gill cell into the blood.

Conversely, Kormanik and Cameron (1981) proposed that ammonia excretion (free base, NH3 ) in adult blue crabs occurs via passive

diffusion in blue crabs adapted to 28 to 30roo salinity. These investigators demonstrated that the net excretion rate of ammonia was

unchanged by sodium transport inhibitor amiloride or by transfer

to Na+- and K+-free artificial seawater. Cameron (1979) reported

that 33 to 50% of sodium effluxes and 50 to 70% of chloride

effluxes from adult blue crabs in freshwater resulted from exchange diffusion, with sodium and chloride transport pr.)bably

occurring via largely independent, electroneutral processes.

Hemolymph composition

Gleeson and Zubkoff (1977) estimated hemolymph volume

(25.4± 1.4 ml at 95% confidence intervals) of 9 penultimate

instar females weighing 98.2 ± 6.2 g (standard deviation) in the

intermolt-white sign molt stages (C4 -D\).

An ultrastructural study of hemocytes prompted Bodammer

(1978) to suggest that granulocytes, hyaline cells, and intermediate

cells represent different stages of cytogenesis along a path of

cellular differentiation. Granulocytes were broadly oval, less

elongate (13.4 to 15.7 /Lm), and wider (6.8 to 9.3 /Lm) than intermediate cells. Hyaline cells were slightly round to oval, 7 to 13/Lm

diameter with centrally located nuclei. Intermediate cells were 13.5

to 19.5 /Lm long and 6.3 to 9.8 /Lm wide with eccentric nuclei.

Lynch and Webb (l973a) found that serum protein of intermolt.

adult males (20-80 mg/ml) and females (35-100 mg/ml) did not

differ significantly. Temperature and salinity did not appear to

affect serum protein concentration. Total serum protein increased as

ovary weight increased. After egg transfer from the ovaries to the

pleopods, a slight decrease in serum protein occurred.

Serum glucose did not differ between males (2.3 to 125.5 mg/

100 ml) and females (1.3 to 305.3 mg/ 100 ml), nor did it vary with

reproductive stage of females (Lynch and Webb 1973b). However,

molt status of the crabs examined was not monitored. Serum

glucose levels were lowest in late summer/early fall in males and

"old year class" females. Glucose concentrations rose to yearly

high levels during spring and early summer (Lynch and Webb

1973b).

Serum zinc and copper of adult blue crabs have been found to be

unrelated to environmental temperature and salinity (Colvocoresses

and Lynch 1975). Serum copper (ranging from 35 to 150 ppm) was

directly related to total serum protein; probably as a result of copper

being a component of hemocyanin. Copper content of blue crab

serum ranged from 10.7 to 16.2 mg/ 100 ml (N = 6 at 22 roo sal inity)

(Mangum and Weiland 1975). Serum zinc (6 to 22 ppm) was also

strongly related to total serum protein, possibly as a result (of zinc

being a cofactor for carbonic anhydrase.

Serum calcium concentrations of females in their penultimate

instar were found to be similar regardless of salinity (10, 20, or

30%.) within anyone molt stage (Haefner 1964). Late premolt

females had 500 to 700 ppm serum calcium, early postmolt females

had 250 to 300 ppm serum calcium, and intermolt females had 420

to 450 ppm serum calcium (Haefner 1964).

6

1.37 Internal anatomy

The digestive, muscular, reproductive, circulatory, respiratory,

excretory, and nervous systems, and sense organs of the blue crab

were described by Pyle and Cronin (1950). Additionally, Cochran

(1935) described the skeletal musculature. Descriptions of ovarian

tissue at various stages of sexual maturity include those by Cronin

(1942), Hard (1942), and Johnson (1980). A detailed account ofthe

male reproductive system was presented by Cronin (1947). Anatomy and neurophysiology of the fifth pair of pereopods (swimming

legs) were described by White and Spirito (1973). Johnson (1980)

presented a detailed account of the histology of various blue crab

tissues and cell types including connective tissue, cuticle, epidermis, gills, antennal gland and bladder, central nervous system,

the gut, circulatory system, hemopoietic tissue and hemocytes,

phagocytes, and male and female reproductive systems. Ultrastructural features of the myocardium were described by Hawkins

et al. (1977). Special features discussed were components of the

myocardial cell, structure of the intercalated discs, and organization

of the neuromuscular junctions.

Gill morphometry ofthe blue crab was described by Aldridge and

Cameron (1982). Two types of gill epithelial surface included the

site presumed to be for respiratory exchange (0.5 /Lm in thickness)

and the primary site for active ion transport (averaging 10 /Lm

thickness).

An anatomical and cytological description of the mandibular

gland of the bl ue crab was presented by Yudin et al. (1980). The

authors believe that this gland has an endocrine function, being

actively involved in steroid production.

2 DISTRIBUTION

2. 1 Total area

According to Williams (1974), the original range of C. sapidus

was from Nova Scotia, Maine, and northern Massachusetts to

northern Argentina, including Bermuda and the Antilles. However,

the species has been introduced into several parts of Europe-

.Dresund. Denmark; the Netherlands and adjacent North Sea,

southwest France; Gulf of Genoa; northern Adriatic; Aegean,

western Black, and eastern Mediterranean Sea. Recently, the species has also been reported from Lake Hamana-Ko, central Japan

(summarized by Williams 1974, in press).

Williams (1974) recognized 14 species of Callinectes, nine

distributed in the western Atlantic, three in west Africa (only two of

these species are different than those occurring in the western

Atlantic), and three in the eastern tropical Pacific. Since that time

another species has been described from northern Brazil (FaustoFilho 1980) and Manning and Holthuis (1981) have revised the

African species, disclosing older names as well as separating the

amphi-Atlantic species into closely related eastern and western

Atlantic species.

Currently, there are 16 recognized species of Callinectes as listed

under geographic regions below. Almost all records are from

shallow coastal waters of continents or islands.

Western Atlantic

Callinectes affinis Fausto-Filho 1980. Coco River, Fortaleza,

Ceara, Brazil (Fausto-Filho 1980).

Callinectes bocourti A. Milne Edwards 1879. New Hanover Co.,

N.C., Mississippi, and British Honduras through West Indies
and Caribbean Sea to Estado de Santa Catarina, Brazil (Williams

1974; Williams and Williams 1981).

Callinectes danae Smith 1869. Bermuda and New Hanover Co.,

N.C.; southern Florida and eastern side of Yucatan Peninsula,

West Indies, and Caribbean Sea to Estado de Santa Catarina,

Brazil (Williams 1974; Perschbacher and Schwartz 1979).

Callinectes exasperatus (Gerstaecker 1856). Bermuda, northeastern Florida, Florida Keys, and eastern Yucatan through West

Indies and Caribbean Sea to Estado de Santa Catarina, Brazil.

with exception of no records from the Guianas and northern

Brazil (Williams 1974, in press).

Callinectes larvatus Ordway 1863. Bermuda and North Carolina,

through Florida Keys, West Indies, and continental perimeter of

Caribbean Sea to Sao Francisco, Brazil (Williams 1974; Manning

and Holthuis 1981).

Callinectes maracaiboensis Taissoun 1969. Lake Maracaibo and

to unknown extent in West Indies region, including Jamaica

(Williams 1974; Norse 1978).

Callinectes ornatus Ordway 1863. Bermuda and off Cape Charles,

Va., through southwestern Florida, northwestern Yucatan, West

Indies, and southern Caribbean Sea to Estado de Sao Paulo,

Brazil (Williams 1974, in press).

Callinectes rathbunae Contreras 1930. Gulf of Mexico from mouth

of Rio Grande, Texas-Mexico border to southern Veracruz

(Williams 1974).

Callinectes sapidus Rathbun 1896. See beginning of section 2. I.

Callinectes similis Williams 1966. Eastern United States and Gulf

of Mexico from eastern Long Island Sound, Conn., to off

Campeche, Yucatan, Mexico, and Jamaica (Williams 1974,

in press).

Eastern Atlantic

Callinectes amnicola (De Rochebrune 1883) (= C. latimanus

Williams 1974). West Africa from Baie de Saint-Jean, Mauritania, to at least as far south as a lagoon north of Luanda, Angola

(Manning and Holthuis 1981).

Callinecles marginatus (A. Milne Edwards 1861). West Africa

from Cape Verde h., and Port Etienne, Mauritania, tu central

Angola (Williams 1974; Manning and Holthuis 1981).

Callinectes pallidus (De Rochebrune 1883) (= C. gladiator

Williams 1974). West Africa from Baie de Saint-Jean, Mauritania, to Baia do Lobito, Angola (Williams 1974; Manning and

Holthuis 1981).

Eastern Pacific

Callillecles arcuatus Ordway 1863. Los Angeles Harbor, Calif., to

southern Peru and Galapagos Islands (WilliJms 1974).

Catlinecte~ betlicosus (Stimpson 1859). San Diego, Calif., to

southern Magdalena Bay; La Paz Harbor around Gulf of

California to Topolobampo, Sinaloa, Mexico; apparently absent

from extreme southern tip of Baja California (Williams 1974).

Callinectes toxotes Ordway 1863. Cape San Lucas, Baja Calif.

(historical record only) to extreme northern Peru; extraterritorial,

Juan Fernandez Islands (Williams 1974).

2.2 Differential distribution

2.21 Larvae and juveniles

Early zoeal stages of the blue crab are located in surface waters

7

(Tagatz 1968a; Dudley and Judy 1971; Sandifer 1973; Dittel and

Epifanio 1982) of high salinity, usually 20Yoo or greater (Perry

1975). Zoeae are planktonic and positively phototropic (Costlow et a!. 1959). According to Norse (1977), "the occurrence of

[various] Callinectes [sp.] in a geographic area [usually] implies

settlement from plankton, rather than adult immigration." However, juvenile migrations from high salinity to low salinity areas are

important in determining annual recruitment to brackish water areas

( Van Engel 1958; Sulkin 1977). Nichols and Keney ( 1963) sampled

Callinectes sp. larvae offshore between Cape Hatteras, N.C., and

Jupiter Light, Fla. Large numbers of early stage zoeae were collected nearshore with progressively later zoeal stages found 20 to

40 mi (32 to 64 km) offshore. Highest frequency of megalopae

occurred beyond 40 mi (64 km) offshore. During a 6-mo sampling

period (May to October 1978), first stage blue crab zoeae were most

abundant in 5 or 11 m depths in Delaware Bay as opposed to 20 m

(Dittel and Epifanio 1982). Sandifer (1973) reported Callinectes

sp. larvae from daytime samples taken from Chesapeake Bay I m

below the surface and near bottom from June to November, in

salinities ranging from 15.8 to 32.4%". Most of the larvae (85'1c)

were collected in salinities of 20 to 30%" just outside the mouth of

Chesapeake Bay. Most zoeae occurred in the near surface (I m

depth) samples, whereas megalopae were found only in a few

bottom samples (Sandifer 1975). Conversely, Williams (1971)

and Smyth (1980) reported that megalopae of Callinectes sp.

were common in surface samples at night. An extensive monthly

quantitative sampling of portunid larvae in Mississippi coastal

waters from July 1974 to September 1979 indicated that blue crab

megalopae were about equally distributed in surface and bottom

waters (Stuck and Perry 1981).

Factors that affect vertical distribution of zoeae and megalopae

include light intensity, swimming rate, sinking rate, and barokinetic

and geotactic behavior. Also, vertical di~tribution of blue crab

megalopae may be dependent upon base activity levels which may

be rhythmic due to exogenous or endogenous control (Sulkin et al.

1979; Sulkin and Van Heukelem 1982). Smyth (1980) suggested

that diel increases in abundance of megalopae of Callinectes sp. in

surface waters at night is a negative phototropic response. Laboratory studies with blue crab megalopae have indicated a low

threshold in barokinetic response (0.4 atm), such that megalopae

would swim a short dbtance from the bottom when subjected to

artificial pressure increases from above (Naylor and Isaac 1973).

These authors speculated that such a barokinetic response would

inhibit swimming in surface waters of stratified estuaries where

net flow often is seaward, while promoting occurrence in deeper

waters which often have a net flow upstream. However, data from

previously described sampling studies (Williams 1971; Smyth

1980), suggest that this phenomenon of vertical distribution, with

megalopae ab,ent from surface waters, may only occur during

daylight. Sulkin et a!. (1980) evaluated swimming rate and

geotactic and barokinetic responses of blue crab zoeal stages I, IV,

and VII. Negative geotaxis (upward movement) occurred in stage I

zoeae, a period of transition between negative and positive geotaxis

(downward movement) was indicated in stage I V zoeae, and positive geotaxis was indicated in stage VII zoeae. Geotactic behavior

may explain why Sandifer (1975) did not obtain any zoeal stages

later than stage IV in surface samples (I m depth) during daylight.

Although sinking rate increased 3.2 fold between the first and

seventh zoeal stages, swimming rate increased 4.4 fold over the

same developmental period; therefore, Sulkin et al. (1980) concluded that sinking rate was not responsible for the geotactic

behavior that was measured.
Distribution and recruitment of blue crabs is also affected by

temperature-salinity interactions on the megalopal stage. For

megalopae maintained at 15°C, average duration of the megalopa

stage was found to increase as salinity increased (e.g., 15°C,

20%, = 34 d before metamorphosis; 15°, 40Y", = 58 d before

metamorphosis) (Costluw 1967). Therefore, Costlow and Bookhout (1969) suggested that transport of megalopae to waters below

20° C and high salinities may retard further development to the first

crab stage, until megalopae are transported to lower salinities,

possibly as an adaptive behavioral mechanism to increase recruitment. Perry (1975) reported the occurrence of Callinectes sp.

megalopae in salinities as low as 4Y"" while most megalopae

occurred in salinities> 20Y",.

Following larval development, early juvenile stages migrate to

lower salinity and shallow waters during summer months; later they

move to slightly deeper channels or hibernate during colder months

when growth ceases or decreases appreciably in Chesapeake Bay

(Van Engel 1958) and Delaware Bay (Cronin 1954). Juvenile blue

crab distribution in Mississippi waters was as follows: 1) First and

early crab stages (3 to 10 mm) occurred most often in 15 to 20Y""

2) 10 to 20 mm juveniles were most frequently found in salinities

< lOY"" and 3) maximum number of crabs (20 to 40 mm) were

sampled from salinities below 5Y", (Perry and Stuck 1982). Juvenile

and/or adult blue crabs occur in much higher densities in areas

covered by eelgrass, Zostera marina, than on unvegetated bottoms

(Heck and Orth 1980).

2.22 Adults

See section 3.51.

2.4 Hybridization

2.41 Hybrids

None reported.

3 BIONOMICS AND LIFE HISTORY

3. I Reproduction

3.11 Sexuality

Blue crabs are heterosexual and shuw obvious st'xual jimor·

phism. Immature females have a triangular-shaped abdomen,

whereas mature females have a broader, semicircular-~.hapcd

abdomen (Fig. 2). Males have a T-shaped abdomen readily distinguishable from either female stage (Fig. 2). Typical culoration

differences between adult males and females ar~ d~scribed in

section 1.21. Histology and external morphology of two bisexual

crabs (bilateral gynandromorphs) from separate areas of Chesapeake Bay have been described by Cargo (1980) and Johnson and

Otto (1981). Johnson and Otto (1981) indicated that few sp~rm

were in the seminal receptacle of their specimen, indicating that

copulation had not been completely successful.

3.12 Maturity

Male blue crabs reach maturity after 18 or 19 postlarval molts,

whiIe females reach maturity after 18 to 20 postlarval molts (Van

Engel 1958). The abdomen of immature males is tightly sealed to

8

the ventral surface, whereas the abdomen of mature males hangs

free or is held in place by a pair of "snap-fastener-like" tubercles

(Van Engel 1958). Tagatz (1968a) used the followi ng criteria to

identify immature and mature males: Immature males have vasa

deferentia that are small and the middle vas deferens is white, while

mature males have vasa deferentia with large prominent ducts and

the middle vas deferens is bright pink. However, Johnson (1980)

observed that the middle vas deferens may be white or only pale

pink following copulation, while many large, mature male crabs

sampled during winter have a white middle vas deferens. Gray and

Newcombe (1938b) approximated the average size at which males

attain maturity as 89 mm carapace width, since males attain their

highest growth increment per molt at this size. As the terminal molt

(maturity molt) in females approaches, dark coloration of the

mature, semicircular abdomen shows through the translucent,

triangular shaped, immature abdomen within 6 d of the terminal

molt (Gleeson 1980). Cronin (1942) and Johnson (1980) described

histological changes in female ovarian tissue as maturation approaches, and Hard (1942) described ovarian tissue changes before,

during, and after the first and second spawnings.

Time to maturity from hatching is reduced by longer growing

seasons varying from 10 to 12 mo in Mississippi (Perry 1975) or

Florida (Tagatz 1968a) to 12 to 20 mo in Chesapeake Bay (Van

Engel 1958; Lippson 1973). Due to semihibernation in winter,

crabs hatched in early summer in Chesapeake Bay reach sexual

maturity within 12 to 16 mo, while individuals that are hatched in

late summer and early fall reach sexual maturity within 18 to 20 mo

(Lippson 1973).

3.13 Mating

Unlike males. which may mate several times, female blue crabs

mate only once, while in the soft shell stage (AJ-see section 3.44

for molt stage descriptions) following their terminal molt (maturity

or pubertal molt). According to Van Engel (1958), mating may

occur day or night and may last for 5 to 12 h. Ablation of the outer

flagella of antennules of males in laboratory studies indicated that

these portions of the antennules are probably primary chemoreceptors for a fcmale pheromone that triggers male reproductive

behavior (Gleeson 1980). The pheromone is believed to occur in the

uri ne of pubertal females. Further work indicated that the aesthetasc

tuft on the outer flagellulll of the antennule ofthe male is responsible

for detection of pheromones produced by females (Gleeson 1982).

Olfactory stimuli are of primary importance in the ability of a red

sign pubertal molt female to discriminate sexes prior to mating

(Teytaud 1971). Still, a visual stimulus (e.g., a male model)

combined with olfactory stimuli (culture water from a mature male)

provided more rapid sex recognition. White-sign, pubertal molt

females did not respond to the aforementioned stimuli. Also,

Teytaud (1971) observed mating activity in mature females in the

intermolt stage if they had not mated at the time of their pubertal

molt in the laboratory. The female is protected by a male before

(Stage D4 ), and after (Stages A and B), her terminal molt by

being grasped by the male's first pair of walking legs, and held right

side up in a "cradle-carry" position under the male (Van Engel

1958; Gleeson 1980; Johnson 1980). The first pair of pleopods of

the male are the functional intromittent organs, each receiving

spermatophore~ and semen from the respective penis. The second

pair of pleopods are inserted into the posterior foramen of the first

pleopods, forcing the spermatophores and semen through the tubelike first pleopods. During copulation, semen and spermatophores

are passed from the first pleopods of the male into the paired
oviducts and eventually the seminal receptacles (spermathecae) of

the female (Cronin 1947). The majority of mating occurs in low

salinity waters since males usually remain in brackish areas during

the adult stage.

3.14 Fertilization

Eggs are fertilized when passing from the ovaries to the seminal

receptacles before being extruded onto the female's pleopods. Egg

extrusion onto endopodites of the female's pleopods may be complete within 2 h (Van Engel 1958).

3.15 Gonads

The male reproductive system consists of paired testes, vasa

efferentia, vasa deferentia, external penes, and highly modified first

and second abdominal pleopods (Cronin 1947). The anterior vas

deferens is the most important storage region for completed

spermatophores. Spermatophores that transport sperm are ovoid

with an average size of 300 x 225 fLm (Cronin 1947). Testes of

immature males < 5 em carapace width contain germi;Jative areas

and spermatocytes, but spermatids and mature sperm are not present

(Johnson 1980). Males larger than 6.5 em carapace width have

spermatids and mature sperm in some testicular lobes (Johnson

1980). Sperm can survive for at least 1 yr in the female's seminal

receptacles (Van Engel 1958).

The female reproductive system consists ofthe paired ovaries and

oviducts, part of which serve as the spermatheca. The most ventral

portion of each ovary joins the spermatheca. An opening in the

posterodorsal surface of the spermatheca permits the passage of

eggs and the oviduct leads to a genital pore on the ventral surface of

the sixth thoracic somite (Pyle and Cronin 1950). Hard (1942j used

histological techniques to verify stages of ovarian growth and

maturation which could be recognized by gross, external appearance. Immediately after copulation the ovary is small and white,

whereas the large, pinkish spermathecae are distended with sperm.

Following copulation, ova require at least 2 mo to develop,

resulting in an increased ovarian size and orange color. Females

may ovulate more than once. Before the first ovulation, follicles are

expanded. After the first ovulation, the ovary remains large and

orange colored and egg shell remnants are on the pleopods after

hatching. After the second ovulation, the ovary is collapsed, gray or

tan colored, and egg shell remnants again appear on the pleopods

after hatching.

3.16 Spawning

Spawning of blue crabs is initiated progressively earlier in the

spring at lower latitudes. Typically, blue crabs from Chesapeake

Bay spawn in Mayor June. followed by a second spawning in

August (Van Engel 1958). Spawning in North Carolinian waters

occurs from mid-March to October (Williams 1971), with peak

spawning in Beaufort Inlet, N.C., from .June to August (Dudley and

Judy 1971). Ovigerous females occurred from March through

September in southern South Carolina waters, with peak occurrence

in April (Eldridge and Waltz 1977). In St. Johns River, Fla.,

spawning occurs from February through October, with peak

occurrence from March to September (Tagatz 1968a). In Lake

Pontchartrain, La., two peak spawning periods for blue crabs are

February and March, followed by August and September (Jaworski

1972), with similar spawning periods in Mississippi (Perry 1975).

9

The primary spawning ground along the Gulf coast of Florida is

located in Apalachicola Bay (Oesterling 1976).

Sulkin et al. (1976) successfully induced ovarian maturation and

spawning in laboratory cultured female crabs following their

capture during hibernation in winter in Chesapeake Bay. Ovarian

development progressed appreciably more rapidly over a 2.5 mo

period at 19° C than at 15°C.

3.17 Eggs

Blue crabs generally produce between 1.75 x 106 and 2x 106

eggs per spawning (Churchill 192 I; Graham and Beaven 1942; Pyle

and Cronin 1950; Van Engel 1958). Truitt ( 1939) reported that egg

production during a single spawning ranged from 723,500 to

2,173,300, but total number of females examined and possible

reasons for variation of fecundity were not discussed. No publ ished

data are available concerning fecundity as a function of female

carapace width, length, or body weight. Additionally, no data have

been reported on relative amounts of eggs spawned from the first

and second spawnings from the same individual in the laboratory.

The fertilized eggs extruded from the oviduct are distributed on the

setae of the endopodites of all four pairs of pleopods (Pyle and

Cronin 1950). Approximate ages of blue crab egg masses (sponges)

according to color have been assigned as follows: Yellow to orange

= I to 7 d since extrusion; brown to black = 8 to 15 d since extrusion (Bland and Amerson 1974). According to Roberts and Leggett

(19~0), a female blue crab produces approximately 30 g (wet

weight) of eggs in a single spawning. Tagatz (1965) recorded egg

mass (sponge) weights of females ranging from 24 to 98 g with an

average value of 37 g.

Total sterol concentration of eggs does not change appreciably

during their development; however, the percent esterified f0rm of

,terol decreases appreciably Whitney 1969). Cholesterol was

determined to be the dominant sterol in both free and esterified

forms. Palmitic acid and stearic acid were the predominant free fatty

acids of blue crab eggs. Steryl esters contained large amounts of

palmitic, palmitoleic, and oleic acids. Whitney (1970) concluded

that the blue crab requires an exogenous source of squalene and

sterols, since they were not synthesized from radioactive labelled

acetate-I-t. Cor mevalonate-2-J4 C by ovaries or eggs.

Lipid-rich blue crab eggs might serve as a clearance route for

toxic, lipophilic compounds. Eggs of blue crabs from selected areas

near the James River, Va., had two to three times more Kepone (0.1

to 0.15 fLg Kepone per g eggs) than that occurring in backfin muscle

(Roberts and Leggett 1980). Nevertheless, further research should

examine comparative amounts of Kepone in the hepatopancreas of

males and females, before conclusions can be drawn concerning the

importance of eggs as a clearance route for toxic, lipophilic

compounds. lt is unknown to what degree hatchability of eggs or

viability of larvae may be affected by this contaminant (Roberts and

Leggett 1980).

3.2 Preadult phase

3.21 Embryonic phase

Hatching occurs between 14 and 17 d following spawning (egg

extrusion) onto the pleopods at 26° C and between 12 and 15 d at

29°C (Churchill 1921). Hatching occurs in high salinity waters

(ca. 23 to 301.,.) near river mouths, inlets, and coastal areas.

Extruded eggs in the early stages of development are 273 i< 263

fLm, whereas, just before hatching, eggs are larger (320 x 278 fLm)
(Davis 1965). In laboratory studies, successful hatching never

occurred at the next lowest experimental salinity (15.6Yoo) below

20.1 %" (Costlow and Bookhout 1959). All successful hatchlings

were first lOeal stage larvae and no prezoeae were observed

(Costlow and BookhoUI 1959). Similarly, successful hatching

occurred in salinities of 18 and 26%. in a separate study (Davis

1965). Davis (1965) observed blue crab larvae escaping the egg's

inner membrane as prezoeae, with subsequent prezoeal ecdysis

within I to 3 min. Davis (1965) suggested that hatching is promoted

by osmotic swelling of the inner membrane which ruptures the

chorion, followed by rupture of the inner membrane through

mechanical action of the prezoea.

3.22 Larval phase

Development and environmental requirements

Newly hatched blue crab larvae normally develop through

seven lOeal stages before transforming to megalopae (Costlow et al.

i 959; Costlow and Bookhout 1959), but have successfully metamorphosed to the megalopal stage after only six lOeal stages (Sulkin

et al. 1976) or occasionally after an eighth lOeal stage in the

iaboratory (Costlow and Bookhout 1959; Costlow 1965). Sandoz

and Rogers ( 1944) described prezoeae as lacking a dorsal spine,

retaining an embryonic cuticle, having an undeveloped telson, and

,ettling to the bottom without successful metamorphosis. Duration

of lOeal development (7 stages) in the laboratory at 25°C and a

salinity of 26Yoo , ranged from 31 to 49 d (Costlow and Bookhout

1959), whereas the average duration of lOeal development that

included only six lOeal stages before the megalopa was 35.7 d

(Sulkin et al. (976). Average duration for complete lOeal development (7 stages) ranged from 32 to 43 d in control groups of four

separate chemical contaminant studies (Bookhout and Costlow

1975; Bookhout and Monroe 1977; Bookhout et al. 1976, 1980).

External morphology of the first and second weal stages and third

and fourth lOea: stages was described by Hopkins (1943, 1944,

respectively). Thereafter, description of all seven lOeal stages plus

a supernumerary eighth weal stage and the megalopa of hatcheryraised crabs was recorded by Costlow and Bookhout (1959) (see

Figs. 3A-1OA, 3B-1OB). Mortality of lOeallarvae in the laboratory

is usually highest during the first two lOeal stages. The optimal

salinity and temperature combination for lOeal development was

30%, and 25°C, respectively (Costlow and Bookhout 1959; Sulkin

and Epifanio 1975; Bookhout et al. 1976). Optimal salinity and

temperature for the megalopal stage of development of laboratory

reared blue crabs was 30%0 and 25°C, with an average duration of

8.4 d and a range of 6 to 12 d (Costlow 1967). Mean total length

and selected morphological characteristics for the various larval

stages are Iisted in Table I.

Costlow (1965) observed a high degree of variability in several

weal stages of laboratory reared crabs. During zoeal stage~. [-IV,

morphological characteristics were the same as those reported by

Costlow and Bookhout (1959). Costlow (1965) found that in later

stages some molts were not accompanied by any morphobgical

change, a molt stage was omitted entirely, or a molt resulted in a

larva with some characteristics from the previous stage that are

usually not retained, as well as some new morphology. Stage~, V- VI

and VI-VII were the most frequent combined stages with abdcminal

segments and development of pleopod buds more similar to the

previous stage and the more advanced characters usually in the

anterior portion of the larva.

10

Laboratory studies demonstrated that if a megalopa lost a

cheliped prior to day 4 following the final lOeal molt, then a

completely regenerated appendage usually appeared after metamorphosis from megalopa to first crab stage (Costlow 1963a).

Chelipeds lost after the fourth day of the megalopal stage were not

regenerated until the first or second postlarval molt. Removal of

both eyestalks from megalopae within 12 h following the final weal

molt accelerated metamorphosis to the first crab stage and increased

the size of postlarval crabs (Costlow 1963b). Removal of both

eyestalks I to 5 d following metamorphosis did not affect duration

of the megalopal stage. Therefore, molt inhibiting hormone of

the X-organ may not be secreted in megalopae more than I d old

(Costlow 1963b),

A comparative description of larval development ofspecimens of

the blue crab and the lesser blue crab, Cullinectes similis. from

North Carolina waters indicated several minor differences in

morphometry between the species which could aid in identification

of larval field samples (Bookhout and Costlow 1977). The dorsal

spine of C. similis was longer than that of C. sapidus for each weal

stage. The rostrum and antennae were usually significantly longer

in later lOeal stages of C. similis compared with those of C.

sapidus. Usually, there were more short spines on the telson from

lOeal stages IV-VIII in C. similis. Also, there were usually more

natatory setae on the exopodites of the maxillipeds in stages V to

VIII of C. similis. The megalopa of C. similis had a shorter

carapace and longer rostrum and antennae than those of C, sapidus.

These characters were not useful for identification of C. sapidus

and C. similis from Mississippi waters (Perry and Stuck 1982),

possibly due to morphometric variation in different geographical

regions.

Laboratory culture

Several proven laboratory methods exist for successful hatching

of blue crab eggs. Lochhead and Newcombe (1942) reported 90%

hatching success when 8 eggs/cm2 were reared in white, enamel

pans (20 x 26 x 6 cm) in natural water (19 to 21 Yoo salinity, 24° to

27°C). Sandoz and Rogers (1944) observed prezoeae at hatching in

salinities of 10 to 22%" whereas no prezoeae were observed in crabs

hatched in 23.4 to 32%. salinity. Also, eggs hatched successfully in

temperatures from 19° to 29°C. Costlow and Bookhout (1960) cut

with fine scissors eggs from previously detached pleopods maintained in seawater of 30%. salinity and 200

to 30°C, and further

dissociated the eggs with glass needles into groups of 100 to 1,000.

Groups of eggs were placed in compartmented boxes (9 cm2 )

containing 20 ml of freshly filtered seawater treated with penicillin

(200,000 units/I). The boxes were then placed on a variable speed

shaker (110 to 120 oscillations/min) until hatching. More recently,

Bookhout and Costlow (1975) placed setae bearing black eggs (eyes

and a visible heartbeat) into plastic compartmented boxes (32.5 x

227 cm) in seawater of 30%0 salinity and 25°C, on a variable speed

shaker for 60 oscillations/min until they hatched. Sulkin et al.

(19801 removed "black eyespot stage" eggs from female blue

crabs, culturing sets of approximately 500 eggs in 50 ml ofseawater

(30%" salinity, 20 0-3U°C) which also contained penicillin (60

mg/I) plus streptomycin (50 mg/I) or chloramphenicol (5 mg/I) to

improve egg and larval survival. Maximum survival rate for blue

crabs reared from newly hatched first stage lOeae to first crab stage

in the laboratory has been 40% (Costlow4 ).

IJ. D. Costlow. Duke University Marine Laboratory, Pivers Island, NC 28516,

pers. commun. November 1982.
3A

t------i

0.1 Il1l1

>---------l

0.1 Il1l1

Figure 3.-Side (A) and front (B) views of first zoeal stage of Callinectes sapidus (from Costlow and Bookhout 1959). Figure 4.-Side (A) and

front (B) views of second LOeal stage of Cal/inectes sapidus (from Costlow and Bookhout 1959). Figure 5.-Side (A) and front (B) views of third

zoeal stage of Cal/mectes sapid"s (from Costlow and Bookhout 1959).

11
Chemical toxicity

'From Bookhout and Costlow (1977).

2 From Costlow and Bookhout (1959).

Figure 6,-Side (A) and front (B) views of fourth zoeal stage of Calliuectes

sapidus (from Costlow and Bookhout 1959). Figure 7,-Side (A) and front (B)

views of fifth zoeal stage of Calliuectes sapidus (from Costlow and Bookhout

19591, Figure S,-Side (A) and front (B) views of sixth zoeal stage of Calliuectes

sapidus (from Costlow and Bookhout 1959).

salinity (10, 20, 30, or 40;:0). Also, average duration of the

megalopal stage was significantly longer for individuals exposed to

10 and 20 ppb Hg compared with crabs not exposed to Hg.

Methoxychlor (1, 1, l-trichloro-2, 2-bis (p-methoxyphenyl)

ethane) concentrations above 1 ppb (e.g., 1.3 and 1.9 ppb) were

acutely toxic to blue crab zoeae, with survival being rare after the

third zoeal stage (Bookhout et al. 1976). Average duration from

hatching to first crab increased in individuals exposed to 0.7 or 1.0

ppb methoxychlor compared with larvae in the control treatment.

Blue crab zoeae exposed to 20 ppb malathion (0, O-dimethyl

phosphorodithionate of diethyl mercaptosuccinate) had reduced

survival rate (33%) to the first crab stage and longer average

duration (48.6 d) to the first crab stage compared with larvae not

exposed to malathion (54% survival and 44.5 d average duration

to first crab stage) (Bookhout and Monroe 1977). Acute toxic

concentrations (80 ppb or more) caused 10o/c or less survival

of blue crab larvae to the first crab stage and increased duration

of zoeal and megalopal development. Mirex (dodecachlorooctahydro-I, 3, 4-metheno-2H-cyclobuta (cd) pentalen-2-one) concentrations of I and 10 ppb were acutely toxic to blue crab zoeae,

wh~reas 0.01 and 0.1 ppb mirex were sublethal concentrations

(Bookhout and Costlow 1975). Kepone (decachlorooctahydro-l,

3, 4-metheno-2H-cyclobuta (cd) pentalen-2-one) concentrations

of 0.5 and 0.75 ppb were sublethal to blue crab zoeae (increased

duration between hatching and metamorphosis), whereas 1.0 ppb

caused a reduced survival rate of 5% to the first crab stage compared with 22% in the control group (Bookhout et a1. 1980). A

juvenile hormone mimic, MONO-585 (2,6-di-t-butyl-4-(a,adimethyl benzyl) phenol) was acutely toxic (100% mortality) to blue

crab megalopae exposed to 10 ppm (Costlow and Bookhout 1979).

Also, concentrations of I ppm MONO-585 reduced ~urvival to the

first crab stage from 100 to 40% at a constant temperature of 25°C

combined with a salinity of 20 or 35Yoo • Culturing megalopae in

5°C, 24 h cyclic temperature regimes (20° to 25°c' 25° to 30°C,

or 30° to 35°C) did not result in any significant change in survival

of the control group or megalopae exposed to O. I ppm MONO585, whereas some reduction in survival occurred in mcgalopae

exposed to 1.0 ppm MONO-585 in each cyclic temperature

regime. Another juvenile hormone mimic, methoprene (isopropyl

ll-methoxy-3, 7, II-trimethyl-dodeca-2, 4-dienoate), showed no

toxicity to megalopae exposed to 0.1 or 0.01 ppm in salinities of

15,25, and 35Y"" combined with either 25° to 30°C or 30° to 35°C,

24 h cyclic temperatures (Costlow and Bookhout 1979). Survival

was reduced slightly for megalopae exposed to 0.1 ppm methoprene at a cyclic temperature of 20° to 25° C at all salinities.

Costlow (1979) evaluated the effects of Dimilin (diflubenzuron),

an insect growth regulator, on average duration of the megalopal

stage of blue crabs and survival rates of megalopae to the first crab

stage. Concentrations of 0, 0.1, 0.3, 0.5, 1.0, 3.0, and 6.0 ppb

were combined with one of six different temperature (20°, 25°, or

30°C) and salinity (20 or 30%,) combinations. Percentage survival

for megalopae reared in 0.5 or 1.0 ppb Dimilin combined with

20° C plus 20 or 30Y"" was similar to that in the control series.

However, 3 or 6 ppb Dimilin combined with 20°C plus 20 or

30/;~ caused 95 or 100% mortality of megalopae. All salinities

Characteristics 2

Table 1.-Selected characteristics of blue crab larval stages of development.

Mean total

length

(mm)'

Zoea 1.0 Five abdominal segments plus telson. Eyes are not

stalked.

II 1.3 Eyes stalked. Third segment of endopodite of first

maxilliped with one additional spine.

III 1.4 Mandible with several small teeth in addition to broad

cutting surface. Sixth abdominal segment is present.

JV I.7 Slight swelling in basal region of antenna indicates

beginning of endopodite bud.

V 2.2 Developing endopodite bud larger than in stage IV.

Buds of3rd maxilliped, chela, and pereopods visible

beneath carapace.

VI 2.3 Pleopod buds appear on abdominal segments 2 through

6 for first time.

VII 2.8 Terminal aesthetes of antennule increase to 7;

5 subterminal aesthetes added. Basal portion

of antennule is swollen.

Megalopa 3.0 Rostrum pointed, longer than antennule, but shorter

than antennae. Dactylopod of first pereopod chelate.

Developmental

stage

Toxicities of the dissolved phase of cadmium and mercury,

several chlorinated hydrocarbons, and juvenile hormone mimics or

insect growth regulators have been determined for several blue

crab larval stages. Megalopae reared at optimal temperature and

salinity survived for I to 6 d with no successful metamorphosis to

the first crab stage when exposed to 250 ppb cadmium in tht; form

of Cd(N03h (Rosenberg and Costlow 1976). Also, a ;;ignificant

delay in development from the megalopa to third juvenile crab

stage and reduced survival rate occurred within each salinity (10,

20, 30, and 40%0) at 50 ppb cadmium compared with 0 ppb

cadmium. Blue crab megalopae had similar survival rates when

cultured at 25°C in salinities of 20,30, or 40%, containing either 5

or 10 ppb mercury (HgCI 2 ) (McKenney and Costlow 1981).

Significant reduction in survival occurred in megalopae exposed to

5 or 10 ppb Hg in a salinity of lOY"". Megalopae exposed to 20 ppb

Hg experienced significant increases in mortality regardless (If

Results of several studies examining the effects of various live

food organisms separately or in combination on survival rate and

duration of blue crab larval development were reviewed by Millikin

(1978). Briefly, it was recommended that the first two blue crab

zoeal stages be fed sea urchin, Arbacea punctulata, embryos, plus

freshly hatched brine shrimp, Artemia salina, nauplii, preferably

< 12 h old. Since energy content decreases and ash increases over

time in starved Artemia nauplii (Claus et al. 1979), feeding freshly

hatched nauplii to crab zoeae prevents the need to culture Artemia

nauplii over prolonged periods. Subsequent zoeal stages and the

megalopa can be reared entirely on brine shrimp nauplii. Sulkin

(1978) recommended feeding rotifers, Brachionus plicatilis, to

blue crab larvae for the first 14 d of zoeal development, followed by

brine shrimp nauplii beginning with the 15th day of development.

Mass culture of blue crab zoeae (approximately 1,000) has been

ac,~omplished in large finger bowls (19.4 em diameter) containing

700 ml of freshly filtered seawater (3D%. and 2YC) which was

replaced daily (Bookhout and Costlow 1975). Daily rations consisted of one medicine dropper full of A. punctulata embryos for

the first two zoeal stages and a similar volume of brine shrimp

nauplii for the la;;t five zoeal stages.

12
13
o.lrrm

9A

O.lrrm

Figure 9.-Side (A) and front (B) views ofseventh weal smge of Callinectes sapidus (from Costlow and Bookhout 1959). Figure lO.-Side (A) and

front (B) views of megalopal stage of Callinectes sapidus (from Costlow and Bookhout 1959).

14
combined with 30°C resulted in 100% mortality at 0.5 to 1.0 ppb

Dimilin. Average duration of the megalopal stage decreased as

temperature increased from 20° to 30°C in the control group (JI

concentrations of 0.1 and 0.3 ppb Dimilin.

3.23 Juvenile phase

Temperature acclimation

Thermal tolerance limits of blue crabs are dependent on

acclimation temperature and salinity. Tagatz (1969) evaluated

maximum and minimum median thermal tolerance limits (48 h) of

juvenile blue crabs (40 to 60 mm carapace width) and adult blue

crabs from St. Johns River, Fla., acclimated in 35 or 7%. salinities

combined with 6°, 14°,22°, or 30°C. Adult females and both sexes

in the juvenile stage were less tolerant than adult males to

temperature extremes at 7%. salinity. At both low and high

salinities, the upper and lower tolerance limits increased as

acclimation temperature increased. The upper thermal tolerance

limit was about 37°C for juveniles and adults, while the lower

median thermal limit was 3°C when crabs were acclimated to 7%.

salinity and 30°C. Adult blue crabs from South Carolina were

found to have an upper thermal tolerance limit of 35.2°C in 36X~

and a lower thermal tolerance limit of 3.2°C in 8.6100 over 96 h

(McKenzie 1970). Blue crabs were less tolerant to low saHnities at

high temperatures and high salinities at low temperatures.

Laboratory culture

Culture of juvenile stage blue crabs has been highly successful

when crabs resulted from individuals hatched in the laboratory and

juveniles were assigned separate compartments to prevent cannibalism. Blue crabs, starting with the first juvenile crab stage,

attained 94% survival (375 of 400) to a size of 25 mm carapace

width (1.25 g) and 88% survival from the first crab stage to 40 mm

carapace width (6.0 g) in the laboratory (Millikin unpubl. data).

For crabs beginning with the first juvenile stage, culture conditions

included individual rearing in plastic urine cups (100 ml) containing artificial seawater of 25100 salinity at 23°C, an 8-h light, 16-h

dark photoperiod, ad libitum feeding three times daily over an 8-h

period with live, adult brine shrimp from San Francisco or San

Diego, but not Utah (see Bookhout and Costlow 1970), and daily

replacement of 100% of the culture water (Millikin unpubl. data).

Culture conditions for juvenile blue crabs between 40 and 70 mm

carapace width were similar to methods for smaller juveniles

except for reduced salinity (20100 or less) and larger rearing

containers (Millikin et al. 1980).

Juvenile blue crabs (initial carapace width ranging from 5 to 40

111m) sampled from Galveston Bay showed maximum weight gain

at 29° to 30°C in the laboratory over a 45-d period (Holland et al.

1971). Temperatures above 30°C (up to 35° C) reduced growth of

juvenile crabs in the same study. Salinity values < I %. caused high

mortality rates of juvenile blue crabs (5 to 40 mm carapace width),

at a temperature of 29° C, but not at 15°C (Holland et al. 1971).

Design and construction of closed recirculating seawater systems for juvenile blue crabs during intermolt are described by Ogle

et al. (1982). These researchers recommended that research be

conducted concerning excretion rates, nutrient requirements, and

respiration rates of crabs to improve design of closed systems for

rcaring juveniles and adults.

15

Chemical toxicity

Several pesticides and other chemicals have been tested for

toxicity to juvenile stages of the blue crab. Lowe (1965) reported in

a preliminary stutly that intlividuals averaging 27 mm carapace

width died within a few days of exposure to DDT (I, I, l-trichloro2, 2-bis (p-chlorophenyl) ethane) concentrations :> 0.5 jJ.g/l,

while exhibiting extreme irritability, increased sensitivity to

external stimuli, and finally, paralysis. Mirex was determined to be

a more potent stomach poison than DDT for juvenile crabs (35-50

mm carapace width), since lower internal concentrations of mirex

(0.02 to 0.2 ppm) than DDT (0.8 ppm) were required to increase

metabolic rate (Leffler 1975b). Internal concentrations of 0.02

ppm mirex were accompanied by inhibition of the autotomy reflex.

Also, juvenile crabs were slightly more sensitive to mirex when

reared in dilute (50 mOsm/l) or concentrated (1,000 mOsm/l)

media compared with individuals reared in 200 or 600 mOsm/1.

No acute effects were elicited by four weekly feedings of 0.53 jJ.g

DDT over a 35-d period, whereas three weekly feedings of 0.14 jJ.g

mirex over a 35-d period produced acute poisoning in juveniles (ca.

35 to 45 mm carapace width). In a separate study, mirex leached

from fire ant, Solenopsis richteri. bait caused significant mortality

of juvenile blue crabs (21 to 75 mm carapace width) over a 28-d

period at average concentrations of 0.06 jJ.g/1 (fall) or 0.12 jJ.g/l

(summer) (Tagatz et al. 1975). Growth (percent increase of carapace width) was not affected by concentrations ranging from 0.04

to 0.12 jJ.g/1 of mirex over a 28-d period. Juvenile blue crabs

(mean carapace width = 34 mm) were shown to have a 96 h

LCso to Kepone of> 210 jJ.g/I (nominal concentration), since this

was the highest of a set of experimental concentrations, none of

which were toxic (Schimmel and Wilson 1977). Juvenile blue

crabs (mean carapace width = 37 mm) fed 0.5 g eastern oyster,

Crassostrea virginica, muscle contaminated with 0.25 jJ.g

Kepone/g tissue twice weekly, for 4 wk, accumulated O. I jJ.g

Kepone/g in muscle (Schimmel et al. 1979). No Kepone was

detected in muscle or whole body tissue of blue crabs exposed to

0.3 jJ.g/I Kepone for 28 d. Juvenile blue crabs (mean carapace

width = 28 mm) fed 0.5 g of oyster muscle contaminated with 1.9

jJ.g Kepone/g tissue twice weekly for 8 wk had an 80% mortality

rate. Gross external signs of Kepone poisoning were characterized

as excitation, followed by several days of lethargy, then death

(Schimmel et al. 1979). Cadmium and chromium toxicity in terms

of LCso's at 24, 48, 72, and 96 h to juvenile blue crabs (mean

carapace width = 1.5 cm) decreased appreciably with increasing

salinity (I, 15, or 35100 salinity) as shown in Table 2 (Frank and

Robertson 1979). Juvenile blue crabs (mean carapace width = 2.6

mm) evaluated for toxicity to bromine chloride had a 48-h LCso

(median concentration) of 1.2 mg/I and a 96-h LCso (median

concentration) of 0.8 mg/I (Burton and Margrey 1978). Median

time to death for the same size crabs at 1.6 mg hromine chloride/l

was 36 h. Fluoride concentrations of 20 mg/l decreased growth

increment per molt of juvenile crabs (mean carapace width = 70

mm) by 4.5% (Moore I<)7 I). Juvemle blue crabs weighing between

25 and 40 g :l~similated portions of radiolabelled paraffinic anti

aromatic hydrocarbons such as benzo( a)pyrene and naphthalene,

chiefly in the hepatupancreas, which contailled metabolites in

the form of dihydroxy eOIl1!,nunds anti their conjugates (Lee et al.

1976). Highly polar metabolites were fc)und in the green gland,

probably in readllless for excretion. Juvenile blue crabs (I to cm

carapace width) exposetl tu I ppm benzene had signiticantly longer

intermolt periods (50 vs. 33 d) and slower rates of limb regenera-
Table 2.-Cadmium and chromium toxicity to juvenile blue crabs

in several salinities.]

LC50 LCoo

Time (h) Salinity (cadmium. mg/l) (chromium, mg/I)

24 1 1.25 48

15 13.8 142

35 34.1 171

48 I 0.9 39

15 9.4 126

35 23.8 130

72 1 0.65 36

15 7.4 98

35 17.8 114

96 I 0.32 34

15 4.7 89

35 11.6 98

'From Frank and Robertson (1979).

tion following induced autotomy compared with individuals not

exposed to benzene (Cantelmo et al. 1981).

3.3 Adult phase

Laboratory culture

Adult blue crabs are easily cultured in the laboratory in either

freshly filtered seawater or artificial seawater, if crabs are reared

individually to prevent cannibalism. Maximum and minimum

median thermal tolerance limits (48 h) of adult crabs were reported

by Tagatz (1969) (see section 3.23). Designs for closed recirculating culture systems include those for peeler crabs (Perry et al.

1982) and intermolt crabs (Biddle et al. 1978; Ogle et al. 1982).

Descriptions of shedding tanks in natural waters and closed

systems include those by Haefner and Garten (1974) and Perry et

al. (1982), respectively (see section 7.5).

Chemical toxicity

Several pesticides and toxic inorganic contaminants have been

evaluated with respect to their effects on adult blue crabs. Adult

blue crabs (120 to 167 mm carapace width) that were exposed to

either 0.01, 0.1, or 1.0 ppm DDT (dissolved) for 12 h, showed a

pattern of gill absorption, hemolymph transport, and biotransformation of DDT to DDD (I, l-dichloro-2, 2-bis (p-chlorophenyl)

ethane) and DDE in the hepatopancreas over a 240-h period

(Sheridan 1975). Exposure to either 0.1 or 1.0 ppm of DDT for

12 h. resulted in hepatopancreatic accumulation levels of DDT

ranging from 100 to 800 ppb (wet weight) over 12 to 240 h following exposure, 70 to 300 ppb of DDD, and 60 to 280 ppb of DDE

over the same time period. High mortality rates of blue crabs in

salt marshes off Alligator Harbor, Fla., in November and December 1973, were attributed to reduced temperatures (below 18°C)

and high body burdens of DDT (Koenig et al. 1976). Crabs from

salt marshes that were highly contaminated from DDT and its

metabolites had hepatopancreatic concentrations ranging from

approximately 1.4 to 39 ppm total DDT (DDT plus its metabolites).

Schoor (1974) suggested that dissolved mirex is absorbed through

the gills of adult blue crabs because hemolymph had traces of

mirex-14C after 5 min of exposure and the hepatopancreas had

labelled mirex after 15 min. Mean Kepone concentration of backfin muscle in ovigerous blue crabs sampled from the lower James

River, Va., was < 0.05 f.Lg/g (Roberts and Leggett 1980). Eggs

16

had approximately three times more Kepone than in muscle connected to the fifth pair of pereopods. Further study indicated that

adult male crabs had higher Kepone concentrations in backfin

muscle than females (Roberts 1981). Higher Kepone concentrations usually occurred in the ovary (~ 0.2 f.Lg/g) compared with

backfin muscle « 0.1 f.Lg/g) of ovigerous and nonovigerous

females from the lower James River. Nonovigerous females

generally concentrated more Kepone into the ovary than ovigerous

females. Two cadmium-binding proteins (molecular weight =

10,600 and 9,400) were isolated from the hepatopancreas of adult

blue crabs following successive laboratory injections of cadmium

chloride over 3 d of 10, 50, and 100 f.Lg cadmium/d (Wiedow et al.

1982). Adult blue crabs exposed to either I X 10- 6 M, 5 X 10- 5

M. or 5 x 10- 3 M of pentachlorophenol or 2,4-dinitrophenol had

reduced respiration rates at the highest concentration of either

pesticide (Cantelmo and Ranga Rao 1978). Several enzyme

activities of mature blue crabs were measured following injection

with either sodium pentaclorophenol (Na-PCP) or dinitrophenol

(DNP) at a rate of 6 f.Lg/g body weight, or the hepatopancreas was

removed and subjected to Na-PCP or DNP in in vitro experiments

(R"o et al. 1979). Fumarase, malate dehydrogenase, succinate

dehydrogenase, pyruvate kinase, and lactic dehydrogenase were

inhibited by Na-PCP and DNP under both in vivo and in vitro

conditions. Microsomal Ca++ -ATPase from blue crab hepatopancreas was also inhibited by Na-PCP and DNP under in vivo and

in vitro conditions.

3.31 Longevity

Tagatz (1968a) reported that few blue crabs survive more than I

yr after reaching maturity (2 yr of age) and estimated a maximum

age of 4 yr for individuals from the St. Johns River, Fla. Van Engel

(1958) estimated maximum age of blue crabs in Chesapeake Bay to

be 2 to 3 yr.

3.32 Hardiness

Oxygen

Abnormally high local mortalities of blue crabs in commercial

pots have been observed in Chesapeake Bay (Carpenter and Cargo

1957) and Mobile Bay (Tatum 1979), probably due to low dissolved oxygen concentrations. Several experiments by Carpenter

and Cargo (1957) evaluated the effects of low dissolved oxygen

concentrations on blue crabs off the western shore of Chesapeake

Bay, slightly north of the mouth of the Patuxent River. No

mortality was observed in 34 blue crabs exposed for 24 h to 1.8

mg/l of dissolved oxygen at 30 m, while 70% mortality occurred

when 32 crabs were exposed to a mg/I of dissolved oxygen for

13 h. The authors concluded from these and several other studies,

that at 28° to 30°C, 25% or greater mortality occurred with oxygen

concentrations of 0.6 mg/l or less over 24 h and at 24° to 26°C,

significant mortality occurred at oxygen concentrations of O. 5 mg/l

or less over 24 h.

3.33 Competitors

Occurrence of other species of Callinectes in the same areas as

C. sapidus implies competition among them for food. In the St.

Johns River. Fla., Callinectes simi/is and C. ornatus occurred up

to 13.5 km (25 mi) upstream along with C. sapidus in the main

river (as low as 15')100 salinity) and in some tributaries (as low as I ~~
salinity) (Tagatz 1967). A general description of total distribution

of the various Callinectes species is in section 2.1.

Several other crab species such as the mud crab, Panopeus

herbstii, the stone crab. Menippe mercenaria, and the shore

crab, Carcinus maenus, cohabit with the blue crab in certain

geographical regions and prey on similar organisms (Williams and

Duke J979).

3.34 Predators

Predators of blue crabs include numerous fish species as well as

other blue crabs. Blue crabs are especially subject to predation

during molting and following molting while still in the soft shell

stage. Lack of sufficient cover, such as eelgrass, to provide refuge

may have an appreciable effect on rate of predation upon the blue

crab. Jaworski (1972) reported that black drum, Pogonias cromis,

red drum, Sciaenops ocellata, and the American eel, Anguilla

rostrata, are important predators of blue crabs in the Lake

Pontchartrain estuary of Louisiana. Forty-one percent of stomachs

examined from young (60-110 Col total length) sandbar sharks,

Carcharhinus plumbeus, from Chincoteague Bay, Va., contained

blue crabs (Medved and Marshall 1981). Blue crabs occurred in

11.1 % of 131 small (95 to 198 mOl SL (standard length) ) Atlantic

croakers, Micropogonias undulatus, and 4.6% of 119 large (200 to

350 mOl SL) Atlantic croakers from Mississippi Sound (Overstreet

and Heard 1978b). Blue crabs first appeared in stomach contents of

red drum at 40-49 mOl SL, assuming importance in the diet of red

drum of70-79 mOl SL (Bass and Avault 1975). Blue crabs were the

predominant species of crustaceans (42% occurrence) in red drum

of 250-932 mOl SL (Boothby and Avault 1971). Blue crabs

occurred in 31 % of the digestive tracts of 16 red drum (43 to 102

Col SL) from Sapelo Island, Ga., and 17% of 104 red drum (190 to

780 mm SL) examined from Mississippi Sound (Overstreet and

Heard 1978a). Soft blue crabs are a major prey item of the

American eel (Wenner and Musick 1975). The blue crab was a

major food item of the American eel (300 to 700 mOl SL) sampled

in the James River, Va. (33.3% of total food volume), York River,

Va. (68.2% of total food volume), and the Rappahannock River,

Va. (35% of the total food volume). Oyster toadfish, Opsanus tau,

American eel, and white catfish, Icta/urus catus, were the three

major predators of softshell blue crabs captured in habitat pots and

peeler pots in the Wando River estuary, S.c. (Bishop5). Spotted

gar, Lepisosteus oW/alUS, prey heavily upon blue crabs along the

Mississippi Gulf Coast, especially in salinities of 2 to lOY""

(Goodyear 1967). Adult striped bass, Morone saxatiiis, prey

heavily upon blue crabs during May in Albemarle Sound, N.C.

(42.2% occurrence) (Manooch 1973). Several freshwater fishes

including alligator gar, Lepisosteus spatula, spotted gar, and

largemouth bass, Micropterus sa/moides (Lambou 1961), in

waters adjacent to brackish water feed upon blue crabs.

Herring or menhaden species which consume zooplankton

during some life stages are probably important predators of blue

crab larvae. Van Engel (1958) speculated that chief predators

of blue crab larvae might include jellyfishes, comb jellies. and

fish larvae.

3.35 Diseases, parasites, and commensals

Diseases

Viral infections

Eight viruses have been found in the blue crab (Table 3). Several

are pathogenic singly or in combination. Reolike virus (RLV),

singly or in combination with Rhabdolike virus-A (Rh VA), causes

fatal infections characterized by weakness, decreased clotting

ability of the hemolymph, and eventual paralysis (Johnson 1977d,

1978). Herpeslike virus (HLV) also causes fatal disease. The crab

becomes progressively weaker, with no external or internal signs

until shortly before death, when the hemolymph becomes a chalky

white (Johnson 1977b, c). Laboratory-reared juvenile blue crabs

with HLV had several external signs shortly before death (usually

24 h or less) including: 1) Darkened and/or retracted eyestalks,

2) extension of chelae, 3) anorexia, and 4) loss of equilibrium

(anterior facing down) (Biddle et al. 1978). Hemolymph was often

chalky white with lack of normal clotting and the hepatopancreas

'J. M. Bishop, South Carolina Marine Resources Research Institute, PO. Box

12559, Charleston, SC 29412, pers. commun. October 1982.

Table 3. - Viruses of blue crabs.

Probable Size (nm) Main tissues

taxonomic (including envelope Lucatiun Jnd cel"

Name affinities if present) Shape Enveloped in ('ell "ttacked References

RLV (Reolike virus) Reoviridat' 55-60 Icosahedral No Cytoplasm Many types, Johnson and Bodammer (1975)

esp_ hemocytes, Johnson (1977b,d)

hemopoietic

tissue, gl ia,

epidermis

CB V (Chesapeake Bay VlfUS) Picormlviridae 30 Icosahedral No Cytoplasm Ectodermally Johnson (1978)

denved, esp.

nervous tissues;

also hemocytes

Rh VA (Rhabdol ike virus·A) Rhabdoviridae 20-30 x 110-170 Bacilliform Yes Cytoplasm Many meso- and Jahromi (1977)

and and ectodermal Iy Yudin and Clark (1978, 1979)

20-30 x 600 elongate deriv~d tissues

RhVB (Rhabdolike virus-B) Rhabdoviridae 50·70 X 100·170 Bacilliform Yes Cytoplasm Mandibular organ Yudin "nd Clark (1978)

EH V (Enveloped helical virus) Uncertain 105 X 194 Bacilliform Yes Cytoplasm Hemocytes, Johnson and Farley (1980)

and and hemopoietic

105-300 elongate tissue

HLV (Herpeslike virus) Herpetoviridae 185 )( 214 Icusahedral Yes Nud~us Hemocytes Johnson (1976b.c)

Baculo-A (Baculovirus-A) Baculoviridae 70 X 285 Bacilliform Yl",\ NUlleU> Epithelium of Johnson (197R)

hepatopanl rea;

Baculo·B (Baculovirus-B) Baculoviridae 100 x 335 Bacilliform Yes Nucleus Hemocytes Johnsun, unpub!.

17
was creamy colored with little texture. Chesapeake Bay virus

(CB V), w'li~h is related to the Picornaviridae, attacks neurosecretory and l"her cells of the nervous 5ystem and has a predilection for

ectodermal ti~sues (Johnson 1978). Infected crabs show behavioral

abnormalities and may be blinded. Signs of CBV disease may be

present at least 2 mo before death ensues. Baculovirus A (BaculoA) does not cause serious disease, although up to 50% of a crab

population may be infected (Johnson 1978). Nothing is known

regarding pathogenicity of the remaining viruses listed in Table 3.

The importance of viral disease in natural populations of the blue

crab has not been ascertained. With the exception of HLV and

Baculo-A, all information on blue-crab viruses is based on study

of captive populations.

Bacterial infections

Shell disease of the blue crab is characterized by the presence of

cratered lesions or softened and darkened areas on the exoskeleton.

As shell disease progresses, the lesions become confluent (Rosen

1967; Cook and Lofton 1973; Sandifer and Eldridge 1974; Sindermann 1977a). Chitinoclastic species of Vibrio, Pseudomonas, and

Beneckea have been isolated from the lesions (Cook and Lofton

1973). Shell dise2se is contagious. It occurs in natural populations

and may become epizootic in shedding tanks and other artificial

enclosures. especially during periods of high water temperatures.

It is not fata~, and the cuticle apparently is not completely

perforated. However, extensive erosion may lead to secondary

invasion of the soft tissues by pathogenic bacteria and fungi. Since

molting rids a crab of shell disease, older crabs have a prevalence

due to longer intermolt periods.

I-!emocoelic bacterial infection is caused by various gramnegative bacteria, often species of Vibrio (Krantz et al. 1969;

Johnson 1976b, 1977e). Signs of disease are weakness, nonclotting

hemolymph, prominent aggregations of hemocytes and bacteria in

th~ gills, and pre.,ence of gram-negative bacteria in the hemolymph

(Johnson 1976c; 1977e) . Hemocoelic bacterial infection may

cause extensive mortalities in wounded or otherwise stressed crabs

being held in sheJding tanks, especially during times of high water

temperatures (Johnson 1977e).

Fungal disease of eggs and larvae

Blue crab eggs are attacked and killed by the marine phycomycete, Lagenidium callinectes (Couch 1942; Rogers-Talbert 1948;

Sindermann 1977b). Zoospores of L. callineCles settle on blue

crab eggs, germinate on the surface, and extend germ tubes

which develop into a branched, septate mycelia (Sinderllann

1977b). More than 25% of an egg mass may be destroyed and,

in the laboratory, newly hatched larvae can also be killed by

L. callinecles. fungal invasion is limited to the periphe;y of

the egg mass. Diseased portions are brownish in yellow to orange

egg masses and grayish in the more mature brown to black egg

masses. Incidence of infection in natural populations varies from

year to year, according to locality. and prevalence may be lip to

95% of ovigerous females sampled (Bland and Amerson 1974).

Lagenidium callinectes is capable of infecting blue crab egg

masses in salinities ranging from 5 to 30:1'00, occurring on the east

and gulf coasts of the United States (Sindermann 1977b).

Paramoebiasis

Sprague and Beckett (1966) discovered paramoebiasis (gray

18

crab disease) in blue crabs often characterized by grayish-colored

appendages and ventral surface and hemolymph containing

amoeboid cells with two nucleusl:ke bodies having different

morphology. Mortalities of 20 to 30% attributed to this disease

occurred in several shedding tanks on the lawer seaside shore of

Maryland (Chincoteague Bay) and Virginia (Wachapreague) in

1965 (Sprague and Beckett 1966). The crabs suffering from the

infection were lethargic with grayish-colored appendages and

ventral surface. These amoeboid cells were later identified as a

Paramoeba sp. The amoebae range from 3 to 35 Mm and contain a

well-defined nucleus and a secondary nucleus (amphosome) which

is a major diagnostic character (Couch and Martin 1982). Hepatopancreas, gonad, muscle, gills, and blood are heavily infected

during advanced stages of the disease. Newman and Ward (1973)

reported that hemolymph smears containing 95 amoebae/100 cells

usually occurred in blue crabs exhibiting grayish-colored appendages (Sprague and Beckett 1968). Johnson (1977a) extended the

known geographical range of Paramoeba perniciosa, detecting the

organism only from high salinity areas of Sandy Hook Bay, N.J.,

and Long Island Sound near Milford, Conn., as well as Chesapeake

Bay aild Chincoteague Bay. Paramoeba perniciosu generally

occupies connective tissues and hemal spaces during early stages

of infection followed by invasion of hemolymph and replacement

of hemocytes (Johnson 1977a, f). Sawyer et al. (1970) reported

that paramoebiasis occurs in blue crabs only in salinities> 25100.

Paramoebiasis has not been reported to occur in blue crabs from the

gulf coast of the United States (Tatum 1979; Couch and Martin

1982).

Gas bubble disease

Gas bubble disease has been demonstrated in laboratory cultured

blue crabs (Johnson 1976a). Intermolt blue crabs (9.5 to 14 cm in

width) were held in flow-through systems and exposed to water

supersaturated with air resulting in 38% mortality (25 of 66) within

the first 2 d after exposure (Johnson 1976a). Exposure to water

supersaturated with air caused formation of gas emboli in the hemal

system, which caused local ischemia. Hemal spaces of many gill

lamellae and stems were filled with gas. Connective tissues were

mechanically disrupted by gas bubbles, but ischemic injury was

rare. Areas of thickened lamellar epithelia (salt glands) that occur

basally on some gills were not affected. Gas bubbles in the heart

frequently displaced muscle fibers, causing partial hemal stasis,

and blood vessels were often distended by large gas emboli. Johnson (1976a) suggested that focal degeneration of the heart muscle

and associated pacemaker nerves may contribute to hemal stasis.

Parasites, egg predator and commensals

Microsporidian infections

A microsporidian protozoan originally referred to as Nosema

michaelis (Overstreet 1975), and more recently called Ameson

michaelis (Overstreet 1978), may cause heavy mortalities of blue

crabs in natural populations. Infection often occurs when a crab

ingests tissues containing spores from an infected crab that is being

cannibalized or eaten after death (Overstreet 1975). Vegetative

growth occurs in hemocytes located in sinuses in connective tissue

surrounding the midgut. According to Weidner (1970) and Overstreet (1975), this is followed by sporogenesis in the sarcoplasm of

striated muscle cells and dissolution of infected and adjacent

un infected muscle fibers. Crabs with this so called "sick crab
disease" usually die under any additional stress, such as high water

temperature or suboptimal water quality. Gross signs of the disease

in the terminal stages incluC:c sluggish movement and coarse white

appearance and fibrous texture of mu:,cles (Sprague 1965; Overstreet 1977). Preliminary results by Overstreet and Whatley (1975)

and Overstreet (1975) indicated that incorporation of Buquinolate

(ethyl 4-hydroxy-6, 7-diisobutoxy-3-quinoline-carboxylate) into

food offered to blue crabs 48 h before or after infection with

A. michaelis spores retarded and often ,terminated "sick crab

disease." Oval spores of A. michaelis are characteristically 2.2 x

1.7 IJ-m and are known to occur in blue crabs from the Atlantic and

Gulf States (Sprague 1965; Sprague and Couch 1971). Also, the

closely related microsporidian, Ameson sapidi (Overstreet 1978),

occurs in muscles of the blue crab in North Carolina waters. The

oval spores of this protozoan are 3.5 x 2.1 IJ-m. Biochemical

changes in blue crabs infected with A. michaelis include: I) Lactic

acid accumulation in the hemolymph, thoracic muscle, and hepatopancreas, 2) increases in most free amino acids (taurine, aspartic

acid, threonine, proline, glycine, arginine) detected in the hemolymph except glutamic acid, 3) lower muscle protein content, 4)

hypoglycemia (66.3 mg glucose/l00 ml hemolymph declining to

51 mg glucose/IOO ml hemolymph), and 5) reduced lr and

Na+ ion levels in the blood (Findley et al. 1981).

A Minchinia-like haplosporidian was observed in hemolymph

from a blue crab from Chincoteague Bay, Va., as well as ar. individual from Beaufort, N.C. (Newman et al. 1976). Uninucleate

cells ranged from 3.4 to 7.3 IJ-m greatest diameter.

Sprague (1966) found the protozoan PlelStophora cargoi (=

Plistophora) in muscles of an infected male blue crab from the

Patuxent River, Md. The spores are ellipsoidal (3.3 by 5.1 IJ-m)

and give rise to many spores (32 to 100). Infected host tissues

include skeletal and cardiac muscle. The vegetative stages and

significance of mortalities of the blue crab in natural populations

is unknown.

Nemertean egg predator

The nemertean Carcinonemertes carcinophila infests gills and

eggs of adult female blue crabs. The nemertean produces a mucus

capsule that cements two gill lamellae together (Overstreet 1982).

Then, C. carcinophila migrates from the gill cavity into the blue

crab egg mass, soon after egg extrusion onto the pJeopods (Davis

1965). The nemertean feeds on the host's eggs and becomes

sexually mature in the blue crab egg mass (Hopkins 1947). Female

C. carcinophila withdraw from their mucus tubes after mating,

leaving their eggs behind whilt: they migrate back to the gills (Pyle

and Cronin 1950). Ciliated nemertean larvae hatch and either stay

in the mucus tube or swim within the egg mass. Following

metamorphosis into a creeping stage, individuals may remain in

the original host or infest other crabs (Davis 1965). Large, bright

red adult C. carcinophila occur on the gills of female blue crabs

that have ovulated, serving as a useful indicator of previous

spawning (Pyle and Cronin 1950). Hopkins (1947) noted that

nemertean infestations of both the sponge and gills were greatest

during August and September in blue crabs in Chesapeake Bay

following the peak fall spawning period. Infestations of C.

carcinophila on blue crabs are limited to high salinity regions,

therefore gills of male blue crabs are seldom infested.

Parasitic dinoflagellates

Newman and Johnson (1975) found a new disease c?used by

parasitic dinoflagellates (Hematodinium sp.) in blue crabs exam19

int:d from North Carolina, Georgia, and Florida in 1968 and 1969,

whereas individuals from South Carolina did not exhibit the

disease. According to Couch and Martin (19~Q). Hematodinium

sp. infections have been reported in blue crabs from Maryland to

northeas.tern Gulf of Mexico. Both male and fer,lale crabs ranging

from 70 to 170 mm carapace width were sampled from salinities of

oto 36%.. Crabs infected with these parasitic dinoflagellates were

only found in salinities> 11 %,. In advanced stages, when crabs

were moribund, the internal tissues were milky white and the

hemolymph was milky white with few, if any, hemocytes. Also, in

advanced infections, total lysis of hepatopancreatic tubules and

partial destruction of muscle occurs (Couch and Martin 1982). The

parasites were unicellular with a mean diameter of 8. I IJ-m and the

cells were nonmotile.

Hyperparasitism in the blue crab

Metacercariae of two digenetic trematodes (Microphallus basodactylophallus and Megalophallus sp.) being hyperparasitized by

the haplosporidian Urosporidium crescens have been isolated

from blue crab musculature and hepatopancreas (Couch 1974;

Overstreet 1978). The metacercariae of the trematode encyst in the

blue crab which serves as an intermediate host, with a mammal

(e.g., racoon) often serving as a definitive host. Harm is incurred

by the blue crab only in the case of very heavy infestations, which

cause a localized muscle weakening. Metacercariae of M. basodactylophallus infect thoracic muscles, hepatopancreas, and

ventral ganglion of blue crabs from Chesapeake Bay to Texas

(Overstreet 1982). llninfected and Iightly infected metacercariae

are white to cream colored and range from 189 to 269 IJ-m in

diameter, whereas infected metacercariae are easily recognized

because they become enlarged (410 to 654 IJ-m), darkly pigmented, and easily ruptured. The single layered cyst wall of the

metacercariae decreases in thickness from 2.3 to 0.8 IJ-m with

heavy infection, and most of the cell parenchyma is replaced by the

hyperparasite. Normally, the hyperparasite is released upon death

of the blue crab as the metacercarian cell wall ruptures. Couch

(1974) suggested that release of the hyperparasites through death of

the metacercariae in live blue crabs would probably result in death

of U. crescens due tu dt:struction by hemocytes or other defense

mechanisms of the blue crab.

Metacercariae of the microphallid trematode Levinseniella

capitanea were isolated from the hepatopancreas and gonads of

several adult blue crabs from Cat Island, Miss., (4 of 92 crabs) and

I of 41 crabs sampled near Raccoon Island, La. (Overstreet and

Perry 1972). Hyperparasitism of the metacercariae was not noted.

Peritrichous ciliates

Couch (1966) found two peritrichous ciliates on the gills of blue

crabs from Chincoteague Bay and Chesapeake Bay. These crabs

were sampled from salinities ranging from II to 30:1"". The loricate

genus Lagenophrys callinectes proliferated in high concentrations

in the gill lamellae of several crabs in a shedding operation in

Franklin City, Va., when water temperatures exceeded 27°C.

Once the gill surface was heavily occupied, gill lamellae became

fastened together and respiratory current was apparently retarded

by L. cullinectes. Also, the stalked genus Epistylus sp. was found

in low amounts on the gills of 32 of 41 individuals examined.

Although a commensal relationship, Couch (1966) theorized that

high density rearing conditions, higher water temperature (i.e.,

lower oxygen solubility), and an increase of peritrich ciliates may

cause anuxia. Couch (1967) later reported that L. callineetes
as well as other gill fauna remained on the "old" gill cuticle of

blue crabs following ecdysis. However, crabs that had rea:heJ

their terminal molt no longer have such a mechanism for removing these organisms. Peritrichs feed on bacteria (Sawyer and

MacLean 1978) and adhere to the gill lamellae by the lorica (i.e.,

Lugenophrys callinectes) or by a single basal holdfast (i.e.,

Epistylis sp.). These species have not been reported to occur in

gulf coast blue crabs (Tatum 1979). Lagenophrys callinectes has

been reported on some blue crabs sampled from Beaufort, N.C.,

and Trechard's Inlet, S.C., Maryland, and Virginia waters

(Couch 1967).

Barnacles as ectocommensals and parasites

A gooseneck or pedunculate barnacle, Octolasmis muelleri,

is often found on the efferent side of gills of blue crabs as well

as other decapods in high salinity waters (i.e., generally above

15~oo). This ectocommensal does not gain nutrition from crabs,

but it may interfere with respiration during heavy infestations due

to decreased activity of the host's scaphognathite (Walker 1974;

Overstreet 1978).

The acorn barnacle, Chelonibia patula, is an ectocommensal

that often occurs on the carapace of blue crabs, especially those

reaching their final postlarval molt. Acorn barnacles on blue

crabs have been reported from Delaware Bay (Williams and

Porter 1964), South Carolinian waters (Eldridge and Waltz

1977), Mississippi (Overstreet 1978), and St. Johns River, Fla.

(Tagatz 1968a).

The cypris larva of the parasitic sacculinid barnacle, Loxothylacus texanus, settles on early postmolt juvenile blue crabs and

enters into the interior of the crab in toto (Overstreet 1978). Growth

of the blue crab is often retarded to a maximum size of 3 to 8 cm

carapace width (often called button crabs), thereby making the

individual unavailable to the fishery. Most infected crabs appear

similar to miniature adult females, with males often transforming

secondary sexual characteristics to those of females, possibly as a

result of alteration of the androgenic gland. Parasitized males often

have 6 distinct abdominal segments, whereas normal males only

have 4 abdominal segments (Reinhard 1950). Also, parasitized

males have abdominal musculature that is similar to that of females

and parasitized females have a broadened abdomen similar to adult

females. After formation of an extensive "root system" inside the

crab. externae consisting of a brood pouch for larvae and gonads of

the parasitic barnacle protrude between the crab's abdomen and

thorax. Occurrence of L. texanus is rare in low salinity waters

(Darnell 1959) and appears to be restricted to gulf coast waters 5uch

as Louisiana (Adkins 1972), Mississippi (Perry 1975), and Texas

(Reinhard 1950). Loxothylacus texanus has not been reported on

blue crabs in Chesapeake Bay (Lawler and Van Engel 1973).

A leech (Myzobdella lugubris Leidy) occurred on several blue

crabs (from Bulow Creek, Volusia County, Fla.) which had

suffered recent mortality (Hutton and Sogandares-Bernal 1959).

The leeches were located inside the crab near perforations on the

carapace and articulations of the pereopods. It is not clear from

their report whether the leeches infested the crabs before or after

death. Further study indicated that M. lugubris derives nutrition

primarily from fishes, but uses the carapace of blue crabs as a

substrate for its cocoons (Daniels and Sawyer 1975). The cocoons

were equally distributed on the posterolateral regions (both dorsal

and ventral) of the carapace and occurred on blue crabs from the

20

Ashley River, S.C.. (0 to 14/",,) from March to November with

peak occurrence in October and November.

3A Nutrition and growth

3Al Feeding

Dietary protein concentration requirements of juvenile blue

crabs have been examined in several laboratory feeding studies.

Juvenile male and female blue crabs (mean initial weight = 1.2 g,

mean carapace width = 2.5 cm) achieved better growth when fed

44 or 60% crude protein diets (as percent of dry weight) compared

with individuals fed 27% crude protein over a 105-d period

(Millikin et al. 1980). Adult, live brine shrimp, Artemia salina,

fed as reference diet to a separate group of juvenile blue crabs,

promoted better growth than any of the laboratory formulated

diets. Apparently, there were important nutritional differences

other than dietary protein concentration between the high protein

(60%) laboratory formula diet and the live brine shrimp diet (58 to

64% crude protein, dry weight). In a separate study, juvenile blue

crl'bs of larger initial size (mean initial weight = 7.8 ± 3.9 g SD, n

= 62) gained significantly more weight per molt on a 5-d postmolt

basis when fed 25 or 40% crude protein than when fed 15% crude

protein (Fortner 6 ). Therefore, laboratory reared juvenile blue

crabs ranging from approximately I to 10 g (25 to 55 mm carapace

width) require a minimum of 40% dietary protein in terms of

growth rate, while slightly larger juvenile blue crabs require

dietary protein concentrations of 25% or more, but < 40%.

Winget et al. (1976) reported no significant differences in growth

of juvenile blue crabs (initial mean carapace width = 59 mm) fed

26,46,62, or 75% crude protein on a dry weight basis over a 60-d

period. However, growth differences in the previously described

protein requirement studies (Millikin et al. 1980; Fortner footnote

6) were not observed until after 105 and 63 d, respectively.

Therefore, a longer experimental period by Winget et al. (1976)

possibly would have demonstrated differential growth due to

dietary protein concentration.

Salinity is not an important factor governing food conversion

in laboratory reared juvenile blue crabs. Holland et al. (1971)

reported that juvenile blue crabs (initial weight = 0.2 to OA g)

had similar food conversion rates regardless of salinities ranging

from 6 to 21 ~"" .

Blue crabs require an exogenous source of sterol. Whitney

(1970) demonstrated that juvenile and adult blue crabs at intermolt,

premolt, and soft shell stages were incapable of synthesizing either

squalene or sterols from acetate-1}4C, or meva10nate-2}4C.

Biddle et al. (1978) concluded that 3 or 9% dietary fi ber (dry

weight basis) was satisfactory for juvenile blue crabs of two size

groups (2.5 to 6.0 g and 6.1 to 36A g initial weight) in terms of

survival, growth, and food conversion over a 9-wk period.

However, crabs fed diets containing 27% fiber had increased

mortality due to a Herpeslike virus. Also, crabs fed 27% dietary

fiber had poorer food conversion ratios and greater feed intake,

possibly due to less availability of nutrients per unit weight of

food ingested.

Glycogen content of the hepatopancreas of blue crabs has been

evaluated according to molt stage, season, and sex. Blue crabs (5 to

15 cm carapace width) sampled from Delaware Bay had highest

"A. R. Fortner, Southeast Fi5heries Center Charleston Laboratory. National

Marine Fisheries Service, NOAA, P.O. Box 12607. Charleston, SC 29412, pers.

commun. September 1982.
hepatopancreatic glycogen concentration (8.1 mg/g tissue) during

premolt (stage D), followed by 6.6 mg glycogen/g hepatopancreas

(midgut gland) during soft shell stage (stage A), and approximately

3.0 mg glycogen/g tissue during early postmolt (stage B) and

intermolt (stage C) (Rouse 1972). Glycogen concentrations of the

hepatopancreas were significantly lower in summer, with highest

concentrations during winter and decreasing concentrations during

spring. Also, hepatopancreatic glycogen concentrations were

higher for females in molt stages A, B, and D than comparative

stages in males. Lower hepatopancreatic glycogen concentrations

for females than males in stage C might be the result of mobilization of glycogen reserves to aid in production of ova (Rouse 1972).

Midgut gland glycogen content appears to have an annual cycle in

blue crabs, according to a study from Indian River Bay, Del.

(Winget et al. 1977). Crabs ranging from 11 to 150 g, had 8.4 mg

glycogen/g tissue during November-December, \0.4 mg during

February-March, 5.3 mg during April, and 3.3 mg during JuneSeptember (Winget et al. 1977). Sampling during each season for

several years is necessary to verify these preliminary results.

Mean whole body protein values of intermolt juvenile blue crabs

3 to 7 cm carapace width ranged from 34.7 ± 0.27 % (SE) on a dry

weight basis for individuals fed 27% dietary crude protein over 18

wk to approximately 40% whole body protein for crabs fed 44 or

60% dietary crude protein (Millikin et al. 1980). Crabs fed adult

brine shrimp over the same time interval had significantly higher

whole body protein (46.1 ± 1.8%) than individuals fed the laboratory formula diets containing 27, 44, or 60% crude protein. Whole

body glucosamine values were approximately 4% on a dry weight

basis for juvenile blue crabs regardless of dietary treatment

(Millikin et al. 1980).

Ash composition of juvenile blue crabs reared in the laboratory

that were from 8 d postmolt to late premolt stage (3 to 7 cm

carapace width) consisted of 30 to 40% of total, whole-body dry

matter, while calcium was 9 to 13% of whole body dry matter and

phosphorus was 0.7 to 1.5% of whole body dry matter (Millikin

unpubl. data). Whole-body dry matter of juveniles 8 d postmolt to

late premolt ranged from 20 to 32%. Blue crabs that were 2 to 6 d

postmolt generally had lower whole body ash as a percent of dry

matter (24-28%), lower calcium as a percent of dry matter (5 to

6%), lower whole-body dry matter content (17 to 20%), and

similar phosphorus concentrations as a percentage of dry matter

(1.3 to 1.4%).

3.42 Food

Data are scarce on food organisms consumed by larval stages of

the blue crab in natural waters. Although phytoplankton may be

consumed, plant material alone is believed to be deficient in

protein content (Costlow and Sastry 1966). Various phytoplankton

species fed to early zoeal stages in the laboratory without concurrent feeding of zooplankton, prolonged time before death

compared with survival time of starved larvae (Costlow and

Bookhout 1959), but unicellular algae fed alone did not result in

successful molting of first stage zoeae to second stage zoeae.

According to Sulkin and Epifanio (1975) and Sulkin (1978),

rotifers, Brachionus plicatilis. ranging from 45 to 80 fLm, promote

high survival and molting success during the first 14 d of

development, whereas recently hatched nauplii of Artemia salina

(250 fLm) fed alone to first and second stage zoeae produce low

survival and molting success (Costlow and Bookhout 1959).

Therefore, zooplankton in the size range of 45 to 80 fLm probably

~I

are among the chief sources of food organisms for blue crab larvae,

especially in the early stages.

Juvenile and adult stages of blue crabs feed chiefly upon

molluscs, crustaceans, and fish. According to Tagatz (1968a),

crustaceans and molluscs are the chief food items of juvenile blue

crabs < 21 mm carapace width. Juveniles larger than 21 mm

carapace width fed primarily upon clams, fish, and crustaceans.

Jaworski (1972) reported that crustaceans and molluscs are equally

important in terms of percentage stomach volume in blue crabs

between 30 and 74 mm carapace width. Molluscs comprised an

appreciably greater stomach volume than crustaceans in crabs >

125 mm carapace width (Jaworski 1972). Laughlin (1982) divided

blue crabs from Apalachicola Bay into three trophic groups based

upon stomach contents. Smallest juveniles « 31 mm carapace

width) fed mainly upon bivalves, plant matter, ostracods, and

detritus. The second trophic group (31 to 60 mm carapace width)

consumed mostly fishes, gastropods, and xanthid crabs, while

the third trophic group of blue crabs (> 60 mm carapace width)

fed on bivalves, fishes, xanthid crabs, and blue crabs. Van Engel

(1958) listed occurrence of several food items in juvenile and adult

blue crabs, such as fresh and decaying fish; juvenile oysters

(Crassostrea virginica); hard clams, Mercenaria mercenaria; sea

lettuce, Viva sp.; roots, shoots, and leaves of eelgrass; and salt

marsh grass, Spartina alternifiora. The coot clam, Mulinia

lateralis. is an important prey item of blue crabs in the York River,

Va., estuary (Virnstein 1977). Other food items of blue crabs

include small oysters (Menzel and Hopkins 1956; Menzel and

Nichy 1958), three species of mussels: Congeria leucophaeta.

Mytilopsis leucopheata. and Brachidontes exustus (Darnell 1961;

Odum and Heald 1972), the soft shell clam, Mya arenaria

(Dunnington 1956), and the wedge clam, Rangia cuneata (Darnell

1961). Hamilton (1976) observed predation by juvenile blue crabs

(25 to 65 mm carapace length) on marsh periwinkles, Littorina

irrorata (9 to 15 mm shell length). Laboratory studies indicated

that juvenile blue crabs ranging from 10 to 15 cm carapace width

consumed cultchless oyster (C. virginica) up to 4 cm shell length,

while blue crabs from 65 to 80 mm carapace width could not feed

upon oysters> 25 mm shell length (Krantz and Chamberlin 1978).

Large blue crabs (15 to 16 cm) consumed Atlantic ribbed mussel,

Geukensia (= Modiolus) demissa, < 8 cm shell length (Seed

1980). Medium-sized crabs (10 to II cm) and small crabs (6 to 7

cm) consumed Atlantic ribbed mussels up to 5.5 and 4.8 cm,

respectively, in the laboratory. Blue crabs > 80 mm carapace

width were capable of breaking open the largest Mya arenaria

shells (90 mm dorsoventral axis) tested according to compression

measurements of chelae force at the dactyl tip (Blundon and

Kennedy 1982a). The same investigators reported that only large

(> 40 mm shell height) wedge clams were capable of withstanding crushing capabilities of large blue crabs (100 to 165 mm

carapace width). In a separate study, M. arenaria (30 to 40 mm

shell length) buried at either 5, \0, 15, or 20 cm were consumed

less frequently at either 15 or 20 cm depths than at 5 or 10 cm

depths by large blue crabs (121 to 180 mm carapace width)

(Blundon and Kennedy 1982b).

3.43 Growth and morphometry

Growth of blue crabs is dependent upon temperature, molting

frequency, food availability and nutritional quality, and life stage

of the individual. Growth primarily occurs during molting,

although sma!1 weight increases occur through relative changes

in tissue content during the intermolt period (relative increases
of whole body protein compared with moisture), Generally, as

temperature decreases, food consumption rate uecreases, ~specially between 34° and 13°C (Leffler 1972). Tagatz (l968b)

observed that growth per molt of blue crabs remained similar

regardless of temperature (summer vs. winter) in the St. John~

River. Fla., but that intermolt intervals were three to four times

longer in winter. Therefore, low temperatures that prevent molting

altogether or increase intervals between molts significantly reduce

growth rate. Churchill (1921) reported that females cease feeding

activity in Chesapeake Bay below 10°C. According to Van Engel

et al. (1973), growth occurs in blue crabs in Chesapeake Bay from

late April to mid-October when temperatures are generally above

15°C and blue crabs undergo semihibernation when temperatures

fall to 5°C or less.

Salinity values ranging from 6 to 30roo do not differentially affect

growth of juvenile and adult blue crabs. Growth of female crabs

during and following their terminal molt was not differentially

affected by salinities of 9, 16, or 27 roo (Haefner and Shuster 1964)

or 10, 20, or 30roo (Haefner 1964). Holland et al. (1971) reported

that salinities ranging from 6 to 21 roo (6, II, 16, or 21%.) did not

affect growth or food conversion of laboratory reared juvenile

crabs (5 to 40 mm carapace width).

Growth rate of juvenile blue crabs up to at least 7 cm carapace

width does not vary between sexes. Only one of eight groups of 40

juveniles of equal age fed various diets over an 18-wk period

showed differential growth between sexes (Millikin et al. 1980).

Size variation of the first seven juvenile instars reared in the

laboratory is described in Table 4 (Newcombe, Sandoz, and

Rogers-Talbert 1949).

Table 4.-Size variation of first seven juvenile stages of blue crabs.'

Mean carapace

Carapace width width

Juvenile instar (range-nun) (mm) SD

I 2.2 to 3.0 2.5 0.14

2 3.010 4.2 3.7 0.25

3 4.1 to 6.0 5.1 0.42

4 5.5 to 7.4 6.6 0.46

5 7.2 to 10.0 8.6 0.66

6 8.7 to 12.4 10.2 0.85

7 9.71013.0 11.0 0.81

IN = 50 for first 6 instars, N = 35 for seventh instar (Newcombe.

Sandoz, and Rogers-Talbert 1949).

After early juveniIe stages, considerable overlap in sizes

between instars occurs. Tagatz (1968a) reported that fully grown

crabs range from 100 to 240 mm carapace width. Tyler and Cargo

(1963) showed considerable overlap in carapace width between the

penultimate instar (81 to 135 mm, N = 254) and the ultimate instar

(118 to 195 mm, N = 88) for females from several sampling sites

of Chesapeake Bay. Tagatz (1968b) reported mean carapace

widths and ranges for adult males (mean = 147 mm, ranging from

117 to 181 mm), mature females (mean = 148 mm, ranging from

128 to 182 mm). immature females (mean = III mm, ranging

from 100 to 130 mm), and ovigerous females (mean = 152 mm,

ranging from 116 to 180 mm). Tagatz (l968b) also found a

considerable range between the smallest mature female (99 mm

carapace width) and the largest immature female (177 mm

carapace width) sampled from St. Johns River, Fla. Tagatz (1965)

reported a maximum carapace width for males sampled from

commercial catches in St. Johns River to be 246 mm carapace

width. while the heaviest male (225 mm carapace width) weighed

550 g. Van Engel et al. (1973) reported a maximum size for male

22

bl ue crabs from Virgi nia as 241 mm carapace width. The largest

immature female sampled from commercial catches during a 3-yr

period from live estuaries in South Carolina was 154 mm carapace

width (Eldridge and Waltz 1977).

Variation of growth increments per molt «postmolt weight -

premolt weight) -;- (premolt weight) x 100) may be largely the

result of both dietary nutritional quality and temperature. Tagatz

(1968b) reported that temperatures between winter (IYC) and

summer (25° to 30°C) in St. Johns River were not reflected in

significant seasonal alteration in growth increments of blue crabs.

However, Leffler (1972) reported that blue crabs of 20 to 69 mm

carapace width, demonstrated an average increase in growth per

molt as temperature decreased from 34° to 15°C. Juvenile blue

crabs ranging in initial weight from 3.5 to 23.7 g increased

33 ± 8.4% (n = 18) in weight when fed a 15% crude protein diet,

51.9±9.6% (n = 19) increase in weight when fed a 25% crude

protein diet, and 57.6± 12.2% (n = 23) increase in weight when

fed a 40% crude protein diet for individuals weighed on a 5-d

postmolt basis in the laboratory (Fortner footnote 6).

Growth per molt of blue crabs is highly variable in natural

waters. Growth increments per molt in terms of carapace width

varied from 7.8 to 50% in juvenile and adult blue crabs held in

floating tanks in St. Johns River (Tagatz 1968b). Percent growth

increase averaged 32% for females during their terminal molt.

Mean percent increments in growth in 10 mm width intervals

varied from 21 to 34% for crabs from St. Johns River (Tagatz

1968b). Mean percent increments in carapace width ranged from

II.9% for small juvenile females (10 to 19.9 mm) to 33.7% for

large females (100 to 109.9 mm) while mean percent increments

ranged from 14.4% for small juvenile males (10 to 19.9 mm) to

32.9% for males ranging from 80 to 89.9 mm carapace width (Gray

and Newcombe 1938b). Growth per molt (percent increase in

carapace width) during the terminal molt of females from St. Johns

River averaged 34.4% (n = 85) in salinities> 5roo and 30.2% (n

= 85) in salinities < 1%. (Tagatz 1968b). Average values of

growth per molt (percent increase in length) during the terminal

molt of females from three sampling sites in Chesapeake Bay

ranged from 23.4 to 25.6% (Haefner and Shuster 1964).

Average size (carapace width) at maturity for females was

appreciably smaller for Chincoteague Bay crabs (5.3 in = 13.5

cm) compared with Delaware Bay crabs (6.3 in = 16 cm) and

Chesapeake Bay crabs (6.5 in = 16.5 cm) (Porter 1955).

Autotomy and regeneration of appendages are common among

many crustacean species (Weis 1976), including blue crabs. If

an appendage is firmly imprisoned or seriously injured, a break

occurs along a fracture plane located on the distal end of the basis,

with eventual replacement of the appendage by regeneration (Pyle

and Cronin 1950). The regenerating limbs grow in a folded

position and are often functional following the next molt (Weis

1976). Laboratory studies showed that mortality among blue crab

megalopae was low when chelae were induced to autotomize,

compared with megalopae with chelae cut at a site other than the

basi-ischiopodite fracture plane (Costlow 1963a). If the chelae

were lost prior to day 4 of the megalopal intermolt period, then the

completely regenerated appendage normally appeared at metamorphosis. Although 100% of hatchery-raised crab~ had a right

"crusher" cheliped (larger) and a left "cutter" cheliped, only

79% of 1,156 crabs sampled from natural waters showed the same

morphological character (Hamilton et al. 1976). A trend existed for

larger crabs to have a larger percent occurrence of a left" crusher"

cheliped and right "cutter" cheliped, which Hamilton et al. (1976)

felt to be due to reversed cheliped laterality through autotomy and

regeneration. Shuster et al. (1963) attributed most of the bizarre
claw deformities of blue crabs sampled from the field to new

growth in the form of duplication at the site of injury rather than

autotomy followed by regeneration.

Male blue crabs are generally heavier than female blue crabs for

a given carapace width. Pullen and Trent (1970) reported carapace

width-total weight relations for crabs from Galveston Bay, Tex.,

> 25 mm carapace width, excluding gravid females: Male crabs

(log weight = - 3.74 + 2.775 log carapace width) and female

crabs (log weight = -3.54 + 2.639 log carapace width).

Newcombe, Campbell, and Eckstine (1949) reported carapace

width-total weight relationships for blue crabs from Chesapeake

Bay (Virginia portion) using untransformed data as follows: Male

crabs (weight = 0.00026 width 2.

67 ) and female crabs (weight = 0.00034 width 2.

57 ). A graphic presentation of carapace width vs.

weight also demonstrated that male blue crabs are generally

heavier than females from St. Johns River for a given carapace

width between lOO and 200 mm (Tagatz 1965).

Length to width ratio (in the form of a percentage) steadily

decreases as carapace width increases in blue crabs. Tagatz

(l968b) reported values of 50% for males and females from St.

Johns River between 10 and 19 mm carapace width and 40.7% for

males and 38.8% for females for individuals between 140 and 149

mm carapace width. Newcombe, Sandoz, and Rogers-Talbert

(1949) reported values equivalent to length:width ratio as a

percentage as being 53.2% for males and 53.9% for females from

Chesapeake Bay between 10 and 19 mm carapace width. Carapace

length was 44.9% of carapace width for males and 41.7% for

females from Chesapeake Bay between 140 and 149 mm carapace

width.

Mature males from Ashley River, S. c., had a greater average

weight than immature males having the same carapace width,

while immature females were heavier than mature females for a

given carapace width (Olmi and Bishop 1983). According to Olmi

and Bishop (1983), the lighter weight of adult females compared

with immature females of the same carapace width, may result

from the maturity molt transformation of females into long spined

forms as observed by Tagatz (1965) and Gray and Newcombe

(l938a).

Due to the problem of variation in lateral spine lengths for a

given size crab (e.g., carapace length or width at the base of the

spine), Olmi and Bishop (1983) suggested that measurement of

carapace length or width at the base of the lateral spines might

be more accurate in developing a regression analysis with closer

agreement between observed and predicted values of body weight.

Carapace length was originally suggested by Gray and Newcombe

(1938a) as an alternative to carapace width for prediction of body

weight, whereas Williams (1974) suggested the use of carapace

width at the base of the lateral spines (rather than from tip to tip) to

predict body weight.

3.44 Metabolism

Oxygen consumption in blue crabs is a function of size,

temperature, and molt cycle. Leffler (1972) reported greater

increases in standard metabolic rate in blue crabs (20 to 69 mm

carapace width) reared in 34° vs. 2rC compared with crabs reared

in 27° vs. 20°C. Oxygen consumption increases as temperature

increases from 10° to 25°C (Laird and Haefner 1976). Lewis and

Haefner (1976) demonstrated that premolt female blue crabs in the

penultimate instar had higher oxygen consumption rates (J..d 02/g

wet weight per h) than molting or postmolting crabs. Twelve hours

or less after completion of molting, weight-specific oxygen

23

consumption was depressed, whereas oxygen uptake increased

after 12 h. Also, total oxygen consumption increased with higher

wet weights of females in the terminal molt stage, but weightspecific oxygen consumption decreased as total wet weight of the

blue crabs increased (Lewis and Haefner 1976). Laird and Haefner

(l976) also found that oxygen uptake decreased with increases of

total wet weight of blue crabs. Oxygen uptake evaluated in a 3 x 3

factorial (10°, 17°, and 25°C; 10, 20, and 30'}'"" sal inities) resulted

in no differences in oxygen consumption between sexes (Laird and

Haefner 1976). Lower average oxygen consumption occurred at a

salinity of 20%" than at 10%". Also, blue crabs acclimated to cold

temperatures (IO°C) had a higher mean oxygen uptake when

acclimated to a salinity of 10%" than when acclimated to a salinity

of 30%".

Oxygen consumption rates generally increase as sal inity decreases. King (1965) showed that oxygen consumption increased

53% in juvenile blue crabs reared in salinities of7 to 9A'~ compared

with crabs reared in full strength seawater, regardless of the salinity

from which the experimental animals were captured. Excised gills

from adult blue crabs demonstrated higher respiration rates when

crabs were reared in 300 mOsm seawater than individuals reared in

940 mOsm seawater (Cantelmo and Ranga Rao 1978). Furthermore, Findley et al. (1978) demonstrated that adult blue crab

respiration was significantly higher at salinities of 10 and 20A'~ than

at 30'Y"". Oxygen consumption rates decreased with increase in

salinity from 5 to 35'Yoo among adult male and female crabs and

immature crabs during winter (Engel and Eggert 1974).

The mixed function oxygenase, arylhydrocarbon hydroxylase,

was shown to have highest specific activity in the pyloric stomach

and antennal gland (green gland) of males and females (Singer and

Lee 1977). Since mixed function oxygenases function in the

metabolism of bile acids, fatty acids, and steroid hormones as well

as hydroxylation of the aromatic ring of petroleum hydrocarbons,

the authors suggested that exposure to sublethal concentrations of

petroleum could conceivably interfere with metabolism related to

molting (i.e., growth).

3.45 Molting

Stages of the molt cycle of the blue crab were described by

Johnson (1980) as modified from Passano (1960) as shown in

Table 5.

Intermolt periods increase with age of blue crabs, but are

generally shortened by ample food availability, high dietary

nutrient quality, and higher water temperatures. Based upon

laboratory studies, blue crabs are believed to molt 8 times (seven

lOeal stages plus a megalopal stage) (Costlow and Bookhout

1959), followed by as many as 18 to 20 postlarval molts for females

and 21 to 23 postlarval molts for males (Van Engel 1958). Blue

crabs ranging in carapace width from 10 to 160 mm attained

maximum expansion of their new exoskeleton within 6 h following

molting (Gray and Newcombe 1938b). Graham and Beaven (1942)

wggested that blue crabs cease molting when temperatures are

depressed to about 16°C from approximately November to the

middle of April in Chesapeake Bay. Ecdysis has been observed at

temperatures as low as 3.8°C in blue crabs from St. John, River,

Fla. (Tagatz 196Rb) Leffler (1972) observed appreciably shorter

intermolt intervals for blue crabs (20 to 69 mm carapace width)

reared at 27° or 34° ethan indi viduals reared at 13°C. How·

ever, growth per molt frequently decreased as intermolt pel iod

decreased. Similarly, blue crabs molted more frequently at 30°C

than at 20°C, but growth per molt was greater at 20° C (Winget et
Table 5.-Molt cycle stages of the blue crab, Callinectes sapid"s (from Johnson

1980).

Stage

Stage A

AI

A,

Stage B

BI

B,

Name

Newly molted

Soft

Papershell

Characteristic

Continued waler absorption and initial mineralizltion

Exocuticle mineralization

Endocuticle secretion begins

Active endocuticle formation, chelae hard: tiSSI e

growth begins

and Bishop (in press) reported capture of an adult female in

proecdysis (pink sign) that consequently died during ecdysis in

the laboratory.

. The absolute amount of calcium in blue crabs (size not reported)

Increased steadily in individuals up to at least 10 d postmolt with

a 2.5-fold increase in total protein and a 4-fold increase in

chitin in the cuticle from ecdysis to 10 d postmolt (Vigh and

Dendi nger 1982).

Stage CHard

CI

C,

C3

or

Main tissue growth

Tissue growth continue,!)

Completion of exoskeleton: membranous layer

formed

"Intermolt"; major accumulation of organic reserves

3.5 Behavior

3.51 Migrations and local movements

Table 6.-lntermolt periods during early juvenile development of blue crabs

reared individually in the laboratory,I

7 L. E. Cronin, Chesapeake Research Consortium, Shadyside, MD 20764, pers.

commun. August 1982 and March 1983.

ITemperature = 23.5 ± I.soC, salinity = 25%. with gradual reduction to 20::1..

after 160 d, photoperiod = 8 h Iight/16 h dark. (Millikin, unpub!. results,)

'Mean ± standard deviation of number of days between molt instars of juvenile

development. Number of crabs for each intermolt period are in parentheses,

Instar numbers of Mean intermalt Instar numbers of Mean intermolt

juvenile stage period (d)' juvenile stage period (d)

1-2 72±2.7 (104) 8-9 16.0+4.6 (156)

2-3 7.0± 3U (157) 9-111 19.7±7.1 (143)

3-4 8.0± 2.5 (157) 10-11 22.9± 8.0 (119)

4-5 9.2± 3.8 (157) 11-12 27.6±7.7 (75)

5-6 1/.1 ±4.2 (157) 12-13 27.7±7.1 (38)

6-7 126± 28 (157) 13-/4 27.7±6.1 (32)

7-8 147:!'5.1 (150)

After hatching and larval development of blue crabs in high

salinity waters (Costlow and Bookhout 1959; Costlow 1967),

megalopae are recruited into estuarine waters (Boicourt 1982;

Sulkin and Van Heukelem 1982) followed by migration of early

juvenile stages of both sexes upstream to lower salinity waters

(Van Engel 1958). Early juvenile stages feed in shallow waters

(Van Engel 1958; Tagatz 1968a; Jaworski 1972). During fall

migrations in Chesapeake Bay, concentrations of females in the

Virginia portion of the Bay occur in deep channels, while males

generally overwinter in lower salinity waters, with larger males

occurring in deeper waters than smaller males (Van Engel 1958).

Especially in spring, immature females approaching their terminal

molt migrate to lower salinity waters to seek out mature males for

mating. Soon after mating, females initiate migrations back to

higher salinity waters that are favorable for larval development.

This pattern of migration has been observed in many geographical

regions, such as Delaware Bay (Cronin 1954), Chesapeake Bay

(Van Engel 1958), Newport and White Oak Rivers, N.C. (Judy

and Dudley 1970), South Carolina (Eldridge and Waltz 1977),

Barataria estuary, La. (Jaworski 1972), Lake Pontchartrain, La.

(Darnell 1959), Mobile Bay (Tatum 1979), Mississippi Sound

(Perry 1975), and to some degree in St. Johns River, Fla. (Tagatz

I 968a) . Migration of adult females along the gulf coast of Florida

is generally in a northward direction alongshore following mating

(Oesterling 1976). Most tagged adult males that were recaptured

had remained in the estuary of initial capture. Adult blue crabs do

not migrate appreciably between adjacent estuaries in South

Carolina (Fischler and Warburg 1962).

Most migrations of adult female crabs were southward from

four release sites within Delaware-Maryland-Virginia seaside bays

(north to south: Assawoman Bay, Isle of Wight, Sinepuxent Bay,

and Chincoteague Bay, Maryland and Virginia) (Cargo 1958).

This occurred dcspite lack of a continuous increase in salinity from

north to south across several inlets (Cargo 1958).

Several studies have been conducted to determine optimal

tagging methods for tracking movements of juvenile and adult blue

crabs. Cronin ( 1949) reported that Nesbit-type tags wired across

lh~ carapace and looped over the lateral spines of adult blue crabs

were a better technique for tracking movements than are tags

placed in the branchial chamber through a slit in the carapace,

external tags inserted into slots on the carapace, or a Nesbit modification of a Petersen tag on the lateral spine. Fannaly (1978)

reported high rat~s of retention of a spaghetti-type tag by juvenile

and adult blue crabs ~80 to 150 mm carapace width) following

molting in the laboratory. The tag consisted of a monofilament

leader (30 mm) and anchor to which plastic tubing was connected.

The tags were insert~d through the thick membrane at the dorsoposterior articulation of the fifth pereopod.

Epidermal and hepatopancreas aClivat ion

Epicuticle formed and spine formation begins

Exocuticle secretion begins

Major portion of skeletal resorption

Ecdysial sutures open

Rapid water uptake and exuviation

Terminal stage in adult female; no further growth

About to molt

Molt

Peeler

Permanent

anecdysis

Stage D Proecdysis

Do

DI

D,

D3

D,

Stage E

a!. 1976). Molting frequency of juvenile blue crabs was greater

when individuals were fed live, adult brine shrimp, Artemia

salina. than when fed laboratory formula dicts over an 18-wk

period (Millikin et a!. 1980). Importantly, limited data from the

same study indicated that growth per molt did not decline in blue

crabs that molted more frequently.

Intermolt periods during early juvenile development of blue

crabs that occurred in the laboratory are listed in Table 6 (Millikin

unpub!. data). Mean duration of the first and second crab stage was

longer in a salinity of 25%0 and 23.5 ± 1.5°C (Millikin unpub!.

data) than in studies conducted at a salinity of 30%0 and 25°C

(Rosenberg and Costlow 1976; McKenney and Costlow 1981).

Mean duration of the first crab stage in control groups was

3.0±OA d (SD) or 4.5±0.25 d (SE) (Rosenberg and Costlow

1976; McKenney and Costlow 1981).

Female blue crabs rarely, if ever successfully, molt again

following their maturity molt (terminal molt) (Churchill 1921; Van

Engel 1958). An adult female in the process of molting (buster

stage) was reported by Abbe (1974) and four different mature

females in the buster stage have been observed by Cronin7. Olmi

24
Blue crabs frequently move inshore during rising tid , forage in

the intertidal zone, and move offshore during ebb tide (Nishimoto

and Herrnkind 1978). According to both field and wave tank

experiments on "eye-capped" individuals, blue crabs are capable

of using the beach slope and wave surge for directional orientation.

3.52 Schooling

According to Van Engel (\958), schools of adult female crabs

have been observed migrating downstream in tributaries of the

Chesapeake Bay. Also, schools of "sea run" or "ocean" crabs

appear in late July or early August in the Lynnhaven Roads area

(Virginia), consisting of older, adult females with carapaces

heavily encrusted with barnacles (Van Engel 1958). However, it is

more accurate to describe these groups as aggregations rather than

schools (see Partridge 1982).

3.53 Responses to chemical stimuli

Detection of seawater solutions of freeze-dried clam extract by

blue crabs was assessed by noting rates of antennular flicking and

gill bailing activity (Pearson and alia 1977). Blue crabs detected

the extract solution at 10- 15 gil, while feeding behaviors were

elicited at concentrations of 10- 1 to 10- 2 gil. Blue crabs were

reported to detect naphthalene concentrations of 0.12 mg/l

(Pearson and alia 1979). Further study revealed that the threshold

concentration at which 50% of the crabs detected naphthalene was

8.5 x IO- R mgll (Pearson and alia 1980). Lack of information

regarding toxicity of naphthalene to blue crabs, especially sublethal effects, prevents comparison of detection threshold concentrations with concentrations which may be toxic.

4 POPULATION

Blue crab populations or stocks in the fishery management

context have not been adequately defined for adjacent estuarine

areas along the Atlantic coast or gulf coast of the United States.

Recently, a workshop was convened to attempt to develop methods

for analysis of local blue crab stocks, particularly in Chesapeake

Bay (Cronin footnote 7). Results of this workshop are not yet

available. Miller et al. (1975) suggested as a result of a 2-yr

sampling study that the Chesapeake and Delaware Canal provides

little, if any, access for crab migration from Delaware Bay

westward to upper Chesapeake Bay. Cole and Morgan (1978a)

suggested that observed genetic similarity (gene frequencies of

polymorphic enzymes, percentages of polymorphic enzymes, and

average heterozygosity) between Chesapeake Bay and Chincoteague Bay populations is the result of larval intermixing of zoeae

just outside Chesapeake Bay. However, Porter (1955) reported that

adult female crabs in Chincoteague Bay are significantly smaller

than those in either Delaware or Chesapeake Bay populations.

Possibly, adult females that occur in the Chincoteague Bay are

smaller from spending more of their life cycle in waters containing

less available food.

4. I Structure

4.11 Sex ratios

Sex ratios of adult blue crabs differ spatially with respect to

salinity, and temporally with respect to peak occurrence of mating,

and female migrations to high salinities for spawning. Truitt (1939)

25

reported that 64% of all (adult) crabs captured in the Capes section

(Cape Charles to Cape Henry, Va.) of Chesapeake Bay during June

and July were egg-bearing females. Crabs taken from areas below

the York River and Cape Charles averaged about 9.8% males over

a I-yr period, whereas males may reach 90% of the total adult

population in headwaters of the Maryland portion of Chesapeake

Bay. Sex ratios (male to female) for crabs sampled in the

Chesapeake and Delaware Canal combined with immediately

adjacent waters in Chesapeake Bay and Delaware Bay (in salinities

of 0.6 to 2.3Yoo) ranged from 1.0 (June 1972) to 5.8 (August 1971)

for recruitment stage classification (\-59 mm carapace width), 0.4

(June 1972) to 4.7 (September 1971) for growth stage classification (60 to 119 mm carapace width), and I. 9 (August 1971) to 16.0

(August 1972) for mature stage (> 12U mm carapace width)

(Miller et al. 1975). Much lower male to female sex ratios varied

from 0.3 to 1.8 for crabs sampled by the same methods in Tangier

Sound, Md., a higher salinity area than the Chesapeake and

Delaware Canal and adjacent waters (Miller et al. 1975). Tagatz

(1968a) reported that 60 to 90% of crabs sampled in the lower 24

km of the SI. Johns River, Fla., from January through September,

were females while only 10 to 25% of crabs sampled from October

through December were female. In the next 24 km upstream, 75%

of the crabs sampled from December through February were

females, whereas from March through November 15 to 55% were

females. Males dominated the lower salinity areas (farther upstream than 48 km from the mouth of the river). According to

Eldridge and Waltz (1977), incidental crab catch contained 84%

females during shrimp trawling operations conducted primarily

during winter in high salinity areas (e.g., sounds) within 0.8 to 4.8

km offshore of South Carolina. Crab pot catches, which are

primarily in lower salinity waters, contained 72% m~Jes and 28%

females in southern estuaries of South Carolina (Eldridge and

Waltz 1977). Van EngelS reported that 85 to 95% of the winter

dredge crab fishery of Virginia is adult females.

4. 13 Size composition

Tagatz (1965) monitored size composItIOn of commercial

catches of the St. Johns River, Fla. (80 samples from crab pot

catches yielding 11,620 individuals and 12 samples from trawl

catches yielding 1,655 individuals). The modal 10 mm carapace

width interval for both gear types was 160 to 169 mm. Mean

carapace widths from three different areas along the 51. Johns

River ranged from 155 to 166 mm.

Mean carapace widths of males from commercial crab pot

catches from five estuaries in South Carolina ranged from 135.5 to

137.9 mm, while mean carapace widths of females ranged from

148 to 154 mm (Eldridge and Waltz 1977).

4.2 Abundance and density (of population)

4.22 Changes in abundance

Variations in abundance of blue crabs can be estimated to some

extent by examining fluctuations of annual commercial catches,

although factors other than abundance such as demand and exvessel price also affect blue crab landings. Among many factors

that cause wide t1uctuations in annual population sizes of blue

crabs are: I) Maturation at an early age (I to 2 yr), 2) short Iife span

'w. A. Van Engel. Virginia Institute of Marine Science. Gloucester Point. VA

23062, pers. commun. August 1982.
(3 to 4 yr), and 3) natural catastrophes that may cause higher than

average mortality rates during hatching or winter (Dintaman 9 ).

In the mid-1930's, the decline of the Rhode Island blue crab

fishery began (Jeffries 1966). By the early 1940's, commercial

landings were no longer recorded (Table 6).

In a 3-yr sampling study in Core Sound, N.C., juvenile crabs :s

4 cm reached greatest numbers between December and April in

creeks, followed by migrations to deeper waters, causing low

annual amounts of this size range in May (Dudley and Judy 1973).

These authors proposed using the mean catch of juvenile blue crabs

from creeks in this area as an annual index of juvenile abundance

and predictive measure for subsequent adult abundance.

4.24 Changes in density

Miller et a!. (1975) reported catch density values (number of

crabs/m 2 x 10- 3) of three size classes of crabs (recruitment: I-59

mm carapace width, growth: 60-119 mm carapace width, and

mature: 2: 120 mm carapace width) in Delaware Bay and

Chesapeake Bay (Elk River) near the Chesapeake and Delaware

Canal, the Chesapeake and Delaware Canal itself, and T~lngier

Sound. For "recruitment size," values changed from 1.27 in June,

to 0.88 in August, to 0.28 in September in the Elk River, and 0.55

in June, to 0.08 in August, to 0.07 in September in upper Delaware

Bay. For the "growth" size class, catch density vaJue~ varied from

1.99 in June, to 2.49 in August, to 1.04 in September in the Elk

River and from 0.65 in June, to 1.56 in August, to 0.16 in

September in upper Delaware Bay. Catch density values were at

least two to three times greater for crabs in each size class in

Tangier Sound than in the Elk River sampling area for similar

sampling periods. Also. catch density values in the Chesapeake

and Delaware Canal were 0.8 or lower for the growth size class,

0.1 or lower for the recruitment size class, and 0.3 or lower for the

mature size class.

4.3 Natality and recruitment

4.33 Recruitment

Time to recruitment to blue crab fisheries varies according to

geographical region, mostly as a function of peak spawning

periods and length of growing season. In Chesapeake Bay, crabs

hatched in May may reach 5 to 6 cm (carapace width) in November

and l2.7 cm the following August (15 mo to maturity), or crabs

hatched in August to September may reach 1.3 cm in November,

7.5 to 10 cm the following November, and 12.7 cm in May (20 mo

to maturity) (Van Engel 1958). In southern estuaries of South

Carolina, most recruitment to the blue crab pot fishery occurs

in September and October for males and Octuber and November

for females (Eldridge and Waltz 1977). In Texas and the SI.

Johns River, Fla., recruitment occurs approximately 1 yr following spawning; therefore peak recruitment in Texas is in May to

July and September to October (More 1969), while in the SI.

Johns River, peak recruitment occurs from March to September

(Tagatz 1968a).

Recruitment into the Maryland portion of Chesapeake Bay is

determined by reproductive success of blue crabs spawned in the

Virginia portion of the Bay and the extent of migrations of

"R. C. Dinlaman, Tawes Stale Office Building C-2, 580 Taylor Ave., Annapolis,

MD 21.j1J I. From "The status of the blue crab populalion in Maryland's oceanside

bays:' 21 p. (mimeogr.).

26

juveniles (Sulkin 1977). Year class strength in the Maryland

portion of Chesapeake Bay is determined primarily from sampling

areas in Tangier and Pocomoke Sounds, which are important

nursery areas for early juvenile instars (Dintaman 1982). Abundance of crabs in the size range of 61 to 120 mm carapace width that

are sampled in September and October provide a good indication of

the following spring's harvest in the Maryland portion of Chesapeake Bay and Maryland's oceanside bays (Dintaman footnote 9).

4.4 Mortality and morbidity

4.42 Factors affecting mortality rates

Factors affecting natural mortality rates of blue crabs include

predation. disease, extreme weather conditions, environmental

contaminants, and lack of food availability. Presumably, lack of

food availability is especially important for blue crab larval stages.

Difficulty in quantitating these factors is reflected in the small

amount of data present in the literature. Van Engel (1982b)

reported that extreme temperature declines during January 1977

and February 1978 in Chesapeake Bay were correlated with high

mortalities in Virginia's winter crab-dredge fishery. Also, during

winter low pressure centers accompanied by high winds in southern Chesapeake Bay often cause strong onshore surface currents

cuupled with strong offshore subsurface currents that result in

abrasion of blue crabs swept along the bottom in shallow waters

(Van Engel J982b). Gray crab disease was prevalent (percent

occurrence not reported) in samples of dead blue crabs during 1966

and 1967 from South Carolina (McKenzie] 970).

4.43 Factors affecting morbidity rates

See section 3.35.

5 EXPLOITATION

5.1 Crabbing equipment

5.11 Gear

In the early years of the Chesapeake Bay fishery, trotlines were

the principal gear used for catching hard blue crabs, but eventually

the crab pot (patented by Lewis in 1938) became the chief gear for

hard blue crabs except during winter (Van Engel 1962). The primary gear in winter in the Virginia fishery is the crab dredge, a

heavy, rectangular frame, bearing a 1.8 m (6-ft) toothed drag bar

on its lower edge, followed by a mesh bag made of ri ngs, cotton,

and twine (Van Engel 1962). Descriptions of commercial crabbing

gear include those by Wharton (1954) and Cargo (1954).

Typical Chesapeake Bay crab pots are commonly 0.6 m (2 ft)

square, top and bottom, and 53.3 cm (2l in) high, made of 18

gauge steel wire of 2.9 cm mesh (l'/a in) specially treated with zinc

(Warner ]976). Crab pots consist of two to four conical funnels

~erving as entry ports, a partition separating the "upper" and

"lower" ;,eetion, which utilizes a blue crab's tendency to swim up

and away from the bottom if alarmed, and a cylindrical bait "box"

in the center of the lower section (Warner 1976). Crab pots are set

about 30.5 m (100 ft) apart at the edges of river or bay channels of

Chesapeake Bay in depths of ] .8 to 18 m (6 to 60 ft), with actual

fishing time averaging 2.5 h/IOO pots (Van Engel 1962). The

nunlber uf pots used by commercial crabbers in Mississippi Sound

ranged trom 65 to 400 (Perry ]975).
Trotlines are plain cotton twine consisting of bait (eel, other

fishes, or chicken parts) tied intermittently to the line OA to 1.6 km

(0.25 to I mi) set in water depths of 1.5 to 4.5 m (5 to 15 ft). Heavy

metal objects, such as engine blocks, are used as anchors at either

end, attached to 1.2 to 1.5 m (4 or 5 ft) of chain, a rope pennant.

and several plastic (bleach) bottles serving as marker buoys

(Warner 1976).

Otter trawls used in North Carolina are generally 10.6 m (33 £I)

across the mouth and have 10 cm stretched mesh at the cod end

when used primarily to harvest crabs and a 3. I cm stretched mesh

at the end when used primarily to harvest shrimp (Fischler 1965).

Crab dredges and crab scrapes are described briefly in section 5AI.

5.12 Boats

Dredge boats in the Virginia winter dredge fishery are usually

9.7 to 12.2 m (32 to 40 £I) long, with two dredges sometimes towed

simultaneously. Boats usually contain a captain plus a crew of two

or three persons (Van Engel 1962).

5.2 Blue crab areas

5.21 General geographic distribution

See section 2.1.

5.22 Geographical ranges

See section 2.1.

5.23 Depth ranges

See section 5.1.

5.24 Conditions of the grounds

Pollution by Kepone contamination (Roberts and Leggett 1980)

has forced the closure of the blue crab fishery in the James River,

Va. Total blue crab catch from the James River estuary declined

90% from 1972 through 1975 (Schimmel et al. 1(79), whereas

landings from the adjacent Rappahannock River estuary did not

decline appreciably over the same periC'd.

5.3 Crabbing seasons

Georgia and Gulf Coast States (Florida, Alabama, Mississippi,

Louisiana, and Texas) have no closed season for harvesting blue

crabs (Bearden 1978). In South Carolina it is unlawful to trawl

specifically for blue crabs in specified offshore areas and sounds

from April through November. In Virginia and North Carolina ir is

unlawful to catch blue crabs with crab scrapes or dredges between

I April and I December and 1 April and 30 November, respectively

(Bearden 1978). Capture of hard crabs is unlawful in Maryland

waters between I January and I April. North Carolina and Virginia

have designated sanctuary areas in which crabbing is unlawful

during certain seasons (Bearden 1978).

27

5.4 Crabbing operations and results

5AI Effort and intensity

Catch per unit effort (pounds/pot day = total pounds caught on a

given day 7 number of pots used on a given day) in Mississippi

Sound averaged over monthly periods ranged from 1.3 to 7.2 Ib

(0.6 to 3.3 kg) over a 3-yr period for commercial size crabs (Perry

1975). According to Perry (1975), rises in catch per unit effort

accompanied I) migration periods of mature females into the

Sound, 2) fall-winter arrival of females from Lake Borgne, La.,

and 3) maturation of both sexes in early spring with warming

temperatures.

Several studies have evaluated catch efficiencies of various

sampling gears, both modified and unmodified, for several size

classes of blue crabs. A crab dredge divided equally into lined (1.3

em mesh) and unlined (7.6 cm mesh) portions had significantly

better catch efficiency of crabs < 60 mm carapace width in the

lined section (Sulkin and Miller 1975). Also, a lined crab dredge

(1.3 cm mesh) had significantly better catch efficiency than a lined

oyster dredge (1.2 cm mesh) for two size classes (I-59 and 60- I 19

mm carapace width). For experimental sampling purposes, no

single gear type effectively samples all Juvenile stages and adults,

all seasons, depths, and bottom types considered (Miller et al.

1980). Comparison of a push net (81.3 cm wide, 60.9 cm high, 0.6

cm stretch mesh bag), a crab scrape (96.5 cm wide, 38.1 cm high,

0.6 cm stretch mesh net), and an otter trawl (working width of 3.6

m, cod end lined with 0.6 cm stretch mesh netting) indicated that

the crab scrape had highest catch efficiency (number of crabs/m 2 )

in each of three carapace width size classes (I to 20 mm, 20.1 to 40

mrn, 40.1 to 60.0 mm). Also, Miller et al. (1980) mentioned that a

crab scrape can be operated by one person and is cheaper than an

otter trawl.

Sampling studies in the York River, Va. (0 to 15 mi upstream),

evaluated the effects of tow duration and attachment or removal of

a tickler chain (20 mm link) with a 9.1 m semiballoon !Iawl on

catches of blue crabs (Chittenden and Van Ellgel 1972). Fiveminute tows produced mean catch values similar to either 10- or

15-min tows. Catches almost tripled when the tickler chain was

attached to the trawl.

Annual sampling of blue crabs (all sizes) In Maryland from MilY

through October is conducted with a 4.9 m (16 ft) semiballoon olter

trawl for 6 min at a speed of 4 kn (Dintaman 1982, footnote 9).

Average catch values were lower for 3.6 m (12 ft) and 7.6 m (25 ft)

trawls conducted on an experimental basis in the mid-1970's

(Dintaman 1982).

5A2 Selectivity

An experimental self-culling crab pot containing two escape

rings (6.35 cm diameter) in the top and'one (6.35 cm diameter) in

the bottom, trapped only 9.5% (72 of 761 crabs caught) sublegal

crabs « 127 mm carapace width) compared with 37.60/, sublegal

crabs (398 of 1,059) caught in commercial crab pots without

escape rings (Eldridge et al. 1979). According to Eldridge et al.

(1979), advantages of a self-culling crab pot (3 escape rings, each

6.35 cm diameter) include: I) Less culling required by law abiding

crabbers, 2) less injury to sublegal crabs, and 3) reduced Jaw

enforcement problems and fewer deliveries of illegal crabs to

processors. Similar studies for ideal escape ring size may be

necessary in various regions having a blue crab fishery if length to
width relationships of crabs are appreciably different from those in

South Carolina (Eldridge et al. 1979).

5.43 Catches

Commercial blue crab landings thard crabs) along the Atlantic

and gulf coasts of the United States are Iisted in Tables 7 through 9.

Also, commercial fisheries for the peeler and soft crab stage of blue

crabs are listed in Table 10. Only Maryland, Virginia, Louisiana,

and North Carolina currently have important fisheries for soft shell

crabs. Suggested methodology for estimating annual total harvest

for blue crabs using simple or stratifieu random sampling of

licensed crabbers via questionnaires was described by Summers,

Hoffman, and Richkus (1983) and Summers, Richkus, Hoffman,

Bonzek, King, and Burch (983).

Crab pots and otter trawls are principal gear types for hard crab

harvesting in several states. The Georgia blue crab fishery consists

mainly of otter trawls and crab traps or pots (Palmer 1974). Over a

5-yr period (1972-76) a mean of 13.4± 5.8% of the annual hard

crab harvest resulted from shrimp otter trawls and 0.9 to 12.8% of

the harvest ill different years resulted from crab otter trawls

(Wheeland 1975; Pileggi and Thompson 1976, 1978, 1980; Wise

Table 7.-Commercial blue crab landings (hard crabs) of New England, Middle

Atlantic, and Chesapeake Bay fisheries (in thousands of kg).'

Rhode Conn~r· ew New

Year 1,land ticut Yorl- Jersey Delaware Maryland Virginia

1939 715 17 18.8 2.174.8 348.7 10,934.4 12,243.0

1940 53.6 29.0 364.3 125.9 6,824.2 10,449.5

1941 5,436.8 7,135.4

1942 r.Y 01 333.5 177.1 6,377.9 8,464.4

1943 4.8 176.9 10.9

1944 132.9 10.9 7,788.4 10,863.7

1945

1946 U.I 11,557.8 12,196.6

1947 1.7 286.0 566.6 11,608.0 15,587.4

1948 1.4 6.7 299.6 662.4 9,361.7 19,382.7

1949 15 905.4 1,013.8 10,048.0 18,456.3

1950 05 3.8 967.7 2,002.9 12,495.1 21.063.6

1951 0.8 1.2 474.7 2,109.3 12,338.1 17,061.7

1952 1.7 25.8 484.2 567.5 12,484.6 15.225.9

1953 0.4 ::!.2 340 271.5 784.5 11,9711 14,677 .4

1954 13.6 0.9 2U.4 395.4 1.322.0 8,659.1 14,741.0

1955 0.9 19.5 286.0 1,276.6 6,915.3 12,206.7

1956 I.X 4.1 314.1 1,706.1 9,628.4 11,688.2

1957 I 4 4.1 551.1 2,234.6 12,879.5 11,295.5

1958 14 8.2 394.1 I, 114.1 12,3011 8,060.3

1959 U.9 09 446.3 749.1 9,618.9 9,601.2

1960 1.4 0.4 695.9 947.5 12,288.9 17,828.6

1961 1.8 0.4 2933 344.6 12,102.7 19,965.1

1962 09 683.3 855.3 12,558.1 24,366.6

1963 04 390.9 236.9 7,688.0 20.946.7

1964 0,4 258.3 142.1 10,233.2 23,413.7

1965 0.4 410.9 247.9 14,527.1 22,955.6

1966 310.5 259.2 13,789.3 28,933.9

1967 21U.5 130.8 11,162.9 24,890.1

1968 61.3 101.2 .4,242.6 20.357.8

1969 282.4 231.5 10,448.3 15 272.6

1970 0.4 340.9 276.0 11,320.5 195269

1971 0.4 500.8 460.4 11,8380 21.704.4

1972 652.4 1,158.6 10,660.8 22,043.5

1973 1,167.7 1,077.3 8,870.7 ]6.682.7

1974 1.2462 1,02U.6 11,195.6 18.545.9

1975 1,302.9 1,612.1 II,UI5.9 15807.8

1976 1,223.') 1,618.5 8.8212 116'15.5

'1939-67 = Fishery Statistics of the United States, U.S. Dep. Inter" Fish ',vildl

Serv., Stat. Dig. 1-61. 1968-76 = Fishery Statistic, of the United State" U.S. De~.

Cummer., NOAA, NMFS, Stat. Dig. 62-70.

28

and Thompson 1977). Hard crab harvest 1rom pots pi us traps

averaged 79.9 ± 11.5% over the same 5-yr period. In Alabama,

hard crab harvest from 1972 through 1976 averaged 95.9 ± I.9%

from crab pots with the remaining catch resulting from shrimp otter

trawl operations (Wheeland ]975; Pileggi and Thompson 1976,

1978, 1980; Wise and Thompson 1977). Likewise, in Mississippi

Sound, most of the commercial hard crab catch is harvested with

crab pots (Perry 1975). Crab pots produced 95.4± 2.3% of the

total annual hard crab harvest in Texas from] 972 through 1976.

Crab otter trawls in North Carolina contributed 15.6± 6.6% of the

total hard crab catch from 1972 through 1976, whereas crab pots

produced 76.7 ± 7.2% of the hard crab catch. Over a 10-yr period

(1965-75). Eldridge and Waltz (1977) estimated that 12% of South

Carolina's commercial crab landings were from trawl catches,

while most of the remaining catch resulted from crab pots.

The hard crab fishery of Maryland results mainly from crab pots

(62.1 ± 3.5%) and trotlines t37.4± 3.5%) as indicated by the 1972

to 1976 period (Wheeland 1975; Pileggi and Thompson ]976,

1978, 1980; Wise and Thompson 1977). This represents a larger

contribution (total kilograms and percentage of total catch) from

tro!lines than in any other blue crab fishery. Estimates of blue

crabs harvested in 1981 based upon simple and stratified random

sampling of licensed crabbers via questionnaires indicated that

commercial crab potters and trotliners accounted for 51 and 36% of

the annual harvest, respectively (Summers, Richkus, Hoffman,

Bonzek, King, and Burch 1983).

Table 8.-Commercial blue crab landings (hard crabs) of

South Atlantic fisheries (in thousands of kg).'

North South

Year' Carolina Carolina Georgia Florida' ...__._--

1939 1,295.7 1,133.5 999.8 2,038.9

1940 1,819.5 1,879.5 1,065.8 2,561.7

1949 '933.3

1950 3,032.9 1.302.6 2,282.5 2,799.6

1951 3,551.3 1,156.9 2,963.0 3,956.4

1952 2,797.4 ],182.5 4,294.0 3,692.6

1953 4,761.1 1,709.8 4,206.6 4,303.9

1954 4,416.1 1,129.5 4,830.6 4,462.9

1955 4,303.9 2,108.4 4,878.2 5,737.2

1956 3,743.2 863.5 3,878.1 5,346.7

1957 5,253.7 1,627.1 4,071.5 5,373.0

1958 5,685.4 2,196.9 4,623.0 7,577.3

1959 6,69\.5 2,166.5 5,727.6 9,310.6

1960 6,781.0 3,232.9 7,157.8 11,626.9

1961 7,2095 2,121 I 5,589.6 11,175.6

1962 5,54~.3 2,877.0 5,585.6 8,274.1

1963 8,551.1 4.012.0 6,583.0 9,851.3

1964 IO,~37.8 4,283.9 5,235.5 9,542.0

1965 10,139.6 3.368.7 4,657.6 12,058.7

1966 8,586.9 2,598.7 3,884.4 10,836.9

1967 6,479.5 2,382.1 3,857.1 10,576.4

1968 8.703.2 1,753.3 1,665.7 7,092.8

1969 10,0602 3,745.5 2,336.7 7,857.8

197U 9.479.5 3,155.3 3,219.8 10,244.5

1971 6,572.1 3,408.6 3,829.5 9,720.6

1972 6,119.5 3,369.6 4,112.8 7,700.3

1973 5,431.2 3,610.2 3,624.7 6,134.9

1974 5,976.5 3,426.8 4,599.5 7,993\

1975 5,026.7 2,896.5 4,024.7 7,7 ]4.4

1976 5,326.3 2,605.9 2,666.3 7,296.7

'1939-67 = Fishery Statistics ufthe United States, U.S. Dep.

Inter., Fish Wild!. Serv., Stat. Dig. 1-61. 1968-76 = Fishery

Statistics of the United States, U.S. Dep. Commer., OAA,

NMFS, Stat. Dig. 62-70.

'No data tor 1941-48.

'Includes Atlantic and gull coast catches, combined.

'Gult co"t catch only.
Table lO.-Peeler and soft shell blue crab fisheries in the United States (in

thousands of kg),1 ------

1 1939-67 = Fishery Statistics of the United States, U.S. Dep. Inter., Fish Wild!.

S·~rv., Stat. Dig. 1-61. 1968-76 = Fishery Statistics of the United States, U.S. Dep.

Cammer., NOAA. NMFS. Stat. Dig. 62-70.

Since 1957, oceanside blue crabs (hard and soft crabs) have

comprised between I and 7% of the total annual catch of blue crabs

in Maryland on an annual basis (Dintaman footnote 9). During

April, the oceanside bays contribute the majority of the total catch

of blue crabs in Maryland waters, most of which are intermolt, hard

crabs, The primary month for harvest of soft/peeler crabs is usually

May in the oceanside bays, followed by peak total blue crab

catches (hard, peeler, and soft crabs combined) in Chesapeake Bay

from June through September.

Peak harvest months for blue crabs in Alabama (Mobile Bay) are

July, August, and September (Tatum 1979). Blue crabs (hard) are

harvested primarily from March through October in North Carolina

waters, while soft shell crabs are harvested primarily from April

through July (Pearson 1951). Peak months for hard crab catches in

Louisiana are July and August, whereas lowest monthly yields

occur from December through March (Roberts and Thompson

1982). Peak crab pot catches occur from June to November in

South Carolina and the trawl fishery for blue crabs operates mainly

from December through March (Eldridge and Waltz 1977).

Though recreational fishing for blue crabs is substantial in

locations where important commercial fisheries exist, recreational

catch statistics for blue crabs are generally not available. Sport

crabbers were estimated to land about 22,500 kg (50,000 lb)

Winter crab dredge fisheries exist in New Jersey, Delaware, and

Virginia. Over the 1972 through 1976 period, crab dre 'ging

operations contributed an average of 18.6 ± 6.3% of New Jersey's

hard crab catch and 21.2±5.1% of Virginia's total annual hard

crab catch (Wheeland 1975; Pileggi and Thompson 1976, 1978,

1980; Wise and Thompson 1977). From 1974 through 1976,

Delaware's crab dredging operations contributed 15.3 ± 5.1 % of

that state's hard crab harvest. The large remainder of hard crabs

harvested in each of these states was from crab pots or traps

(81.3 ± 6.3% for New Jersey and 78.6 ± 5.1 % for Virginia).

Since 1960, usage of trotlines in Louisiana for hard crab capture

has steadily decreased from 75% of the total catch to 1% in 1977,

while hard crabs harvested by crab traps has increased from I to

98% over the same period (Roberts and Thompson 1982).

Generally, increases or decreases in catches of hard crabs from

one year to the next occur simultaneously for New Jersey and

Delaware (Table 6), probably due to dependence on the mouth of

the Delaware Bay as a major spawning site for several river

populations in both states. Similarly, hard blue crab landings in

Virginia and Maryland generally increased or decreased in the

same years due to the dependence of crab fisheries in both states on

the spawning area at the mouth of Chesapeake Bay (Table 6).

Peeler and soft crabs in Maryland were harvested mainly by crab

scrapes (77.8 ± 4.0%), pound nets (10.1 ± 2.2%), and peeler pots

(8.2± 2.0%) over a 5-yr period from 1972 through 1976. Over the

same time interval, 55.1 ± 10% of Virginia's peeler and soft crab

harvest resulted from peeler pots, 15.8 ± 8.7% resulted from crab

scrapes, and 28.9± 11.9% resulted from pound nets.

Table 9. -Commercial blue crab landings (hard crabs) of

Gulf Coast States (in thousands of kg).'

Year' Alabama Mississippi Louisiana Texas

1939 253.1 667.1 5,097.8 184.5

1940 626.8 675.7 6,384.1 1146

1948 1,077.1 2,498.2 9,583.8 238.9

1949 966.1 1,890.0 8,114.7 169.8

1950 271.7 1,834.2 5,950.3 175.8

1951 503.5 736.9 3,954.2 127.0

1952 297.5 783.6 3,329.6 153.4

1953 493.5 641.0 3,691.4 196.1

1954 441.3 570.2 3,216.6 172.1

1955 732.3 800.4 4,908.2 161.6

1956 329.2 898.5 4,268.5 88.5

1957 663.7 1,089.6 3,885.8 91.2

1958 5366 964.3 4,238.5 258.8

1959 496.2 1,363.3 4,344.8 541.2

1960 226.5 1,276.6 4,562.7 1,301.6

1961 380.5 1,137.3 5,407.1 1,305.3

t'l62 287.8 411.8 4,323.4 2,030.7

1963 588.8 504.8 3,623.8 1,352.9

1964 799.9 5838 2,584.0 1,127.0

1965 822.6 768.2 4,214.0 1,644.4

1966 991.1 661.5 3,625.0 1,261.2

1967 1.068.3 461.8 3,431.8 1,191.8

1968 898.9 515.7 4,336.2 1,854.1

1969 871.7 78'l.9 5,267.3 2,879.7

1970 638.8 920.2 4,655.3 2,508.3

1971 906.6 571.6 5,532.4 2,637.7

1972 732.3 618.3 6,847.7 2,934.7

1973 952.5 824.0 10,478.3 3,123.9

1974 829.0 756.8 9,370.6 2,763.9

1975 744.6 516.2 7,783.3 2,720.4

1976 589.8 6061 6,905.8 3,027.3

' 1939-67 = Fishery Statistics of the United States, U.S. Dep.

Inter., Fish Wild!. Serv., Stat. Dig. 1-61. 1968-76 = Fishery

Statistics of the United States, U.S. Dep. Commer., OAA,

NMFS, Stat. Dig. 62-70.

'No data for 1941-47.

29

New

Year Jersey Delaware Maryland Virginia

1939 278 11.5 1,468.1 1,263.5

1940 24.9 'l.9 812.9 897.6

1941 379.7 776.1

1942 24.3 4.2 746.9 656.2

1943 9.9 2.9

1944 9.6 2.9 5048 1,099.8

1945

1946 1,173.0 745.4

1947 0.2 1.4 1,366.7 1,109.0

1948 1.4 4.0 845.5 1,257.8

1949 5.8 26.2 1,058.3 1,139.4

1950 6.2 4.3 1,315.7 1,466.6

1951 4.6 3.0 412.7 663.1

1952 3.6 20 7208 1,002.2

1953 2.3 0.9 865.3 1,171.8

1954 3.2 1.4 361.8 585.2

1955 4.'l 3.6 545.2 820.8

1956 4.1 0.9 830.4 824.5

1957 2.3 0.4 718.7 7328

1958 2.3 0.4 1,482.3 611.'l

1959 OA 772.7 689.6

1960 77 18.2 1,265.8 721.9

1961 25.9 24.5 1.222.1 711.9

1962 70.S 11.8 1,766.'1 611.5

1963 15.0 1.4 957.0 430.8

1964 4.5 09 1,588.1 453.1

1965 15.0 5.4 1,2231 4899

1966 1.8 01 8553 466.7

1967 2.3 14 992.4 552.5

1968 0.4 I) 9 4549 365.9

1969 4.1 41 1,021.0 894.8

1970 15.G 45 717.3 413.0

1971 5.9 ~.2 694.6 314.6

1972 6.8 33.1 715.1 389.5

1973 10.4 15.4 686.9 446.3

1974 572 386 8272 369.6

1975 17.7 750.9 3423

1976 409 6692 345.4

North

Carolina

78.5

1298

'14.8

75.9

56.4

76.2

43.1

118

32.2

29.1

345

59.3

41.3

45.9

44.5

37.7

31.8

107.6

572

3'1.0

38.1

42.2

27.2

22.2

22.7

20.4

15.0

9.1

91

Louisiana

97.5

1143

3999

206.7

164.8

1590

203.4

221.6

206.6

263.8

272.4

2502

261.9

274.7

233.4

281.5

156.2

149.4

90.8

92.6

5H.1

663

128.9

89.4

40.9

57.7

46.3

54.0

43.6

49.9

40.0
annually in Mississippi waters (Herring and Christmas 1974, cited

in Perry 1975). Tatum (1979) estimated that the recreational catch

equals about 209~ of the commercial catch in Mobile Bay, Ala.

6 PROTECTION AND MANAGEMENT

6. I Regulatory measures

6.11 Limitation or reduction of total catch

abunJance in Chesapeake Bay as well as methods of restoration of

eelgrass beds and evaluation of limiting factors for growth of

eelgrass. Orth (1977) induced significant increases in biomass of

eelgrass in fertilized (5 or 10% NH4 N0 3 , 10% P2 0 S , 10% K2 0)

plots compared with unfertilized, adjacent plots in Chesapeake

Bay off Church Neck on the "Delmarva" Peninsula, Va.

6.5 Artificial stocking

Table ll.-Minimum size limits and laws regarding protection of females for

various slales having blue crab fisheries.'

Minimum sizes allowed for harvest of hard crabs, peekrs, and

soft crabs in various states are listed in Table II. Since the

minimum harvestable size for hard crabs is 10.2 cm (4 in) in

Alabama, percentage of immature females compared with mature

females harvested should be evaluated to determine if the minimum size is permitting capture of too many immature females.

State

, Delaware

Maryland

Virginia

North Carolina

S,'uth C..rolina

GeorglJ.

Hurida

MisSissippi

Alabama

Louisiana

Minimum size limils

Hard crabs -5 in

Hard crab,-5 in'

Peelers-3 in

Soft crabs-·-3.5 in

Hard crabs-5 in

(Adult females, peelers

Jnd sutt crabs exempt'

Hard crabs-5 in

(Peeler> exempt)

Hard crabs·- 5 in

(Crabs in commen':lai

shedding floats exempll

Hard crabs-5 in

Peelers--3 in

Soft crabs- 3 in

Hard crabs- 5 in

Nunt"

Hard crabs-4 in

Hard crabs-5 in

Soft crabs-4.5 in

Laws protecting females

Possession of egg bearing females

caught in Delaware waters unlawful

Possession of egg bearing females

caught in Maryland waters unlawful

Unlawful to catch female crabs within

a designated sanctuary area from

15 May to 15 September

None

Possession of egg bearing females

caught in South Carolina waters

unlawful

Unlawful to retain spawning females

Mayor June

Unlawful 10 sell egg bearing blue crabs

Unlawful to harvest sponge crabs

from south of intracoastal waterway,

commencing at Alabama-Mississippi

boundary and running west 10 Gulfport-Ship Island Channel

None

Retention of berried females unlawful

An attempt to supplement a natural population of blue crabs near

Crisfield, Md., by means of ovigerous females in shedding floats,

failed because of low survival of the females, possibly as a result of

transport from the areas of capture (Graham and Beaven 1942).

Additionally, larval survival rate was low since salinity was

probably below the minimum level of l8A'. for successful hatching

(Davis 1965) and below the minimum level of 20Y", for early larval

development (Costlow and Bookhout 1959). Dramatic increases

oc,:ur in survival rate of blue crabs in the laboratory beginning with

the first crab stage (Millikin 1978; Millikin et al. ]980) compared

with larval stages reared in the laboratory (see section 3.22). This

suc:gests that any attempts to supplement natural populations

would more likely be successful if early juvenile stages were used.

7 SOFT CRAB SHEDDING OPERATIONS

7. I Use of cultured blue crabs

Blue crabs are often harvested in the peeler stage (I to 3 d

premolt) in Maryland, Virginia, North Carolina, and Louisiana

for subsequent culture until ecdysis in shedding floats in natural

waters or land-based flow-through shedding tanks located near

natural waters. Blue crabs still in thl' soft shell stagl' (A 1),

especially < 3 to 6 h after molting, are removed from water te,

prevent further shell hardening (exocuticle mineralization) and

packed in ice for shipment or stored frozen for eventual human

consumption. Also, peeler crabs of a minimum size of 3 in (7.6 cm)

are often sold as bait for sport fishing in Chesapeake Bay (Haefner

and Garten 1974).

7.2 Procurement and mai ntenance of stock

I From Bearden (1978).

'From Miller (1~7n)

'Mature females m..y be harvested in Maryland regardless of size.

6. ]2 Protection of portions of population

Laws protecting gravid (i.e., berried, sponge, egg bearing)

females in various states are listed in Table II. Eldridge and Waltz

(1977) reported that 85'!c of immature females captured in

commercial crab pots were less than the minimum legal size of 5 in

(] 27 mm) carapace width in several South Carolina estuaries.

Therefore, the authors suggested that retention of immature

females, regardless of actual size, should be discouraged_

6.4 Control or alteration of the biological features of the

environment.

6.41 Control of aquatic vegetation

The importance of eelgrass as a habitat for blue crabs has

prompted investigations to evaluate the causes for its decline in

30

Blue crabs are harvested in the peeler stage by several gear

types. mainly a, a function of mating season and location of

vegetated areas which can provide refuge for crabs from predators

during the soft shell stage. Peeler pots are larger versions of

commercial crab pots but with smaller mesh (2.5 cm). Usually,

peeler pots are "baited" with two or three live, adult male crabs

which may attract as m~ny as 20 or 30 female peelers approaching

their tcrminal molt into the peeler pots (Warner 1976). Use of bare

pOh (without bait or large male crabs) is less common and used in

areas dcvoid of bottom vegetation, where the pot may appear to be

a refuge to peeler stage crab,. Peeler pounds consist of a wire lead

(2.5 cm mesh) oriented perpendicular to the shore which channels

peelers into the head section which is 0.08 x 0.11 x 0.14 m (3 x

4 x 5 ft) which is not completely covered at high tide (Otwell

and Cato 1982)_ Crab scrapers in Chesapeake Bay harvest the

largest portion of peelers and soft crabs, catching as many as 200 to

500/d during peak occurrence of mating, mostly in seagrass beds

I Warner I ':176). The first peak occurrence of peelers in Chesapeake

Bay occurs in the third or fourth week of May (water temperature

approaching 18°C) near a full moon (Warner 1976). Experimental

work in South Carolina, designed to determine the best methods for
7.6 Foods: feeding

Blue crabs in the peeler stage or buster stage do not require food,

since feeding activity temporarily ceases until after molting (Van

Engel 1958; Haefner and Garten 1974; Johnson 1980). Observed

final feeding activity prior to ecdysis occurred at variable time

intervals for 79 molts undertaken by 68 crabs (2.9 to 6.9 cm

carapace width) reared in the laboratory (Millikin unpubl. data)

as follows:

are often about 3.7 m long x 1.1 m wide x 0.3 m to 0.45 m deep

with 23 cm of water depth in the float (Beaven and Truitt 1940).

Nicking, a process whereby crabbers push in the free or movable

segment (dactyl) of the cheliped until it snaps and becomes

nonfunctional, is often performed to reduce cannibalistic behavior

in shedding operations. However, excessive bleeding or swelling

may result in lowered rates of successful molting (Beaven and

Truitt 1940; Haefner and Garten 1974) or potentially harmful

bacterial infection (Johnson 10).

Although blue crabs occur in a wide range of salinities (see

section 1.36), recently harvested peelers should not be placed in

shedding tanks having a salinity more than 10%0 different from the

area of harvest (Haefner and Garten 1974).

Eyestalk removal from the megalopal stage of blue crabs in the

laboratory promotes molting (Costlow 1963b), since the moltinhibiting hormone occurs in the X-organ/sinus gland complex

within the eyestalks. Still, removal of eyestalks of blue crabs

of peeler size or larger is not recommended because of high

mortality rates of subadults and adults following this procedure

(Van Engel 1982a).

harvesting peelers in that area, indicated a peak run of female

peelers preparing for their terminal molt during the waxing and

waning of the full moon in April (Bishop et al. 1982, 1983).

Optimal methods for peeler crab capture in South Carolina were

evaluated by comparing hard crab pots, fyke nets, bush lines,

habitat pots (similar to hard crab pots except with 2.5 cm mesh,

covered with interwoven plastic flagging tape and unbaited), and

peeler pots (Bishop et al. 1983). Crab scrapes were not evaluated

due to the lack of subtidal grass beds in South Carolina. Bush lines

and fyke nets were unsuccessful because of the strong tidal currents (> 1.8 km/h) and large tidal amplitude (1.5 m = mean) in

South Carolina. Large intertidal areas of marsh may also have

rendered bush lines less effective. Habitat pots captured 2.5 times

as many male peelers as female peelers and 2.5 times as many late

premolt crabs (6 d or less premolt) as early premolt crabs (7 to 14 d

premoIt). Hard crab pots captured the highest amount of peelers/

gear-day, but culling and damage to peelers from intermolt crabs

reduces this gear's effectiveness. Peeler pots only captured good

numbers of peelers during the full moon of April. Bush lines

are unique to the Louisiana soft crab fishery and consist of six or

seven branches of wax myrtle, Myrica cerifera, bOtind together

at the bases and tied to the crabbing line at regular intervals

(Jaworski 1972).

Soft shell crab production has historically been low in South

Carolina, Georgia, Florida, Alabama, Mississippi, and Texas.

According to Perry et al. (1982), minimal growth of the soft crab

industry in these states is primarily the result of lack of a reliable

source of supply of peeler crabs.

Initial investigations have suggested that heated effluents in

discharge canals from power plants may provide a feasible culture

medium for soft shell crab production during winter (Reimer and

Strawn 1973; Parker et al. 1976). The main season for soft shell

crab production is April to October in Chesapeake Bay and March

to October in Louisiana (Reimer and Strawn 1973).

7.4 Spawning

See sections 3.17, 3.21, and 3.22.

Last day offeeding activity

prior to molting (d)

7

6

5

4

3

2

1

Frequency (number

ofmolts)

3

9

6

7

14

26

14

7.5 Shedding operation management

Molting signs on the translucent penultimate and ultimate

segments of the fifth pair of pereopods (swimming paddles) of

juvenile and adult blue crabs include the following: 1) White sign

or line (7 to 14 d), 2) pink sign or line (3 to 6 d), and, 3) red sign or

line (I to 3 d premolt) (Van Engel 1958; Warner 1976; Perry et al.

1982). External signs often associated with the red sign stage are

possession of well developed, limb buds due to regeneration of

autotomized pereopods or changes in the color of the abdomen of

an immature female approaching her pubertal molt from creamy

white to reddish purple (Perry et al. 1982). Other external

characters related to molt stage that are commonly used within

commcfcial shedding operations or by crabbers are: 1) Papershell

(slightly stiff exoskeleton, 12 h postmolt), 2) buckram (crinkly

hard exoskeleton, 24 h postmolt), and 3) external carapace similar

to intermolt (72 h postmolt) (Warner 1976).

At least four or five shedding floats or tanks are desirable to

separate various molt stages, such as white line, pink line, red line,

and busters (crabs that are molting) (Van Engel 1958) into different

containers (Bearden et al. 1979). Pine wood shedding floats

constructed with laths on the sides and ends for water circulation

31

It is noteworthy that these crabs were acclimated to laboratory

culture conditions, whereas recently harvested peelers that are

placed in shedding tanks do not have time to acclimate before

ecdysis. Nevertheless, these data strongly suggest that blue crabs

in natural waters derive nutritional benefit from feed consumption

up to and including part of the "red sign stage." Therefore,

laboratory-acclimated crabs should be offered food at least through

the white sign and pink sign stages prior to molting.

7.7 Diseases

See section 3.35. Shell disease and paramoebiasis are two

diseases that sometimes occur in shedding operations (Sindermann

1977a; Johnson I977f). Also, if early premolt crabs that are still

feeding are mistakenly placed in shedding tanks with late premolt

peeler stage crabs, bacterial infection in late premolt peelers may

result from wounds inflicted by the early premolt crabs.

'0 P. T. Johnson, Northeast Fisheries Center Oxford Laboratory, National Marine

Fisheries Service, NOAA, Oxford. MD 21654, pers. conunun. July 1982.
ACKNOWLEDGMENTS

We thank Phyllis T. Johnson for contributing Table 2 and

portions of '. Viral infections" and "Bacterial infections" in

section 3.35. This manuscript benefitted from critical reviews by

L. Eugene Cronin, Phyllis T. Johnson, Charles Biernbaum, Peter

J. Eldridge, James M. Bishop, Patricia H. Fair, Alan R. Fortner,

and Lowell V. Sick. Sections of the manuscript were also reviewed

by Bruce Collette, John D. Costlow, and C. G. Bookhoul. We are

grateful to John Costlow and C. G. Bookhout for permission to use

their figures on blue crab larval stages of development which were

previously published in the Biological Bulletin. We thank Sandra

West for typing several drafts of the manuscript. M. Millikin is

especially grateful to Charles Wenner for his encouragement

during preparation of earlier drafts of this synopsis.

LITERATURE CITED

ABBE, G R.

1'174 Second terminal molt in an adult female blue crab, Callinecte, mpid"s

Rathbun. Trans. Am. Fish. Soc. 103:643-644.

ADKINS, G.

1972. otes on the occurrence and distnbution of the rhizocephalan parasite

(Loxo/hylacl" /exan"s Boschma) of blue crabs tCallineele' sapid"s Rathbun)

in Louisiana estuaries. La. Wildl. Fish. Comm. Tech. Bull. 2, 13 p.

19~2. The blue crab fishery of Louisiana. In H. M. Perry and W. A. Van Engel

(editors). Proceedings Blue Crab Colloquium, October 18-19.1979. p. 15-16.

ALDRIDGE, 1. B., and J. N. CAMERON.

1982. Gill morphometry in the blue crab, Callinectes ,apid", Rathbun

(Decapoda Brachyura). Crustaceana 43:297-305.

BASS, R. J., and 1. W. AVAULT. .Ir.

1975. Food habits. length-weight relationship. condition factor, acd growth

of juvenile red drum-, Sciaenops ocella/a. in Louisiana. Trans. Am. Fish.

Soc. 104:35-45.

BEARDEN, C M.

197B. Blue crab laws and regulations of the South Atlantic and Gulf states In

1977. In R. J. Rhodes and W. A. Van Engel (editors), Report of the Ndtional

Blue Crab Industry Workshop, September 10-11,1977, p. 53-70. Spec.

Rep. Applied Mar. Sci. Ocean Eng. 185, Va. Inst. Mar. Sci. and Spec. Rep.

S. C Mar. Resour. Cent.

BEARDEN. C. M., D. M. CUPKA, C H. FARMER, J. D. WHITAKER, and J. S.

HOPKINS.

1979. Information on establishing a soft shell crab operation in South Carolina.

S.C. Wildl. Mar. Resour. Dep., Educ. Rep. 10,11 p.

BEAVEN, G. E, and R. V. TRUITT.

[1940.J Crab mortality on Chesapeake Bay shedding floats. Chesapeake

BioI. Lab. Contrib. 33, 14 p.

BIDDLE, G. N.. M. R. MILLIKIN, P. H. FAIR, and A. R. FORTNER.

1978. The effects of dietary tiber on survival, growth and feed efficiency of

juvenile bluc crabs (Callinectes sapid",). In R. Nickelson (editor), Proc. 3rd

Annu. Trop. Subtrop. Fish. Tech. ConL Americas, p. 285-296. Tex. A&M

Univ., College Stallon.

BISHOP, J. M, E J OLMlllI, and J. D. WHITAKER.

1982. South Carolina's softshell program. In H. M. Perry and W. A. Van

Engel (editors). Proceedings Blue Crab Colloquium, October 18-19, 1979,

p. 159-165.

BISHOP, 1. M., E J. OLMIIII, J. D. WHITAKER, and G. M. YIANOPOULOS.

1983. Capture of blue crab peelers in South Carolina: An analysis of techniques. Trans. Am. Fish. Soc. 112:60-70.

BLAND, C E., and H V. AMERSON.

1974. Occurrence and distribution in North Carolina waters of La~enidi",n

callinec/es Couch, a fungal parasite of blue crab ova. Chesapeake SCI.

15:232-235

BLUNDON.1. A., and V. S KENNEDY.

1982a. Mechanical and behavioral aspects of blue crab, CallineeleS sapidllI

(Rathbun), predation on Chesapeake Bay bivalves. 1. Exp. Mar. BioI. Ecol.

65:47-65

1982b. Refuges for infaunal bivalves from blue crab, Callinec/es sapidus

(Rathbun), predation in Chesapeake Bay. 1. Exp. Mar. BioI. Ecol. 65:

67-81.

32

BODAMMER. J. E.

1978. Cytological observations on the blood and hemopoietic tissue in the

crab, Callinec/l's !iopidus. I. The line structure of hemocytes from intermolt

animals. Cell Tissue Res. 187:79-96.

BOICOURT, W. C.

19H2. Estuarine larval retention mechanism> on two scales. In V. Kennedy

(editor), Estuarine comparisons, p. 445-457. Acad. Press, N.Y.

BOOKHOUT. C G.. and J. D. COSTLOW, Jr.

1970. Nutritional effects of Artemia from different locations on larval development of crabs. Helgol. Wiss. Meeresunters. 20:435-442.

1975. Effects of mirex on the larval development of blue crab. Water, Air,

Soil Poilu!. 4: 113-126.

1977. Larval development of Callinectes simi/i, reared in the laboratory.

Bull. Mar. Sci. 27:704-72~.

BOOKHOUT. C G., J. D. COSTLOW, Jr., and R. MONROE.

1976. Effects of methoxychlor on larval development of mud-crab and bluecrab. Water, Air, Soil Poilu!. 5:349-365.

1980. Kepone effects on larval development of mud-crab and blue-crab.

Water, Air, Soil Pollut. 13:57-77.

BOOKHOUT, C G.. and R. J. MONROE.

1977. Eftects of malathion on the development of crabs. In F. J. Vernberg,

A. Calabrese, F. P. Thurberg, and W. Vernberg (editors), Physiological

responses of marine biota to pollutants, p. 3-19. Acad. Press, N.Y.

BOOTHBY, R. N., and 1. W. AVAL'LT, Jr.

1971. Food habits, length-weight relationship, and condition factor of the red

drum (Sciaenops ocella/a) in southeastern Louisiana. Trans. Am. Fish.

Soc. 100:290-295.

BOSC, LA. G.

[1802.J Histoir< naturelle des Crustaces, contenant leur description et leurs

moeurs; avec figures des>inee, J'apres nature. Paris, J: 1-258.

BURTON, D. T., and S. L. MARGREY.

1978. An evaluation of acute bromine chloride toxicity to grass shrimp

(Palaemone/es spp.) and juvenile blue crabs (Callinecles sapidus). Bull.

Environ. Contam. Toxicol. 19: 131-138.

CAMERON, J. N.

1979. Effects of inhibitors on ion fluxes, trans-gill potential and pH regulation

in freshwater blue crabs. CallinecleS sapid"s (Rathbun). 1. Compo Physiol.

133:219-225.

CANTELMO. A. C.. R. 1. LAZELL, and L H. MANTEL

1981. The effects of benzene on molting and limb regeneration in juvenile

Callinec/es sapid", Rathbun. Mar. BioI. Lell. 2:333-343.

CANTELMO, A. C, "nd K. RANGA RAO.

1978. The eflects of pentachlorophenol (PCP) and 2,4-dinitrophenol (DNP)

on the oxygen consumption of tISsues from the blue crab, Callinec/es

supidus, under different osmotic conditions. Camp. Biochem. Physio!.

"llC.215-219.

CARGO,D.G

1954. Maryland commercial fishing gears. III. The crab gears. Chesapeake

BioI. Lab., Educ. Ser 36, 18 p.

195~. The migration of adult temale blue crabs, Callinectes sapidus Rathbun,

in Chincoteague Bay and adjacent waters. J. Mar. Res. 16: 180-191.

19~U. A bisexual blue crab, Callmeete, sapidu, Rathbun, from the Chesapeake Bay. Am. MidI. Nat. 104:37H-3~2.

CARPENTER, J. H.. and D. G. CARGO

1957. The oxygen requirement and mortality of the blue crab in the Chesapeake Bay. Chesapeake Bay Ins!.. Johns Hopkins Univ., Tech. Rep. XIII,

22 p.

CHACE, E A.. Jr.. and H. H. HOBBS. Jr.

1969. The freshwater and terrestrial decapod crustaceans of the West Indies

"ith special reterence to Dominica. U.S. Natl. Mus. Bull. 292, 258 p.

CHITTENDEN, M. E.. Jr., and W. A. VAN ENGEL

1972. Effect of a tickler chain and tow duration on trawl catches of the blue

crab, Callinee/es sapidus. Trans. Am. Fish. Soc. 101:732-734.

CHL:RCHILL, E. P., Jr.

1921. Life history of the blue crab. Bull. U.S. Bur. Fish. 36:95-128.

CLAUS, C., E BENIJTS, G. VANDEPUTTE, and W. GARDNER.

1979. The biochemical composition of the larvae of two strains of Anemia

salina (L.) reared on two different algal foods. J. Exp. Mar. BioI. Ecol.

36: 171-183.

COCHRAN,D.M.

1935. The skeletal musculature of the blue crab, Callinec/es sapidus Rathbun.

Smithson. Misc. Collect. 92(9), 76 p.

COLE, M. A., and R. P. MORGAN II.

1978a. Genetic variation in two populations of blue crab, CallineCles sapidus.

Estuaries 1:202-105.
1978b. Muscle, eye and serum proteins in the blue crab, Callinecles sapidus

Rathbun. Camp. Biochem. Physiol. 59B:25-26.

COLVOCORESSES, J. A., and M. P. LYNCH.

1975. Variations in serum constituents of the blue crab, Callinecles sapidlls:

Copper and zinc. Camp. Biochem. Physiol. 50A: 135- 139.

COLVOCORESSES, J. A., M. P. LYNCH, and K. L. WEBB.

1974. Variations in serum constituents of the blue crab, Calline,",es sapidlls:

Major cations. Camp. Biochem. Physiol. 49A:7g7-803.

CONTRERAS, F.

1930. Contribucion al conocimiento de las jaibas de Mexico. An. Inst. BioI.

Univ. Nac. Auton. Mex. 1:227-241.

COOK, D. W, and S. R LOFfON.

1973. Chitinoclastic bacteria associated with shell disease in Pen,leus shrimp

and the blue crab (Callinecles sapidlls). .I. Wildl. Dis. 9: 154-159.

COSTLOW, J. D., Jr.

1963a. Regeneration and metamorphosis in larvae d the blue crub, Callineues

sapidlls Rathbun. J. Exp. Zool. 151:21'1-227.

1963b. The effect of eyestalk extirpation on metamorph<lsis 01 megalops at

the blue crab, Callillecles sapidus Rathbun. Gen. Com". E.ndorrino!.

3: 120-130.

1965. Variability in larval stages of the blue crab, Callinectes sapid'/I. Bioi.

Bull. (Woods Hole) 128:58-66.

1967. The effect of salinity and temperature on ,urvival and metamorphosis of

megalops of the blue crab Callinecles supidus. Helgo!. Wiss. Meeresunters.

15:84-97.

1979. Effect of Dimilin on development of larvae at the stone crab Mellippc

mercellaria, and the blue crab, CullineCles sapidlls. In W B. Vernberg,

A. Calabrese, F. P. Thurberg, and F. J. Vernberg (editors), Marine pollution:

Functional responses, p. 355-363. Acad. Press. N.Y.

COSTLOW, J. D., Jr., and C. G. BOOKHOUT.

1959. The larval development of Cal/inecle.\" saph!lIs R:tthbun reared in Ihe

laboratory. BioI. Bull. (Woods Hole) 1/6:373-396.

1960. A method for developing brachyuran eggs in ~llru. Limnol. Ocennogr

5:212-215.

1969. Temperature and meroplankton. Chesapeake Sci. 10:253-255.

1979. Second generation pesticides and crab development. In F. S . .!acuff

(editor), Advances in Marine Environmental Research, Proc of a Symp.,

US ERA/ORD/ERL, Narragansett, R.l., p. 320-336.

COSTLOW, J. D., Jr., G. H. REES. and C. G. BOOKHOUT.

1959. Preliminary note on the complete larval development of Callillecles

sapidlls Rathbun under luboratory conditions. Limnol. Oceanogr. 4:

222-223.

COSTLOW, J. D., Jr., and A. N. SASTRY.

1966. Free amino acids in developing stages of two crabs, Callinecles sapidlls

Rathbun and Rhithropallopeus harrisii (Gould). Acta Embryo!. Morpho!.

Exp. 9:44·55

COUCH, J. A.

1966. Two peritrichous ciliates from the gills of the blue crab. Chesapeake

Sci. 7:171-173

1967. A new species of Lagenophrys (Ciliatca: Peritnchida: LagenophryidJe)

from a marine crab. Callinectes sapidus. Trans. Am. Microsc. Soc. 86:

204-211.

1974. Pathological effects of Uru.lporidillm (Haplosporida) infection in

microphallid metacercariae. J. Invertebr. Pathol. 23:389-396.

COUCH, J. A., and S MARTIN.

1982. Protozoan symbionts and related diseases of the blue crab, Callinecles

sapidlls Rathbun from the Atlantic ami Gulf coasts of the United States.

III H. M. Perry and W A. Van Engel (editors), Proceedings Blue Crab

Colloquium, October 18-19,1979, p. 71·80.

COUCH, J N.

1942. A new fungus on crab eggs. J. Elisha Mitchell Sci. Soc. 58: 158-162.

CRONIN, L. E.

1942. A histological study of the develo~ment of the ovary and accessory

organs of the blue crab, Cul/ill'ctes sapid".s Rathbun. M.S. Thesis. Uni •.

Maryland, College Park, 2b p.

1947. Anatomy and histology of the male reproductive system of Callinectes

sapidl/s Rathbun. J. Morphol. 81 :209·219.

1949. Comparisor. of methods of tagging the blue crab. Ecology 30:

3<){)-394.

1954. Blue crab studie,. Biennial report 1"53 and 1954, Mar. Lab .. Univ.

Ddaw,lfe, Newark and L.ewes, Publ. 2, p. 65-70

CRONIN, L. E , W A VAN ENGEL, D. G CARGO, and F. J. WOJCIK.

1957 A partial bIbliography of tile genus Callillecte,·. V1. Fish. Lab., Spec.

Sci. Rep. 8; and Md. Del'. P.es. Educ .. Ret. 57··26, 21 p.

DANIELS, B A., and R. T. SAWYER.

1975. The biology of the leech MJ'<obriellG II/gubns infesting blue crabs aad

33

catfish. BioI. Bull. (Woods Hole) 148: 193·198.

DARNELL, R. M.

1959. Studies of tl'e Iife history of the blue crab (Cullillecles sapidl/s Rathbun)

in Louisiana waters. Trans. Am. Fish. Soc. 88:294-304.

1961. Trophic spectrum of an estuarine community, based on studies of Lake

Pontchartrain. Ecology 42:553-568.

DAVIS, C. C.

1965. A study of the hatching process in aquatic invertebrates: XX. The blue

crab, Callinec/es sapidus, Rathbun, XXI. The nemertean, Carcinonemerft!f

curcillophila (Kolliker). Chesapeake Sci. 6:201·208.

DeKAY. J. E

1841. Zoology of New-York, or the New·York fauna; comprising detailed

descriptions of all the animals hitherto observed within the state at New-York,

with brief notices of those occasionally found near its borders, and accum··

panied by appropriate illustrations. 6, Crustacea, p. 1-70, plates 1-13.

Carroll and Cook, Albany, N.Y.

DENDINGER, J. E.

1980. Electrophoretic separation of three dehy(irogenases from various lissues

of the Atlantic blue crab, Callillectes sapidl/". Com~ Bi"chem. Physiol.

66B:431·433.

DESMA.REST, A. G.

Ig23. Malacostraces. In Dictionnaire des sciences naturelles... etc. 28·

118-425. Paris.

DINTAMAN, R. C.

1982. Blue crab management project annual report, January 1992. Maryland

Department of Natural Resources Tidewater Administration, Annapol is.

DITTEl R, A. I., and C. E. EPIFANIO.

1982. Seasonal abundance and vertical distribution of crab larvae in Delaware

Bay. Estuaries 5: 197-202.

DUDLEY, D. L., and M. H. JUDY.

197!. Occaerence of larval, juvenile, and mature crabs in the vicinity of the

Beaufort Inlet, North Carolina. U.S. Del'. Cammer., NOAA Tech. Rep.

NMFS SSRF-637, 10 p.

197.3. Seas0nal abundance and distribution of juvenile blue crabs in Core

Sound, N.C. 1965·68. Chesapeake Sci. 14:51·55.

DUNNINGTON, E. A.

1956. Blue crabs observed to dig soft shell clams for food. Md. Tidewater

News 12(12):1,4.

ELDRIDGE, P. J., V. G. BURRELL, Jr., and G. STEELE

1979. Development of a self-culling blue crab pol. Mar. FISh. Rev. 41(12):

21-17.

ELDRIDGE, P. J., and W WALTZ.

1977. Observations on the commercial fishery for blue crabs, Culllnecte.1

sapidl/s, in estuaries in the southern half of South Carolina. S.C. Mar.

Resour. Cent., Tech. Rep. 21, 35 p.

ENGEL, D. W

1977. Comparison of the osmoregulatory capabilities of two portunid crabs.

Callinecles sapidus and C. similis. Mar. BioI. (Berl.) 4 I:275-219.

ENGEL, D. W, and L. D. EGGERT.

1974. The effect of salinity and sex on the respiration rates of excISed gill'

of the blue crab, Callillecles sapidlls. Camp. Biochem. Physiol. 47A:

1005·101 I.

ETTl GER, W S, and R. W. BLYE, Jr.

1981. Occurrence of the blue crab Callinectes sapidus in the tidal freshwater

reaches of the Delaware and Schuylkill Rivers in 1976. J. Crustacean Bio!.

1: 177·182.

FANNALY, M. T.

1978. A method for tagging immature blue crabs (Callinecles sapidus

Rathbun). Northeast Gulf Sci. 2: 124-126.

FAUSTO-FILHO, J.

1980. Callinecles a/finis a new species of crab from Brazil (Decapoda,

Portunidae). Crustaceana 39:33-38.

FINDLEY, A. M., B. W BELISLE, and W B STICKLE.

1978. Effects of salinity nuctuations on the respirution rate of the southefll

oyster drill Thais haemastoma and the blue crab Callinecles .,-apidlls. Ma'

BioI. (Berl.) 49:59-67.

FINDLEY, A. M., E. W BLAKENEY, Jr., and E. H WEIDNER

1981. Ameson michaelis (Microsp0rida) in the blue crab, Cl1l1inectes sapldu.. :

Parasite-induced alterations in the biochemical composition of hosl tissue,.

BioI. Bull. (Woods Hole) 161: 115-12'1

FISCHLER, K. J.

1965. The use of catch-effort, catch··sampling, and tagging data to estimate a

population of blue crabs. Trans. Am FISh. Soc. 94:287·310.

FISCHLER, K. J., and C. H. WALBURG.

1962. Blue crab movement in coas!al South Carolina, 1959·54. Truns Am.

Fish. Soc. 91:275-278.
FRANK, P. M., and P. B. ROBERTSON.

1979. The influence of salinity on toxicity of cadmium and chrolTium to the

blue crab, Cal/inectes sapid,,,. Bull. Environ. Contam. Toxicol. 21:74-78.

GERARD, J. F., and R. GILLES.

1972. The free amino-acid pool in Callinectes sapidus (Rathbun) tissues and

its rok in the osmotic intracdlular regui,tion. J. Exp. Mar. BioI. Ecol.

10: 125-136.

GERSTAECKER, A.

1856. Carcinologische Beitrage. Arch. NalUrgesch. 22(1):101-162.

GILLES, R.

1973. Oxygen consumption as related iO the amino-acid metabolism Juring

osmoregulation in the hlue crab Catiinecte.1 sapid"s. Neth J. Sea Res.

7:280-289.

GILLES, R., and J F. GERARD.

1974. AmJnv-acid metabolism dUllng \lsmotic !oltres:; in i~lllated axon~ or

Callinectc\ sap,d"s. Life Sci. 14:1221-1229

GLEESON, R. A.

1980. Pheromone communication in the reprodu~tive behavinr l,f the blue

crab, Cal/inrctes ,apidus. Mar. Behav. Physiol 7: 119-134

1982. MorphologIcal and behavioral identification of the se"sory structures

mediallng pheromone reception in the blue crab, Callillec/I'< sapiJrs. BioI.

Bull. (Woods Hole) 163:162-171.

GLEESON, R. A., and P. L. ZUBKOFf.

1977 The determination of hemolylnph 'ulume in tLe bille crah, Callinec[n sapidtH, utilizing 14C-thiol:yani.lte. Compo Hiochem. Physiol. 56A:

411-413.

GOODYEAR, C. P.

1967. Feeding habits of three species of gars, Lepisosteus, along the Mississippi Gulf Coasl. Trans. Am. Fish. Soc. 96:297-300.

GOULD, A. A.

1841. Report on the ",vertebrata of Massachusetts, comprising the Mollusca,

Crustacea. Annelida, and Radiata. Folsom, Wells, and Thurston, Cambridge, Ma;>., 373 p.

GRAHAM, J. G.. "nd G. F. BEAVEN.

1942. F.xperimental sponge-crab planting' "nd crab larvae distributioll in the

region at Crisfield. Md. Chesapeake BioI. Lab. Cuntrib. 52, 18 p.

GRAY, E. H., and C. L. 'EWCOMBE.

1938a. The relative growth of part> in the blue cmb Cal/mectes sapid",

Rathbun. Growth 2:235-246.

1938b. Studies of moulting in Cal/int'cIPl sapid".r Ratheun. Growth 2:

285-296.

GUNTER,G.

1938. The common blue crab in fresh waters. Science (Wash., D.C.)

87:87-88.

HAEFNER, P. A.. Jr.

1964. Hemolymph calcium fluctuations as related to environmental salinity

during ecdysis of the blue crab, Callinectes sapidus Rathbun. Physiol. Zool.

37:247-258.

HAEFNER, P. A, Jr., and D. GARTEN.

1974. Methods of handling and shedding blue crab, Callinectes rapid,'s. Va.

Ins!. Mar. Sci. Mar. Resour. Advis. 8, and Spec. Rep. Applied Mar. Sci.

Ocean Eng. 52, 14 p.

HAEFNER, P. A., Jr., and C. N. SHUSTER, Jr.

1964. Length increments during terminal molt of the female blue crab,

Cal/inec/es sapidus, in different salinity environments. Chesapeake SCI.

5:114-118.

HALL, W. R., Jr.

1976. Delaware's blue crab. Univ. Delaware Sea Grant Program, Celaware

Marine Advisory Services Publ. II.

HAMILTON, P. V.

1976. Predation on Littorina irrorata (Mollusca: Gastropoda) by Callinec/es

sapidus (Crustacea: Portunidae). Bull. Mar. Sci. 26:403-409.

HAMILTON, P. v., R. T. NISHIMOTO, and J G. HALUSKY.

1976. Cheliped laterality in Cal/inectes sapidus (Crustacea: Portunidae).

BioI. Bull. (Woods Hole) .150:393-401.

HARD, W. L.

1942. Ovarian growth and ovulation in mature blue crab, Cal/inectes sapidus

Rathbun. Chesapeake BioI. Lab. Contrib. 46, 17 p.

HAWKINS, W. E., H. D. HOWSE, and V. J. FERRANS.

1977. Fine structure of the myocardium of the blue crab Callinectes sapidus

Rathbun. Chesapeake Sci. 18:197-207.

HAY, W. P.

1905. The life history of the blue crab (Cal/inectes sapidl/s). Rep. U.S. Bur.

Fish., 1904, p. 395-413

HECK, K. L., Jr., and R. J. ORTH.

1980. Structural components of eelgrass (Zostera marina) meadows in the

lower Chesapeake Bay-Decapod Crustacea. Estuaries 3:289-295.

34

HOLLAND, J. S., n. v. ALDRICH, and K. STRAWN.

1971 Effects of temperature and salinity on growth, food conversion, survival and temperature resistance of juvenile blue crabs, Callinectes sapidus

Rathbun. Texas A&M Univ., College Station, Tex., Sea Grant Publ. '0.

TAMU-SE-71-222, 166 p.

HOLMES, F. S.

1858. Post-Pliocene fossils of South-Carolina. No. I & 2, p. 1-28, plate,

1-4. Russell and Jones, Charleston, S.C.

HOLTHUIS, L. B.

1962. Report on a collection of Crustacea Decapoda and Stomatopoda from

Turkey and the Balkans. ZooI. Verh., Rijksmuseum van Natuurlijke

Historie, Leiden, 47,67 p.

HOPKI S, S. H.

1943. The external morphology of the first and second zoeal stages of the blue

crab, Callinectes sapidus Rathbun. Trans. Am. Microsc. Soc. 62:85-90.

1944. The external morphology of the third and fourth zoeal stages of the blue

crab, Callinectes sapid/,s Rathbun. BioI. Bull. (Woods Hole) 87: 145-152.

1947. The nemertean Carcinonemertes as an indicator of the spawning history

of the host, Callinectes sapidus. 1. Parasitol. 33: 146-150.

HUTTON, R. J., and F. SOGANDARES-BERNAL.

1959. Notes on the distribution of the leech, Myzobdella II/gubris Leidy,

and its association with mortality of the blue crab, Callinectes sapiJus

Rathbun. J. Parasito!' 45:384, 404, 430.

INTERNATIONAL COMMISSION ON ZOOLOGICAL NOMENCLATURE.

1964. Opinion 712. Forty-seven genera of decapod Crustacea: placed on the

official lis!. Bull. Zool. Nomenc!. 21:336-351.

JAHROMI, S, S.

1977. Occurrence of rhabdovirus-like particles in the blue crab, Cal/ineCles

sapidus. J. Gen. Virol. 36:485-493.

JAWORSKI, E.

1972. The blue crab fishery, Barataria Estuary, Louisiana. La. State Univ..

Sea Grant Publ. LSU-SG-72-0I, 112 p.

1982. History and status of Louisiana's soft shell blue crab fishery. Itl H. M.

Perry and W. A. Van Engel (editors), Proceedings Blue Crab Colloquium,

October 18-19,1979, p. 153-157.

JEFFRIES, H. P.

1966. Internal condition of a diminishing blue crab population (Cal/ineCl~s

sapidus). Chesapeake Sci. 7: 164-170.

JOHNSON, P. T.

1976a. Gas-bubble disease in the blue crab, Cal/inectes sapidus. J. Invertebr. Patho!. 27:247-253.

1976b. A herpeslike virus from the blue crab, Callinectes sapidus. J. Invertebr. Pathol. 27:419-420.

1976c. Bacterial infection in lhe blue crab, Callinectes sapidl/s: Course of

infection and histopathology. J. Invertebr. Patho!. 28:25-36.

1977a. Paramoebiasis in the blue crab, Callinectes sapidus. J. Invertebr.

Pathol. 29:308-320.

1977b. A viral disease of the blue crab, Callinectes sapidl/s: Histopathology

and differential diagnosis. J. Invertebr. Pathol. 29:201-209.

1977c. Herpes-like virus of blue crabs. In C. J. Sindermann (editor), Disease diagnosis and control in North A:nerican marine aquaculture, p. 100-

102. Elsevier Sci, Publ. Co., N.Y.

1977d. Reovirus-like virus of blue crabs. In C. J. Sindermann (editor),

Disease diagnosis and control in North American marine aquaculture,

p.103-105. Elsevier Sci. Publ. Co., N.Y.

1977e. Bacterial disease of blue crabs. In C. 1. Sindermann (editor),

Disease diagnosis and control in North American marine aquaculture,

p.I06-108. Elsevier Sci. Publ. Co., N.Y.

1977f. Paramoebiasis of blue crabs. In C. J. Sindermann (editor), Disease

diagnosis and control in North American marine aquaculture, p. 122-124.

Elsevier Sci. Publ. Co., N.Y.

1978. Viral diseases of the blue crab, Cal/inectes sapidus. Mar. Fish. Rev.

40(10):13-15.

1980. Histology of the blue crab Cal!inectes sapidl/s. A model for the

Decapoda. Praeger Sci. Publ. Co., N.Y., 440 p.

JOHNSON, P. T., and 1. E. BODAMMER.

1975. A disease of the blue crab, Cal/inectes sapidl/s, of possible viral

etiology. J. Invertebr. Pathol. 26:141-143.

JOHNSON, P. T., and C. A. FARLEY.

1980. A new enveloped helical virus from the blue crab, Callinectes sapidus.

J. Invertebr. Pathol. 35:90-92.

JOHNSON, P. T., and S. V. OTTO.

1981. Histology of a bilateral gynandromorph of the blue crab, Callinectes

sapidus Rathbun (Decapoda: Portunidae). BioI. Bull. (Woods Hole) 161:

236-245.

JUDY, M. H., and D. L. DUDLEY.

1970. Movements of tagged blue crabs in North Carolina waters. Commer.
Fish. Rev. 32(11):29-35.

KING, E. N.

1965. The oxygen consumption of intact crabs and excised gills as a function

of decreased salinity. Compo Biochem. Physiol. 15:93-102.

KOENIG, C. C., R. J. LIVINGSTON, and C. R. CRIPE.

1976. Blue crab mortality: Interaction of temperature and DDT residues.

Arch. Environ. Contam. Toxicol. 4:119-128.

KORMANIK, G. A., and J. N. CAMERON.

1981. Ammonia excretion in the seawater blue crab (Callineetes sapidus)

occurs by diffusion, and not Na+/NH, + exchange. J. Compo Physiol.

141 :457-562.

KRANTZ, G. E., and J. V. CHAMBERLIN.

1978. Blue crab predation on cultchless oyster spat. Proc. Natl. Shellfish.

Assoc. 68:38-41.

KRANTZ, G. E., R. R. COLWELL, and E. LOVELACE.

1969. Vibrio parahaemolytiells from the blue crab Cullineetes sapidlls in

Chesapeake Bay. Science (Wash., D.C.) 164: 1286-1287.

LAIRD, C. E., and P A. HAEFNER, Jr.

1976. Effects of intrinsic and environmental factors on oxygen consumption

in the blue crab, Callineetes supidlls Rathbun. J. Exp. Mar. BioI. Ecol.

22:171-178.

LAMBOU, V. W.

1961. Utilization of macrocrustaceans for food by freshwater fishes in louisiana and its effects on the determination of predator - prey relations. Prog.

Fish-Cult. 23: 18-25.

LATREILLE, P. A.

1825. In Genre de Crustad:s. Encyclopedie methodique. Histoire naturelle.

Entomologie, ou histoire naturelle des Crustaces, des Arachnides et des

Insects, Paris, 9, 10, 832 p.

LAUGHLIN, R. A.

1982. Feeding habits of the blue crab, Callineetes sapidus Rathbun, in the

Apalachicola estuary, Aorida. Bull. Mar. Sci. 32:807-822.

LAWLER, A. R., and W. A. VAN ENGEL.

1973. Triple regeneration of the fifth pereiopod of a blue crab, Callineetes

sapidlls Rathbun. Chesapeake Sci. 14: 144-145.

LEE, R. F., C. RYAN, and M. L. NEUHAUSER.

1976. Fate of petroleum hydrocarbons taken up from food and water by the

blue crab Callineetes sapidus. Mar. BioI. (Berl.) 37:363-370.

LEFFLER, C. W.

1972. Some effects of temperature on the growth and metabolic rate of juvenile blue crabs, Callineetes sapidus, in the laboratory. Mar. BioI. (Berl.)

14: 104-110.

1975a. Ionic and osmotic regulation and metabolic response to salinity ofjuvenile Callineetes sapidus Rathbun. Compo Biochem. Physiol. 52A:545-549.

1975b. Effects of ingested mirex and DDT on juvenile Callineetes sapidus

Rathbun. Environ. Pollut. 8:283-300.

LEWIS, E. G .• and P. A. HAEFNER, Jr.

1976. Oxygen consumption of the blue crab, Callineetes sapidlls Rathbun,

from proecdysis to postecdysis. Compo Biochem. Physiol. 54A:55-60.

LlNNAEUS, C.

1767. Systema naturae per regna tria naturae, secundum classes, ordines,

genera, species, cum characteribus, differentiis, synonymis, locis, 12th

ed. 1:1-1327.

LlPPSON, R. L.

1973. Blue crab. In A. J. Lippson (editor), The Chesapeake Bay in Maryland. An atlas of natural resources, p. 26-27. Johns Hopkins Univ. Press,

Baltimore.

LOCHHEAD, M. S., and C. L. EWCOMBE.

1942. Methods of hatching eggs oFthe blue crab. Va. J. Sci. 3:76-86.

LOWE, J.l.

1965. Chronic exposure of blue crabs, Callineetes sapidus, to sublethal concentrations of DDT. Ecology 46:899-900.

LYNCH, M. P, and K. L. WEBB.

1973a. Variations in serum constituents of the blue crab, Cullineetes sapidus:

Total serum protein. Compo Biochem. Physiol. 44A:1237-1249.

1973b. Variations in serum constituents of the blue crab, Callineetes sapidus:

Glucose. Compo Biochem. Physiol. 45A: 127-139.

MANGUM, C. P, and L. M. AMENDE.

1972. Blood osmotic concentration of blue crabs (Callineetes sapidus Rath·

bun) found in fresh water. Chesapeake Sci. 13:318-320.

MANGUM, C. P, S. U. SILVERTHORN, J. L. HARRIS, D. W. TOWLE, and

A.R.KRALL.

1976. The relationship between blood pH, ammonia excretion and adaptation to low salinity in the blue crab Callineetes sapidus. J. Exp. Zool.

195:129-136.

MANGUM, c., and D. TOWLE.

1977. Physiological adaptation to unstable environments. Am. Sci. 65:

35

67-75.

MANGUM, C. P., and A. L. WEILAND.

1975. The function of hemocyanin in respiration of the blue crab Callineetes

sapidus. J. Exp. Zool. 193:257-263.

MANNING, R. B., and L. B. HOLTHUIS.

1981. West African brachyuran crabs (Crustacea: Decapoda). Smithson.

Contrib. Zool. 306, 379 p.

MANOOCH, C. S., Ill.

1973. Food habits of yearling and adult striped bass, Marone saxarilis

(Walbaum), from Albemarle Sound, North Carolina. Chesapeake Sci.

14:73-86.

McKENNEY, C. L., Jr., and J. D. COSTLOW, Jr.

1981. The effects of salinity and mercury on developing megalopae and early

crab stages of the blue crab, Callineetes sapidus Rathbun. In F. J. Vernberg,

A. Calabrese, F. P. Thurberg, and W. B. Vern berg (editors), Biological

monitoring of marine pollutants, p. 241-262. Acad. Press, N.Y.

McKENZIE, M. D.

1970. Fluctuations in abundance of the blue crab and factors affecting

monalities. S.c. Mar. Resour. Div. Tech. Rep. 1,45 p.

MEDVED, R. J., and J. A. MARSHALL.

1981. Feeding behavior and biology of young sandbar sharks, Careharhinus

plumbeus (Pisces, Carcharhinidae), in Chincoteague Bay, Virginia. Fish.

Bull., U.S. 79:441-447.

MENZEL, R. w., anJ S. H. HOPKINS.

1956. Crabs as predators of oysters in Louisiana. Proc. Natl. Shellfish.

Assoc. 46: 177-184.

ME ZEL, R. w., and F. E. NICHY.

1958. Studies of the distribution and feeding habits of some oyster predators

in Alligator Harbor, Florida. Bull. Mar. Sci. Gulf Caribb. 8: 125-145.

MILLER, R. E., D. W. CAMPBELL, and P. J. LUNSFORD.

1980. Comparison of sampling devices for the juvenile blue crab, Callineetes

sapidus. Fish. Bull., U.S. 78:196-198.

MILLER, R. E., S. D. SULKIN, and R. L. LIPPSON.

1975. Composition and seasonal abundance of the blue crab, Callineetes

sapidus Rathbun, in the Chesapeake and Delaware Canal and adjacent

waters. Chesapeake Sci. 16:27-31.

MILLER, R. J.

1976. North American crab fisheries: Regulations and their rationales. Fish.

Bull., U.S. 74:623-633.

MILLIKIN, M. R.

1978. Blue crab larval culture: Methods and management. Mar. Fish. Rev.

40(11): 10-17.

MILLIKIN, M. R., G. N. BIDDLE, T. C. SIEWICKI, A. R. FORTNER, and

P. H. FAIR.

1980. Effects of various levels of dietary protein on survival, molting frequency and growth of juvenile blue crabs (Cullineetes sapidus). Aquaculture 19: I49-161.

MILNE EDWARDS, A.

1861. Etudes zoologiques sur les Crustaces recents de la famille des Portuniens.

Archives du Museum d'Histoire Naturelle, Paris, 10:309-428.

1873-1881. Etudes sur les Xiphosures etles Crustaces de la Region Mexicaine.

In Mission Scientifique au Mexique et dans I' Amerique centrale, part 5,

p. 1-368, plates 1-61. (Published in numbers cited as follows: 3, p. 57-120,

plates 16-20, 1875; 4, p. 121-184, plates 21-27, 29-30, 1878; p. 185-224,

plates 32-39, 1879; 7, p. 265-312, plates 49-54, 1880.)

MILNE EDWARDS, A., and E. BOUVIER.

1900. Crustaces Decapodes. Premiere partie. Brachyures et Anomures. In

Expeditions sCientifiques du Travailleur et du Talisman pendant les annees

1880,1881,1882,1883. Masson et Cie, ed., Paris, 396 p.

MOORE, D. J.

1971. The uptake ancl concentration of fluoride by the blue crab, Callineeres

sapidlls. Chesapeake Sci. 12:1-13.

MORE, W. R.

1969. A contrtbution to the biology of the blue crab (Callineetes sapidus

Rathblln) in Texas, with a description of the fishery. Tex. Parks Wildl.

Dep., Tech. Ser. 1,31 p.

NAYLOR, E., and M. J. ISAAC.

1973. Behavioural significance of pressure responses in megalopa larvae of

Cullineetes supidlls and Maeropipus sp. Mar. Behav. Physiol. 1:341-350.

NEWCOMBE, C. L., F. CAMPBELL, and A. M. ECKSTINE.

1949. A study of the form and growth of the blue crab Callineetes sapidlls

Rathbun. Growth 13:71-96.

NEWCOMBE, C. L., M. D. SANDOZ, and R. ROGERS-TALBERT

1949. Differential growth and moulting characteristics of the blue crab,

Callinectes sapidus Rathbun J. Exp. Zool. 110: 113-152.

NEWMAN, M. w., and C. A. JOHNSO

1975. A Jisease of blue crabs (Collineetes sapidus) caused by a parasitic
dinoflagellate, Hematodinium sp. J. Parasitol. 61:554-557.

NEWMAN, M. w., C. A. JOHNSON III, and G. B. PAULEY.

1976. A Minchinia-like haplosporidan parasitizing blue crabs, Callinectes

sapidus. J. Invertebr. Pathol. 27:311-315.

NEWMAN, M. w., and G. E. WARD, Jr.

1973. An epizootic of blue crabs, Callinectes sophlus, caused by Paramoeba

perniciosa. J. Invertebr. Pathol. 22:329-334.

NICHOLS, P. R., and P. M. KENEY.

1963. Crab larvae (Callinectes), in plankton collections from cruises of M/V

Theodore N. Gill, South Atlantic coast of the United States, 1953-54. U.S.

Fish Wildl. Serv., Spec. Sci. Rep. Fish. 448, 14 p.

NISHIMOTO, R. T., and W. F. HERRNKIND.

1978. Directional orientation in blue crabs, Callinectes sapidu; Rathbun:

Escape responses and influence of wave direction. J. Exp. Mar. BioI. Ecol.

33:93-112.

NORSE, E. A.

1977. Aspects of the zoogeographic distribution of Callinectes (Brachyura:

Portunidae). Bull. Mar. Sci. 27:440-447.

1978. An experimental gradient analysis: hyposalinity as an "upstress"

distributional determinant for Caribbean portunid crabs. BioI. Bull. (Woods

Hole) 155:586-598.

NORSE, E. A., and V. FOX-NORSE.

1982. Geographical ecology and evolutionary relationships in Callinectes spp.

(Brachyura: Portunidae). In H. M. Perry and W. A. Van Engel ~editors),

Proceedings Blue Crab Colloquium, October 18-19, 1979, p. 1-9.

ODUM, H. T

1953. Factors controlling marine invasion into Florida fresh waters. Bull.

Mar. Sci. Gulf Caribb. 3: 134-156.

ODUM, W. E., and E. J. HEALD

1972. Trophic analyses of an estuarine mangrove community. Bull. Mar.

Sci. 22:671-738.

OESTERLING, M. J.

1976. Reproduction, growth, and migration of blue crabs along Florida's Gulf

coast. Univ. Fla., Florida Sea Grant Publ. SUSF-SG-76-003, 19 p.

OGLE, 1. T, H. M. PERRY, and L. NICHOLSON.

1982. Closed recirculating seawater systems for holding intermolt blue crabs:

Literature review, systems design and construction. Gulf Coast Res. Lab.,

Tech. Rep. Ser. 3: I-II.

OLMI, E. 1., Ill, and J. M. BISHOP.

1983. Variations in total width-weight relationships of blue crabs, Callinectes

sapidus, in relation to sex, maturity, molt stage and carapace ferm. J.

Crustacean BioI. 3:575-581.

In press. An adult female blue crab, Callinectes sapidus, in proecdysis.

Crustaceana.

ORDWAY, A.

1863. Monograph of the genus Callinectes. J. Boston Soc. Nat. Hist.

7(13):567-583.

ORTH, R. J.

1977. Effect of nutrient enrichment on growth of the eelgrass Zostera marina

in the Chesapeake Bay, Virginia, USA. Mar. BioI. (Berl.) 44: 187-194.

ORTMANN, A.

1894. Die Decapoden-Krebse des Strassburger Museums. VI Theil. Abtheilung: Brachyura (Brachyura genuina Boas) I. Unterabthei lung: Majoidea

und Cancroidea, I. Section Portuninea. Zool. Jahrb. Abt. Syst., Geogr.

BioI. Tiere 7:23-88.

OTWELL, W. S., and 1. C. CATO.

1982. Review of the soft-shell crab fishery in the United States. In H. M. Perry

and W. A. Van Engel (editors), Proceedings Blue Crab Colloquium, October

18-19,1979, p. 129-136.

OVERSTREET, R. M.

1975. Buquinolate as a preventive drug to control microsporidosis in the blue

crab. J. Invertebr. J'athol. 26:213-216.

1977. Microsporidosis of the blue crab. In C. J. Sindermann (editor), Disease

diagnosis and control in North American marine aquaculture, p. 117-121.

Elsevier Sci. Publ. Co., N.Y.

1978. Marine maladies? Worms, germs, and other symbionts from the

Northern Gulf of Mexico. MissiSSippi-Alabama Sea Grant Consortium,

MASGP 78-02 J, 140 p.

1982. Metazoan symbionts of the blue crab. In H. M. Perry and W. A. Van

Engel (editors), Proceedings Blue Crab Colloquium, October 18-19, 1979,

p.81-87.

OVERSTREET, R. M., and R. W. HEARD.

1978a. Food of the red drum, S<"iaenops ocellata, from Mississippi Sound.

Gulf Res. Rep. 6:131-135.

1978b. Food of the Atlantic croaker, Micropogonias undulatus, from Mississippi Sound and the Gulf of Mexico. Gulf Res. Rep. 6: 145-152.

36

OVERSTREET, R. M., and H. M. PERRY.

1972. A new microphallid trematode from the blue crab in the northern Gulf

of Mexico. Trans. Am. Microsc. Soc. 91 :436-440.

OVERSTREET, R. M., and E. C. WHATLEY, Jr.

1975. Prevention of microsporidosis in the blue crab with notes on natural

infections. Proc. World Mariculture Soc. 6:335-346.

PALMER, R. A.

1974. Studies on the blue crab (Callinectes sapidus) in Georgia. Ga. Dep.

Nat. Resour., Contrib. Ser. 29, 59 p.

PARKER, N. C., R. HOLT, and K. STRAWN.

1976. The potential for soft shell crab production util izing heated effluents

from power plants in the Galveston Bay system. Proc. Annu. ConL Southeast. Assoc. Game Fish Comm. 29:215-222.

PARTRIDGE, B. L.

1982. The structure and function of fish schools. Sci. Am. 246(6):114-123.

PASSANO, L. M.

1960. Molting and its control. In T H. Waterman (editor), The physiology of

crustacea, Vol. 1:473-536. Acad. Press, N.Y.

PEARSON, J. C.

1948. Fluctuations in the abundance of the blue crab in Chesapeake Bay. U.S.

Fish Wildl. Serv., Res. Rep. 14,26 p.

1951. The blue crab in North Carolina. In H. F. Taylor (editor), Survey of

marine fisheries of North Carolina, p. 205-218. Univ. North Carolina Press,

Chapel Hill.

PEARSON, W. H., and B. L. OLLA.

1977. Chemoreception in the blue crab, Callinectes sapidus. BioI. Bull.

(Woods Hole) 153:346-354.

1979. Detection of naphthalene by the blue crab, Callinectes sapidus. Estuaries 2:64-65.

1980. Threshold for detection of naphthalene and other behavioral responses

by the blue crab, Callinectes sapidus. Estuaries 3:224-229.

PERRY, H. M.

1975. The blue crab fishery in MiSSissippi. Gulf Res. Rep. 5:39-57.

PERRY, H. M., J. T. OGLE, and L. C. NICHOLSON.

1982. The fishery for soft crabs with emphasis on the development of a closed

recirculating system for shedding crabs. In H. M. Perry and W. A. Van

Engel (editors), Proceedings Blue Crab Colloquium, October 18-19, 1979,

p. 137-152.

PERRY, H. M, and K. C. STUCK.

1982. The life hi,tory of the blue crab in Mississippi with notes on larval

distribution. In H. M. Perry and W. A. Van Engel (editors), Proceedings Blue

Crab Colloquium, October 18-19,1979, p. 17-22.

PERSCHBACHER, P. w., and F. J. SCHWARTZ.

1979. Recent records of Callinectes danae and Callinectes marginatus

(DecapoJa: Portunidae) from North Carolina with environmental notes.

Fish. Bull., U.S. 76:879-880.

PILEGGI, 1., and 8. G. THOMPSON.

1976. Fishery statistics of the United States, 1973. U.S. Dep. Commer.,

NOAA, Natl. Mar. Fish. Serv., Stat. Dig. 67,458 p.

1978. Fishery statistics of the United States, 1975. U.S. Dep. Commer.,

NOAA, Natl. Mar. Fish. Serv., Stat. Dig. 69,418 p.

1980. Fishery statistics of the United States, 1976. U.S. Dep. Commer. ,

NOAA, Natl. Mar. Fish. Serv., Stat. Dig. 70, 419 p.

PORTER, H. J.

1955. Variation in morphometry of the adult female blue crab, Callinectes

sapidus Rathbun. M.S. Thesis, Univ. Delaware, Newark, 69 p.

PULLEN, E. J., and W. L. TRENT

1970. Carapace width-total weight relation of blue crabs from Galveston Bay,

Texas. Trans. Am. Fish. Soc. 99:795-798.

PYLE, R., and E. CRONIN.

1950. The general anatomy of the blue crab Callinectes sapidus Rathbun. Chesapeake BioI. Lab. Publ. 87, 40 p.

RAO, K. R., F R. FOX, P. 1. CONKLIN, A. C. CANTELMO, and A. C.

BRANNON.

1979. Physiological and biochemical investigations of the toxicity of the

pentachlorophenol to crustaceans. In W. B. Vernberg, A. Calabrese,

F. P. Thurberg, and F. 1. Vernberg (editors), Marine pollution: Functional

responses, p. 307-339. Acad. Press, N.Y.

RATHBUN, M. J.

1896. The genus Callinectes. Proc. U.S. Natl. Mus. 18 (for 1895):349-375.

RATHBUN, R.

1884. Crustaceans. In G. B. Goode, The fisheries and fishery industries of

the United States, section I, part 5, p. 759-830. U.S. Comm. Fish Fish.,

Washington, D.C.

REIMER, R. D., and K. STRAWN.

1973. The use of heated effluents from power plants for the winter production

of soft shell crabs. Proc. World Maricu!ture Soc. 4:87-96.
REINHARD, E. G.

1950. An analysis of the effects of a sacculinid parasite on the external

morphology of Callinectes sapid"s Rathbun. BioI. Bull. (Woods Hole)

98:277-288.

ROBERTS, K. J., and M. E. THOMPSON.

1982. Economic elements of commercial crabbing in Lake Pontchanrain and

Lake Borgne. Louisiana Sea Grant Publ. LSU-TL-82-001, 19 p.

ROBERTS, M. H., Jr.

1981. Kerone distribution in selected tissues of blue crabs, Callinectes

sapid"s, collected from the James River and lower Chesapeake Bay. Estuaries 4:313-320.

ROBERTS, M. H., Jr., and A. T. LEGGETT, Jr.

1980. Egg extrusion as a Kepone-clearance route in the blue crab, Callineetes

sapid"s. Estuaries 3:192-199.

ROCHEBRUNE, A. T. de.

1883. Diagnoses d'Arthropodes nOuveaux propres a la Senegambie. Bull.

Soc. Philom. Paris. Ser. 7,7:167-182.

ROGERS-TALBERT, R.

1948. The fungus Lagenidi"m callineetes Couch (1942) on eggs of the blue

crab in Chesapeake Bay. BioI. Bull. (Woods Hole) 95:214-228.

ROSEN,B

1967. Shell disease of the blue crab, Ca/lineetes sapid"s. J. Invertebr.

Pathol. 9:348-353.

ROSENBERG, R., and J. D. COSTLOW, Jr.

1976. Synergistic effects of cadmium and salinity combined with constant

and cycl ing temperatures on the larval development of two estuarine crab

species. Mar. BioI. (Berl.) 38:291-303.

ROUSE, A. M.

1972. Hepatopancreas glycogen concentrations and their relationship to

ecdysis in the blue crab, Callinectes sapid"s Rathbun. M.S. Thesis, Univ.

Delaware, Newark, 85 p.

SANDIFER, P. A.

1973. Distribution and abundance of decapod crustacean larvae in the

York River estuary and adjacent lower Chesapeake Bay, Virginia, 1968-

1969. Chesapeake Sci. 14:235-257

1975. The role of pelagic larvae in recruitment to populations of adult decapod

crustaceans in the York River estuary and adjacent lower Chesapeake Bay,

Virginia. Estuarine Coastal Mar. Sci. 3:269-279.

SANDIFER, P. A., and P. J. ELDRIDGE.

1974. Observations On the incidence of shell disease in South Carolina blue

crabs, Callinectes sapid"s. In R. L. Amborski, M. A. Hood. and R. R. Miller

(editors). Proceedings of Gulf Coast Regional Symposium on Diseases of

Aquatic Animals, p. 161-184. Publ. LSU-SG-74-05, Louisiana State

University.

SANDOZ, M., and R. ROGERS.

1944. The effect of environmental factors on hatching, moulting, and survival

of zoea larvae of the blue crab Cal/inettes sapid"s Rathbun. Ecology

25:216-228.

SAWYER, T. K., R. COX, and M. HIGGINBOTTOM.

1970. Hemocyte values in healthy blue crabs. Callinectes sapid"s, and crabs

infected with the amoeba, Paramoeba perniciosa. J. Invertebr. PathoJ.

15:440-446.

SAWYER. T. K., and S. A. MacLEA

1978. Some protozoan diseases of decapod crustaceans. Mar. Fish. Rev.

40(10):32-35.

SAY, T.

1817-1818. An account of the Crustacea of the United States. J. Acad. at.

Sci. Phila. 1(1).(1817):57-63,65-80,97-101,155-169; (2) (1818):23'i-253,

313-319, 374-401. 423-444, 445-458.

SCHIMMEL, S. C.• J. M. PATRICK, Jr.• L. F. FAAS. J L. OGLESBY, and

A. J. WILSON, Jr.

1979. Kepone: Toxicity and bioaccumulation in blue crabs. Estuaries

2:9-15

SCHIMMEL, S. c., and A. J. WILSON, Jr.

1977. Acute toxicity of Kepone to four estuarine animals. Chesapeake Sci.

18:224-227

SCHOOR. W. P.

1974. Accumulation of mirex-"C in the adult blue crab (Callinectes

sapid"s). Bull. Environ. Contam. Toxicol. 12:136-137.

SEED, R.

1980. Predator-prey relationships between the mud crab Panopeus herbstii.

the blue crab. Callinectes sapidus and the Atlantic ribbed mussel Ge"kensia

(= Madia/us) demissa. Estuarine Coastal Mar. Sci. 11:445-458.

SHERIDAN, P. F.

1975. Uptake, metabolism. and distribution of DDT in organs of the blue crab,

Cal/inectes sapid"s. Chesapeake Sci. 16:20-26.

37

SHOLAR, T. M.

1982. Blue crab fisheries of the Atlantic coast. In H. M. Perry and W. A.

Van Engel (editors). Proceedings Blue Crab Colloquium, October 18-19,

1979. p. 111-127.

SHUSTER. C. N., Jr.• D. H. BULMER. Jr., and W. A. VAN ENGEL.

1963. A commentary on claw deformities in the blue crab. Estuarine Bull.

7(2-3): 15-23.

SINDERMANN, C. J.

1977a. Shell disease of blue crabs. In C. J. Sindermann (editor), Disease

diagnosis and control in Nonh Amencan marine aquaculture. p. 109-

112. Elsevier Sci. Publ. Co., N.Y.

1977b. Fungus disease of blue crab eggs and larvae. In C. J. Sindermann

(editor), Disease diagnosis and control in North American marine aquaculture,

p. 113-116. Elsevier Sci. Publ. Co., .Y.

SINGER, S. c., and R. F. LEE.

1977. Mixed function oxygenase activity in blue crab, Ca/lineetes sapid"s:

Tissue distribution and correlation with changes during molting and development. BioI. Bull. (Woods Hole) 153:377-386.

SMITH, S. I.

1869. Notice of the Crustacea collected by Prof. C. F. Hartt on the coast of

Brazil in 1867. Trans. Conn. Acad. Arts Sci. 2:1-41.

SMYTH. P. O.

1980. Callinectes (Decapoda: Portunidae) larvae in the Middle Atlantic Bight,

1975-77 Fish. Bull., U.S. 78:251-265.

SPRAGUE. Y

1965. Nosemu sp. (Microsporida, Nosematidae) in the musculature of the crab

Callinectes sapid"s. J. Protozoal. 12:66-70.

1966. Two new species of P/istophora (Microsporida, Nosematidae) in

decapods, with particular reference to one in the blue crab. J. Protozool.

13: 196-199.

SPRAGUE, Y, and R. L. BECKETT.

1966. A disease of blue crabs (Callineetes sapidus) in Maryland and Virginia. J. Invertebr. Pathol. 8:2~7-289.

1968. The nature of the etiological agent of "gray crab" disease. 1. Invertebr. Pathol. 11:503.

SPRAGUE. Y, and J. COUCH.

1971. An annotated list of protozoan parasites. hyperparasites, and commensals of decapod crustacea. 1. Protozoal. 18:526-537.

STEPHENSON, W.

1962. Evolution and ecology of ponunid crabs, with especial reference

to Australian species. In G. W. Leeper (editor). The evolution of living

organisms, p. 311-327. Melbourne Univ. Press. Aust.

STEPHENSON. w.. and B. CAMPBELL.

1959. The Australian panun ids (Crustacea: Portunidae). III. The genus

Portunus. Aust. J. Mar. Freshw. Res. 10:84-124.

STEPHENSON, W., W. T. WILLIAMS. and G. N. LANCE.

1968. Numerical approaches to the relationships of certain American swimming crabs (Crustacea: Portunidae). Proc. U.S. atl. Mus. 124 (3645), 25 p.

STIMPSON, W.

1859. Notes On North American Crustacea, No. I. Ann. Lyceum Nat. Hist.

N.Y. 7(1862):49-93.

1860. Notes on North American Crustacea, in the Museum of the Smithsonian

Institution. No. II. Ann. Lyceum Nat. Hist. N.Y. 7: 176-246.

STUCK, K. C, and H. M. PERRY.

1981. Observations on the distribution and seasonality of portunid megalopae

in Mississippi coastal waters. Gulf Res. Rep. 7:93-95.

SULKIN, S. D.

1977. Factors influencing blue crab population size: Nutrition of larvae and

migration of juveniles. Chesapeake Biological Laboratory, Completion

Report. Volumes I and II. UMCEES Ref. No. 77-16 HPEL. 86 p.

1978. Nutritional requirements during larval development of the portunid crab,

Cal/ineetes sapid"s Rathbun. 1. Exp. Mar. BioI. Ecol. 34:29-41.

SULKIN. S. D.• E. S. BRANSCOMB, and R. E. MILLER.

1976 Induced wInter spawning and culture of larvae of the blue crab. Cal/ineL·tes sapid"s Rathbun. Aquaculture 8: /03-113.

SULKIN, S. D., and C. E. EPIFANIO.

1975. Comparison of rotifers and other diets for rearing early larvae of the blue

crab, Callineetes sapidlls Rathbun. Estuarine Coastal Mar. Sci. 3: 109-113.

SULKIN. S. D., and R. E. MILLER.

1975. Modihed commercial crab and oyster dredgcs as sampling devices for

the blue crab Callineetes sapidlls Rathbun. Chesapeake Sci. 16: 137- 139.

SULKIN, S. D., I. PHILLIPS, and W VA HEUKELEM.

1979. On the locomotory rhythm of brachyuran crab larvae and its significance

in vertical migration. Mar. Ecal. Prog Ser. 1:331-335.

SULKIN, S. D., and W. VAN HEUKELEM.

1982. Larval recruitment in the crab Cul/ineoes sapidlls Rathbun: An

amendment to the concept of larval retention in estuaries. In Y Kennedy
Cal/ineetes sapidus

J. Exp. Mar. BioI.

Atlantic Monthly Press Book, Little Brown and

(editor), Estuarine comparisons, p. 459-475. Acad. Press, NY

SULKI ,S. D., W VAN HEUKELEM, P. KELLY, and L. VAN HEUKELEM.

1980. The behavioral basis of larval recruitment in the crab Callineeres

sapidlls Rathbun: A laboratory investigation of ontogenetic changes in

geotaxis and barokinesis. BioI. Bull. (Woods Hole) 159:402-417.

SUMMERS, J. K., H. W HOFFMAN, and W A. RICHKUS.

1983. Randomized sample surveys to estimate annual blue crab harvests by a

multi-gear fishery in the Maryland waters of Chesapeake Bay. North Am.

J. Fish. Manage. 3:9-20.

SUMMERS, 1. K., W A. RICHKUS, H. v.. HOFFMAN, C. BONZEI', H. H

KING. and M. BURCH.

1983. Application of random sample sU:'vey deSIgns to estimate the ""mmercial blue crab harvest in Maryland. North Am. J. Fish. Manage. 3:21-25.

TAGATZ, M. E.

1965. The fishery for blue crabs in the St. Johns River, Florida, with special

reference to fluctuation in yield between 1961 and 1962. U.S. Fi,h Wildl.

Serv., Spec. Sci Rep. Fish. 50 I. lip.

1967. Noncommercial crabs of the genus Callineeks in St. Johns River,

Florida. Chesapeake Sci. 8:202-203.

1968a. Biology of the blue crab, Callineetes sapidlls Rathbun, in the St. Johns

River, Florida. U.S. Fish Wildl. Serv., Fish. Bull. 67:17-33.

I968b. Growth of juvenile blue crabs, Cal/ineetes sapidtls Rathbun, in the St.

Johns River, Florida. U.S. Fish Wildl. Serv., Fish. Bull. 67:281-288.

1969. Some relations of temperature acclimation and salinity to thermal

tolerance of the blue crab, Cal/ineetes sapidlls. Trans. Am. Fish. Soc.

98:713-716.

1971. Osmoregulatory ability of blue crabs in different temperature-salinity

combinations. Chesapeake Sci. 12: 14-17.

TAGATZ, M. E.. P. W BORTHWICK, and 1. FORESTER.

1975. Seasonal effects of leached mirex on selected estuarine animals. Arc·h.

Environ. Contam. Toxicol. 3:371-383.

TAGATZ, M. E., and A. B. HALL.

1971. Annotated bibliography on the fishing industry and biology 01 the blue

crab, Cal/ineeres sapidlls. U.S. Dep. Commer., NOAA Tech. Rep. NMFS

SSRF-640, 94 p.

TAISSOUN N., E.

1969. Las especies de cangrejos del genern "Callinectes" (brachyura) en

el Golfo de Venezuela y Lago de Maracaibo. Bol. Cent. Invest. BioI.,

2, 103 p.

TAN, E.-C., and W A. VAN ENGEL.

1966. Osmoregulation in the adult blue crab, ('allineetes sapia 115 Rathbun. Chesapeake Sci. 7:30-35.

TATUM, U. M.

1979. The blue crab fishery of Alabama. In H. A. Loyacano and J. P. Smith

(editors), Symposium on the natural resources of the Mobile Bay estuary,

Alabama, p. 211-220.

TEYTAUD, A. R.

1971. The laboratory studies of sex recognition in the blue 'Tab Callineetes

sapidlls Rathbun. Univ. Miami. Sea Grant Tech. Bull. 15.63 p.

TRUITT, R. V.

1939. The blue crab. In Our water resources and the'r consenation, p.

10-38. Chesapeake BioI. Lab. Contrib. 27.

TYLER, A. v., and D. G. CARGO

1963. Sile relations of two inst"rs of the blue crab. CallineC/<'.\ mpidus.

Chesapeake Sci. 4:52-54.

VAN ENGEL, W A.

1958. The blue crab and it, fishery in the Chesapeake Bay. Part I - Repwduction, early development, grow,h, and migration Commer. Fish. Rev

20(6):6-17.

1962. The blue crab and its fishery in Chesapeake Bay. Part 2 - Types 01 gear

for hard crab fishing. Commer. Fish. Rev. 24(9): 1-10

1982a. Biological aspects of blue crab shedding. 11/ D. Cupk.l and W. A. Van

Fngel (editot». Proc. Workshup on softshell blue crah,. September 22, 1979,

p.25-32. S.c. Mar. Resour. Cent., Tech. Rep. 48, and Va. Illst. M:H. Sci

('ontnb 1003.

1992b. Blue l:lab morta!ities a~~oCJated with pestil:ldes, herbi('idl'~. temlJ~ra+

ture. salinity, and dissolved ,oxygen. In H. M. Peflv and \Y. A. Van Engel

(editors), Proceedings Blue Crab Collo~uium, October 18-19, 1974. p. 89 92.

VAN ENGEL, W A., D. G. CARGO, ami F. J. WOJCIK.

1973. The edible blue crab: Abundant crustacean. Marine Resources of tht

Atlantic coast. Leafl. 15,8 p. Atlantic States Marine Fisheries Commission.

VIGH, D. A., and J. E. DE DINGER.

1982. Temporal relationships of postmolt deposition 01 calCium. O'agn('siulTJ,

chitin and protein in the cuticle of the AtlantiC blue ,rab, Cal/inpet" sapidlls

Rathbun Compo Biochem. Physiol. 72A:365-369.

VIRNSTEIN, R. W

1977. The importancc of predation by crabs and fishes on benthic infauna in

38

Chesapeake Bay. Ecology 58: I 199- 1217.

WALKER, G.

1974. The occurrence, distribution and attachment of the pedunculate barnacle

Oetolasmis mul/eri (Coker) on the gills of crabs, particularly the blue crab,

Callineetes sapidlls Rathbun. BioI. Bull. (Woods Hole) 147:678-689.

WALKER, W W, A. R. LAWLER, and W D. BURKE.

1979. Acute toxicity of 3-chloro-4-methyl benzenamine hydlOchloride to

shrimp and crabs. Bull. EilViron. Contam. Toxicol. 21:643-651.

WARNER,WW

1976. Beautiful swimmers.

Company, Boston, 304 p.

WEIDNER, E.

1970. Ultrastructural study of microsporidian development. I. Nosema sp.

Sprague, 1965 in Cal/ineetes sapidlls Rathbun. Z. Zellforsch. 105:33-54.

WEILAND, A. L., and C. P. MANGUM.

1975. The influence of environmental salinity on hemocyanin function in the

blue crab, Callineetes sapidus. J. Exp. Zool. 193:265-273.

WEIS, J. S.

1976. Effects of environmental factors on regeneration and molting in fiddler

crabs. BioI. Bull. (Woods Hole) 150: 152-162.

WENNER, C. A., and 1. A. MUSICK.

1975. Food habits and seasonal abundance of the American eel, Anguil/a

rostrata, from the lower Chesapeake Bay. Chesapeake Sci. 16:62-66.

WHARTON, J

1954. The Chesapeake Bay crab industry. U.S. Fish Wildl. Serv., Fish.

Leafl. 358, 17 p.

WHEELAND, H. A.

1975. Fishery statistics of the United States, 1972. U.S. Dep. Commer. ,

NOAA, Natl. Mar. Fish. Serv., Stat. Dig. 66, 517 p.

WHITE, A. Q., and C. P. SPIRITO.

1973. Anatomy and physiology of the swimming leg musculature in the blue

crab, Cal/ineetes sapidus. Mar. Behav. Physiol. 2: 141-153.

WHITNEY, J O'C.

1969. Sterols, fatty acids and sterol content in eggs and hepatopancreas of

the blue crab Cal/ineetes sapidlls (Rathbun). Acta Embryol. Exp. 1969:

111-121.

1970. Absence of sterol biosynthesis in the blue crab

Rathbun and in the barnacle Balanuf nubilus Darwin.

Ecol. 4:229-237.

WIEDOW, M. A., T. J. KNEIP, and S. J. GARTE.

1982. Cadmium-oinding prOleins from blue crabs (Ca/lineetes sapidu»

environmentally exposed to cadmium. Environ. Res. 28: 164-170.

WILLIAMS, A. B.

1965. Marine decapod crustaceans of the Carolinas. U.S. Fish Wildl. Serv.,

Fish. Bull. 65:1-298

1966. The Western Atlantic swimming crabs Callineetes ort/all/S, C. danae,

anrl a new, related species (Decapoda, Portunidae). Tulane Stud. Zool.

13.~3-93.

1971. A ten-year study uf meroplanktol1 in North Carolina estuaries: Annual

"ccurrence of sume brachyuran developmental stages. Chesapeake Sci.

12:53-61.

1974. Tile swimming crabs of the genus Cal/ineetes (Decapod a: Portunidae).

Fish. Bull .. U.S. 72.685-798.

1478. True crabs. In W Fischer (editor), FAO species identification sheets for

fishery purposes. Western central Atlantic (Fishing area 31), Vol. VI. Food

A~ric. Organ. U.N , Rome, Italy.

In pres<. Shrimps, lobsters, and crabs of the Atlantic coast of the eastern United

State,. Maine to Florida. Smithson. inst. Press, Wash., D.C.

WILLIAMS, A. B.. and T. W DUKE.

1979. Crabs (ArthropodJ: Crustacea: Decapoda: Brachyura). In C. W Hart,

Jr. an,1 S. L. H. Fuller Ieditors), Pollution e,'ology of estuarine invertebrates,

p. 171-233. Acad. Press. NY

WILLIAMS, A. BoO and H. J PORTER

1964. An u~us"ally large turtle barnacle (rhelonibia p. pall/la) on a blue crab

from Delaware Bay. Chesapeake Sci. 5: 150-151.

WILLIAMS. A. 8., and D. MeN WILLIAMS.

j 981 C:Jfolinian recordo;; for Americ:m lobster, Hnmarus amer;canus, and

tropical !-wimming ~rab, Cullinn;lrs h()('ourti. Po~tulated means of dbpersal Fish. Hull , US. 'i9: 192·198

WINGET, R. R., C. E. EPIFANIO, T. RUNNELS, and P. AUSTIN.

1976. Effects of diet and temperature on growth and mortality of blue crab,

Callineetes sapidus, maintained in a reCirculating culture system. Proc.

Natl. Shellfish. Assoc. 66:29-33.

WINGET, R. R., A. M. ROUSE, and D. MAURER.

1977. Seasonal fluctuations in brachyuran nutrient storage: Glycogen in

Col/ineetes midgut glands. Compo Biochem. Pllysiol. 56A:69-71.
WISE, J. P., and B. G. THOMPSON.

1977. Fishery statistics of the United States, 1974. U.S. Dep. Commer.,

NOAA, Natl. Mar. Fish. Serv., Stat Dig. 68,424 p.

YOUNG, C. G.

1900. The stalk-eyed Crustacea of British Guiana, West Indies, and Bermuda. John M. Watkins, Lond., 514 p.

YUDIN, A. I., and W. H. CLARK, Jr.

1978. Two viruslike particles found in the ecdysial gland of the blue crab,

39

Callinecres sapidlls. J. (nvenebr. Pathol. 32:219-221.

1979. A description ofrhabdovirus-like panicles in the mandibular gland ofthe

blue crab, Callineeres sllpidlls. J. Invenebr. Pathol. 33: 133-147.

YUDIN, A. I., R. A. DIENER, W. H. CLARK, Jr., and E. S. CHANG.

1980. Mandibular gland of the blue crab, Callilleeres sapidlls. BioI. Bull.

(Woods Hole) 159:760-772.