Reproductive Biology and Embryology of the ... - Seaturtle.org

32B 

EMBRYOLOGY OF MARINE TURTLEB 

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Voeltzkow, A. (1903). Beitrage zur Entwicklungsgeschichte der Reptilien. VI. Gesichtsbildung 

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Wassersug, R. J. (1976). A procedure for differential staining of cartilage and bone in whole 

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Wiedersheim, R. (1890b). Uber die Entwicklung des Urogenitalapparates bei Krokodilen und 

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CHAPTER 

5 

Reproductive Biology and 

Embryology of the 

Crocodilians 

MARK W. 0. FERGUSON 

Department of Basic Dental Sciences, Turner Dental School, University of 

Manchester, Higher Cambridge Street, Manchester, M 1 5 6FH England

INTRODUCTION 

331 

I. INTROOUCTION 

II. 

REPRODUCTIVE BIOLOGY 

A. General, 333 

B. 

C. 

CONTENTS 

Sexual Maturity and Adult Sex Ratios, 333 

Courtship, Spermatogenesis, Ovulation, Copulation, 

the Breeding Season, Fertilization, and Egg Laying, 336 

D. Nesting Biology, 349 

E. Maternal Behavior, 355 

F. 

Captive Breeding, Egg Collection, and Artificial 

Incubation, 356 

G. Hatching, 360 

331 

333 

P. Endocrine Glands, 445 

Q. Thymus and Immune System, 446 

R, Limbs and Tail, 446 

S, Integument and Its Glands, 450 

T. Caruncle, 450 

VIII. DEVELOPMENTAL ABNORMALITIES 

451 

IX. SHELL·LESS, SEMI.SHELL.LESS, AND IN VITRO 

CULTURE TECHNIGUES 460 

X. CONCLUSIONS 462 

REFERENCES 464 

III. 

THE EGGSHELL AND SHELL MEMBRANES 

A. General, 363 

363 

B. Egg Banding, 363 

C. Structure, 367 

D. Chemical Composition, 374 

E. 

Water and Gas Conductance and Embryonic 

Metabolism, 376 

I. INTRODUCTION 

IV. 

V. 

VI. 

VII. 

THE EGG CONTENTS AND EXTRA.EMBRYONIC 

MEMBRANES 

EARLY EMBRYONIC DEVELOPMENT [BEFORE EGG 

LAYING) 

STAGES OF EMBRYONIC DEVELOPMENT [AFTER EGG 

LAYING) 

ORGANOGENESIS 

A. Branchial Arches, 41 6 

B. Face and Nose, 41 9 

C. 

Palate and Nasopharyngeal Duct, 421 

D. Tongue, 427 

E. Ear, 428 

F. Eye, 428 

G. 

Chondrocranium and Osteocranium, 431 

H. Teeth, 431 

I. Central Nervous System, 432 

..J. Vertebrae and Ribs, 435 

K. Respiratory System, 436 

L. Cardiovascular System, 436 

M. Diaphragm, 438 

N. Gastrointestinal System, 438 

O. 

Urogenital System and Sex Determination, 440 

377 

381 

390 

416 

I" 

t 

I' 

The living crocodilians, represented by 26 living species placed in three 

subfamilies (see Table III later in chapter), are the end products of a relatively 

conservative lineage, which arose from thecodont ancestors approximately 

230 million years ago (Carroll, 1969; Walker, 1972). The evolution 

and taxonomy of the order have been the subject of considerable debate 

involving paleontological (Steel, 1973), neontological (Wermuth and Mertens, 

1977; Groombridge, 1982), immunological, and biochemical approaches 

(Perutz et al., 1981; Le Clercq et al., 1981; Densmore, 1981; Coulson 

and Hernandez, 1983). Densmore's and Groombridge's classifications 

are used here. All types of data indicate that birds are the closest living 

relatives of crocodilians. However, several aspects of crocodilian embryogenesis, 

for example, palatogenesis (Section VII, VIII), resemble mammalian 

phenomena, and variations in hemoglobin amino acid sequences place 

crocodilians closer to mammals than to snakes (Perutz et al., 1981; Le 

Clercq et al., 1981; Densmore, 1981). Thus, further studies of crocodilian 

embryogenesis not only may shed light on fundamental aspects of organogenesis, 

but also exploit the potential of these vertebrates as models 

for experimental investigations, which are difficult or impossible to perform 

in other amniotes (Ferguson, 1981a, b, 1984a). 

Crocodilian embryology has received very little attention. Some general 

accounts are available for Crocodylus niloticus (Rathke, 1866; Voeltzkow, 

1899, 1901, 1903a), Alligator mississippiensis (Clarke, 1891; Reese, 1908, 

1910a-c, 1912, 1915a, b, 1921, 1936), C. palustris, and C. porosus (Derani 

330

332 REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS 


333 

yagala, 1934, 1936, 1939), whereas Wettstein (1937, 1954) included some 

embryological data in his general review of crocodilian biology. The present 

chapter reviews the older, incomplete data and a special effort is made 

to correct ambiguous or misleading statements or concepts. My studies on 

shell structure (Ferguson, 1981c, 1982a), sex determination (Ferguson and 

Joanen, 1982, 1983), and craniofacial development (Ferguson, 1979a, b, 

1981a, b, 1982b, 1984a) tend to give the chapter a bias, but an attempt has 

been made to review all available data in the hope that neglected topics will 

be pursued by others. 

Many of the studies on which the present chapter is based have been 

carried out on the population of Alligator mississippiensis at the Rockefeller 

Wildlife Refuge in Louisiana, and it might be argued that these animals are 

not representative of all alligator populations nor of crocodilians in general. 

However, observations on other populations have revealed no significant 

differences with respect to basic reproductive biology and embryology. 

Furthermore, analyses of protein electrophoretic patterns have shown that 

A. mississippiensis is one of the most genetically homogeneous vertebrate 

species; protein variations are so low that animals from populations separated 

by more than 1000 miles cannot be identified as to their geographic 

source (Gartside et al. 1977; Menzies et al. 1979; Adams et al. 1980). In 

addition, I have made limited observations on some aspects of the embryology 

of Crocodylus johnsoni, C. porosus, C. niloticus, C. cataphractus, and C. 

novaeguineae (Ferguson, 1984a). The basic events during the period of organogenesis 

(first half of development) are remarkably similar and may be 

regarded as crocodilian, whereas species-specific differences are more 

geometrical, for example, variations in the relative sizes and proportions of 

structures such as the tail, limbs, snout, and pigmentation pattern, and 

they appear principally during the second half of development. Therefore 

this account is likely to be representative of crocodilians in general. 

A recent, dramatic increase in the literature on ecology, behavior, natural 

history, management, and farming of crocodilians across the world 

offers hope not only for the survival of these reptiles, but also for the future 

availability of embryonic study material. Such are particularly exciting 

prospects not only for the study of neglected species such as the gharial, 

but also for determining general principles pertaining to crocodilians, and 

for establishing species-specific variations. 

Much of this natural history and farming literature is relevant to embrylogical 

studies. Because it is extremely unlikely that investigations on the 

"T stages of embryogenesis can be conducted on anything but captive 

1

TABLE I 

The Size and Age at Sexual Maturity for Some Crocodilian Species, Including Data for the Same Species Inhabiting Different 

Geographical Locations 

Sex 

Age at 

(M = male Age at Sexual 

F = female Size Sexual Maturity 

grouped = no at Sexual Maturity in 

distinction Maturity in Wild Captivity 

III 

III 

" Species reported) 

(meters) (years) (years) 

Alligator mississippiensis 

M 

1.8 

9-10 

6 

Louisiana 

F 

1.8 

9-10 

6 


North Carolina 

Caiman crocodilus crocodilus 

Caiman crocodilus fuscus 

Gavialis gangeticlIs 

M 

F 

1.8 

1.8 

15 

18 

References 

Joanen (1969), Joanen and 

McNease (1973, 1975a, b, 

1978, 1979a, 1980) 

Klause et al. (1984a, b) 

~} 

1.3 4 Staton and Dixon (1977) 

~} 

1.08 Staton and Dixon (1977) 

~} 

3 8-12 Acharjyo et al. (1975), 

Bustard (1979, 1980c, 

1984), Singh and Bustard 

(1977), Singh (1979)

 

138 REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCOOILIANB 

tains and with its normal growth rate. Forms that mature at smaller sizes, 

r example, Caiman crocodilus fuscus, are obviously well suited for captive 

eeding and developmental study. 

Although sexual maturity may be reached at the sizes specified in Table 

fIl 

social order favors the mating of larger animals, for example, male Al- r: 

.8 

Igator mississippiensis that are longer than 2.7 m (McIlhenny, 1935; Giles r; 

< 

(lj 

(/'J 

trong evidence that for maximum reproductive success (i.e., total number 

-0 

(lj 

f viable hatchlings produced) in captive breeding, the sex ratio ought to 1:: 

o 

ary according to the age/size of the animals (Ferguson, unpublished). 

l:lo 

(lj 

ounger (smaller) animals should be stocked in ratios approaching 1: 1 ~ 

(lj 

particularly if the males are young) and as they mature so the percentage 

o:S 

f females should be increased. 

oS 

fIl 

r: 

C. CourtBhip. Spermatogenesis. Ovulation. Copulation. .8 

the Breeding Season, Fertilization, and Egg Laying o~- ... 

III 

;> 

here is detailed information on the reproductive biology of Alligator mis 

,issippiensis Goanen, 1969; Joanen and McNease, 1970, 1973, 1975a, b, 1978, 

979a, 1980; Lance, 1983, 1984) and Crocodylus niloticus (Cott, 1961; 

raham, 1968) with fragmentary information available for other species, 

~ ..>

338 

REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS 

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340 

REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCDDILIANS 

mone biosynthesis) are present (Lance, 1983, 1984). However, mature 

sperm do not appear in the seminiferous tubules until late April/early May. 

Living sperm first appear in the penial groove in early May and are present 

for only a 6-week period, with maximum output (coincident with female 

ovulation) during a 2-week period in late May/early June (Joanen and 

McNease, 1970, 1972, 1973, 1975a, b, 1979a, 1980). Jenkinson (1913) states 

that crocodilian sperm are small (20-30 jJ-m), but Lance (1983) reported a 

mean length of 60 to 70 jJ-m for alligators. Mature males have sperm concentrations 

of 0.5 to 1 billion per ml of semen (Larsen, 1981). Sperm storage 

occurs principally in the ductus deferens as the epididymis is poorly developed 

(Lance, 1983). Dead, immotile sperm were first noted in mid-June, 

and by June 20th, 90% of spermatozoa were dead (Joanen and McNease, 

1970, 1973, 1975a, b, 1980). Sperm production varies significantly amongst 

reproductive size classes (i.e., 1.8 m and above), and large males (over 2.7 

m) produce living spermatozoa for a longer time period than do smaller 

males (Joanen and McNease, 1980). This may explain the reproductive 

superiority of the large bulls, and why social order favors breeding by 

males over 2.8 m in length. During testicular regression, the diameter of 

the seminiferous tubules decreases, they contain progressively fewer spermatozoa, 

~5-3J3-HSD activity is minimal and the interstitium is composed 

of connective tissue (Lance, 1983). There is no evidence for over winter 

sperm storage in the males; indeed, the epididymides (the usual site of 

sperm storage in postnuptial reptiles, Lofts, 1977; Moll, 1979) are not enlarged 

even at the height of the breeding season (Lance, 1983). 

Plasma testosterone levels show a similar seasonal fluctuation to that 

observed for testicular weight (Fig. 2) with a peak of 90 ng/ml in April/May 

(Lance, 1983, 1984) and rapidly decreasing thereafter. The association of 

elevated testosterone levels with the peak of testicular weight, spermatogenic 

activity, bellowing intensity, and mating behavior (Fig. 2) suggests 

that these functions are androgen dependent, although this remains to be 

tested experimentally. The mechanism initiating gonadal regression is unknown 

although Lance (1983) discusses several possibilities. Clearly the 

topic is worthy of investigation not only for its biological interest, as it 

occurs at a time when temperature and photoperiod appear to be optimal, 

but also for its importance in alligator farming. The plasma testosterone 

rises slightly in September, when the alligator testes are fully regressed 

(Lance, 1983). This is difficult to explain, but it does not coincide with 

mating behavior, spermiogenesis or renewed spermatogenesis. The profile 

is, however, typical of postnuptial spermatogenic cycles (Lance, 1983) and 

may represent the curtailing of another breeding cycle imposed by the 

climatic habitat of Alligator mississippiensis, the most northerly ranging 

crocodilian. In this regard, comparison between annual testosterone levels 

in this species with those in more tropically located crocodilians (particularly 

those suspected of having two breeding seasons, e.g., Crocodylus 

niloticus, C. palustris, C. porosus, and C. johnsoni) would be profitable. 


The reproductive system of females shows seasonal variation. Each 

ovary contains at least three size classes of follicles:

342 

REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCOOILIANS 

taken up by the follicles (Lance, 1983). Immature alligators of either sex can 

be induced to produce vitellogenin by injection of estradiol (Dessauer, 

1974; Lance, 1983). The oviduct is also stimulated by ovarian oestrogen and 

hypertrophies during the follicular growth phase (Lance, 1983). After 

oviposition, plasma estradiol levels remain low to nondetectable, and there 

is no evidence for a prehibernation onset of vitellogenesis (Lance, 1983). 

Nonbreeding females have low to nondetectable estradiol levels (Lance, 

1983) and show nO evidence of follicular growth (Joane n and McNease, 

1980). Likewise, immature females have low estradiol levels and small 

ovaries that consist of clear cortical tissue with numerous clusters of 00 

cytes distributed evenly throughout the organ. Reproductively senescent 

or "barren" females (usually over 2.7 m in length) have few growing 00 

cytes, no corpora lutea, black hemoglobin deposits, and their ovaries frequently 

contain medium-sized resorbing ovarian follicles and retained 

oviducal eggs undergoing resorption (both in Alligator mississippiensis, 

Joanen and McNease, 1980; Lance, 1983 and in Crocodylus niloticus, 

Graham, 1968). The occurrence of senescence in the Crocodilia suggests 

that there is a limit on the number of oocytes a female can produce, despite 

the fact that oogonial proliferation continues throughout much of adulthood. 

Macroscopic changes similar to those detailed above have been described 

in the male and female reproductive systems of Crocodylus niloticus 

(Cott, 1961; Graham, 1968). However, the breeding season of Alligator 

mississippiensis is shorter and more clearly defined than is that of the more 

tropical C. niloticus (Graham, 1968), C. porosuS (Webb et al., 1983b), and C. 

palustris (R. Whitaker, personal communication). This may reflect environmental 

temperature; the cold winters in Louisiana make the alligator inactive 

from October through February, and it must complete its reproductive 

cycle within a restricted time frame. ThuS for C. niloticus in Kenya, sperm 

production extends from April through December, and although there is a 

peak breeding season (October-December), follicular development and 

some reproduction occur throughout the year (Graham, 1968). Moreover, 

15% of the sexually active females of C. niloticus have two sets of follicles 

developing concurrently (Graham, 1968). This means that maturation of 

one batch of ova may be accompanied by development of a younger batch 

that will near maturity soon after oviposition of the first; the second set can 

then be fertilized and two clutches of eggs may be produced in one season. 

This "double clutching" has also been reported for some C. palustris (in 

captivity, R. Whitaker, personal communication) and is suspected in some 

C. porosus (Webb et al., 1983b). The controlling mechanism remains uninvestigated. 

However, given the amount of yolk protein necessary to produce 

two clutches of approximately 40 eggs weighing apprOXimately 60 

70 g each, not to mention the energy expended in mating behavior, nest 

building, and maternal care, it may be assumed that only females in an 

REPROOUCTIVE BIOLOGY 

343 

optimal environment (in terms of temperature and food) may pursue this 

strategy. 

The data on the peak nesting times of various species presented in Table 

III generate many unanswered questions: What environmental factors trigger 

ovulation, spermatogenesis, reproductive behavior, and egg laying, 

and how do they operate? 

Numerous reports for various species imply a possible correlation 

among breeding times and changing water levels in the various habitats. 

The data do not reveal consistent trends across all species and the interested 

reader is referred to the original publications (Cott, 1961, 1975; 

Modha, 1967; Graham, 1968; Pooley, 1969a; Staton and Dixon, 1977; Deraniyagala, 

1939; Webb et al., 1977, 1983e; Joanen, 1969; Joanen and 

McNease, 1975a, 1979b, 1980; Kushlan and Kushlan, 1979). 

A strong relationship exists between peak nesting times in Alligator 

mississippiensis and average air temperatures. An analysis of nesting data 

for ten years reveals a highly significant correlation between the time of 

nesting and the ambient temperatures for March-May (Joanen and 

McNease, 1979b). Nesting occurs earliest when these temperatures are 

highest. Rainfall is significant only when nesting effort is reduced during 

extremes of accrued surface water levels; females apparently respond to 

higher water levels by laying eggs at higher levels within the nest (Kushlan 

and Kushlan, 1979). Oviposition occurs on the longest days of the year 

(Joanen and McNease, 1979b). Temperature appears to be the major regulator 

of male and female reproductive cycles (Lance, 1983); as in other 

reptiles photoperiod plays only a minor, or at best synergistic, role (Fischer, 

1974; Crews and Garrick, 1980). It is known that males in an 

artificially heated pond in Georgia had spermatozoa in the penial groove 

one month earlier than their counterparts inhabiting ponds at ambient 

temperatures (Murphy, 1980). Moreover, motile sperm have been recovered 

in January from males kept in artificial hot houses (Cardeilhac, 

1981). Whereas adult A. mississippiensis do respond to experimentally altered 

photoperiods (Lang, 1976), such experiments have not been conducted 

in relation to the cueing of reproductive events. Experiments involving 

altered photoperiods at constant high temperatures would be 

interesting, especially for tropical species in regions with minimal annual 

variations of temperature. 

It is currently impossible to state categorically which of the potential 

extrinsic regulating factors, such as drought, temperature, water level, 

photoperiod, or the psychological effect of available breeding grounds, 

stimulate ovulation and spermatogenesis, and whether these factors are 

similar in all crocodilians. Clearly more information is required, not only 

on the stimuli themselves, but also on their perception and the way in 

which they achieve their effects. 

The hormonal regulation of the reproductive cycle is poorly understood.

TABLE III 

Comparative Data on the Nests, Eggs, and Hatchlings of the Crocodilians"'C 

~cies 

ligator lineage 

Type of Nest 

(from Greer 

1970, 1971; 

Campbell 1972) 

Average SizC' 

of Nest 

(Breadth x 

Height/ 

Dep'h) (em) 

Peak Times 

of Egg 

Laying 

Incubation 

Period 

(days) 

Average 

Number 

of Eggs 

per Clutch 

Range of 

Egg 

Numbers 

per Clutch 

Average 

Length 


(em) 


Egg 

Lengths 

(em) 

Average 

Breadth 


(em) 

Range 


Breadths 

(em) 

Average 

Weight 


(g) 

Range 


Weights 

(g) 

Average 

Temperature 


Cavity in 

Nest ('C) 

Range 


Cavity 

Temperatures 

(0C) 

Average 

Humidity of 

Egg Cavity in 

Nest (c/o ReI. 

Humidity) 


d 972 

Mound Av = 181.6 June 63-65 38.9 2-68 7.4 6.1-88 4.3 3.1-5.0 84 51-108 30 233-35.7 

x 60.2 

(45-799, 

Range = 

moisture 

112-304.8 content) 

x 33-121.9 


Humidities 

(% Rel. 

Humidity) 

Average 

Total 

Length of 

Hatchling 

(em) 

Range 

of Total 

Hatchling 

Lengths 

(cm) 

Average 

Weight of 

Hatchling 

(g) 


Hatchling 

Weights 

Ig) 

94-99.8 26.4 22-29 676 58-77 

A. sinensis e Mound 106 x 29.2 June-Aug 70-80 26 10-40 68 5.6-75 3.4 3.1-3.6 51.7 40.3-56 21 30.2 

Caiman CTOCQdi/us Mound ? x 30 75-90 30 20-50 6.2 3.8 

apapoTjen~·s-f 

C. cTocodilus 

rrocodilus g Mound 117 x 44 Aug.-Oct 73 28 17-40 6.5 4.3-7.2 4.0 2.5-43 59 48.7-774 30 28-32 90 85-95 21.5 191-23.8 41.5 31-51.2 

C. crocodilus 

yacare" 

Mound 150 x 40 Dec.-April 84 31 21-40 6.8 5.6-7.5 4.2 3.3-4.7 75.3 59-83 28 25-32 24 20-25 492 46-62 

C. CTocodilus } 

fusnd and 

C. crocodilus 

Mound April-July 75-80 15-30 

chiapasius ' 

C. 1oliTastris} Mound 163 x 41 Aug.-Mar. 63-86 40 20-60 6.6 4.6 84 

.\Vlanosucnus Mound 210 x 100 Sept.-Jan. 40-90 40 18-75 8.0 4.4-9.7 5.0 3.5-5.6 100+ 

nigeri: 

Pa/eosuchu5 

palpebrosus l Mound 125 x 39 Aug 90-92 13 6.6 6.1-7.1 4.2 4.1-5.1 687 66.1-74.5 28-31 95 90-96 23.6 202-24.5 45.5 39.8-50.0 

p, trigonafus Mound 

aviallineage 

Gavialis 

gangt'ficus m Hole ? x 5.1 April 71-94 40 16-61 8.6 8.5-10.2 6.7 6.5-7.0 31 25-37 35 32-40 75 

Tomistoma 

St'hlegelii!f 

rocodHe lineage 

Osteolaemus tetraspis 

osborn; 

Osteolaemus tetraspis 

tetraspiso 

Crocodylus 

acutusP 

Mound ? x 60 75-90 20-60 10.1 9.7-12.0 7.6 6.4-8.0 30 28-33 

Mound 

Mound 75 x 41 June 82-120 13 6-19 6.3 5.5-8.0 3.7 3.4-4.4 51.9 38.5-70.7 28.8 22.6 19.7-25 34.3 262-445 

Hole/mound 

of sand 

230 x 33 April-May 85-111 44 19-81 72 6.3-·7.6 4.4 4.2-5.1 30 265-341 24.1 

C. cataphracfu;;q Mound 50-80 x Mar.-July 90-98 19 13-27 8.3 8.1-8.5 5.2 

305 27-34 100 

120-220 

(37% mois 

303 28.3-35.6 

ture content) 

C. intermedius r Hole Jan.-Feb. 60 1.5-70 

C. ;ohnsoni s Hole 

19 x 13 August 63-98 13 10-24 6.6 6.1-7.3 4.2 

3.7-4.6 

C. mindormsis 1 Mound 150 x 40 April-July 85 12 

7-14 6.5 61-7.2 4.0 

3.6-42 

68.2 498-85.8 29.4 28-34 24.4 21-26 42 36-54 

30 

26-31 

87 85-90 21.5 16-23 

C niloticus F Hole 

60 x 40 Jan.-Dec. 84-98 55 25-95 7.5 5.5-9.0 4.8 40-5.5 110 85-125 31 

22-34 

depending 

28 27-37 80 

C. moreletii U 

Mound 300 x 100 July 

78-89 

20-45 10 

on rains, 

sometimes 

two seasons 

C. Hovaeguineae Mound 

novaeguineaei.l' 

122 x 61 Nov.-Jan. 83-87 26 12-40 7.6 6.4-8.8 4.3 3.6-5.8 85 63-111 32 30-38 77% H20 28 24-33 

C pa/ustri~ 

by weight 

Hole;n sand 30 x 50 Feb.-Aug. 60-80 26 6-41 7.5 60-8.8 4.6 3.7-5.2 83.7 47-120 32 

or? mound 

26-41 

70 25 22-30 70 60-110 

C. porosus Ceylon; Mound 170 x 53 Dry season 70-90 42 25-72 8.2 61-9.6 5.1 4.2-5.7 113 105-135 32 25-37 

July-Aug. 

30 28-32 61.7 82-95 

C. porosus 

AustraliaY 

Mound 

200-350 x 

80-100 

Wet season 

Nov.-Mar. 

C. rhomb~rerz Hole/mound April 68 20 7.6 51 

C. siLlmensis llll Mound April-July 68-80 20-48 7.6 5.1 

~able Footnotes: see page 346. 

80-98 50 16-71 7.7 6.6-89 5.2 4.2-5.7 113 65-143 30.1 25-37 32 31-33 92 82-95 

344 

346

348 

REPRODUCTIVE BIOLOGY AND EMBRYOLOGY DF CRDCODILIANS 

No available data indicate that the hypothalamus can respond to changing 

blood temperatures by the secretion of releasing hormones. However, two 

distinct glycoproteins similar in molecular weight and amino acid composition 

to mammalian follicle stimulating hormone (FSH) and luteinizing hormone 

(LH) have been isolated from the pituitaries of Alligator mississippiensis 

(Licht et al., 1974, 1976). Both of these gonadotropins stimulate 

testosterone production by ovarian and testicular tissues, with LH being 

slightly more effective (Licht and Crews, 1976; Tsui and Licht, 1977). There 

are no data on circulating gonadotropin levels in any crocodilian. Follicular 

development can be stimulated artificially with pregnant mare serum 

gonadotropin, but artificial induction of ovulation has not been achieved 

(Cardeilhac et al., 1982). Alligators (unlike turtles) show a marked response 

to the synthetic decapeptide luteinizing hormone releasing hormone (LH 

RH) (Lance, 1983, 1984). Alligator LH-RH is similar to that of mammals but 

differs from it in having at least one amino acid substitution in the 8 

position of the molecule (Lance, 1983, 1984). This is similar, if not identical, 

to avian LH-RH (King and Millar, 1982). Tissue levels of between 816 and 

2780 pg/mg protein have been estimated for the alligator hypothalamus 

(Lance, 1983, 1984). 

Footnotes for Table IfI 

UThc figures quoted are poolt'd maximum, minimum, ilnd i'l\Trage villul's frpm Ihe \'i1riou~ publications cited Older accounts \,",'ith 

obViously exaggerated figures, e.g., Boake 11870) have been omitted. 

'Species classification b.:lscc! on Densmore (1981) and Groombridge (1982) 

dClarke (I88Ba, b, 1891); Reese (1915b); McIlhenny 0934, 1935); Giles and Childs (1949): POpt> (1956); Jnolnen (196'J\; Ch,lDred< (197:3, 

19(5); Joanen dlHj McNease (1975a, 1978, 1979f, :980); GoodWin .'lnd :\1anon (1978). 

loanen (personal communication); Chu (1957); Chu-Chcn (1982); Behler dnd Joanen (19tl2). 

tCasas and Guzman (1970); Del Toro (l974). 

gHagmdnn (1906, 1909); Schmidt (1928); Del Taro (1969); Hunt (1969); Neill (1971); Staton dnd Dixon (1977); Corzula (197R) 

hFort Worth Zoo (personal communication); Mcdem (1960); Cuy;gisber~ (l\i72); Crawshay.,' and Schaller (980). 

'Neill (1971); Del Taro (1974) 

'Neill (l q 71); Groombridge (1982). 

kHagmann (1902a, h, 1909); Recse (1923); Mellem (1963); Croornbridge (19~2). 

IRec5e (1931 a); Medcm (1971, 1972). 

m Ander50n (lB7S); Butler (1905); Werner (1933); Singh and Bustard (1977); Singh (1979); Bu"tard (1lJ79, 1980a, 1984); Acharjyo et cd 

(1975). 

TlMiil1er (1838); De Rooij (1915); Butler (1905): Sudharfna (1976) 

°Guib(' (19701); Neill (1971); Tr"'on (980) 

PDescourtilz (1809); Schmidt (1924); Werner (143.1); Rand (196H); Neill (1971); Carrick and Lln/; (1977); Ogden (1978); CasaS' .'lnd Cuzrn,1n 

(1970). 

QWaitkuwail (1982). 

TBlohm (1982); Medem (1976). 

'Dunn (1977); Webb (1977); Webb e, al. (1983e). 

I Croombridge (1982). 

UHunt (1973,1975); Del Toro (1975); Pcrcz-IJigdredd (1%0). 

l'VoeHzkow (1899); Bigalke (1931); Cott (1961, 1969, 1(75); Pooley 0962, 19fil}il, 1971. 1977); Modh


lay in August. Thus the interval between copulation and egg laying could 

be as short as one month. Detailed autopsies to check for the state of 

ovarian development in different species are required to determine 

whether the interval between copulation/ovulation and egg laying varies 

as widely as the present data suggest. 

Because crocodilians range over wide areas with drastically varying 

climatic conditions, a prolonged interval between copulation and egg laying 

may offer some advantages (Webb et aI., 1983e). During these time 

intervals, there may be (1) sperm storage in the female and delayed fertilization 

or (2) a very slow rate of embryonic development while the eggs are 

in the oviducts (embryonic hibernation). Sperm storage occurs in birds (R. 

Bellairs, 1971), in many lizards and snakes (Saint Girons, 1962; Cuellar, 

1966; R. Bellairs, 1971), and in some turtles (Barney, 1922; Ewing, 1943; 

Smith, 1956). In artificial insemination experiments, sperm from Alligator 

mississippiensis has been stored successfully for many days (Joanen and 

McNease, 1973, 1975b; Larsen, 1981; Larsen et aI., 1982). The length of time 

that sperm remain viable within the female reproductive tract is unknown. 

No sperm storage structures can be identified in the female reproductive 

tracts of A. mississippiensis or Crocodylus Iziloticus from Zimbabwe. 

Embryos are stored in some turtles (Oliver, 1955; Ewert, 1979) and in the 

tuatara (Sharell, 1966), but this phenomenon is undocumented for crocodilians. 

Embryos of Alligator mississippiensis cannot be arrested experimentally 

(or naturally) in their development at any time after fertilization. It is 

possible to slow development by cooling the eggs but temperatures below 

26°C for prolonged periods are usually lethal (Ferguson, unpublished 

data). Likewise, freshly laid eggs of Crocodylus niloticus from Zimbabwe 

apparently die at temperatures below 26°C (Hutton, personal communication). 

Embryos of A. mississippiensis, C. johnsoni, C. niloticus, and C. porosus 

are laid at the same stage of development (Ferguson, 1984a), which is 

considerably advanced (see Section VI) as compared to avian or turtle eggs, 

which are laid at the late blastula and late gastula stages, respectively 

(Ewert, 1979). The stage of embryonic development at laying is not invariant, 

and egg retention within the body of females faced with adverse 

environmental conditions (e.g., drought, flooding, temperature variations) 

at the time of normal egg laying has been reported (McIlhenny, 1934, 1935; 

Reese, 1907, 1915a, 1931a). Females kept in artificial pens at high stocking 

densities exhibit elevated plasma levels of corticosteroids. These levels 

inhibit ovulation and hence delay egg laying; however, stressed females 

also retain eggs within their oviducts for abnormally long times. Such 

freshly laid eggs may contain embryos that are considerably advanced and 

have already become attached to the inner aspect of the shell membrane, 

so that there is an opaque band, or it develops shortly thereafter (see 

Section III). Such eggs are frequently deposited with the embryo on the 

underside which causes death (see Section II.F.). How long crocodilians 

can retain their eggs, the rate of development during this period as com- 


349 

pared to the "normal" rate within the nest, and whether egg retention 

affects the viability of the embryo are all unknown. 

The stimuli for egg laying are presumably extrinsic factors, such as temperature, 

humidity, water levels, and photoperiod. Precisely how they are 

involved, how they are detected and how they produce their effects are all 

unknown. Turtles may be induced to deposit eggs prematurely by injections 

of one to three units of synthetic oxytocin per 100 g body weight 

(Ewert, 1979). An analysis of the circulating levels of this hormone in 

crocodilians around the time of egg laying would be useful, as would 

similar experimental injections into gravid females. If this technique were 

successful in "forcing" premature egg laying, it would facilitate obtaining 

the earlier stages of embryonic development, particularly in the rarer, endangered 

species of crocodilians. 

D. Nesting Biology 

An extensive literature on crocodilian nesting biology has accumulated, 

and detailed accounts are available for Alligator mississippiensis (Feilden, 

1810, Clarke, 1888a, 1891; Reese, 1915a, 1931a; McIlhenny, 1934, 1935; 

Pope, 1956; Joanen, 1969; Joanen and McNease, 1971, 1973, 1975a, b, 1980; 

Neill, 1971; Fogarty, 1972, 1974; Goodwin and Marion 1978; Deitz and 

Hines, 1980), Crocodylus acutus (Ogden, 1978), C. cataphractus (Waitkuwait, 

1982), C. jolmsOlzi (Webb, 1977a, b; Webb et aI., 1983e; Compton 1981). C. 

novaeguineae (Graham, 1981), C. niloticus (Voeltzkow, 1891, 1892, 1893, 

1899; Cott, 1961, Guggisberg, 1972; Modha, 1967, 1968; Graham, 1968; 

Graham and Beard, 1973; Graham et aI., 1976), C. palustris (Symons, 1918; 

Deraniyagala, 1939; Whitaker and Whitaker, 1976a,b, 1977a, b, c; Yadav, 

1969, 1979), C. porosus (Deraniyagala, 1936, 1939; Webb, 1977a, b; Webb 

et aI., 1977, 1983b; Magnusson et aI., 1978; Magnusson, 1980), Caiman 

crocodilus crocodilus (Del Toro, 1969; Hunt, 1969; Staton and Dixon, 1975, 

1977), and Osteolaemus tetraspis (Tryon, 1980). The present account only 

highlights areas of embryological interest; strictly ecological data are 

omitted. 

Some crocodilians construct mound-like nests out of vegetation, 

whereas others dig holes in the ground (Table Ill). The phylogenetic (Greer 

1970, 1971; Campbell, 1972) and ecological (Webb et aI., 1983e) implications 

of these two strategies have been debated. There is no well-documented 

example of both strategies occurring naturally in the same species; the 

behavior is maintained even in sympatric species (such as Crocodylus johnsoni 

and C. porosus in which a change of strategy from hole to mound by C. 

jolmsoni might decrease its embryonic mortality due to flooding). 

Eggs normally are laid during the night or early morning (Voeltzko w , 

1899; Cott, 1961). The female positions her cloaca over the egg cavity and 

deposits the entire clutch at one time. Presumably, the eggs move down 

the oviducts and through a shell gland one at a time, so that the first

350 REPROOUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS 

b!l 

OJ I-< 

shelled eggs are stored longer than the last. However, they all appear to be ..c OJ 

'O.S .... 0 ~ ~ 

c"1:S III en 

Lf) N c OJ 

at comparable stages of development at the time of laying (see Section VI). 

If': .- III 0... 

0'"-< 

£! 6 '" .-< .-< o .-< 

When first laid, the eggs are coated with a slimy gel (Goodwin and Marion, i3b c .... 0 +1 

~ 

"' N 

..... 1 

'y; til 00» +1 +1 +1 500 

1978). On average, 3 to 15% of eggshells are cracked during the laying "" 

t ~u::;a15..c: o Lf) Lf) 

.-< Ei x 

.~ 

o 

OJ 0 

>"'~6.B I-< I-< 

process (Joanen, 1969; Pooley, 1969a; Webb et al., 1977; Goodwin and 

-< ~ ~..2 

::l D 

.~ ~ u D 

Marion, 1978). Cracked eggs develop normally if the shell membrane re 'J> 

'J> OJ~ 

'" '" 

'y; 

mains intact, but if it is torn, the eggs rot or are eaten by ants (Joanen, 1969; 

Ei '" 

'J> 

OJ 

.~ '" 

Ei..c 

Goodwin and Marion, 1978). The number of eggs present in each nest 

If) 

r--.. o ~ 

If': 

.. 

~ {3 0 ọ I-< ::l 

-< c:i o '"--l III ~ 0 ~en -- '"d ....... 

I:Q Ei 

;; > "8 '[ Ei 

_ ("lj lr) 

maternal age and clutch size, egg size, and egg quality in Alligator mississip ., -< 

piensis (Table IV). "Middle-aged" females are obViously the most "success QI 

~~~ 

00 

'w;:n "0 

ONe 

ful" breeders in terms of both quantity, size, and quality of the eggs pro 

00 6 

~ 

-O~00 '" 

;::J CJ 

duced. Interestingly, old females are nearly as successful, although their '1:l 6..c: C· ..... -£ 

~ ..0 

C .- 'w 

Lf) 

clutch size is usually smaller and more malformed embryos result, whereas III Ei § c 

~ 6 0 N ;:l ..... "O"'~ ~ 

-< 

'" 

Z 0 ~ +1 

'D o 

.5 N ~ N OJ OJ '" 

males (which may mate with either large or small males). Similar relation 0... 

'" c w 

"" 

a ~ '" 

I:I.. til W"'"O 

season. 15 ~ '" ~ I ;:l 6"1:S ~ gg c 

o~ bO~~_'""Ol-log 

OJ OJ '" 

For several species, weight of eggs correlates positively with that of §~:g~8..8:;c ~ &3 

~:< 

eno 

.- ro 0-,; bJ ........ rc 

hatchlings (Staton and Dixon, 1977; Deitz and Hines, 1980; Webb et al., 

E5 gg 

>

3152 

REPROOUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS 

1983b, e). In mammals, fetuses with a low birth weight are frequently 

malformed, and the same is true of the small light eggs laid by young 

female alligators (Table IV). The alligator is an excellent animal model in 

which to study such malformations. The eggs of middle-aged females are 

also useful for teratogenic studies as they show a lower basal incidence of 

spontaneous malformations than avian eggs (Ferguson, 1981a). 

Possible factors that determine the relationships of egg fertility and rate 

of embryonic malformation with maternal age include the following: (1) 

reduced ovulation of young and old females, (2) the immaturity of young 

ova, (3) slight asynchrony between the times of ovulation and copulation 

in young females, (4) the lower sperm numbers and quality in males near 

the end of the breeding cycle (when young females mate), and (5) the 

poorer quality of the large ova released by old females. There is some 

evidence that prior to senescence old females either resorb their ova or 

their eggs, or both. Possibly, young and very old females contribute little to 

population recruitment. 

Nest temperature and humidity are important factors for embryonic 

survival, development, growth, and sex determination (Table III). Only 

Alligator mississippiensis (McIlhenny, 1934, 1935; Joanen, 1969; Chabreck, 

1973), Crocodylus porosus (Webb, 1977a; Webb et a1., 1977, 1983b; Magnusson, 

1979c), Caiman crocodilus (Staton and Dixon, 1977) and Crocodylus 

acutus (Lutz and Dunbar Cooper, 1982, 1984) have been studied in detail. 

In general, the positioning and construction of nests ensure fairly stable 

thermal and humidity conditions within the egg cavity. 

Crocodilian nests may obtain three sources of heat: solar heating, decomposition, 

and metabolism. The temperature of the nest cavity is 

equilibrated; the outer layers of the nest absorb solar heat at times of 

maximum incidence, but slow its passage to the central cavity; some of this 

heat later dissipates to the central cavity. However, solar energy is unlikely 

to be the sole source of heat, as crocodilian mean nest temperatures are 

often 1° to 5°C above the mean air temperature (McIlhenny, 1935; Joanen, 

1969; Chabreck, 1973; Modha, 1967; Webb et a1., 1977, 1983b; Staton and 

Dixon, 1977). 

The temperature of mound nests may be raised by the decomposition of 

the nesting vegetation, particularly during the early stages of incubation 

(McIlhenny, 1935; Joanen, 1969; Chabreck, 1973; Webb et a1., 1977a, 1983b; 

Staton and Dixon, 1977; Magnusson, 1979c). Such decomposition may be 

enhanced by females urinating onto the nest during and after its construction 

(Pooley, 1962, 1969a, b; Chabreck, 1975; McIlhenny, 1934, 1935; Reese, 

1915a; Deraniyagala, 1936, 1939; Whitaker and Whitaker, 1976b, 1977b; 

Tryon, 1980; Webb et a1., 1983b). However, decomposition should be 

minimal during the latter stages of incubation in mound nests and nonexistent 

in hole nests. Embryonic metabolism has also been postulated to represent 

a source of heat (Modha, 1967; Magnusson, 1979c; Webb et a1., 

1983b). Clearly, the extent of such heating varies with the proportion of the 


3153 

dutch containing living embryos. The assumption (Staton and Dixon, 1977; 

Webb et a1., 1983b) that the rate of heating increases up to the end of 

incubation implies that heating relates more closely to the size of the embryo 

(Drent, 1967) than to its metabolic rate. Indeed, the peak weightspecific 

metabolic rate in C. porosus occurs during the period of organogenesis 

(i.e., during early/middle incubation; G. Grigg, personal 

communication). This assumption correlates with the fact that nests of A. 

mississippiensis are often warmer during the second to fourth week of incubation 

than later in their 9-week incubation period (Chabreck, 1973). Moreover 

the hole nests of C. acutus (Lutz and Dunbar Cooper, 1982, 1983) and 

C. joJmsoni (Webb and Smith, 1984) become progressively warmer during 

incubation, and this temperature rise may accelerate development and 

improve survivorship (Webb and Smith, 1984). 

Although critical for successful embryonic development, nest humidity 

and the water relations of crocodilian eggs are poorly understood. ,All 

crocodilian nests studied to date have exhibited very high humidities 

(Table III). Precise data on the mechanisms of humidity maintenance are 

lacking, but there have been reports of maternal behavior involving urination 

and splashing of the nest (e.g., Whitaker and Whitaker, 1976b, 1977b). 

If this is deliberate, it would imply that the mother can monitor nest temperature 

and/or humidity, and it has been suggested that this might be the 

function of sensory receptors on the jaws (Ferguson, 1979b, 1981b, 1982b; 

see also Section II.E). However, during abnormally hot dry years, the 

humidity within nests of A. mississippiensis decreases, and the eggs begin to 

form abnormal air spaces (Ferguson and Joanen, 1983) due to cleavage of 

the shell membrane away from the calcified eggshell. If the air space is 

large, it causes embryonic death, representing a significant source of mortality, 

particularly among eggs near the top of alligator nests during hot dry 

years. 

Nest temperatures and humidities outside the normal range lead to 

malformed or dead embryos (Modha, 1967; Bustard, 1969, 1980a, c; Pooley, 

1962, 1969a, h; Whitaker and Whitaker, 1976a, b, 1977b; Joanen and 

McNease, 1975a, b, 1977, 1979a, 1980, 1981; Staton and Dixon, 1977; Ogden, 

1978; Ferguson and Joanen, 1983; Webb et a1., 1983a,b,e). Within the 

range of viable temperatures, temperature and incubation time are inversely 

correlated (Reese, 1915a; McIlhenny, 1934, 1935; Pooley, 1962, 

1969a,b; Joanen and McNease, 1975a, b, 1977, 1979a, 1980; Packard et a1., 

1977; Webb et a1., 1977, 1983a,b,e; Ferguson, 1982b; Ferguson and Joanen, 

1982, 1983). Thus, there are only mean incubation periods (Table III), and 

field incubation times are difficult to interpret even with scant information 

on nest temperatures. Consequently, developmental stages must be related 

to an arbitrary (normal) time scale calculated for controlled environmental 

conditions (Ferguson, 1982b; Webb et a1., 1983a). 

The gaseous conditions within crocodilian nests have been assayed 

twice, once for the hole nests of Crocodylus acutus (Lutz and Dunbar

354 REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCODILIANS 

REPROOUCTIVE BIOLOGY 

355 

Cooper, 1982, 1984) and again for mound nests of C. porosus (Grigg, unpublished 

data). The data do not differ significantly. Shortly after egg 

laying, the pOz in the nest is 147 torr (ambient =: 150), and the pC02is 5.0 

torr (ambient =: 0.3). These values reflect the properties of the nest as the 

same values are obtained whether or not eggs are present (Grigg, unpublished 

data). Embryonic arterial blood is approximately 86% saturated and 

venous blood about 40%: arterial pOz is 52 torr (range 42-57), arterial pC02 

is 12 torr (range 8-14), and venous pOz is 22 torr (Grigg, unpublished 

data). Clearly, the eggshell and shell membrane represent a significant 

barrier to gaseous diffusion (see Section III.E). As incubation proceeds, 

nest p02 falls and pCOz rises (Lutz and Dunbar Cooper, 1982, 1984; Grigg, 

unpublished data). Crocodilian embryos can tolerate extremely low p02 

levels and recover; furthermore, they are very resistant to raised pC02 

levels. Even pCOz levels as high as 60 torr do not inhibit the embryonic 

metabolic rate (Grigg, unpublished data). 

Flooding of crocodilian nests has long been known to cause embryonic 

death (Reese, 1915a; McIlhenny, 1935; Joanen, 1969; Joanen et aI., 1977; 

Fleming et aI., 1976; Hines et aI., 1968; Goodwin and Marion, 1978; Nichols 

et aI., 1976; Voeltzkow, 1891, 1892, 1893, 1899; Webb, 1977a; Webb et al., 

1977, 1983b and e; Magnusson, 1982; Deraniyagala, 1939; Cott, 1961, 1969; 

Pooley, 1962, 1969a). Joanen et al. (1977) tested the effects of simulated 

flooding on hatchability by immersing the eggs of Alligator mississippiensis 

for single periods of two, six, 12, and 48 hours at different stages of development. 

Immersing eggs of any age for two to six hours had little effect on 

subsequent hatchability, nor did submerging eggs for 12 hours if done 

before day 30 (after laying). Thereafter, 12 hour submergence killed all 

embryos. Submergence of the eggs for 48 hours killed all the embryos at all 

ages. Similar results are recorded for experiments on Crocodylus porosus 

eggs (Magnusson, 1982). Any moisture on the surfaces of crocodilian 

eggshells drastically reduces their gas conductance and may totally inhibit 

oxygen diffusion. It therefore seems likely that flooding inhibits gaseous 

diffusion, and thus causes embryonic death: a similar mechanism operates 

in chicken eggs (Kutchai and Stean, 1971). All embryos could utilize air 

within the eggshell and so withstand short periods of flooding; whereas 

younger embryos would have a lower O 2 requirement and relatively more 

air within the eggshell, so that they could withstand longer periods of 

flooding than older embryos. 

Factors that influence the site selected by females for nest construction 

are complex and incompletely understood but probably include social interactions 

with other animals, vegetation type at site, proximity of the site 

to water, its temperature, degree of exposure to sunlight, and height above 

water level. Whereas investigations of these parameters are, by definition, 

ecological and behavioral, there can be no doubt that these factors affect 

fecundity within a population. Further studies are likely to contribute not 

onlv to our understanding of crocodilian reproductive strategies, but may 

shed light on the role of incubation temperature in determining the sex 

ratios of hatchlings (Section VILO). 

E. Maternal Behavior 

Those behavioral repertoires of female crocodilians that improve reproductive 

efficiency are not restricted to nest selection and construction (Section 

11.0), but include subsequent nest protection and opening, egg opening, 

transport of hatchlings, and parental care. They have been described for 

Crocodylus acutus (Campbell, 1973; Garrick and Lang, 1977), C. cataphractus 

(Waitkuwait, 1982), C. johnsoni (Dunn, 1981; Webb et aI., 1983e), C. 

moreletii (Hunt, 1975, 1980), C. niloticus (Cott, 1961, 1909, 1971, 1975; Garrick 

and Lang, 1977; Pooley, 1962, 1969a, 1974a, b, 1976, 1977; Pooley and 

Gans, 1976; Voeltzkow, 1891, 1892, 1893, 1899; Bohme, 1977; Guggisberg, 

1972; Neill, 1971), C. novaeguineae (Neill, 1971), C. palustris (Symons, 1918; 

Deraniyagala, 1939; Whitaker and Whitaker, 1977a, b), C. porosus (Deraniyagala, 

1936, 1939; Webb, 1977a; Webb et aI., 1977; Magnusson, 1980; 

Bustard and Kar, 1981), Alligator mississippiensis (Reese, 1915a, 1931a; 

McIlhenny, 1935; Giles and Childs, 1949; Lee, 1968; Joanen, 1969; Campbell, 

1973; Kushlan, 1973; Herzog, 1975; Garrick and Lang, 1977; Garrick et 

aI., 1978; Kushlan and Kushlan, 1980; Kushlan and Simon, 1981; Deitz and 

Hines, 1980; Hunt and Watanabe, 1982), Caiman crocodilus crocodilus (Del 

Toro, 1969; Campbell, 1973; Garrick and Garrick, 1978; Staton and Dixon, 

1975, 1977), Osteolaemus tetraspis (Tryon, 1980), and Cavialis gangeticus 

(Singh and Bustard, 1977; Bustard, 1980b,c,d; Bosu and Bustard, 1981). 

Maternal and prehatching vocalization of crocodilians are reminiscent of 

avian behavior (Vince, 1969, 1973; Gottlieb, 1973; Oppenheim, 1973). In 

birds, they are supposed to accelerate and synchronize late embryonic 

development in the clutch of eggs. The same effects have been suggested 

for crocodilians (Lee, 1968). However, a single experimental study (Magnusson, 

1980) showed that advanced embryos of Crocodylus porosus called 

in response to a tape recording of wild hatchling vocalizations, but neither 

developed faster nor hatched more nearly synchronously than did control 

eggs. How embryos, enclosed within the fluid environment of an egg, emit 

such sounds is unknown. Vocalizations are also involved whenever the 

females excavate nests, open eggs, or transport hatchlings in their cavernous 

jaws. All these activities require a high degree of oral sensitivity and 

muscular control. Numerous domed sensory receptors on the palate and 

jaw margins (Figs. 3A and B) may facilitate these and other (e.g., courtship 

and feeding) behaviors (Ferguson, 1981b, 1982b). All available reports are 

phenomenologic, but it is obvious that further investigations of the neuralendocrine 

mechanisms underlying these complex behaviors are warranted, 

not merely for their intrinsic interest, but also for their potential 

contribution to our understanding of the evolution of parental care in 

birds and mammals.

356 

I 

REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS REPRODUCTIVE alOLOGY 357 

--:-""J;:;;iF'P:'" 

Fig. 3. Alligator mississippiellsis. (A) The palate of an adult photographed under oblique 

incident illumination to highlight the numerous low sensory papillae. (B) Histological section 

of the sensory papillae on the palate. Note the highly specialized sensory nerve ending and 

dermal Merkel cells. 292 x. 

F. Captive Breeding, Egg Collection, and 

Artificial Incubation 

Data pertaining to egg collection, artificial incubation, hatchling culture, 

and captive breeding are available for several species (Bustard, 1980c), but 

principally for Alligator mississippiensis (Joanen and McNease, 1971, 1973, 

1974a, b, 1975a, b, 1976, 1977, 1979a, b, 1980, 1981; Chabreck, 1977, 1978; 

Nichols et al., 1976), and Crocodylus niloticus (Pooley, 1969a, b, 1971; Blake, 

1974; Blake and Loveridge, 1975). Other reports describe laboratory maintenance 

of alligators (Coulson et al., 1973, Coulson and Hernandez, 1964) 

and give detailed information on the construction of egg incubators, controlled 

environmental chambers and suitable breeding enclosures (Joanen 

and McNease, 1974a, b, 1976, 1979a; Pooley, 1971). Successful reprOduction 

has now been achieved for many species (Chaffee, 1969; Del Toro, 

1969; Hunt, 1969, 1973, 1975; Legge, 1969; Yadav, 1969, 1979; David, 1970; 

Yangprapakorn et al., 1971; Joanen and McNease, 1971, 1975a, 1979a, 1980, 

1981; Honegger, 1971, 1975; King and Dobbs, 1975; Dunn, 1977, 1981; 

Whitaker, 1979; Whitaker and Basu, 1983; Bustard, 1980c, 1984). Thus far, 

artificial insemination has proved difficult (Joanen and McNease, 1973, 

1975b) and has only once been achieved, the major problem being induction 

of ovulation in the female (Cardeilhac et al., 1982). 

In most management (and embryological) programs, eggs are collected 

from wild nests and artificially incubated. Documentation of recommended 

collecting methods and their effect on embryonic viability is available 

for several species (Blake and Loveridge, 1975; Chabreck, 1977, 1978; 

Joanen and McNease, 1977, 1979a, 1980, 1981; Pooley, 1969a, b, 1971; 

Whitaker and Whitaker, 1976a). The exact time of collection and possible 

deleterious effects of inversion are particularly important; similar problems 

have been reported with respect to testudinian eggs (Ewert, 1979; Parmenter, 

1980; Blanck and Sawyer, 1981). 

At oviposition, the embryonic disk, covered by a thin layer of albumen, 

floats freely on top of the yolk. Its precise location depends upon the 

position of the egg in the nest. If the egg is moved during the first 24 hours 

after laying, the embryoniC disk moves to the new highest point of the yolk 

without detrimental effects. However, after 24 hours, the disk attaches to 

the inner aspect of the shell membrane, and the egg shows an opaque spot 

in this area (see Section III. B). The embryo remains attached, hence movement 

or turning may kill it. After inversion, attachment to the shell keeps 

the embryos from rising above the yolk maSSi they may be crushed or 

drowned by the superjacent heavy yolk. Moreover, in the first few days 

after laying, the embryo has not undergone torsion (Section VI) and so lies 

at apprOXimately right angles to the shell surface, whereas small blood 

vessels proliferate within both it and the extraembryonic membranes, 

which are also attached to the shell membrane. Any movement tends to 

shear the embryo off the embryonic disk and rupture these delicate blood 

vessels, thereby causing death. Thus eggs should be collected before the 

embryo attaches to the shell membrane (within 24 hours after laying) or 

much later, after the fourth week of incubation, by which time the embryo 

and extraembryonic membranes are less susceptible to damage. Similar 

considerations apply to turtles (Blanck and Sawyer, 1981). Always mark 

the top surface of the egg prior to removal and transport and incubate eggs 

in their original nest orientation or the nearest horizontal position. 

Other reasons for collecting eggs within 24 hours of laying include the

35B 

REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF' CROCODILIANS 


3551 

following: (1) Reduction of losses from predation, flooding, or physical 

damage to the nest (e.g., by another nesting crocodile); (2) control and 

optimization of the incubation environment; (3) control of the sex and 

possibly the future growth of hatchlings (Ferguson and Joanen, 1982, 

1983); (4) monitoring development by observing changes in eggshell banding 

(see Section III. B) and detecting (in order to discard) infertile, 

malformed, dead, or infected eggs; (5) acceleration or delay of development 

by altering the incubation temperature and synchronization of hatchling 

emergence; (6) correction of the orientation of eggs laid in a detrimental 

position. Thus embryos of eggs laid upright (i.e., with their long axes at 

right angles to the nest base) frequently die or are malformed; they develop 

normally if the eggs are reoriented so that the embryonic disk becomes 

positioned beneath the top center of the egg. 

The death of embryos following egg inversion may explain why viviparity 

never evolved in the Crocodilia or Testudines. During the evolution of 

viviparity, there is usually an intermediate stage in which the embryo 

develops inside an egg retained within the oviducts of the female. However, 

in taxa in which the embryo attaches to the shell membrane and 

turning or inversion result in embryonic death, this intermediate stage is 

unlikely to be achieved. The absence of crocodilian viviparity may also be 

related to increased dependence of embryos on eggshell calcium as compared 

to the very slight dependence of snakes and lizards (Packard et aI., 

1977), the absence of biological advantages for prolonged egg retention, 

and the evolution of nest guarding (Tinkle and Gibbons, 1977). 

Artificial incubation of the eggs of "hole nesters" has been effected by 

reburying the eggs in suitable sites (Pooley, 1969a, b, 1971; Blake and 

Loveridge, 1975; Bustard, 1980c). Earlier attempts at incubating the eggs of 

mound nesters, e.g., Alligator mississippiensis, involved placing them in 

buckets of nesting material maintained at the appropriate temperature and 

humidity (Reese, 1901a, 1931a; Table III). Currently eggs are incubated 

either in incubators or in environmental chambers (Joanen and McNease, 

1974a,b, 1976, 1977, 1979a, 1980, 1981). Tests relating to the incubation of 

alligator eggs, including stacked versus nonstacked, wild eggs versus eggs 

from captive breeding programs, oxygenated chambers versus nonoxygenated 

chambers, washed eggs versus nonwashed eggs, exposed eggs 

versus eggs covered with nesting material, and eggs set over water versus 

eggs set over dry concrete (Joanen and McNease, 1977) showed no appreciable 

effects, except for a 13% decrease in hatching success from stacked 

eggs, and a lower hatching rate (72%) for captive produced eggs than for 

those of wild animals (94%). About 15% of embryos from captive produced 

eggs died around hatching, and others had to be assisted from the shell 

(Joanen and McNease, 1975a, 1977). This may indicate weakness of the 

hatchling (compacted yolks are more common in this group) or abnormal 

toughness of the eggshell and its membranes. Decreased hatching success 

in farmed turtles may be associated with replacement of the normal arago- 

nite crystals of calcium carbonate by calcite crystals (Solomon and Baird, 

1979; Baird and Solomon, 1979). Although the eggshells of farmed crocodilians 

may be defective, it seems more likely that decreased fecundity and 

increased abnormality may result from maternal dietary deficiencies, e.g., 

that of vitamin E (Lance, 1982; Lance et a!., 1983; Elsey and Lance, 1983) or 

inappropriate husbandry (Joanen and McNease, 1981; Lance, 1984). 

Incubation of alligator eggs at temperatures ranging from 26° to 34°C, 

produces the best hatching success at 30° to 32°C (Joanen and McNease, 

1977, 1979a, 1980, 1981; Ferguson and Joanen, 1982, 1983). Hatchlings from 

eggs incubated below 28°C often twitch, have balance difficulties and swim 

with their heads constantly under water; eventually they drown. Eggs of 

Crocodylus novaeguineae incubated at 23°, 26°, and 38°C (normal approximately 

32°C) have a low hatching success (Bustard, 1969, 1971). All survivors 

of the high temperature group showed tail and other abnormalities 

and had to be helped from their shell. Hatchlings of C. palustris recovered 

from overheated nests also showed tail abnormalities (Whitaker and 

Whitaker, 1976a,b). 

Crocodilian eggs are susceptible to changes in humidity; at low levels 

the shell membrane dries out and shrivels, thereby killing the embryo 

(McIlhenny, 1935; Deraniyagala, 1939; Pooley, 1962, 1969a, b, 1971; Joanen, 

1969; Staton and Dixon, 1977). The optimum relative humidities for incubating 

Alligator mississippiensis eggs is 90 to 92% (Joanen and McNease, 

1977, 1979a; Chabreck, 1975). Abnormal air spaces form in alligator eggs 

incubated at suboptimal humidities (Ferguson and Joanen, 1983); large air 

spaces are lethal. Intact eggs of Crocodylus acutus are resistant to desiccation, 

losing water at a rate comparable to the eggs of birds (Rahn and Ar, 

1974; Rahn et aI., 1979; Ar and Rahn, 1980; Diamond, 1982), but at a rate 

much lower than that of the leathery-shelled eggs of Iguana iguana (Rand, 

1968). Removal of a 3 x 3 cm piece of shell (leaving the shell membrane 

intact) from the eggs of C. acutus greatly increases the desiccation rate 

(Rand, 1968). The eggs of C. novaeguineae apparently could either lose or 

gain up to 25% water with no adverse effects on the embryos (Bustard, 

1971). The soft-shelled eggs of many squamates and turtles can adsorb 

(and then lose) water up to 300% of their initial weight (Bustard, 1971; 

Packard et aI., 1977); unlike the eggs of crocodilians (and certain other 

turtles) they lack large quantities of hydrophilic albumen, which are likely 

to reduce the rate of water loss under adverse conditions. 

Bustard (1971) has suggested that water uptake is unessential in crocodilian 

eggs, but represents an insurance against lethal levels of water stress 

caused by possible adverse environmental conditions later in development. 

However, Packard et al. (1977) reported that water uptake was critical 

for normal reptilian embryonic development; prevention of water uptake, 

particularly in the early stages of incubation, leads to high rates of 

mortality and developmental abnormalities. Indeed, Tracy et a!. (1978) 

suggested that the only major difference between the eggs of birds and



361 

those of crocodilians (and certain turtles) is the site where they are laid; 

avian nests favor water loss and crocodilian nests favor water gain. However, 

these authors assume that cracks in the alligator eggshell under natural 

nesting conditions indicate water uptake and egg swelling. Alternative 

explanations for such cracks are enlargement and movement of the embryo 

and weakening of the shell to facilitate hatching (Ferguson, 1981c, 1982a). 

Indeed, Lutz and Dunbar Cooper (1982, 1984) reported a net water loss 

from eggs of Crocodyllis aClltlls; the birth weights of hatchlings were 0.64 ± 

0.01 those of the initial egg masses, a value almost identical to that of many 

birds (0.65), (Rahn, 1982; Diamond, 1982). During incubation of avian 

eggs, dry matter is metabolized and metabolic water produced, but water 

concentration is maintained by losing a fixed portion of the water (Ar and 

Rahn, 1980). The fractional loss (F = total weight loss/initial weight) for 

eggs of 81 species of birds was remarkably constant (F = 0.150 ± 0.02580) 

and similar to that of C. aClltlis (F = 0.154) (Lutz and Dunbar Cooper, 1982, 

1984). Preliminary data therefore indicate that crocodilian eggs are structurally 

(Ferguson, 1982a) and functionally (Lutz et al., 1980; Lutz and Dunbar 

Cooper, 1984) similar to avian eggs but dissimilar to those of squamates 

and some testudines (Packard et al., 1977). Their water vapor conductance 

is treated below (Section ULE). 

The water relations of crocodilian eggs require further investigations 

both in the wild and experimentally. In some turtles, the size of hatchlings 

(and hence their survivorship) and the length of egg incubation are both 

related to the hydric environments of egg incubation (Packard et al., 1981, 

1982, 1983). Moreover, eggs of Chrysemys picta exhibit normal embryonic 

metabolism until the pool of egg water is reduced to a threshold level, at 

which time death occurs (in unfavorable hydric conditions) or hatching 

ensues (in favorable conditions), depending on the stage of embryonic 

development (Packard et al., 1983). Thus the water potential of eggs may 

provide the cue for hatching and an explanation of late embryonic deaths 

(often during the week of expected hatching). Moreover, preliminary data 

indicate that the hydric environment of Chrysemys picta eggs influences 

sexual differentiation (Gutzke and Paukstis, 1983); this may also be true for 

crocodilians. 

31 

G. Hatching 

The mechanics of hatching have been described for Alligator mississippiensis 

(McIlhenny, 1934, 1935; ]oanen, 1969), CrocodylliS niloticliS (Voeltzkow, 

1891, 1892, 1893, 1899; Pooley, 1962, 1969a), C. POroSliS (Deraniyagala, 1936, 

1939; Webb, 1977a), and C. pailistris (Deraniyagala, 1939); they appear to be 

similar in all crocodilians (Neill, 1971). The growing and moving embryo 

strains the eggshell, which has already been weakened by mobilization of 

calcium and extrinsic degradation (see Section lII.C), so that it eventually 

cracks. In A. mississippiensis, longitudinal cracks develop about the seventh 

Fig. 4. 

Alligator mississippiensis. Sequence of photographs illustrati]l~ hatching. 

week of incubation (Joanen, 1969), diagonal cracks begin several days later, 

and shell flaking begins prior to hatching. The embryo is thuS surrounded 

by extraembryonic and shell membranes. These membranes are later punc 

T 

tured by the sharp "caruncle" located at the tip of the snout (Section vn. , 

Figs. 26A to Hand 36A to D), thereby pippipg the egg (Fig. 4). ) 8za 

Because crocodilian eggs normally lack an airspace (Ferguson, 19 , 

It ~

3SE! 

REPRODUCTIVE alOLOGY ANO EMBRYOLOGY OF CROCODILIANS 

there is no internal pipping, as seen in birds (Rahn et a1., 1979). Hence, 

crocodilian hatchlings are unlikely to breathe until the shell and membranes 

are broken and the flUids are drained from around the head. The 

time interval between this event and emergence from the egg is relatively 

short (ranging from a few minutes to 24 hours), so that the transition from 

chorioallantoic to pulmonary respiration must be rapid, as in mammals 

and megapode birds (Seymour and Ackerman, 1980), but it is contrary to 

the slow transition in other birds (Rahn et a1., 1979). Details of this transition 

(e.g., the extent of chorioallantoic perfusion during the time between 

first breathing and hatching) are unknown but of considerable interest. 

The problem is even more intriguing when one recalls that for many hours 

prior to pipping embryos vocalize in response to certain stimuli (see Section 

11.E); this suggests that the lungs are functional during this period. As 

alligator eggs progressively lose water during incubation, an air bubble 

forms beneath the shell membrane and this moves around in the albumen, 

so as to always lie beneath the uppermost surface of the egg (Ferguson, 

1982a): similar events OCCur in megapode eggs (Seymour and Ackerman, 

1980). Perhaps this air is used prior to pipping (analogous to the internal 

pipping of avian embryos). This area is significant because late embryonic 

death is common (during both natural and artificial incubation) and could 

be caused by improper hydric conditions. Thus, high humidity may be 

required during the early stages of incubation, but lower humidity may be 

reqUired during the later stages to facilitate water loss and the formation of 

intra-egg air. This in turn may enable internal breathing, the maturation of 

the respiratory system, and provide the extra oxygen required for pipping. 

Indeed, Vocalization may represent a signal that the embryo is mature 

enough to survive outside the egg, and it remains to be determined 

whether it occurs in eggs incubated under conditions of continuous high 

humidity. The changing hydric conditions are likely to be most critical in 

species with long incubation periods (e.g., Crocodylus porosus). 

During the hatching process, the embryo first widens the slit in the shell 

membrane (frequently by pushing its snout into the slit and then opening 

its jaws). Second, it pUShes out the entire head and neck; after a couple of 

minutes, one quick forceful thrust forces the whole bOdy out of the egg 

(Fig. 4). The claws may be used in the hatching process. The ruptured 

extraembryonic membranes progressively detach from the shell membrane 

but remain attached to the umbilical region of the hatchling for about a day; 

they eventually shrivel, dry up, and break (Fig. 21). Hatchlings are COvered 

with slime, presumably derived from the ruptured allantois, and they 

smell of ammonia. This slime dries after about 24 hours. At the time of 

hatching, the abdomen is distended by absorbed yolk and the umbilical 

car is evident (Fig. 21). The yolk serves as a food supply for a few weeks 

ntil the hatchlings begin feeding so that the abdomen becomes less dis 

~ended and the umbilicus closes. In hatchlings kept below 210C, the umilicus 

does not heal correctly, yolk is not utilized, and the animals usually 

e (King and Dobbs, 1975). 


363 

The precise hatching trigger remains unknown. Mechanical stimuli and 

vocalizations may playa role (see Section II.E). Growth and movement of 

the embryo may burst the shell. Increasing levels of waste products, decreasing 

egg water, and the progressive inadequacy of gas exchange may 

also stimulate hatching. Temperature seems to be important; field reports 

note that an inadvertent rise in egg temperature stimulates premature 

hatching, and, conversely, decreases in egg temperature delay hatching 

(McIlhenny, 1934, 1935; Deraniyagala, 1936, 1939; Pooley, 1962, 1969a, 

1971). Hatchlings are sufficiently well developed to be viable if they hatch 

several days prematurely (McIlhenny, 1934, 1935) so that completion of 

development is not the final trigger for hatching. Lengths and weights of 

the hatchlings from different species are summarized in Table III. 

III. 


A. General 

Most crocodilian eggs are elliptical with approximately equivalent ends 

(Figs. 5A to 1). They vary considerably in size and shape, even within 

species (Tables III and IV). Normally there is little variation in size and 

shape within an individual clutch. However, occasional large, infertile, 

double-yolked eggs or very small eggs are seen; these abnormal eggs are 

usually laid at the beginning and end of oviposition and are common in 

clutches laid by young females (Ferguson and Joanen, 1983). Ferguson 

(1982a) described the structure and composition of the eggshell and shell 

membranes of Alligator mississippiensis and provided a comprehensive review 

of the literature. 

B. Egg Banding 

The gradual development of a transverse white band across fertile crocodilian 

eggs has been known for nearly a century (Clarke, 1888a,b, 1891), 

and reports are now available for several species (Ferguson, 1982a; Reese, 

1908, 1912, 1915a, 1931a; McIlhenny, 1935; Deraniyagala, 1936, 1939; Webb 

et aI., 1977, 1983a, b, e; Webb, 1977a; Beck, 1978; Hara and Kikuchi, 1978; 

Deitz and Hines, 1980; Tryon, 1980). 

Within 24 hours of egg laying, the eggs of Alligator mississippiensis show 

a small opaque, chalky white, oval patch on their top surface (Figs. 5A and 

6). The small embryo has attached itself to the innermost surface of the 

shell membrane immediately below this opaque patch. At this and all 

subsequent stages, the white color is associated with changes both of the 

eggshell and shell membrane (which itself appears chalky white in this 

area). Initially the oval opaque patch expands in width around the shell, 

extending approximately half-way around the shell on days 3 to 4, threequarters 

way around the shell on days 5 and 6, and completely around the

1 

microorganisms in the pores of the eggshell and on the surface of the shell membrane. (D) A 

completely opaque, cracked, 58-day egg. (E) Attachment of the embryo and chorioallantois to 

the superior inner surface of the shell membrane in an I8-day egg. Note the two poles of 

albumen (A) at the ends of the egg. The yolk has been removed. (F) Expansion of the 

chorioallantois and the regression of albumen seen in the superior inner surface of a 25-day 

egg. (G) Lateral view. Opaque band in a transilluminated 7-day egg. The band appears black 

by this technique and appears longer toward the top (right) than the bottom (left) of the egg. 

Fig. 5. Alligator mississippiensis. (A) Opaque spot on the eggshell and shell membrane of an (H) View of the opaque band in a I2-day egg. (I) View of the opaque band in a 20-day egg. (J) 

egg 24 hours after deposition. (8) Opaque band on a ten-day eggshell. (C) Opaque band in the 

View of a 40-day egg, which is almost completely opaque. 

eggshell (partIy removed) and shell membrane of a 20-day egg. Note the specks of debris and 

3615

...,... 


o 8 

TO T1 T3 T5 

T7 87 T30 T52 

Fig. 6. Alligator mississippiensis. Diagram depicting the development of the opaque band in 

eggs viewed in transmitted light. T, top of egg; B, bottom of egg; numbers, days after the time 

the egg was laid. 

shell after day 7 (Figs. SA to C, G to J, and 6). This extension in width is not 

matched by a uniform expansion in length, so that the opaque band tapers 

from a maximum length at the top surface of the shell to a minimum length 

on its bottom surface (Figs. 5 and 6). The embryo always lies beneath the 

upper expanded part of the fusiform opaque ring (Figs. 5B, E, F, G-J, and 

6). The opaque band then expands rapidly in length so that it extends over 

approximately 60% of the top surface and 50% of the lower surface of the 

shell between days 8 and 30 (Figs. 5B to 0, G to J, and 6). The top part of 

the band reaches the ends of the egg around day 40 and the bottom part 

around day 50; thereafter the egg is completely opaque and so appears 

unbanded (Figs. 50, J, and 6). The opaque part of the egg never transmits 

light as well as the adjacent translucent regions and is clearly demonstrated 

as a dark zone if the eggs are transiIIuminated (Figs. 5G to J, and 6). 

Eggshell banding represents a useful, external indicator for estimating 

the age of alligator eggs (Ferguson, 1982a) up to approximately day 50; 

subsequently the eggs are completely opaque. The normal expansion of 

the opaque band may be retarded in eggs containing malformed or developmentally 

retarded embryos. Usually, the expansion of the "minimum 

band length" at the bottom of the egg is the most sensitive indicator 

(Ferguson, 1982a, b). Eggs containing embryos that have died but 

that are not infected show an arrest of band development at the stage of 

embryonic death. This may be followed by regression of the band as the 

embryo autolyzes. Infected eggs have the opaque bands vaguely defined; 

opaque blotchy patches may appear and disappear all over the eggshell, 

usually in an erratic fashion. Infertile eggs never become banded and never 

become infected (unless the eggshell or eggshell membrane is damaged or 


367 

malformed), but remain uniformly translucent white (egg contents yellow) 

throughout incubation. They can easily be distinguished from normal, 

completely opaque, post day 50 eggs by transillumination (Figs. 5G to J, 

and 6). Eggs laid at advanced stages of embryonic development, for example, 

by stressed females (see Section II.C), are either banded at the time 

of laying or become so very rapidly. Band expansion is also more rapid, but 

the embryos usually die, at which time the formation ceases. Failure to 

distinguish between unbanded infertile eggs and either those in which the 

embryo has died early and become autolyzed or post day 50 normal unbanded 

eggs has given rise to erroneous accounts of the relationship between 

banding and fertility (Deitz and Hines, 1980; Tryon, 1980). 

That the appearance and regular development of the opaque band indicate 

normal, healthy development of the embryo is of value not only in 

aging eggs but also in identifying abnormal development and monitoring 

teratogenic experiments (Ferguson, 1981a, 1982b). The relationship between 

the development of this band and eggshell hydration, albumen 

metabolism, the formation of the extraembryonic membranes, structural 

changes in the eggshell and changes in the composition of the shell membrane 

is discussed in Sections III.C through E and IV. 

C. Structure 

1. GENERAL 

The combined thickness of the eggshell and shell membrane of Alligator 

mississippiensis is approximately 0.5 to 1.0 mm and consists of five layers 

(Fig. 7). From the surface inwards, there is an outer, densely calcified layer 

(100-200-f.Lm thick), a honeycomb layer (300-400-f.Lm thick), an organic 

layer (8-12-f.Lm thick), a mammillary layer (20-29-f.Lm thick), and the shell 

membrane (150-250-f.Lm thick). Pores run from the egg surface through the 

calcified layers and end in the shell membrane (Fig. 7). The outer openings 

of the pores become modified and numerous erosion craters develop as 

incubation progresses (Figs. 7 and 8). The different layers of the eggshell 

consist of varying amounts of calcite crystals and organic matrix. The 

freshly laid egg is coated with slimy oviducal secretions, which dry and 

then disappear. 

2. OUTER DENSELY CALCIFIED LAYER 

At low magnification (Fig. 9A), the egg surface appears smooth, but at 

higher magnification (Fig. 9B), the granular nature of the numerous calcite 

crystals is evident. There is no detectable organic matrix between the 

rhombohedral crystals in this layer; their form, size, and orientation are 

shown in Figs. 8 and 9, and further details are given in Ferguson (1981c, 

1982a). The calcite crystals are stacked on their ends (or faces) in this layer.

368 REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCOOILIANS 

OPAQUE 

NON 

EC 

1r1:\1 

lltJ] ®a c __ b - 

~ 

00-400f"= 

Fig. 7. Alligator mississippiensis. Diagram of the structure of the eggshell in both the opaque 

and nonopaque zones as seen in cross section. Ee, Erosion crater; ESM, shell membrane; H, 

honeycomb layer (crystals oriented with their c-axis horizontal and containing numerous 

vesicular holes); M, mammillary layer (the mammillary knobs are common in the opaque 

zone, but not so frequent in the nonopaque zone; the eggshell membrane fibers of the latter 

attach directly to a layer of large calcite crystals rather than the mammillary knobs); 0, organic 

layer (a layer immediately superficial to the mammillae containing a higher percentage of 

organic matrix); ODe, outer densely calcified layer (crystals oriented. with their c-axis at right 

angles to the shell surface); P, small pore in the shell membrane; PO, pore (more frequent in 

opaque zone); PP, pore plug (falling out). (From Ferguson 1981c, 1982a.) 

3. HONEYCOMB LAYER 

The honeycomb layer is porous and contains a high percentage of fibrous 

organic matrix (Figs. 9C and E). When compared to those in the outer 

densely calcified layer, the different (horizontal) stacking of the calcite 

crystals in this layer is important in the formation of erosion craters (see 

Figs. 7 to 9 and text below). The numerous matrix-lined holes make the 

honeycomb layer similar to the palisade layer of the eggshells of tropical 

birds (Schmidt, 1957, 1964; Tyler, 1969; Becking, 1975). These holes interconnect 

extensively with each other and with the intermammillary air 

spaces (Fig. 7). 

II iii .OOC.... IIbiId 

CD ® CD CD ® ® 

SD 

.t, 

SD 

ODC 

® CD 

H 

D C C 

C 

·6' . 

o~ dill 

-::) ~~L. c_~ 

H 

0) 

J FillA II 

'~.. 

'r t 

: lB; 

.,........... 

i 

calcified layer (0), the honeycomb layer (H) beneath it, and the mammillae. (B) Higher power 

SEM of the boxed edge seen in (A). Note the granular appearance of the densely packed 

vertical calcite crystals (the diameter of which is 0.5-1 f.Lm) and the absence of any organic 

matrix. (C) Erosion crater in the nonopaque zone of an IS-day egg. Note the stepped concentric 

outline of dissolved crystal layers in the outer densely calcified zone, and the openings of 

numerous vesicular holes in the exposed porous honeycomb layer at the crater base. (D) Pore 

in the central opaque zone of a 40-day eggshell. Note the crater-like concentric stepping of the 

outer densely calcified layers at the pore orifice, remnants of the pore plug and microorganisms 

(bacterial cocci, rods and filamentous fungae). Outer pore diameter approx. 550 f.Lm, 

inner pore diameter approx. 100 f.Lm. (E) Fractured edge of a 4-day eggshell. Note the shell 

membrane (E), mammillary layer (M), and honeycomb layer (H). (F) Innermost (egg content 

side) layer of the shell membrane depicting the numerous blebs caused by the projecting 

mammillary knobs of the mammillary layer. (G) View of the transversely fractured organic 

layer in the nonopaque zone of a 2-day egg. Note the smooth fibers of organic matrix interspersed 

with numerous small calcite crystals. (H) View of the transversely fractured organic 

layer in the opaque central zone of a 56-day egg. Note the scarcity of small calcite crystals (see 

Fig. G) and the blebbing on the fibers of organic matrix. (I) High power view of the fibers in 

the shell membrane of a 3D-day egg. Note the interweaving of the fibers, the numerous blebs 

on the surfaces of the fibers, and the porous nature of the membrane. 

Fig. 9. Alligator mississippiensis. Scanning electron micrographs (SEM). (A) Surface and fractured 

edge of an IS-day eggshell. Note the smooth shell surface (5), the outer, densely

372 

REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCODILIANS 

ment for the calcite crystals, but may be the templates for crystal growth. 

Beginning about the third week of incubation, the number of small calcite 

crystals in the organic layer beneath the opaque band decreases (compare 

Fig. 9G with 9H), and this process continues as the band expands. It 

appears that these crystals, in preference to the larger units in the mammillary 

layer, are mobilized for embryonic calcification. This process steadily 

weakens the calcified shell, and horizontal separation usually begins a few 

days before hatching, so that only the shell membrane and parts of the 

mammillary and organic layers surround the embryo, with the rest of the 

shell either cleaved off or intact (but weakened). 

5. MAMMILLARY LAYER 

The mammillary layer (Figs. 7, 9E, and 9F) comprises numerous cones 

("mammillary knobs") with their bases on the organic layer and their tips 

continuous with the shell membrane at the core of the knobs. The calcite 

crystals of the mammillary knobs are larger than those elsewhere and are 

arranged into regular prism-like subunits, which are clearly evident in 

radial sections of the shell (Fig. 9E). In tangential section, the mammillae 

appear as irregular hexagons. Pores (Fig. 7) occur wherever the irregular 

faces of one or more mammillae fail to contact each other in the horizontal 

plane as has been described for the avian egg (Tullett, 1975). Pores are most 

numerous in the central opaque region of the eggshell. Mammillary outlines 

are much easier to distinguish in the central opaque region than in the 

terminal nonopaque ends, where the shell membrane has flat areas of 

attachment. Shell membrane fibers attach to the central organic core of the 

mammillary knobs. 

6. SHELL MEMBRANE 

The shell membrane consists of randomly oriented fibers (Fig. 7) that interweave 

extensively in a criss-cross pattern, enclosing air spaces (Fig. 91). 

The fibers are approximately 2 j..Lm in diameter and show the characteristic 

surface blebbing of glycosaminoglycans (Fig. 91). The innermost surface of 

the membrane has a thin, amorphous, nonfibrous, surface-limiting layer of 

unknown composition (Fig. 9F). Pores in the latter (Figs. 7 and 9F) are the 

innermost end of a complex system of spaces in the more superficial layers, 

which connect the contents of the egg (e.g., the chorioallantoic membrane) 

to the external environment and are probably important in respiration and 

water regulation. Contrary to some reports (Bigalke, 1931; Pooley, 1962, 

1969a; Bellairs, 1969; Guibe, 1970), crocodilian eggs, unlike those of birds, 

do not have an air space (Reese, 1915a, 1931a; McIlhenny, 1935; Packard et 

al., 1977; Ferguson, 1982a). Abnormal air spaces may form in dehydrated 

eggs where shrinkage of the shell membrane cleaves it from the shell 

through the organic layer (Ferguson, 1982a,b; Ferguson and Joanen, 1983). 

(Also see Section I.G.) 


7. EROSION CRATERS 

373 

Throughout incubation, the egg surface develops erosion craters, which 

appear as stepped concentric rings each corresponding to one crystal layer 

of the outer densely calcified zone (Figs. 7, 8, and 9C). They are caused by 

crystal dissolution, possibly caused by acidic metabolites of nesting microorganisms 

and/or by the hydration of expired carbon dioxide. These 

craters appear to be present in the eggshells of numerous species (Ferguson, 

unpublished data), including fossil crocodilians (Hirsch, 1984). Details 

on their mode of formation are summarized in Fig. 8 (Ferguson, 1981c, 

1982a). Their development confers possible advantages to the embryo in 

terms of enhanced exchanges of respiratory gases and water vapor across 

the shell as well as the facilitation of hatching through eggshell weakening. 

The occurrence of extrinsic degradation is of importance to those incubating 

eggs in embryological, farming, management, or conservation 

projects. Normally eggs should be left in their natural dirty state and incubated 

completely surrounded by damp nesting media in order to facilitate 

the production of bacterial acids. However, clean eggs can be successfully 

incubated without nesting media by maintaining very high 

humidities (circa 90-100%). The high humidity ensures that all of the expired 

carbon dioxide produces carbonic acid, which degrades the shell. 

Presumably, this hydration, possibly in combination with mineral acids 

from the moist nest sand, forms the erosion craters in hole-nesting crocodilians 

(Ferguson, 1981c). The relative importance of humidity, or acids produced 

by microbial fermentation of nesting vegetation, in normal degradation 

of the eggs of mound nesters has not been adequately determined 

(Ferguson, 1981c, 1982a). However, embryos from eggs incubated at 

humidities below 90% without nesting media, frequently die (probably 

from asphyxia or the toxic effects of waste products), whereas the remaining 

living hatchlings usually have to be manually helped out of the tough 

eggshell (Ferguson, 1981c, 1982a). Further investigations are also required 

into which specific microorganisms are involved, whether they derive any 

benefit (e.g., calcium) from the process, the changing pH of the eggshell 

surface in the nest, which acids are involved, and how the system evolved. 

In other systems, various fungi penetrate and degrade calcareous substances 

such as mollusk shells (Kohlmeyer, 1969, 1972). Although thermophilic 

fungi have been isolated from alligator nests (Tansey, 1973), their 

role in eggshell degradation is unknown. 

8. PORES 

Single, unbranched pores run vertically upwards from spaces between the 

mammillae to end at the shell surface (Figs. 7 and 90). They are more 

numerous in the central opaque region of the eggshell, and in freshly laid 

eggs, their openings are capped by an organic plug (Figs. 7 and 90). 

During incubation, the openings of the pores are widened by extrinsic


THE EGGSHELL ANO SHELL MEMBRANES 

375 

acidic dissolution (Figs. 7, 8, and 90), and the plug becomes dislodged. 

Presumably, this widening of the pore orifices facilitates respiratory and 

metabolic exchanges; it also progressively weakens the eggshell, as longitudinal 

cracks often pass through the pores and the erosion craters. 

8. CONCLUDING COMMENTS 

The complex structure of the eggshell reflects its adaptation to crocodilian 

nesting biology. At laying, the eggs are strong and not very porous; 

they are thus protected from physical damage at the time when they are 

deposited one upon another and whenever the mother treads nesting material 

on top of them, as well as from dehydration during early development. 

The complex sequence of changes in the inorganic constituents 

weakens the shell and makes it more porous throughout incubation, so 

that the prehatchling has merely to slit the organic materials with its caruncle. 

Antimicrobial properties of shell or albumen components, known for 

birds (Board and Fuller, 1974) and turtles (Movchan, 1964, 1966, 1967) have 

not been reported in crocodilian eggs but may be related to the development 

of erosion craters and the filter bed arrangement of the porous shell 

and shell membrane. Data on such properties would be valuable, not only 

for their intrinsic interest, but also in view of the practical problems associated 

with artificial egg incubation. 

C. Chemical Composition 

Relatively little is known about the chemical composition of crocodilian 

eggs. The 6 g of calcium carbonate in the eggs is 99% calcite and less than 

1% aragonite (Erben, 1970; Jenkins, 1975), a ratio far closer to that reported 

for birds (Romanoff and Romanoff, 1949; Simkiss, 1967; Rol'nick, 1970) 

than the reported predominance of aragonite in the eggshells of turtles 

(Young, 1950; Erben, 1970; Packard et al., 1977; Solomon and Baird, 1979). 

Eggshells of Crocodylus novaeguineae contain 82.6% calcium carbonate, 

2.82% magnesium, 0.37% phosphorus, and 3.36% organic protein (Jenkins, 

1975). The total protein content of the eggshell is approximately twice 

that for the domestic fowl (Jenkins, 1975). The eggshell of Alligator mississippiensis 

also contains calcium, magnesium, and phosphorus, as well as 

traces of copper, silicon, sodium, aluminum, iron, zinc, and manganese 

(Ferguson, 1982a). 

The thickness of the shell membranes is reflected in the fact that they 

comprise 21.39% of the total weight of dry shell and membrane (Jenkins, 

1975), whereas they only comprise 0.3% in the domestic fowl (Romanoff 

and Romanoff, 1949). The shell membrane acts as an intermediary calcium 

store between that of the eggshell and the calcium in the blood of the 

chorioallantoic vessels (Ferguson, 1982a), a factor which contributes to its 

chalky white coloring beneath the opaque bands. Whether the yolk, albu- 

TABLE V 

Calcium Contents of the Egg and Hatchlings of Various Species" 

Snake Crocodile Turtle Bird 

(Vipera (Crocodylus (Dermocllelys (Gallus 

berus) nouaeguineae) coriacea) domesticus) 

Calcium in 25 124 34 23 

egg contents, mg 

Calcium in 12-16 280 138 120 

hatchlings, mg 

lncrease b 0.8x 2.4x 4x 5.2x 

"From Jenkins, 1975. 

blndex of a mount of shell calcium used by embryo. 

men and extraembryonic membranes also store calcium is unknown for 

crocodilians, but such storage does occur in birds (Simkiss, 1967, 1980). 

The mechanisms of eggshell decalcification and calcium transport are 

unknown; however, carbonic acid, formed from respiratory carbon dioxide, 

apparently attacks the shell of birds (Buckner et al., 1925; Romanoff 

and Romanoff, 1949; Simkiss, 1967, 1980; Rol'nick, 1970). This mechanism 

is in accord with the assumed respiratory function of the opaque zone in 

crocodilian eggs. The levels of calcium in egg contents and hatchlings of 

Crocodylus novaeguineae given in Table V are uncertain due to poor preservation 

and large numbers of infertile eggs (Jenkins, 1975). Because hatchling 

crocodiles contain 2.4 times more calcium than the egg contents, the 

additional calcium must derive from the eggshell. Apparently, the yolk and 

albumen of crocodilians contain more calcium than those of turtles (which 

obtain four times as much calcium from the shell as from the egg contents) 

or birds (which obtain five times as much calcium from the shell), but much 

less than those of snakes, which obtain no calcium from the eggshell (Jenkins 

and Simkiss, 1968). This decreased dependence of crocodilian embryos 

on eggshell calcium permits experimental embryological studies using 

shell-less and semi-sheIl-less culture of alligator embryos; these survive 

to later developmental stages than their avian counterparts (Ferguson, 

1981a, 1982b, 1984a; Fig. 38). 

Available reports on the biochemical constituents indicate general agreement 

on the occurrence of various mucopolysaccharides (Neumeister, 

1895; Simkiss and Tyler, 1959; Kriesten, 1975), but fibrous proteins are less 

well-known. A survey of the amino-acid composition of the material of 

the shell and shell membranes indicated that the former contained sequences 

characteristic of collagen (in contrast to turtles), whereas the latter 

contained sequences reminiscent of a keratin-like protein (Kramptiz et al., 

1974), similar to that suspected for birds and turtles (Romanoff and

37B 

REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCOOILIANB 

THE EGG CONTENTB ANO EXTRAEMBRYONIC MEMBRANEB 

377 

Romanoff, 1949; Young, 1950; Simkiss and Tyler, 1959; Terepka, 1963a, b; 

Masshoff and Stolpmann, 1961; Simons and Wiertz, 1963; Simkiss, 1967; R. 

Bellairs and Boyde, 1969; Board and Fuller, 1974; Kriesten, 1975). 

The kinetics of calcium metabolism of gravid females remains to be 

studied. Whereas the calcium might be mobilized from specialized medullary 

limb bones as in birds (Sturkie, 1965; Simkiss, 1967), from the skeleton 

in general as in turtles (Simkiss, 1967), from specialized stores as in some 

lizards (Simkiss, 1967), and from the diet (Romanoff and Romanoff, 1949), 

it appears to be mobilized from the osteoderms (Ferguson, 1984a, ms) and 

from the diet. There is no medullary bone. 

E. WBter and Gas Conductance and Embryonic 

Metabolism 

We know far less about the water and gas conductance of crocodilian eggs 

(Harrison et al., 1978; Packard et al., 1979; Lutz et al., 1980; Lutz and 

Dunbar Cooper, 1982, 1983), than about those of turtles (Packard et al., 

1977, 1981, 1983; Ewert, 1979; Ackerman, 1980, 1981a, b; Seymour and 

Ackerman, 1980) and birds (Romanoff, 1967; Rol'nick, 1970; Rahn et al., 

1979; Simkiss, 1980; Diamond, 1982). However, these parameters are im 

, portant for embryonic survivorship, metabolic rate, incubation period, sex 

determination (see Section VII.O), and for speculations regarding the evolutionary 

constraints on clutch size and nesting ecology (Seymour, 1979; 

Seymour and Ackerman, 1980). The gas-water relations are also dealt with 

under Section II. F. 

The vapor conductance of the eggs of Crocodylus acutus is 21.0 mg H 0 

1 2 

day-l torr- H 20. This value is approximately two times that for an avian 

egg of equivalent weight (Lutz et al., 1980), even though the shell of the 

crocodilian egg is two times and the shell membrane ten times as thick. The 

low resistance of crocodilian eggs probably represents an adaptation to an 

environment of high humidity (Lutz et al., 1980; Seymour and Ackerman, 

1980; Lutz and Dunbar Cooper, 1982, 1983). At 70% relative humidity, the 

eggsheIl contains 0.92% of its wet weight as water, and the membrane 

contains 22.2%; at saturation, these values increase to 6.8 and 58.6%, respectively 

(Lutz et al., 1980). Permeability to oxygen drops tenfold as 

humidity rises from 70 to 100%. The oxygen diffusion coefficients (K0 ) 

2 

vary between 1.2 x 10- 6 cm 3 STP sec- 1 cm -2 torr- 1 at desiccation to 2.3 x 

7 3 

1 

10- cm STP sec- 1 cm -2 torr- at saturation (Lutz et al., 1980). Apparently 

drying of the sheIl during the expansion of the opaque band facilitates gas 

exchange (Ferguson, 1982a). 

Metabolic rate is maximal during organogenesis at which time it is directly 

proportional to the incubation temperature (G. Grigg, personal communication). 

During the first two months of development, alligator embryos 

produce about 15.3 mg waste nitrogen (0.35 mg per g of embryo 

formed). Throughout development, the proportions are approximately 

46.5% ammonium salts, 46.2% urea, and 7.3% uric acid (Clark et al., 1957). 

The yolk sac functions as an excretory organ before the allantois develops; 

thereafter the urea shifts to the latter. Protein is the preferred energy 

source in early development (Clark et al., 1957). 

IV. 

THE EGG CONTENTS AND EXTRA·EMBRYONIC 

MEMBRANES 

For nearly a century, it has been known that crocodilian albumen is more 

gelatinous than that of birds (Clarke, 1888a, b, 1891; Voeltzkow, 1892, 1899, 

1901). In Alligator mississippiensis, the albumen is either clear and transparent 

or slightly opaque (Clarke, 1888a, b, 1891; Reese, 1915a, 1931a; 

McIlhenny, 1935; Ferguson, 1982a). That of Crocodylus niloticus, C. porosus 

and Paleosuchus palpebrosus has been reported to be green (Voeltzkow, 1892, 

1899; Reese, 1915a, 1931a; Bigalke, 1931; Deraniyagala, 1936, 1939; Wettstein, 

1954), but these observations may have dealt with abnormal (infected) 

eggs. Initially, the albumen completely surrounds the centrally 

positioned yolk, but, as the volume of albumen decreases during development, 

it is increasingly isolated at the poles of the egg (Fig. 10). 

Throughout incubation, as water is assimilated in the yolk sac and embryonic 

body (Agassiz, 1857), the viscosity of the albumen increases so that it 

attains a rubbery consistency. Crocodilian eggs lack chalazae (Clarke, 

1888a, b, 1891; Voeltzkow, 1899; Reese, 1915a; Deraniyagala, 1939; Guibe, 

1970m; Ferguson, 1982a). The detailed chemical constitution of crocodilian 

albumen is unknown, as is its precise fate during development, although 

such information is available for birds (Romanoff and Romanoff, 1949; 

Romanoff, 1967; Rol'nick, 1970). Recently a protease inhibitor, closely resembling 

both the alpha-2 macroglobulin of mammalian serum and avian 

ovomacroglobulin, has been isolated from the albumen of C. rhombijer, C. 

siamensis, and Caiman crocodilus apaporiensis (Ikai et al., 1983). 

The spherical yolk, presumably held in position by the viscous albumen, 

is pale amber/yellow in color and so large that it almost touches the shell 

membrane at the midline (Clarke, 1888a, b, 1891; Reese, 1915a). In the 

freshly laid egg, the yolk is very fluid; it becomes increasingly viscous as 

development proceeds. It is enclosed within a yolk sac in which numerous 

blood vessels develop (Fig. 10). The composition of crocodilian yolk is 

unknown, but it can accumulate organochlorine residues (Hall et al., 1979). 

The yolk is the major nutritional source for the embryo and hatchling, and 

it probably consists of spherical droplets of lipoprotein as does avian yolk 

(Romanoff and Romanoff, 1949; Romanoff, 1967; Rol'nick, 1970). Nothing 

is known of the microscopical, ultrastructural, and biochemical organization 

of the crocodilian yolk and yolk sac, or how these change during 

development (see A. d'A. Bellairs, 1969, and R. Bellairs, 1971, for data on 

other reptiles and birds).

378 

IlA 

OlA 

If. I 

J I", 

ALB \ 7./////, 


BV FZ YS vav Y EEC BV 

' .. 

'\. 

'111.'.'....' 

A 

ALB 

( / I - J I SAC 

Fig. 10. Alligator mississippiensis. Diagrammatic longitudinal section through a 40-day egg. 

At this age the chorion and related blood vessels extend over the entire inner aspect of the 

egg, and the eggshell is entirely opaque. The disposition of the chorionic blood vessels and 

opaque eggshell band illustrated are from an earlier age and for diagrammatic purposes only. 

(A) Amnion (solid black); AC, amniotic cavity; ALC, allantoic cavity; ALB, albumen; BV, 

chorio-allantoic blood vessels; C, chorion (dotted); CE, calcified eggshell (opaque band in 

cross hatched area); EEC, extraembryonic coelom (potential space between yolk sac membrane 

and the chorion, largely occupied by the allantois); ESM, shell membrane (opaque in 

cross hatched area); FZ, fusion zone (fusion of chorion, outer and inner layers of allantois and 

amnion, all of which have a firm attachment to the shell membrane); GL, gut loop (herniated 

out of embryo); ILA, inner layer of the allantois (fused to the amnion); OLA, outer layer of the 

allantois (fused to the chorion); SAC, seroamniotic caVity (potential space between amniun 

and chorion, largely occupied by the allantois); VBV, vitelline blood vessels; Y, yulk; YS, yolk 

sac membrane. 

At the time of laying, eggs have approximately equal volumes of yolk 

and albumen, with the yolk being significantly denser than the albumen. 

Within the first day, these ratios change as water passes across the vitelline 

membrane from the albumen. The vitelline fluid accounts for approximately 

3% of the total egg content weight at day 1, 40/0 at day 2, 7% at day 

3, 15% at day 6, and 27% at day 10. During this period, the volume and 

weight of albumen decreases from 50% of the total egg content weight at 

day 1, to 45% at day 2, 37% at day 3, 31 % at day 6, and finally to 18% at day 

10. Concurrently, the volume and weight of yolk remains fairly constant at 

around 45 to 55%, and the total egg weight changes by only 0.5% (G. 

Webb, personal communication). The most aqueous vitelline fluid, being 

appreciably less dense than the yolk granules, remains separated from 

THE EBB CONTENTS AND EXTRAEMBRYONIC MEMBRANES 

379 

them. Thus the yolk rotates, bringing the embryo up toward the top of the 

egg regardless of its original position. The vitelline fluid remains beneath 

the embryo, and its assimilation is responsible for dispersing the albumen 

to the poles of the egg and also for increasing its viscosity. As a result, the 

vitelline membrane adheres to the inner surfaces of the shell membrane 

and produces the opaque spot. The latter keeps expanding as more and 

more fluid crosses the vitelline membrane (Webb, personal communication). 

Assimilation of vitelline fluid is an active process, which explains 

why infertile eggs never become opaquely banded. The opacity reflects 

dehydration of shell and shell membranes, caused by the loss of the central 

layer of albumen. Chemical changes occur later and include a more permanent 

opacity (Ferguson, 1982a). Thus, the length of the opaque band parallels 

the regression of albumen early in incubation; later it also reflects the 

expansion of the chorioallantois and the movement of minerals from the 

shell to the embryo. Dehydrated or infected eggs (whether fertile or not) 

may become erratically banded or blotchy as albumen regresses and the 

yolk sticks to the shell membrane. 

If an alligator egg is placed with its top surface (as laid in the nest) 

uppermost and the widest (blunt) end of the egg pointing toward the 

investigator, then the head of the embryo lies distally and its snout points 

to the right in 98% of cases. This predictable relationship permits eggs to be 

windowed with accuracy. Because of the advanced stage of development 

at oviposition, it is unknown whether the early embryonic axis of crocodilians 

is formed in a constant fashion relative to egg position in the oviduct 

as in birds (R. Bellairs, 1971). 

After Stage 11, the embryonic gut projects through the body wall into 

the umbilical stalk and contacts the yolk (Figs. 10, 20, 21, and 341) and the 

paired vitelline blood vessels (Figs. 20, 21, and 341). Presumably, these 

structures participate in the breakdown of the yolk and its transport to the 

embryo. The yolk sac encloses the yolk; the paired vitelline vessels are 

continuous with the paired intestinal omphalomesenteric vessels at the 

yolk stalk (Figs. 10, 21, and 341). It is unknown whether there are yolk sac 

septae as in birds (Patten, 1925; Huettner, 1949), or if there are superficial 

and deep layers of yolk as in other reptiles (R. Bellairs, 1971). Between the 

yolk sac and the chorion lies the extra-embryonic coelom, and between the 

amnion and chorion lies the sero-amniotic cavity (Fig. 10). Both are occupied 

eventually by the enlarging allantois. The alligator has two bands of 

firm fusion between the shell membrane, chorion, outer and inner layers of 

allantois, and the amnion (where present) (Fig. 10). These two bands are 

opposite each other and the lower fusion zone adheres less strongly to the 

yolk sac (Fig. 10) than does the upper fusion zone to the amnion. The two 

fusion zones hold the embryo in a fairly constant position and must be cut 

to remove an embryo or yolk from a fixed egg (Ferguson, 1982a). 

Several days before hatching, the embryonic intestines and the small 

yolk sac are withdrawn into the body cavity through the umbilicus (Fig.

380 REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS EARLY EMBRYONIC DEVELOPMENT [BEFORE EGG LAYINGJ 381 

Ectoderm 

fibers in the amniotic mesoderm of Alligator mississippiensis may rock the 

embryo and circulate the amniotic fluid (Schroder and Bautzmann, 1956). 

Entoderm 

Fig. 11. Alligator mississippiensis. Diagrammatic cross section through the yolk (speckled), 

early embryo and forming extraembryonic membranes to show their composition and the 

layers which with further development expand and fuse to produce the arrangements shown 

in Fig. 10. (After Huettner, 1949, in the chick.) AL, Allantois; AM, amnion; C, chorion. 

21). At the time of hatching, the abdomen is considerably distended by 

absorbed yolk and yolk sac (Fig. 21), and the umbilicus is often open. The 

absorbed yolk is used as a food supply over the next couple of weeks 

(traces remain 6 months after hatching, Cott, 1961), and the umbilicus then 

closes. The mechanisms of yolk digestion and utilization are unknown. 

The arrangement of the extra-embryonic membranes has been described 

in Alligator mississippiensis (Ferguson, 1982a), and other data are available 

on their development in this and other species (Rathke, 1866; Clarke, 1891; 

Voeltzkow, 1899, 1901; Reese, 1908, 1912, 1915a; Fisk and Tribe, 1949; 

Ferguson, unpublished). The relevant stages are difficult to obtain as eggs 

are laid at a fairly advanced stage of development (Fig. 18). The development 

of crocodilian extra-embryonic membranes (Fig. 11) is probably similar 

to that of the chick (Patten, 1925; Huettner, 1949). 

Alligators, like other amniotes, have three extra-embryonic membranes: 

the chorion, amnion and allantois, and a yolk sac (Fig. 10). The chorion 

lines the inner aspect of the shell membrane, except in the region of the 

albumen. As development proceeds, the volume of albumen decreases and 

the chorion increases in size. The general organization and relative relations 

of the amnion, chorion and allantois, and the vitelline and chorioallantoic 

circulations (Figs. 10 and 11) are essentially similar to the avian 

condition. No detailed studies are available on the ultrastructure of 

crocodilian extra-embryonic membranes or on the mechanisms of yolk 

mobilization, embryonic excretion, and respiration. The smooth muscle 

V. EARLV EMBRVONIC DEVELOPMENT [BEFORE 

EGG LAVING] 

Little is known about the early development, and we must rely upon 

descriptions nearly 100 years old; Clarke (1891), Voeltzkow (1899) and 

Reese (1908, 1915a). In these studies, early stages were obtained from the 

oviducts after killing gravid females because crocodilian eggs are laid at an 

advanced stage of development (Ferguson, 1984a; see Fig. 18). The earliest 

stage described for any crocodilian is after the appearance of the embryonic 

body folds, neural medullary groove, primitive streak, embryonic shield, 

area opaca, area pellucida, and the early gut (Figs. 12A and B) in Crocodylus 

niloticus (Voeltzkow, 1899). Currently, it is impossible to describe or discuss 

the events that precede this stage. Except for investigations of neural 

crest, palatal, and mandibular development (Ferguson, 1981a, 1982b; Ferguson 

et al., 1982, 1983a, b; Ferguson and Honig, 1984), no experimental or 

in vitro studies have been performed on crocodilians; this description relies 

entirely on old morphological data. 

The present report offers a concise account of the major events; for 

detailed "catalogue type" descriptions of sectioned materials, the reader is 

referred to earlier references. The standardized terminology used here 

does not imply homology with synonymous terms for birds and mammals. 

Shortly after the appearance of the neural folds (Figs. 13A and B), after 

the head fold has begun to sink into the underlying yolk, an anterior 

blastodermic fold forms the amniotic head fold (Clarke, 1891; Voeltzkow, 

1899, 1901; Reese, 1915a). Initially the head fold is crescent-shaped, its free 

edges pointing toward the tail region of the embryo (Figs. 13A, B, 14A, B); 

as development proceeds, the head fold extends craniocaudally. The amniotic 

primordium, derived from somatopleure around the trunk (Fig. 11), 

arises in continuity with that of the head; whether it does so as a posterior 

extension of the head fold or as paired lateral amniotic folds (as in the 

chick, Patten, 1925; Huettner, 1949) is unknown. This combined headtrunk 

amnion reaches the level of the blastopore about the time of egg 

laying (Stage I, Fig. 18). 

Voeltzkow (1899, 1901) recognizes the development of a tail fold (tube) 

of amnion (his Figs. 28A and B), whereas Clarke (1891, 1901), Reese (1908, 

1912, 1915a), and Fisk and Tribe (1949) claim that such never develops. 

Voeltzkow (1899, 1901) also describes a posterior "amniotic duct," but Fisk 

and Tribe (1949) state that this does not resemble the posterior amniotic 

tube of turtles. Attachment of the amnion to the caudal region between 

Stages 1 and 2 (Fig. 18) seems to be effected by extension of the trunk folds, 

as the membrane becomes well developed laterally and caudally between

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'ig. 12. CrocodylllS niloticllS. (A, B) Dorsal and ventral views respectively of an early embryo. 

After Voeltzkow, 1899.) (C, D) Dorsal and ventral views respectively of an embryo of Crocody 

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Stages 1 and 10 (Figs. 18-20). Development of the dorsal amniotic fold 

facilitates craniocaudal separation of the embryo from the blastoderm 

(Figs. 15B, 16B, 17A-E, and 18), but the process is not completed caudally 

until Stage 3 (Fig. 18). With further development of the blastoderm consisting 

of ectoderm and endoderm (Fig. 12E), the neural groove and blastopore 

become clearly demarcated (Figs. 12C-E). The endoderm may form 

"tails" that extend outward and downward into the underlying yolk 

(Voeltzkow, 1899, 1901; Figs. 12C-E), which probably represent presumptive 

extra-embryonic endoderm. The blastopore is relatively large and penetrates 

the entire blastoderm (Figs. 12C-E), and the primitive streak lies 

posterior to the blastopore (Figs. 12C-E). 

With the rapid delineation of the body folds (Figs. 13A and B), the 

boundary between embryonic and extra-embryonic tissues becomes discernible. 

Figures 13A-D show the well-defined head fold bounded anteriorly 

by the proamnion. The beginning of the foregut is evident. The 

notochord extends along the midline from the head fold to the anterior 

edge of the blastopore (Fig. 13C). Earlier explanations (Reese, 1908, 1912, 

1915a, 1926) of the origin of the notochord are dubious in the light of 

current data from other vertebrates. The primitive streak and primitive 

groove lie posterior to the blastopore (Figs. 13A-D); the primitive groove 

[Fig. 14C (6)-(7)] is continuous with its posterior end. The ectoderm bordering 

this groove is thickened, and its two elevations constitute the primitive 

streak (Figs. 13A-C, 14A, B, C (6)-(7)). The primitive streak extends 

about one-third the distance between the head fold and the blastopore 

(Reese, 1908, 1915a). The posterior end of the neural groove is said to be 

continuous with the primitive groove, and the posterior ends of the neural 

folds continuous with the edges of the primitive streak; these structures 

can only be demarcated arbitrarily from the dorsal opening of the blastopore 

(Reese, 1908, 1912, 1915a). This type of gastrulation (blastoporal 

canal, etc.) is found in all reptiles, although specific details may differ. 

Neural folds have a double origin in Alligator mississippiensis (Clarke, 

1891; Reese, 1908, 1912, 1915a) and Crocodlflus niloticus (Voeltzkow, 1899, 

1901). The posterior folds arise as ectode~mal ridges extending forward 

from the blastopore and bounding the neural groove (Figs. 13A,B, and 

14A, B). However, a secondary fold occurs anteriorly in the head region 

(Figs. 13A,B, and 14A,B) and grows posteriorly along the median dorsal 

r;"F 

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A 

Fig. 16. Crocodylus niloticus. Dorsal view of embryos. (After Voeltzkow, 1899). Equivalent to 

Reese (1908, 1915a) stage IV for Alligator mississippiensis. (A) Ventral flexion (V) of the anterior 

end of the neural folds (M). (B) Closure of the neural folds (M). A, the amniotic head fold; B, 

developing brain; PS, primitive streak; Y, yolk. 

line to form a V-shaped process with the apex pointing toward the blastopore. 

The apex of the V-shaped secondary head fold later disappears and 

each of the separate arms becomes continuous with the corresponding 

posterior neural fold. Thus, the secondary head fold forms the anterior 

part of the neural folds (Figs. 13A,B and 14A,B). Several stages in neurulation 

are shown in Figs. 15A-C, 16A,B, and 17A, B). Closure of the folds in 

Fig. 17. Crocodylus niloticus. Embryos. (After Voeltzkow, 1899.) (A. B) Dorsal and ventral 

views illustrating the dorsal fold of amnion and the lateral body folds delimiting the embryo 

from the overlying blastoderm in a craniocaudal sequence. The primitive head region is flexed 

ventrally. EqUivalent to Reese (1908, 1915a) stages V and Vll for A. mississippiellsis. (C, D) 

Dorsal and ventral views illustrating the progressive delineation of the embryo from the 

blastoderm by the caudal fusion of the edges of the amniotic and body folds. The head is 

further flexed ventrally. (E, F) Dorsal and ventral views just prior to egg laying. Note that the 

edge of the dorsal amniotic fold lies at the caudal third of the embryo. The lateral body walls 

are delimiting the primitive gut tube ventrally and the heart has developed as a simple tube. 

The head is flexed ventrally and hangs vertically into the underlying yolk. It is shown rotated 

to one side for diagrammatic purposes only. A, Dorsal amniotic head fold; H, lateral body fold; 

BD, blastoderm; E, caudal edge of dorsal amniotic fold; G, primitive gut; H, head region; HT, 

primitive heart; N, neural tube and notochord; NP, patent posterior neuropore; 0, otic 

placode; PS, primitive streak; S, somite; Y, yolk. 

!:IDCIl

390 REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCDDILIANS 

C. niloticus occurs first in the middle region of the embryo (Fig. 15A), nearer 

the posterior end of the neural groove (Voeltzkow, 1899, 1901), but in A. 

mississippiensis it occurs nearer the cranial end of the neural folds (Reese, 

1908, 1915a). 

The blastoporal or neurenteric canal is visible in all early embryos until 

after the closure of the neural canal. As in earlier development (Fig. 12E), 

the blastoporal canal runs through the embryo (Fig. 14C), backward from 

its cranially located ventral opening [Fig. 14C (3) and (4)], to open dorsally 

into the neural groove at its caudal limit [Fig. 14C (5)]. Some mammals 

show this blastoporal canal as the chordal canal (R. Bellairs, 1971). 

Throughout this period, somitogenesis occurs along the median axis (Figs. 

15A and 17A-E), the first pair developing halfway between the anterior 

and posterior ends. The peripheral somitic cells are compactly arranged, 

but small myocoels lie in the center of the somites [Figs. 15C(i)-(v)]. The 

mesodermal layers cleave, forming the somatic and splanchnic components 

[Figs. 15C(i)-(v)] as the foregut enlarges. 

Early in development the head fold of the embryo projects ventrally into 

'the underlying yolk (Figs. l3C and D). This process is accentuated by a 

ventral bending of the anterior neural folds (Figs. 15B, 16A,B, and 17A-E) 

and still later by cranial flexure. Thus the entire cranial end of the embryo 

cannot be seen from above, because it is pushed down into the yolk (Figs. 

15B, 16A, and 17A,B). Torsion occurs between Stages 3 and 6 (Section VI), 

beginning anteriorly and proceeding posteriorly (Figs. 18-20). Despite earlier 

conflicting reports (Clarke, 1891; Reese, 1908, 1912, 1915a; Deraniyagala, 

1939), the embryos of Alligator mississippiensis, Crocodylus porosus, C. 

johnsoni, and C. niloticus usually come to lie on their left side (Figs. 18-20). 

In Crocodilians, torsion occurs at a more advanced developmental stage 

(Stages 3-6) than in chicks (H. H. Stages 12-15, i.e., 16-20 somites). 

VI. 

STAGES OF EMBRVONIC DEVELOPMENT [AFTER 

EGG LAVING] 

The practical importance of establishing Normal Tables of development for 

vertebrate embryos, and ectotherms in particular, is discussed elsewhere 

(Billett, Cans, and Maderson, Chapter 1, this volume). No staging scheme 

exists for any crocodilian. Clarke (1891), Voeltzkow (1899), and Reese 

Fig. 18. Alligator mississippiellsis. Stages 1 to 4 of embryonic development. Numbers indicate 

the stages. See text for their description. 0, Dorsal view; L, lateral view; V, ventral view. 

Dorsal (20) and ventral (2V) views of a stage 2 embryo illustrate its attachment and vertical 

relationship (i.e., no body torsion) to the blastoderm. Body torsion commences at stage 4 (4V) 

when the cranial end has rotated; it is complete by stage 6 where a ventral view (6V) illustrates 

its relationship to the overlying chorion. Scale bars = 1 mm.


Fig. 19. Alligator mississippiensis. Stages 5 to 14 (numbered) of embryonic development. See 

text for description of stages. Scale bars = 1 mm. 

(1908, 1915a) described some embryos in both surface view and section, 

but the series were incomplete, and chronology and incubation conditions 

were poorly documented. Random embryos of unknown age and 

developmental history have been described by some field biologists (Deraniyagala, 

1934, 1936, 1939; Pooley, 1962; Magnusson and Taylor, 1980); 

such have but limited embryological value. Preliminary information for 

estimating the ages of Crocodylus porosus and C. johnsoni embryos in the 

wild, including some data on the effects of varying incubation temperatures 

is available (Webb et al., 1983a,e). Embryological stages constructed 

for Alligator mississippiensis equated with the number of days after egg 

laying, under known incubation conditions (Ferguson, 1982b), proved 

cumbersome to use because staging older embryos concerned mainly the 

growth of external structures. Moreover, study of C. johnsoni and C. porosus 

has shown that their development is similar to that of A. mississippiensis. 

Timing is the principal variable, particularly in the later stages. Thus, it is 

possible to construct a staging scheme applicable to three species, and 

! ,. 

Fig. 20. Alligator mississ i ppiel1sis. Stages 15 to 21 (numbered) of embryonic development. See 

text for description of stages. Scale bars = 2 mm.

384 


BTAGEB OF EMBRYONIC DEVELOPMENT tAFTER EGG LAYINGJ 385 

presumably to crocodilians in general. This is of greater value than speciesspecific 

stages. 

The approach adopted is twofold (Ferguson and Webb, in preparation). 

First, a detailed morphological staging scheme related to specified incubation 

conditions and chronological age is presented. Species-specific characters 

are noted, but the criteria for designating particular stages are 

"crocodilian" and species-independent. Second, 13 different standard 

measurements, for example, total length, eye length and snout length, 

which were made on embryos of all stages of Alligator mississippiensis, 

Crocodylus johnsoni, and C. porosus, have been supplemented by data on 

egg dimensions, weight, and the percentage of opaque eggshell banding. 

Such morphometric data permit several analyses: determination of the 

relationship between egg size and embryo size within and among species 

(in general, larger embryos come from larger eggs and vice versa); determination 

of the relationship between the percentage of opaque banding and 

embryonic stage; determination of diagnostic ratios (e.g., eye length over 

total length ratios are used to correct for the effect of egg size) for each 

stage in each species; interspecific comparison of similar stages to determine 

species-specific features (e.g., C. porosus embryos are larger and have 

longer tails, C. johnsoni have larger limbs and narrower snouts, A. mississippiensis 

have more abdominal yolk and retarded external genitalia); formulation 

of regression equations of chronological age against particular morphometric 

ratios (at different incubation temperatures and humidities) to 

predict "morphological age" for any embryo of a particular species (d. rat 

embryos, Ferguson, 1978a). This morphological aging is valuable when the 

time interval between stages is long, due either to slow changes in the 

external form (Stages 20-24) or because the embryo is fully formed and 

merely enlarging in size and absorbing yolk (Stages 25-28). The two systems 

are complementary, so that an embryo may be classified as "Stage 24, 

morphological age 58.5 days." In general, staging by external morphological 

criteria is most accurate up to Stage 20, because development is fast, 

and the time intervals between stages are small. After Stage 20, the morphological 

age based on morphometric ratios is most accurate for aging, 

because the time intervals between stages are long, and the embryos are 

large and easily measured. 

In the follOWing summary of the morphological staging system (Normal 

Table), no morphometric data are given. For each stage the most important 

diagnostic features are listed first, and the essential features are illustrated 

in Figs. 18-22. Details of important regions of various stages are illustrated 

in Figs. 23-26. Some stages may be subdivided by reference to certain 

criteria, for instance, Stages 1-6 may be supplemented with a somite count 

(e.g., Stage 1120 s embryo), Stages 17-19 with the percentage of palatal 

closure, and Stages 20-28 by morphometric ratios (e.g., yolk volume, yolk 

scar width, total length) (Ferguson and Webb, in preparation). Initially, the 

stages have not been separated by criteria such as somite counts, because 

these vary in relation to other features, for example, some embryos develop 

by enlarging existing somites before they form new ones, whereas 

others may form new somites before enlargement (such variation is often 

temperature dependent). The development of different structures does 

vary independently (e.g., in a particular specimen branchial arch development 

may lag behind that of the limbs) and appears to be marked at the 

extremes of incubation temperatures. The present staging scheme is based 

on examinations of approximately 1500 embryos of Alligator mississippiensis 

and 300 each of Crocodylus porosus and C. johnsoni. All embryos were fixed 

in 10% formal saline and photographed with oblique incident illumination 

using a Wild M8 stereophotomicroscope. 

Some incubation ages in days are given for embryos within each stage 

(see Table VI). For Alligator mississippiensis, these ages, given for all stages, 

are based on a number of standard series collected within three hours of 

egg laying and artificially incubated at 30°C and approximately 90-100% 

humidity. For Crocodylus johnsoni and C. porosus, the standard series were 

incubated under similar conditions (Webb et aI., 1983a,e; unpublished). 

Ages are not ascribed on the basis of the drawings in Webb et al. (1983a,e), 

which were prepared as a field guide, but on reexamination of the original 

embryos. The timing of development relative to stages is virtually identical 

in all three species up to Stage 20 (i.e., during organogenesis); only later, 

when growth is occurring, do species-specific differences appear. Some of 

these may be due to differing incubation conditions (e.g., whether compensation 

is made for metabolic heat); perhaps the ages given for A. mississippiensis 

up to Stage 24 are similar in all species. 

Table VI also includes "equivalence values" for each described stage to 

the figures in Voeltzkow's (1899) report on Crocodylus niloticus and the 

stages in Reese's (1915a) monograph on Alligator mississippiensis, which 

latter included Clarke's (1891) illustrations. Some of the data in these older 

works are inaccurate, even self-contradictory. In the stage descriptions 

presented here, judgment of such major errors invoked the premise that 

new data for A. mississippiensis, C. porosus, and C. johnsoni are more complete 

and more detailed. Only limited data are available relating chronological 

age and stage at temperatures other than 30°C. In A. lIlississippiensis, 

increasing the incubation temperature (within viable limits) accelerates development 

up to Stage 20 (organogenesis) to a much greater extent than in 

later stages. This is similar to observations on birds (Romanoff, 1967), 

snakes (Zehr, 1962), and turtles (Yntema, 1968; Ewert, 1979). However, it is 

difficult to make precise statements regarding the effects of incubation 

temperature on developmental rates because of tissue specificity, e.g., 

gonadal development is retarded at 34°C compared to stage by stage events 

at 30°C. 

There are also difficulties in adjusting regression equations for variations 

of temperature during incubation in order to predict embryonic age (or 

hatching dates) from morphometric ratios (Webb et al., 1983a,e). Generally, 

development of alligator embryos up to Stage 20 proceeds approximately 

1.2 to 1.8 times as fast at 34° as at 30°C, and approximately 1.6 to 2.0

Stage 

2 

3 

TABLE VI 

Concordance for Staging/Aging of Crocodilian Embryos 

AM" (days) Cr's (days) CP'" (days) V dl , 

W,h 

o I (after egg 0-1 0-1 

laying) 

VI(37a, b) VlII, IX 

2 

3 

2 

3 3 

4 4 4 4 

5 5 5 

6 

6 

7 

7 

9 

10 

11 

12 

13 

14 

IS 

16 

17 

8 8 8 8 

9 

10-11 

12 

13-14 

IS 

16-17 

18-20 

21 

22-23 

9 

10 

12-13 

14 

IS 

16 

18-20 

21-22 

23 

18 24-26 26 25 

19 

20 

27-28 

29-30 

VI(38a, b, c) 

VI(40), 

VlII(5Ia) 

VII(43), 

VlII(5Ia) 

6 6 

7 VII(44), 

9 

10 

VIII(52, 53), 

IX(66 f , 74) 

VII(45), VlII(54), 

IX(67', 75, 76) 

IX(68, 77) 

VII(46), 

VlII(551' 

IX(69) 

12 

13 VII(47), 

VlII(55), 

IS 

17 

19 

21 

29 

30-34 29 

21 31-35 35-39 

22 36-40 40-45 35 

23 41-45 46-54 44 

24 46-50 55-64 55 

25 

26 

51-60 

ABSENT 

65-74 

75-80 

58-70 

71-75 

IX(69)f 

IX(70, 79), 

XV(134) 

VII(48), VIII(56), 

IX(71, 80), X(84A), 

XV(135) 

IX(72, 81) 

VIII(57), IX(82), 

X(85A) 

VII(49), VIII (58), 

X(86A), XI(105) 

X(87A,88A) 

VII (50), VlII(59), 

XI(95, 106), XV(136) 

VIII(60), XI(107A, B) 

VIII(61, 62) 

XI(96) 

XI(97) 

VIII(63), XI(98A-B) 

27 60-63 81-83 75-80 XI(99, 100) 

28 64-70 84-90 81-85 VlII(64), XI(108) XXlII 

X, XI 

XII 

XIII 

XIV 

XV, XVI 

XVII 

XVIII 

"AM, Alligator mississippiensis. dV, Voeltzkow (1899) stages for C. niloticus. VII(4a) = Tafel VII, Fig. 4a. 

bq, Crocodylus johnsani. 'R, Reese (1915a) stages for A. mississippiensis. 

'CP, Crocodylus porasus. fmajor error in the figure. 

sThe numbers of embryos of Cracodylus johnsoni and C. parasus with definite ages are limited, so that the 

days given do not inclUde the range per stage, as provided for Alligator mississippiensis. The values represent 

the known ages of standard embryos available for the stage. 

hThe drawings of Reese (1908, 1915a) contain numerous errors, whereas those of VoeItzkow (1899) tend to 

be accurate. Hence, representative figures selected from the latter are used. Some contain major errors, 

e.g., his Figures 39b and 43; these have either been omitted from this table or, if no other illustration was 

available are listed with the notation! which indicates a major error. The present cross-reference between 

the drawings of whole embryos and those of regions may differ from the original, but is consistent with the 

data of the three species for which recent observations are cited above. Voeltzkow's (1899) ages appear 

inaccurate and are inconsistent within his study (e.g., Figs. 49 and 58 show the same embryo, but are stated 

to represent ages of two months and four weeks, respectively). They have been omitted, although the trend 

suggested is similar to that for the other two species of Cracodylus. 

3BB 

XIX 

XX 

XXI 

XXII 

STA13ES OF EMBRYONIC OEVELOPMENT [AFTER EGG LAYING] 

397 

times as fast at 30° as at 28°C. Thereafter, rates of development at different 

temperatures show less variation; eggs incubated at 34°e hatch after 55-60 

days incubation, at 30 0 e after 62-67 days and at 28°e after 70-75 days. The 

situation may differ in species with longer incubation periods, possibly as a 

result of increased time spent in the egg between Stages 24 and 28. It is likely 

that these later stages of growth and yolk absorption are accelerated by 

increased temperatures. Field nests of Crocodylus johnsoni increase gradually 

in temperature throughout incubation (Webb and Smith, 1984), and it 

would be interesting to know whether this facilitates accelerated development, 

yolk absorption, and hatching. Alligator mississippiensis (and some 

other species) may hatch in the minimum time possible (perhaps as a result 

of enhanced development in the later stages, Webb et al., 1983e) as an 

adaptation to their climatic environment. However, species occupying 

more tropical environments may incur a selective advantage by staying 

longer in the egg. Accurate prediction of hatching date is difficult for any 

species because hatching is initiated by factors other than completion of 

development; viable embryos incubated under constant conditions hatch 

over a range of 5 to 10 days and show varying degrees of yolk absorption. 

The problems of identifying the early embryo have beset numerous 

investigators (Clarke, 1888a, 1891; Voeltzkow, 1892, 1899; Reese, 1907, 

1915a; Wettstein, 1954; Ferguson, 1982b). Statements in the literature, contending 

that development begins at oviposition, reflect the difficulties of 

observing Stage 1 embryos under field conditions (McIlhenny, 1934, 1935; 

Pooley, 1962). For Alligator mississippiensis, Crocodylus johnsoni, and C. 

porosus, numerous embryos that were recovered within a few hours of egg 

laying have always been Stage 1, although with variation in the number of 

somite pairs (9-20; mean 12 ± 2) and in the degree of delineation of the 

dorsal wall. Because development of the earliest laid embryo to the latest 

occurs in 24 hours, Stage 1 embryos have been ascribed an age of 0 to 1 

days. 

Stage 1 

[Fig. 1 Bl 

Blastoderm. Blastoderm and embryo lie on top of the yolk and are not 

attached to the overlying shell membrane. Blood vessels are not evident. 

Somites. 9-20 pairs (16-18 are very common at the time of egg laying) 

lie posterior to the otic placode. 

Delineation of the Embryo from the Blastoderm. Approximately the caudal 

one-third of the dorsal body wall is not delineated from the blastoderm, for 

example, in a ISS embryo the body is delimited to the level of the 7th 

somite, in a 16S embryo to the 8th somite, in an 18S embryo to the 12th 

somite, and in a 20S embryo to the 16th somite. 

Branchial Arches. The first branchial arch is just visible. 

Heart. A simple S-shaped tube in the midline. 

Sensory Placodes, Pits, and Brain. Both the optic and otic placodes are

39B 

REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCOOILIAN9 

evident and may be invaginating. The optic and otic vesicles are also obvious, 

but the brain is not yet regionalized. The optic pIacodes and vesicles 

are more obvious than the otic. 

Flexures and Rotation. The embryo lies at right angles to the yolk surface, 

that is, body torsion has not yet commenced. 

Blastopore and Primitive Streak. Obvious. 

Gut. The gut is incomplete caudally and open ventrally along its entire 

length. 

Somitomeres. Three pairs of cranial somitomeres are visible anterior to 

the otic vesicle. 

Notochord. The notochord is evident. 

Stage 2 [Fig. 1 S) 

Blastoderm. The dorsal surface of the blastoderm is attached to the 

overlying shell membrane, and the embryo remains so tethered throughout 

subsequent development. Blood vessels are now visible; one pair 

emerges from the embryo at the caudal level of the heart, whereas another 

(larger) pair runs down the lateral wall of the embryo to emerge at approximately 

the level of the 20th somite. 

Somites. 21-25 pairs (decreasing markedly in size caudally). 

Delineation of the Embryo. Dorsally the embryo is almost completely 

delineated except for a very small circular area at the extreme caudal tip. 

Ventrally the caudal and caudo-Iateral boundaries of the body wall have 

formed. 

Branchial Arches. 

visible. 

The 1st and 2nd arches and the 1st branchial cleft are 

Heart. An extra vertical loop has developed making a total of three 

loops. The heart lies in the midline of the embryo. 

Sensory Placodes, Pits, etc. The lens placode and optic cup are defined, 

and the otic pit is distinctly patent. 

Brain. Hindbrain discernible as a clear transparent region. 

Flexures and Rotation. The cranial end of the embryo is flexed at approximately 

the level of the heart with the head lying at approximately 45° to 

the plane of the body. No body torsion has occurred. 


Somites. 26-30 pairs. 

Delineation of the Embryo. Complete. 

Branchial Arches. Three branchial arches, the 1st branchial cleft, the 2nd 

branchial groove, and the branchial sinus are all present. 

Tail. Bud present, but lacks somites. 

Brain. Forebrain, midbrain, and hindbrain are now discernible, the 

latter appearing distinctly transparent. 

Sensory Placodes, Pits, etc. The optic cup has an elongated horseshoe 

BTAGEB OF EMBRYONIC OEVELOPMENT [AFTER EGG LAYING) 

399 

shape, which extends below the lens vesicle onto the roof of the primitive 

oronasal cavity. 

Blastopore and Primitive Streak. Not visible. 

Extra-embryonic Membranes. The amnion is attached ventrally to the lateral 

body walls, cranially to the borders of the pericardium about the level 

of the 7th to 8th somite and caudally to the cranial margin of the fold of tail 

bud. 

Flexures and Rotation. Head at approximately right angles to the body. 

No body torsion. 


Somiles. 31-35 pairs. The first is beginning to disappear. 

Tail. Distinct, straight, and contains 3-5 somites in its base; the tip is 

unsegmented. 

Flexures and Rotation. Body torsion has commenced. The cranial half of 

the embryo is rotated so that its right surface is in contact with the shell 

membrane and its left is parallel to the underlying yolk. The caudal half of 

the embryo is not rotated and lies at right angles to the shell and yolk. 

Heart. Displaced to the left side of the embryo and large. 

Allantois. Small elevation is just visible immediately caudal to the craniallimit 

of the ventral tail fold, at approximately somite 27. 

Branchial Arches. Three branchial arches, the 1st branchial cleft, the 2nd 

and 3rd branchial grooves, and the branchial sinus are present. The cranial 

nerves to branchial arches 1-3 are discernible using oblique or transmitted 

illumination. 

Stage 5 

[Fig. 1 gJ 

Somites. 36-40 pairs. Only traces of the first somite can be detected, 

although it is included in this count. 

Tail. The tail tip bends ventrally at right angles to the body of the 

embryo; 6-10 somites in its base, tip unsegmented. 

Flexures and Rotation. Body torsion is complete except for the tail. The 

head is further flexed with the roof of the brain at approximately 25° to 

the plane of the body. 

Allantois. The allantoic bud is distinctly swollen, smaller in height than 

the tail. 

Sensory Placodes, Pits, etc. The otic pit lies dorsal to the junction of the 

2nd and 3rd branchial arches, and its external opening is closing. 

Stage S 

[Figs. 1 S. 1 g. and 23J 

Sensory Placodes, Pits, etc. Nasal placodes present. Otic pit closed. 

Limbs. Hindlimb buds are just visible on each side, the right hindlimb 

~ bud being marginally advanced over the left, but no forelimb buds are 

present (Fig. 23).

400 


Flexures and Rotation. Body torsion complete. The roof of the brain is at 

approximately 45° to the plane of the body, whereas the floor of the brain is 

horizontal and parallel to the plane of the body. 

Brain. The olfactory bulbs, forebrain, and midbrain are distinct. Four to 

six neuromeres are discernible in the transparent hind brain area. 

Tail. The tip is starting to curl but is unsegmented; somites are only 

present in the proximal region of the vertically oriented tail. 

Allantois. The allantoic bud is stilI smaller in height than the tail. 

Gut. The foregut and hindgut are formed, but the midgut is incomplete 

ventrally. 

Extra-embryonic Membranes. Many blood vessels in the vitelline and 

yolk sac membranes; major ones emerge at the level of the 18th somite and 

other smaller ones at the 6th and 11th somite levels. 

Stage 7 

[Figs. 19, 23, and 27AJ 

Limbs. Distinct hindlimb bud; forelimb bud just visible as a sinusoidal 

elevation. The forelimb bud extends over approximately somites 12-15; the 

smaller hindlimb bud extends over somites 26-28. Differences in size between 

forelimb and hindlimb buds are more marked in C. johnstoni. 

Sensory Placodes, Pits, etc. Nasal placode begins to invaginate. 

Brain. Midbrain bulge evident. 

Tail. Tail tip, curled at approximately 90° to the remainder of the tail, 

and contains somites. 

Flexures and Rotation. At the level of the heart, the cranial region is bent 

at approximately 90° to the body. The neck region is also flexed, so that the 

roof of the brain lies at approximately 60° to the body plane and the floor of 

the brain at 45°. This dichotomy in angulation accentuates the midbrain 

bulge. 

Branchial Arches. Three branchial arches, the first branchial cleft and 

groove, the second and third branchial grooves, and the branchial sinus 

are present (Fig. 27A). 

STAGES OF EMBRYONIC DEVELOPMENT (AFTER EGG LAYING) 

Tail. The tail tip is coiled through two 90° turns and flexion of the tail 

base has commenced. Twelve to eighteen somite pairs are present 

throughout the tail. 

Stage 9 

[Figs. 19, 23, 27C and OJ 

Branchial Arches. Four branchial arches, the 1st and 3rd branchial clefts 

and grooves, the 2nd and 4th branchial grooves, and the branchial sinus 

are visible (Figs. 27C and D). Branchial clefts are present only in the dorsal 

halves of the junctions between the 1st-2nd and 3rd-4th arches, the intervening 

ventral tissue is continuous and forms the branchial grooves. 

Facial Processes. The maxillary process is distinct and extends forward 

to the midpoint of the eye. There are elevated rims of tissue around the 

nasal pits. A remnant of the ingrowth of Rathke's pouch is visible in the 

roof of the primitive oronasal cavity. 

Eye. The optic cup and central lens anlage are large and round but 

unpigmented. 

Limbs. There is a distinct apical ectodermal ridge on the hindlimb bud, 

but not on the forelimb bud. The hindlimb bud extends out farther from 

the body than the forelimb bud (Fig. 23). 

Tail. The tail tip is curled through three 90° turns, and the tail base is 

distinctly flexed from the lower lumbar region; the tail contains approximately 

20 somites. 

Allantois. The large allantois is fused to both the amnion and chorion. 

The chorioallantois extends around approximately one-half of the breadth 

of the eggshell. 

Heart. The distinct atria, ventricles, and the lung primordia are visible 

through the transparent pericardial sac. 

Gut and Abdominal Organs. The midgut and body walls are open ventrally 

from the caudal limit of the pericardial sac to two-thirds the way 

down the body. The developing mesonephros and liver are just visible 

through the lateral body walls. 

401 

Stage B 

[Figs. 19, 23, and 27BJ 

Stage 10 [Figs. 19, 23, 27E, 34A and BJ 

Allantois. Now a large ballooning sac, longer than the tail; it contains 

blood vessels, but is not yet fused with either the amnion or the chorion. 

External Genitalia. A small elevated genital tubercle is present. 

Sensory Placodes, Pits, etc. Nasal pits are present on each side of the 

head external to the swellings of the olfactory bulbs; Rathke's pouch is 

invaginating in the middle of the roof of the primitive oronasal cavity to 

approXimate the infundibulum (Fig. 27B). 

Limbs. Forelimb and hindlimb buds both distinct and extending over 

somites 11-16 and 27-32, respectively (Fig. 23). The apical ectodermal 

ridge is developing on the hindlimb bud. 

Eye. Pigment in the iris makes the eye appear light black except for the 

central opaque lens. The right eye is usually pigmented earlier and more 

heavily than the left eye. 

Branchial Arches. Five branchial arches, grooves, and the branchial sinus 

are present. The first cleft lies dorsally, ventral to the otocyst. The third 

branchial cleft (between arches 3 and 4) is closing over. Branchial arches 1 

and 2 are merged together in their ventral halves, and arch 2 has started to 

overgrow arch 3 (Fig. 27£). Branchial arches 4 and 5 are very small. Viewed 

from the frontal aspect, the horseshoe-shaped 1st branchial arch is distinctly 

lobulated in the midline.

402 REPRODUCTIVE SIOLOGY AND EMSRYOLOGY OF CROCODILIANS 

STAGES OF EMSRYONIC DEVELOPMENT tAFTER EGG LAYING) 403 

Facial Processes. The medial and lateral nasal processes are distinct elevations 

on either side of the nasal pits. The maxillary processes extend 

forward as far as the caudal junction of the medial and lateral nasal 

processes, and delimits a distinct groove beneath the eye. 

Limbs. The hindlimb bud is fan-shaped with a distinct apical ectodermal 

ridge (Fig. 23). It extends out further from the body wall than the 

forelimb bud. 

Tail. The tail is coiled through four 90 0 turns. 

Gut and Abdominal Organs. The mesonephros and liver are clearly visible 

through the lateral body walls. 

Stege 11 [Figs. 1 g end 23] 

Facial Processes. The nasal pit slit is starting to form between the medial 

and lateral nasal processes. The club-shaped maxillary processes extend to 

the junction of the medial and lateral nasal processes and is continuous 

with the lateral nasal process. 

Limbs. Both forelimb and hindlimb buds extend out caudally from the 

body wall, and both have distinct apical ectodermal ridges (Fig. 23). The 

forelimb has a distinct constriction demarcating the proximal and distal 

elements, but this constriction is less marked in the hindlimb. 

Gut and Abdominal Organs. A loop of midgut tube is visible at the level 

of the umbilicus. 

Branchial Arches. The 2nd branchial arch continues to overgrow the 

3rd, and arches 4 and 5 are starting to submerge. The 1st branchial cleft is 

immediately ventral to the otocyst. 

Eye. Distinct black pigment present in the iris. 

Extra-embryonic Membranes. The chorioallantois extends two-thirds the 

way around the breadth of the shell membrane. 

Stage 12 [Figs. 1 g, 23, and 27H] 

Branchial Arches. The sinusoidal 1st branchial cleft lies above the otocyst 

and its margins show condensations for the auricular hillocks (Fig. 

27H). The merged conglomerate of arches 1 and 2 is growing caudally and 

has overgrown arch 3 to reach the junction between the 3rd and 4th arches. 

This conglomerate forms the base of the lower jaw, which extends as far 

forward as the middle of the lens of the eye. Arches 4 and 5 are small but 

visible. The branchial sinus is patent. 

Facial Processes. The club-shaped maxillary process extends forward as 

a large shelf of tissue beneath the eye. The nasal pit slits are deepening as 

the medial and lateral nasal processes enlarge. There is a distinct notch and 

furrow in the midline of the face between the medial nasal processes of 

each side. 

Limbs. The forelimb, which is beginning to bend in the region of con- 

striction for the proximal and distal elements, lies closer to the flank of the 

embryo. The elongated hindlimb shows little differentiation into proximal 

and distal elements, and although there is still a distinct apical ectodermal 

ridge, foot plate formation is just discernible (Fig. 23). 

Stsge 13 [Figs. 1 g, 23, 24A-C, 27H, 34C end 0] 

Facial Processes. The nasal pit slits are very distinct (Figs. 24A and B). 

The prominent maxillary processes are continuous with the lateral nasal 

processes (Figs. 24A-C). 

Limbs. The forelimb is now distinctly bent towards the pericardium. 

The distal hindlimb is flattened and enlarged into a footplate primordium 

(Fig. 23). 

Branchial Arches. Arch 3 is almost completely overgrown by arch 2, 

which now reaches the pericardium. Arches 4 and 5 are difficult to see. The 

branchial sinus has closed. The anterior margin of the lower jaw has grown 

forward from the merged conglomerate of arches 1 and 2. The 1st branchial 

cleft is now more horizontally oriented and is hereafter referred to as the 

external auditory meatus. The upper ear flap is sinusoidal with a midline 

bulge formed by the merging of the auricular hillocks (Fig. 27H). A groove 

runs craniocaudally along the basal (dorsal) aspects of the branchial arches 

and lower jaw. 

Extra-embryonic Membranes. The chorioallantois now extends as a ring 

around the inner circumference of the central eggshell membrane. 

Stege 14 [Figs. 1 g, 23, 240, 27F end H] 

Facial Processes. The nasal pit slit has closed due to the merging of the 

medial nasal, lateral nasal, and maxillary processes (Figs. 240 and 27F). 

The medial nasal processes are prominent and have anterodorsal external 

bulges (Figs. 240 and 27F). These will later grow forward to displace the 

external nares dorsally. The medial nasal processes also have two anterior 

intraoral bulges signifying the onset of primary palate development (Figs. 

240 and 27F). The maxillary processes have sinusoidal intraoral margins 

signifying the onset of secondary palate development. The internal nares 

are distinct. 

Limbs. The foot and hand plates are distinct; the former is advanced 

over the latter (Fig. 23). 

Branchial Arches. The 2nd branchial arch has overgrown the 3rd, 4th, 

and 5th, and this merged conglomerate together with the 1st arch forms 

the base of the lower jaw and the neck. A craniocaudal groove is still 

present along the dorsal margins of the merged 1st and 2nd arches. 

Lower Jaw. The lower jaw extends one-fourth the way beneath the 

upper jaw. It is broad and round in A. mississippiensis but more pointed in 

C. johnsoni.

404 


STAGES OF EMBRYONIC OEVELOPMENT [AFTER EGG LAYING] 

405 

Ear. The upper ear flap is overgrowing the external ear opening. White 

opacities, internal condensations for ear development, are evident in the 

region below the ear opening. 

Denticles. One denticle is present on each side of the developing primary 

palate (merged medial nasal processes) margins and one on each side 

of the midline of the lower jaw. 

Flexures and Rotation. The embryonic face rests on the large bulge of the 

thorax (pericardial sac). 

Gut and Abdominal Viscera. A large loop of gut herniates through the 

narrow umbilical stalk and touches the underlying yolk. The abdominal 

viscera (e.g., liver and mesonephros) are visible through the body walls. 

Tail. This is markedly coiled, and the tip has a terminal kink. 

External Genitalia. In A. mississippiensis, the tubercle is still small, but in 

C. porosus and C. johnsoni it is much larger. 

Embryonic Reflexes. Contralateral withdrawal reflexes occur. 

Stage 15 [Figs. 20, 23, 24E, and 2711 

Lower Jaw. This is one-third to one-half the length of the upper jaw 

(Fig. 24E). Dorsal to the lower jaw complex, there are three furrows and 

surface elevations, which appear to be dorsal representations of the branchial 

arches, which have now coalesced ventrally. 

Denticles. Two denticles are present on the anterior upper jaw margins 

of each side, one on the margin of the primary palate, the other on the 

maxillary process behind the closure zone. As before, one mandibular 

denticle is present on each side of the midline. 

Eye. Dark black stria in the iris radiate out from around the lens. The 

anlage for the upper eyelid is an elevated rim of tissue above each eye. 

Limbs. Distinct proximal and distal regions and hand and foot plates 

(but no digit rays) are present on both the forelimb and hindlimb (Fig. 23). 

Facial Processes. The anterodorsal (coalesced) bulges of the medial nasal 

processes have displaced the external nares dorsally (Fig. 24E). The primary 

palate has formed but still retains two anterior bulges (Fig. 24E). 

There is a distinct hollow in the face beneath the anterior one-third of the 

eye. The tectoseptal process is evident in the roof of the primitive oronasal 

cavity. 

Stage 1 6 [Figs. 20 and 231 

Limbs. Faint digital condensations are present in the footplate but not 

in the handplate (Fig. 23). 

Lower Jaw. This is now two-thirds the length of the upper jaw. 

Denticles. There are four on the upper jaw margins of each side, two on 

the primary palate (that closest to the midline may be difficult to see), and 

two on the maxillary process. Two denticles lie on each side of the mandible. 

Facial Processes. The upper jaw is hook-shaped, being moulded around 

the pericardial bulge. Small secondary palatal shelves are present immediately 

behind the posterolateral margins of the primary palate. 

Caruncle. Two tiny, widely spaced thickenings are just discernible on 

the anterior tip of the snout. 

Stage 17 [Figs. 20, 23, 24F, G, 25A, 2SA and B1 

Limbs. Mesodermal condensations for the five forelimb and four hindlimb 

digits are present (Fig. 23). 

Lower Jaw. The lower jaw lies behind the primary palate bulges (Figs. 

24F and G). The furrows and elevations dorsal to the lower jaw have 

disappeared. 

Denticles. At this and later stages, the numbers of denticles discernible 

macroscopically in the upper jaw varies as some appear and others disappear; 

consequently, upper jaw denticle number is of little further diagnostic 

value. Contrarywise, seven denticles occur on each side of the lower 

jaw; three are distinct, three less distinct, and one is disappearing and very 

difficult to see. 

Caruncle. Two small elevations, widely separated, have appeared on 

the anterior tip of the snout (similar to Fig. 26A). 

Palate. The secondary palatal shelves are distinct. They are closest to 

each other anteriorly behind the primary palate, but do not contact (Figs. 

25A, 28A and B). The tectoseptal processes are closing in the posterior half 

of the oronasal cavity. 

Scales, Scutes, etc. The somite pattern is distinct particularly on the 

dorsal aspects of the body and tail. This heralds the onset of scale differentiation. 

Anlagen for the abdominal ribs are visible. 

Flexures and Rotation. The head is extended off the bulge of the pericardial 

sac due to elongation of the neck. 

Ear. The external ear flap is distinct as the adult external ear form is 

established. 

Stage 1 S 

[Figs. 20, 23, 25B, C, 26A, and 2SC-F1 

Limbs. The digit rays on the hand and foot plates are now distinct 

cartilaginous condensations. The hand and foot plates have a crenellated 

appearance (Fig. 23) caused by the straight margins of the interdigital 

tissues. 

Lower Jaw. The lower jaw lies beneath the anterior primary palate 

bulges, and it no longer rests on the pericardial sac. 

Denticles. At the start of this stage, six denticles are visible on each side 

of the mandible, and at the end, eight are discernible. The posterior denticles 

are difficult to see. 

Caruncle. Two widely spaced, white blebs on the anterior tip of the 

snout (Fig. 25A).


Palate. The secondary palatal shelves are one-fourth closed at the beginning 

of Stage 18 and three-fourths closed at the end (Figs. 25B and C). 

This stage can be accurately subdivided by specifying the extent of secondary 

palate closure (Figs. 25B and C, and 28C-E). The upper jaw margin is 

straighter and less hooked than previous. 

Eye. The margins of the upper eyelid anlage extend over the superior 

rim of the iris forming a distinct groove between the eyelids and the eye, 

into which small instruments can be passed. 

Scales, Scutes, etc. Dorsal scalation is now marked. 

PericardiaI Sac. The bulge of the transparent pericardial sac is starting to 

be submerged into the ventral thoracic wall. 

Stage 19 (Figs. 20, 23, 250, 26B, 34E and F] 

Eye. Upper and lower eyelids are distinct. The anterior nictitating 

membrane anlage is discernible in the anterior corner of the eye (Fig. 26B). 

Caruncle. Two elevations of the caruncle have approximated each other 

at the tip of the snout (Fig. 26B), but the tissue between them is thin, 

appearing transparent under incident illumination. 

Lower Jaw. The lower jaw lies behind the anterior margin of the upper 

jaw. Consequently, if the premaxillary bulges are large, the mouth opens 

(as in Fig. 24H). The tongue and floor of the mouth contents sag beneath 

the margins of the lower jaw. 

Limbs. Interdigital clefting has commenced producing slight marginal 

notches particularly in the foot plates (Fig. 23). 

Palate. The palate is almost completely closed (Fig. 250). 

Coloration. White flecks, representing underlying ossifications, are obvious 

on the margins of the upper and lower jaws and around the ears. 

External Genitalia. The end of the external genitalia has developed a 

globular swelling. 

Denticles. There are eight to nine denticles visible on each side of the 

lower jaw. Henceforth, many mandibular denticles appear and disappear 

so that their numbers are too variable to be used in staging. 

Species Differences. In C. johnsoni and C. porosus, the nostrils are elevated 

on a nasal disk. The lateral jaw margins have distinct notches where 

the primary and secondary palates closed (as in Figs. 24H and I); these later 

accommodate the large fourth dentary teeth. 

Stage 20 (Figs. 20, 23, 24H, I, 25E, and 26C] 

Limbs. Nail anlagen develop rapidly in a specific sequence early in this 

stage (Fig. 23). They appear first on the most medial digit of the foot, then 

on the neighboring two digits, then on the most medial digit of the hand 

and finally on the neighboring two medial hand digits. Consequently, nail 

anlagen are present on the most medial three digits of both the hands and 

STAGES OF EMBRYONIC DEVELOPMENT [AFTER EGG LAYING) 

407 

feet, despite the fact that the total number of digits varies between the two 

(Fig. 21). Interdigital clefting now extends along approximately one-fourth 

the length of the digits (Fig. 23). The outer two digits of the hand and the 

outer digit of the foot never develop nails. 

Caruncle. The caruncle is now a solid structure due to consolidation of 

the region between the two initial swellings (Fig. 26C). 

Lower Jaw. The lower jaw is in its adult relationship with the upper jaw 

(Figs. 24H and I). 

External Genitalia. The external genital primordium is now pointed 

with a distinct elevation of its tip. 

Palate. The palate is completely closed but the basihyal valve is not yet 

present (Fig. 25E). 

PericardiaI Sac. The pericardial sac is now one-fourth withdrawn into 

the ventral body cavity. 

Coloration. White flecks of ossification are present along the margins of 

the upper and lower jaws, around the external auditory meatus, and in the 

proximal and distal elements of the limbs. 

Scales and Scutes. Scale formation is marked dorsally and scutes are 

beginning to appear in the neck region behind the skull. 

Stage 21 (Figs. 20, 23, 25F, 260] 

Limbs. Interdigital clefting extends three-fourths of the way along the 

digits. Phalanges can be distinguished in the digits (Fig. 23). 

Scales and Scutes. Scales are now visible on the ventral body wall as well 

as dorsally on the snout, neck, body, and tail. The dorsal neck scutes are 

clearly defined. 

Caruncle. The caruncle is a solid mass on the snout tip, but the tissue 

around the base of the caruncle is not differentiated from the other snout 

scales (Fig. 260). 

Palate. The superior basihyal valve flap is present at the posterior margin 

of the palate (and the inferior flap at the base of the tongue) and a 

plexus of palatal blood vessels is conspicuous (Fig. 25F). 

Pericardial Sac. The pericardial sac is one-half withdrawn into the body 

cavity. 

External Nares. Elevations for the constrictor nares muscles are evident. 

Eye. A white ring in the iris surrounds the outline of the lens of the eye 

and is overlapped by both upper a.nd lower eyelids. 

Stage 22 (Figs. 21, 23, 251] 

Coloration. Pigmentation is first visible on the margins of the upper 

jaw, along the ventral aspect of the flank, and on the proximal and distal 

elements of the limbs, but there is little or no dorsal pigmentation.

STAGES OF EMBRYONIC DEVELOPMENT [AFTER EGG LAYING) 

409 

53 55 58 60 65 

I 50mm I 

12 14 20 25 28 30 35 40 46 48 

DAYS AFTER EGG LAYING- 

Fig. 22. Alligator mississippiensis. Montage at constant magnification, illustrating the growth 

and changing bodily proportions of embryos from 12 days after egg laying until hatching. All 

eggs were collected within 12 hours of egg laying, incubated at a constant 30 Q C and 100% 

humidity and fixed at indicated number of days after deposition. 

-27 

Limbs. Interdigital clefting is at its adult level (Fig. 23). Scalation is 

difficult to see on the limbs. 

Eye. The eyelids are generally closed at this and subsequent stages. 

PericardiaI Sac. The pericardial sac is two-thirds withdrawn into the 

body cavity. 

StBge 23 [Figs. 21. 23. 24.... 25G. 26EJ 

Fig. 21. Alligator mississippiensis. Stages 22 to 28 (numbered) of embryonic development. See Coloration. Pigmentation is more extensive, and the embryos are light 

text for description of stages. Millimeter scales included. 

brown. Dorsally, stripes are present, the pattern of which varies within

STAGES OF EMBRYONIC OEVELOPMENT [AFTER EGG LAYING) 

411 

1ni'm" 

STAGE 6 7 8/9 10 

L:;:::(L~~0-- (\ 

RF~ 

bl:J~ ~ 

1mm 

STAGE 12 13 14 15 

/ 

~0_ 

\ \ 

ffrt 

16 17 18 19 

LHf\~>l~~~~~r 

R' rrmn 

\ 

23 2mm 24 26-28 

STAGE 20 21 22 25 

and between species in association with definitive scale and scute patterns. 

Pigment is also present ventrally and on all limb elements. 

Limbs. Scales are present on the proximal and distal elements. The nail 

tips of the feet have a slight distal elevation (Fig. 23). 

Jaws. Sensory papillae are present along the lateral jaw margins and 

scales are evident on the gular skin (Fig. 24J). 

Caruncle. The caruncle is located on a smooth white base (Fig. 26£). 

Brain. The midbrain is visible as a white bulge at the back of the 

cranium because the overlying skin is poorly pigmented and the osseous 

cranial roof is incomplete. 

Pericardial Sac. The pericardial sac is three-fourths withdrawn into the 

body cavity. 

Palate. An extensive plexus of blood vessels and sensory papillae has 

formed (Fig. 25G). 

Species Differences. In C. johnsoni and C. porosus the scales along the jaw 

margins are triangular with the apex of the triangle toward the jaw margins, 

giving the latter a serrated appearance. The external genitalia are 

larger and more differentiated than in A. mississippiensis. 

St;ags 24 (Figs. 21, 23, and 24KJ 

Coloration. Pigmentation is now denser so that embryos appear blacker 

in color. Various patterns occur both within and between species. 

Limbs. The nails on the hand also have elevations at their tips (Fig. 23) 

and these elevations are starting to form the curves at the tip of the nails. 

Brain. The midbrain, enclosed by bone, is overlain by pigmented skin. 

Pericardial Sac. The pericardial sac is fully withdrawn into the body 

cavity and the ventral thoracic wall is closing in the midline. 

Yolk. A large volume of yolk lies outside the body and the ventral 

umbilical area is large. 

Scales and Scutes. The scales and scutes are now very evident as elevations 

all over the embryo (Fig. 24K). 

l~ 

~. 

~ 

Ai 

Fig. 23. The typical (diagnostic) appearances of crocodilian right fore and left hind limbs 

(including hands, feet and nails) at various stages of development. Stages 6 to 11 are views 

from the dorsal aspect of the embryo and depict the projection of the limb anlage from the 

flank. Stages 12 to 17 are lateral views of the sides of the embryo and depict the proximal and 

distal elements of the limb anlage. Stages 18 to 28 are views of the hands, feet, and nails. Up to 

Stage 17, no structures except the digital mesodermal condensations (at Stages 16 and 17) are 

visible macroscopically in the limbs, thereafter anlage for the limb and digit cartilages, for the 

nails and for the interdigital webbing are evident, hence the change in diagram style at Stage 

18. Based on observation of Alligator mississippiensis, Crocodylus porosus, and C. johnsoni, although 

some (Stages 18 to 28) drawings are adapted from Voeltzkow's (1899) work on C. 

niloticus. AER, Apical ectodermal ridge; 0, mesodermal condensations for the digits; DC, digit 

cartilages; N, nail anlage.

STAGES OF EMBRYONIC OEVELOPMENT (AFTER EGG LAYING] 

413 

Stage 25 [Figs. 21, 23, 26F, 34G, and Hl 

The embryo is like a miniature version of a hatchling, with a considerable 

volume of external yolk and a large umbilical region. The hooking of the 

nails becomes more prevalent toward hatching (Fig. 23). Musk glands are 

visible along the posterior lateral margins of the intergular floor of the 

lower jaw. The caruncle is a bifid elevation on a smooth base (Fig. 26F) 

similar in structure to that at hatching (Figs. 26G, H), although perhaps a 

little less pigmented (this is variable). As few macroscopic changes are 

visible at this and later stages, aging is best estimated by morphometric 

procedures. 

Stage 26 

Stage 26 was intercalated to designate the time of tooth eruption in C. 

johnstoni and C. porosus, an event which occurs around the same time as the 

appearance of sensory elevations on the scales. The teeth of A. mississippiensis 

rarely erupt before hatching, and if they do it is at Stage 28. 

Stage 27 [Figs. 21, 23, 26G, and Hl 

Stage 27 is characterized by the withdrawal of the remaining yolk into the 

abdominal cavity. It commences with the outgrowth of the ventral body 

Fig. 24. Alligator mississippiellsis. Development of the embryonic face. (A) Stage 13. Note the 

eye, club-shaped maxillary process, medial nasal process, lateral nasal process, nasal pit, 

nasal pit slit (arrowed), and notch in the center of the frontonasal process. (B) Stage 13. 

Ventral view. (C) Stage 13. Lateral view. The maxillary process extends beneath the eye and is 

continuous with the lateral aspect of the lateral nasal process. The branchial arches are amalgamated. 

(D) Stage 14. Ventral view. The medial nasal, lateral nasal and maxillary processes 

have merged. Note bulges for the primary palate and the anterior/dorsal extension of the 

nostrils. (E) Stage 15. Ventral view. The development of the anterior/dorsal bulges has displaced 

the external nares dorsally. The primary palate bulges and the maxillary processes are 

evident and the lower jaw is half-way beneath the upper jaw. (F) Early Stage 17 viewed from 

below. The lower jaw is behind the primary palate bulges. (G) Late Stage 17 viewed from 

below. Note the forward growth of the lower jaw. (H) Crocodylus Ililoticus. Stage 20. Lateral 

view illustrating the shift of the lower jaw beneath the primary palate, the notch in the upper 

jaw (arrowed) in the region of junction of the lateral nasal, medial nasal and maxillary processes 

(which will later become the notch for the large fourth dentary tooth), the elevated 

nasal disc, denticles, caruncle, lens, iris, eyelids and external ear flap. (I) Alligator mississippiellsis. 

Face-on view of the embryo shown in E. Note the development of the anterior nictitating 

membrane of the eye. (J) Stage 23. Oblique lateral view of the snout illustrating the 

elevations for the sensory papillae on the jaw margins, the ventral scales on the floor of the 

mouth and the submandibular odoriferous gland. (K) Stage 24. View the top of the head. Note 

the scalation. 0, denticles; E, caruncle; EN, external naris; G, submandibular odoriferous 

gland; L, lateral nasal process; M, medial nasal process; MD, mandibular process; MN, mandible; 

MX, maxillary process; N, notch in the frontonasal process; NB, nostril bulges; NO, 

elevated nasal disc; Nt nictitating membrane; NP, nasal pit; P, primary palate bulges; S, 

sensory papillae; SC, cervical scutes; V, ventral scales on the floor of the mouth.

Fig. 25. Alligator mississippiensis. (A) Stage 17. View of the inside of the mouth (lower jaw 

and tongue removed). Note the closed primary palate, the secondary palatal shelves, which 

have grown out from the maxillary processes on each side but which do not yet contact each 

other, the closing tectoseptal processes, caruncle and denticles. (B) Early Stage 18. Palatal 

view. The secondary palate is half closed. Note the V-shaped margins of the approximating 

palatal shelves. (C) Late Stage 18. Palatal view. The palate is three-quarters closed. (0) Stage 

19. Palatal view. The secondary palate and tectoseptal processes are almost completely closed. 

Note also the distinct anterior nictitating membrane of the eye and lower eyelid. (E) Early 

Stage 20. Palatal view. The palate is completely closed but the basihyal valve has not yet 

developed. (F) Stage 21. Palatal view. Note the development of the palatal plexus of blood 

vessels and the basihyal valve. (G) Stage 23. Palatal view. Note the extensive palatal plexus of 

blood vessels and the outlines of the sensory papillae along the jaw margins. (H) Stage 19. 

View of the tongue and lower jaw. Note the denticles and teeth, epiglottis and swellings of 

Fig. 26. Alligator mississippiensis. Views of the snout to show the development of the caruncle. 

(A) Late Stage 18. Frontal view. Caruncle is represented by two widely spaced elevations. 

The lower jaw has been removed. (B) Stage 19. Ventral view. Note that the two white blobs of 

the caruncle have joined in the midline. (C) Stage 20. The caruncle now appears more solid. 

(0) Stage 21. (E) Stage 23. Note the smooth base of the bifid caruncle and elevations for the 

pigmented sensory papillae (S) on the jaw margins. (F) Stage 25. Note that the smooth base of 

the caruncle is pigmented. (G, H) Ventral and frontal views of Stage 27. E, Caruncle; S, 

sensory papillae. 

the paired hypobranchial eminences which will form the lower flap of the basihyal valve. (I) 

View of the tongue and lower jaw of Stage 22. Note the plexus of blood vessels in the lower 

jaw and the lower flap of the basihyal valve. A, Anterior nictitating membrane; B, basihyal 

valve; E, epiglottis; Ee, caruncle; D, denticles; H, hypobranchial eminences; L, lower eyelid; 

PP, primary palate; P, secondary palate; T, tectoseptal processes.

416 


ORGANOGENESIS 

417 

wall to enclose the embryonic intestines, yolk sac and yolk; it ends with the 

absorption of the latter into the abdominal cavity. Ventral skin forms across 

the yolk scar. 

Stage 28 [Figs. 21 and 23] 

At Stage 28 the yolk scar diminishes in length and width as the absorbed 

abdominal yolk is utilized. There are considerable variations both among 

and within species as to the volume of absorbed yolk and the size of the 

yolk scar at hatching (which occurs at the end of this stage). These parameters 

may be influenced by the temperature of egg incubation (Ferguson and 

Joanen, 1982, 1983). The growth and changing proportions of embryos of 

A. mississippiensis from 12 days after egg laying to hatching are illustrated in 

Fig. 22. 

VII. 

ORGANOGENESIS 

A. Branchial Arches 

In Alligator mississippiensis (Parker, 1883; Clarke, 1891; Reese, 1908, 1912, 

1915a; Ferguson, 1984a), Crocodylus johnsoni (Ferguson, 1984a), C. porosus 

(Ferguson, 1984a), and C. niloticus (Voeltzkow, 1899), the five branchial 

arches appear in a cranio-caudal sequence (described in Section VI), beginning 

as a series of thickened epithelial involutions in the pharynx (Figs. 

27C, E, and G). Initially, these branchial pouches are internal grooves lined 

by thickened epithelium but they soon approximate the external branchial 

grooves. The pharyngeal endoderm fuses with the surface ectoderm to 

form true clefts in the dorsal half of the junctions between the 1st and 2nd 

and the 3rd and 4th arches (Ferguson, 1984a; Figs. 18, 19, 27C, 0, and G). 

In the remaining junctions, thin membranes of ectoderm, mesoderm, and 

endoderm separate the external grooves from the internal pouches (Ferguson, 

1984a; Figs. 18, 19, 27C, 0, and G). 

The first branchial cleft migrates dorsally until it overlies the otocyst near 

which it forms the external meatus of the ear (Ferguson, 1984a, Figs. 27H 

and I). Auricular hillocks develop along its ventral margin, which becomes 

the superior flap of the external ear (Figs. 27H and I). 

The third branchial cleft is transitory. Its epithelia become specialized 

and develop numerous cilia and cell processes, which extend from epithelial 

cells on one side of the cleft to those on the other (Ferguson, 1982b, 

1984a). In this way, the two epithelia oppose one another, fuse, and migrate, 

thus closing the branchial clefts (Ferguson, 1982b, 1984a). In fish, 

patent branchial clefts separate all the arches to form gills, whereas mammals 

lack patent branchial clefts (Sperber, 1981). The limiting membrane 

becomes the tympanum in the remodeling of the first branchial groove and 

pouch in mammals; tympanic development in crocodilians has not been 

reported. The significance of the transitory opening of the third cleft is 

unknown. The branchial sinus forms at the base of the most caudal branchial 

arch (Fig. 270); its specialized epithelium invades the underlying 

pharyngeal mesoderm (Ferguson, 1982b, 1984a). 

Each branchial arch is covered externally by ectoderm and internally by 

endoderm, which extends to the outer border of the mandibular arch (Fig. 

27G) and consists of a central core of mesoderm surrounded by a mass of 

mesenchymal cells derived from the neural crest (Ferguson, 1981a). Preliminary 

experiments, involving surgical excision or chemical destruction 

of alligator neural crest cells and heterotypic grafting of quail neural crest 

cells at different levels of the neuraxis (Ferguson, 1981a, 1982b, 1984a, 

unpubl.) suggest that neural crest cells migrate in temporally distinct 

waves from different levels of the neuraxis. Crest cells destined for the 

mandibular arch and its maxillary processes migrate from the rostral and 

caudal levels of the mesencephalon, respectively (Ferguson, 1981a). It is 

therefore possible selectively to block the migration of various waves of 

neural crest cells to produce alligators with normal maxillary processes and 

therefore a normal upper jaw and palate, but with virtually no lower jaw or 

tongue (Fig. 37E; Ferguson, 1981a, 1984a). This technique, combined with 

shell-less, or semi-shell-Iess culture, has enabled palatal development to be 

viewed as it happens; this is the first such longitudinal study in any animal 

(Ferguson, 1981a, 1982b, 1984a). 

Each branchial arch also contains its own aortic arch artery and appropriate 

cranial nerve (Figs. 18, 19, and 27G). The neural crest cells appear to 

give rise to all of the skeletal and connective tissues of the face, apart from 

striated muscle which is derived from the arch mesoderm; these preliminary 

data (Ferguson, 1981a, 1982b) indicate that the general pattern of 

neural crest derivatives in alligators is comparable to that in birds 

(LeOouarin, 1982). Macroscopic descriptions of branchial arch development 

and closure of the cervical sinus are given in Section VI and in 

Ferguson (1984a). Descriptive accounts of the branchial pouch derivatives 

and the cartilaginous, bony and muscular derivatives of the branchial 

arches exist for several species and include comparative comments with 

other vertebrates (Parker, 1883; Voeltzkow, 1903a,b; Reese, 1908, 1915a; 

Edgeworth, 1907; Goodrich, 1930; Pasteels, 1950; Oalcq and Pasteels, 1954). 

First branchial arches excised from Stage 10 alligator embryos can be 

cultured in vitro at 30°C and 37°C (Ferguson et al., 1981, 1983). In general, 

chemically defined, serumless media at 37°C are useful for short-term (up 

to 14 days) studies, whereas serum supplemented media at 30°C are useful 

for long-term (up to 60 days) studies. Explanted arches consist initially of 

lobulated cylinders of undifferentiated mesenchyme covered by epithelium 

(Fig. 27G). They undergo normal morphogenesis and differentiation 

in culture (Figs. 29A and B; Ferguson et al., 1982, 1983). Paired lingual 

swellings form a tongue-like structure. Blastematae for the Mm. genioglos

ORGANOGENESIS 

419 

sus, hyoglossus, and intermandibularis, the dentary, splenial, and angular 

bones, Meckel's cartilages, lingual lipid, and fibrous tissue appear and are 

patterned comparable to those seen in vivo (Figs. 29A and B; Ferguson et 

al., 1982, 1983). However, anlagen for taste buds and teeth are absent. The 

differentiation of alligator branchial arches in vitro is superior to that for 

any other vertebrate studied to date, so that they are useful models for 

investigation of a variety of developmental phenomena. 

B. Fses snd Noss 

There are accounts of the development of the face and nose for several 

genera: Alligator (Rathke, 1866; Clarke, 1891; Reese, 1901b, 1908, 1915a, 

1925; Bertau, 1935; Wettstein, 1954; Parsons, 1959a, 1970; Ferguson, 1981a, 

b, 1982b, 1984a), Caiman (Rathke, 1866; Parsons, 1970; Saint Girons, 1976), 

Melanosuchus (Bertau, 1935; Parsons, 1970) and Crocodylus (Rathke, 1866; 

Meek, 1893, 1911; Rose, 1893a, b; Seydel, 1896, 1899; Voeltzkow, 1899; 

Bertau, 1935; Wettstein, 1954; Wegner, 1957; Parsons, 1959a, b, 1970; 

Guibe, 1970m; Saint Girons, 1976; Bellairs, 1977; Ferguson, 1984a). That of 

Cavialis is unknown except for a figure in Bruhl (1866), an illustration by 

Bustard (1980c) and a description of the nasal excrescence (Martin and A. 

d'A. Bellairs, 1977). Butler (1905) notes that the snout of either Cavialis or 

Tomistoma is shorter and wider in the embryo than in the adult (but see 

(, 

Fig. 27. Alligator mississippiensis. Embryos. (A-D) and (F-G); Crocodylus lliloticus. (E-I). (A) 

Stage 7. Scanning electron micrograph (SEM) of the cranial aspect. Note the first and second 

branchial arches and first branchial cleft (arrowed). (B) Stage 8. Face-on view. Note the 

midline fissure and facial processes. (C) Stage 9. Scanning electron micrograph. Note the four 

branchial arches, grooves and clefts, nasal pit, limb buds, and tail. (D) Higher power view of 

C illustrating the true 1st and 3rd branchial arch clefts (arrowed), the four branchial arches 

and the branchial sinus. The apparent cleft between the 2nd and 3rd arches is an artifact. (E) 

Stage 10. Note the nasal pits and surrounding nasal processes, the lens and surrounding optic 

cup, the second branchial arch starting to overgrow arch 3, the 1st branchial cleft ventral to the 

otocyst and the brain regions. (After Voeltzkow, 1899.) (F) Stage 14. SEM illustrating closure 

of the nasal pit slits by the medial nasal, lateral nasal and maxillary processes. Note the 

intraoral bulges of the club shaped maxillary processes, signifying the onset of secondary 

palate development, the anterodorsal elevations of the medial nasal processes and the enlarging 

mandible. Compare with Figure D. (G) Stage 9. Horizontal hematoxylin and eosin section 

through the five branchial arches (1-5). Note the thick endoderm, thinner ectoderm, branchial 

grooves and pouches, aortic arch arteries,and branchiomeric nerves. (H, I) Stages 12 and 

15. Two diagrams illustrating the rearrangement of the first branchial cleft to form the external 

ear and superior ear flap. (After Voeltzkow, 1899.) A, Auricular hillocks; AA, aortic arch 

arteries; AD, anterodorsal elevations of nasal processes; BN, branchiomeric nerves; BP, branchial 

pouches; BS, branchial sinus; e, first branchial cleft; 0, denticles; E, eye; Ee, ectoderm; 

EN, endoderm; EX, external surface. F, ear flap; FL, fore limbbud; G, branchial grooves; H, 

hindbrain; HL, hind limbbud; LN, lateral nasal process; M, midbrain; MA, mandibular process; 

MD, mandible; MN, medial nasal process; MX, maxillary process; N, nasal placode; NP, 

nasal pit; 0, optic placode; Oe, optic cup; OT, otocyst; P, palatal shelves; PE, pericardial sac; 

PH, pharynx; 1, 2, 3, 4, branchial arches 1-4.

420 REPROOUCTIVE SIOLOGY ANO EMSRYOLOGY OF CROCOOILIANS 

Bustard, 1980c for a photograph of a Cavialis hatchling with a long thin 

snout). The literature on crocodilian nasal embryology has been reviewed 

by Parsons (1959a, 1970). The buccopharyngeal membrane initially separates 

the surface ectoderm from the pharyngeal endoderm (Parker, 1883). 

After its rupture, the primitive mouth is bounded by the forebrain above 

and the pericardial sac below. The differentiation of Rathke's pouch and 

the pituitary is discussed in Sections VII.I and P and for reptiles in general 

by Pearson, Chapter 9, this volume. 

Facial development in Alligator mississippiensis (Ferguson, 1981a, b, 

1982b, 1984a), Crocodylus johnsoni and C. porosus (Ferguson, 1984a) may be 

summarized as follows (for chronology, see Section VI). The first sign of 

nasal development is the appearance of bilateral epithelial thickenings on 

the anterolateral aspects of the forebrain bulge (Fig. 27A). These nasal 

placodes invaginate toward the roof of the primitive oronasal cavity and 

form the nasal pits (Fig. 27A). As the nasal pits invaginate, two rims of 

tissue are embossed outward around their surface openings to form the 

medial nasal and lateral nasal processes (Figs. 24B, 27B and F). The area 

between the two nasal pits (including the medial nasal processes of each 

side) is known as the frontonasal process and is divided by a medial groove 

(Fig. 24B). 

Bilateral maxillary processes arise from the dorsal ends of the mandibular 

arch (Figs. 24A-C, 27A, B, and F) and grow forward from the angles of 

the primitive mouth cavity to form progressively more of its cranial borders 

(Figs. 19, 24A-D, 27A, B, and F). The club-shaped maxillary processes 

form the lower boundaries of the developing eye (Figs. 19, 24A-D, and 

27F). Mesenchyme in the maxillary processes derives principally from the 

mesencephalic neural crest, whereas that in the medial and lateral nasal 

processes derives principally from the prosencephalic neural crest. The 

maxillary mesenchymal cells migrate along several routes. Extra-orally 

they migrate forward to form an ever increasing portion of the upper 

margin of the stomodeum; also, they pass between the developing eye 

and ear and between the developing eye and lateral nasal process, demarcating 

the naso-optic furrow between the lateral nasal and maxillary processes. 

The furrow eventually closes entrapping an epithelial rod, which 

canalizes to form the naso-Iacrimal duct. Intra-orally the maxillary processes 

have two principal advancing fronts of migrating mesenchymal 

cells, namely the tectoseptal processes and the palatal shelves (Section 

VILC). 

The invaginating nasal pits fuse with epithelium in the roof of the 

oronasal cavity so forming the primitive nasal cavities with anterior and 

posterior choanae. The posterior choanae open into the roof of the mouth a 

few millimeters behind the anterior choanae. By a combination of extremely 

rapid outward proliferation of the medial nasal and lateral nasal 

processes and a caudal extension of nasal pit invagination, the nasal pit 

slits develop between the ipsilateral medial and lateral nasal processes 

ORGANOGENESIS 421 

(Figs. 24A and B). The club-shaped maxillary processes come to lie beneath 

the nasal pit slits, which are eventually closed as the maxillary processes 

unite with the lateral and medial nasal processes (Figs. 240 and 27F). This 

uniting involves limited cell death and extensive migration of epithelial 

cells, which exhibit a characteristic cobblestoned, villous morphology. 

Throughout this period, differential growth of the head brings the nasal 

pits and related processes from their early lateral positions (Figs. 27A and 

B) closer to one another in the midline (Figs. 24A-D and 27F). 

The adjacent medial nasal processes develop paired intraoral projections, 

which merge to form the primary palate and primary nasal septum. 

Differential growth of extraoral caudal elevations of the medial nasal processes 

shifts the anterior nasal choanae from the tip of the snout to the 

adult dorsal position (Figs. 240 and E, 27F). The posterior nasal choanae 

now open behind the primary palate. The development of the secondary 

palate and associated structures is discussed in Section VII. C. 

Later, cartilage, bone, and muscle differentiate in various parts of the 

facial and nasal skeleton, the nasal epithelium becomes regionally specialized, 

the anlagen of the nasal glands and ducts appear, and the complex 

conchae and recesses, which characterize the crocodilian nose, begin to 

form (Bertau, 1935; Parsons, 1959a, 1970; Saint Girons, 1976). 

c. Palate snd Nasopharyngeal Duct 

Unlike any other reptiles, adult crocodilians have a mammal-like secondary 

palate composed of the sutured palatal processes of the maxillary, 

palatine and pterygoid bones (His, 1892; Moll, 1888; Busch, 1898; Gappert, 

1903; Voeltzkow, 1903b; Hoffman, 1905; Fuchs, 1907, 1908, 1910; Sippel, 

1907, Fleischmann, 1910; Thater, 1910; Lakjer, 1927; Barge, 1937; Peter, 

1949; A. d'A. Bellairs and Boyd, 1957; Pasteels, 1950; Guibe, 1970b; Iordansky, 

1973; Ferguson, 1981a, b, 1982b, 1984a). It is even unlike the avian 

palate, which is muscular and permanently cleft (Sippel, 1907; Barge, 1937; 

Pasteels, 1950; A. d'A. Bellairs and Jenkin, 1960). Crocodilian embryos are 

therefore useful in studies of normal and abnormal craniofacial development, 

largely because of their accessibility to experimental manipulations 

inside the egg (Ferguson 1979a, 1981a, b, 1982b, 1984; Ferguson and 

Honig, 1984; Ferguson et al., 1981). 

Some macroscopic drawings of the palates of embryos appear in His 

(1892), Voeltzkow (1899), Gappert (1903), Sippel (1907), and Fleischmann 

(1910), but no histological details of palatogenesis are given. More detailed 

studies of the development of the nasopharyngeal duct by Fuchs (1908) 

and Muller (1965, 1967) contain errors (see below; Ferguson, 1981b, 1984a). 

The following account is based on investigations of Alligator mississippiensis 

(Ferguson 1981a,b,d, 1982b, 1984a; Ferguson et al., 1984; Ferguson and 

Honig, 1984) and of Crocodylus johnsoni and C. porosus (Ferguson, 1984a). 

Description of the early development of the alligator face (Section VILB)

4ee 


indicates that the paired maxillary processes each develop two intra-oral 

migratory fronts of mesenchymal cells, namely the tectoseptal processes 

and the palatal shelves. 

The bilateral tectoseptal processes migrate superiorly between the floor 

of the brain and the roof of the oronasal cavity; eventually merging with 

one another along the midline in a progressive anteroposterior direction 

(Figs. 25A-O and 28B). Anteriorly, the merged tectoseptal processes grow 

downward to form the secondary nasal septum (Fig. 28A). 

The bilateral palatal shelves arise from the maxillary processes as thick, 

blunt-ended structures (Figs. 25A-O, 28A and B), which grow horizontally 

from their first appearance (Fig. 28A), except in the posterior one-fifth of 

the palate where they are more vertically oriented (Fig. 28B). With later 

development, these posterior shelves gradually flow over the tongue to 

become truly horizontal (Figs. 28C and 0). The reorientation involves the 

migration of mesenchymal and epithelial cells, the hydration of palatal 

glycosaminoglycans, the contraction of microfilaments, and differential 

mesenchymal proliferation. The shelves first approximate each other anteriorly 

behind the primary palate and from there closure spreads backward 

(Figs. 25A-O, 28C, and 0). Palatal closure occurs principally during Stage 

18 and is characterized macroscopically by a V-shaped gap along which the 

posterior margins of the opposing shelves approximate each other (Figs. 

25B and C, 28E). Palatal closure has been studied in normal cross-sectional 

sequences and also in ovo in the same embryo (longitudinal sequence) 

developing in a semi-sheIl-less culture (Ferguson, 1981a, 1982b, 1984). 

The process of palatal closure involves contact and adherence of the 

epithelial cells of the two shelves beginning on the oral aspect of the shelf 

margins and spreading nasally (Figs. 28C and 0). Cell death is limited to 

the small area of initial contact, after which the epithelial cells of the two 

shelves migrate nasally and posteriorly out of the region of closure (Figs. 

28C, 0, and E). There is never an extensive epithelial seam (Fig. 28C) and 

epithelial remnants cannot be detected following closure. Numerous small 

blood vessels in the shelf mesenchyme adjacent to the area of closure 

represent the earliest development of the palatal vascular plexus that 

characterizes late embryos (Figs. 24F and G) and adults. Mesenchymal 

continuity is usually established on the oral edges of the palatal shelves 

before the nasal edges are in contact (Fig. 28C). Anteriorly, the shelves 

Fig. 28. Alligator mississippiensis. (A) Stage 17. Mallory stain. Transverse section through 

head. Note the horizontal palatal shelves, bulge from the nasal septum, tongue, Meckel's 

cartilages, and intermandibularis muscle. (B) Stage 17. More posterior section of H & E and 

Alcian Blue stained specimen. Note the vertical secondary palatal shelves, closing tectoseptal 

processes, anlage of the anterior pterygoid muscle, Meckel's cartilages, anlagen for the intermandibularis, 

genioglossus, and hyoglossus muscles. (C) Stage 18. Transverse section 

through the closing secondary palatal shelves (Mallory stained). Shelf contact and mesenchymal 

continuity are established on the oral edges of the shelves, before the nasal edges have 

j~~V 

~:~fi 

..•••..• 

"~" 

F~, .' . 

,,»,,~!,.. ......,. 

.~ 

even contacted each other. Epithelial seam absent. The matrix stains positively for glycosaminoglycans 

and numerous blood vessels are present. (D) Stage 18. Scanning electron 

micrograph of palate. Note the closing palatal shelves, midline palatal elevation, primary 

palatal bulge, denticles, caruncle, and eyes. (E) SEM of the medial edge epithelial cells in the 

region of palatal closure seen in (D). Note the markedly cobblestoned appearance and numerous 

microvilli, both characteristic of cell migration. (F) Stage 18. SEM of the developing tongue 

and lower jaw. Note the paired lingual swellings, tuberculum impar, hypobranchial eminences, 

developing larynx and epiglottis, and denticles. The elevated hypobranchial eminences 

later form from the inferior flaps of the basihyal valve-see Figs. 25H and I. A, 

Anterior pterygoid muscle; B, bulge from the nasal septum; C, caruncle; 0, denticles; E, eye; 

G, genioglossus muscle; H, hyoglossus muscle; HB, hypobranchial eminences; I, intermandibularis 

muscle; L, lingual swellings; LA, larynx; M, Meckel's cartilages; MP, midline palatal 

elevation; P, palatal shelves; PP, primary palatal bulge; T, tongue; II, tuberculum impar; TS, 

tectoseptal process.

424 REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY 01= CROCOOILIANS 

merge with each other and with the bulging downgrowth of the secondary 

nasal septum (Figs. 28A and C) to produce a continuation of the partitioned 

nasopharyngeal duct (Fig. 28A). Posteriorly the palatal shelves merge to 

produce an unpartitioned duct (Fig. 28C), which is subsequently partitioned 

by the fusion of internal midline ductal bulges. Thus, the posterior 

nasal choanae are progressively displaced posteriorly as the secondary 

palate develops. The epithelial cells of the medial edges of the closing 

palatal shelves have characteristic surface topographies (Figs. 280 and E). 

Initially they are flat and rather featureless, but they become progressively 

more bulbous and develop surface microvilli and distinct cell boundaries so 

that by the time of shelf contact they are markedly cobblestoned and villous, 

an appearance characteristic of migrating epithelia (Figs. 280 and E). 

Medial edge, oral, and nasal epithelial differentiation, palatal closure, and 

mesenchymal differentiation occur in palatal shelves cultured in chemically 

defined or serum supplemented media (Figs. 29C,0, and E; Ferguson et 

al., 1984). 

Palatal development in alligators differs from that of mammals and 

birds. Initially, mammalian palatal shelves grow vertically downward, lateral 

to the tongue, and suddenly elevate to a horizontal position above the 

tongue; the opposing medial edge epithelia fuse in a seam and then die, so 

that mesenchymal continuity is established across the definitive secondary 

palate (Ferguson, 1977, 1978a, b). Avian palatal shelves, like those of 

crocodilians, are horizontal from their first appearance. The epithelia of 

Fig. 29. Alligator mississippiensis. Explants. (A) Macroscopic view of first branchial arch explanted 

at Stage 10 and organ cultured for 14 days in serum supplemented media at 30OC. 

Note the mandibular shape and the ventral aspect of the tongue tip. (B) Transverse section 

through an explanted first branchial arch after 36 days culture in serum supplemented media 

at 30°C. Note the tongue, Meckel's cartilages, and osteogenic blastemata for the lower jaw 

bones. (C) Scanning electron micrograph (SEM) of closing palatal shelves explanted from a 

20-day embryo and cultured for three days in chemically defined serumless media at 3TC. (0) 

Higher power view of the area of closure. Note the cobblestoned, villous, migrating medial 

edge epithelia, which are similar to those seen in uitro (Fig. 28E.). (E) SEM of the medial edge 

epithelia of a control recombination of alligator palatal epithelium and alligator palatal mesenchyme 

cultured for three days in media containing serum at 37°C. (F) SEM of a recombination 

of alligator mandibular epithelium on alligator palatal mesenchyme cultured for 3.5 days in 

media containing serum at 37OC. Note the differentiation into stratified squamous oral 

epithelia (0). cobblestoned, villous medial edge epithelia (M) and ciliated columnar nasal 

epithelia (N). (G) SEM of a recombination of mouse palatal epithelium on alligator palatal 

mesenchyme with the medial edges of the two opposed. Cultured for 3.5 days in media 

containing serum at 37°C. Note the oral. nasal. and medial edge epithelia. The latter is 

cobblestoned, villous and exhibits little cell death, a pattern typical of alligator (Fig. E. above), 

not mouse (see Fig. H.). (H) SEM of mouse palatal shelf cultured for 3.5 days in media 

containing serum at 37°C. Note the stratified squamous oral. ciliated columnar nasal epithelia, 

and the massive epithelial cell death along the medial edge (compare with Figs. E. and G.). M, 

Medial epithelium; ME, Meckel's cartilage; N, nasal epithelium; 0, oral epithelium; OB, 

osteogenic blastemata; T, tongue.


their medial edges keratinize, the palatal shelves fail to fuse, and thus 

produce physiological cleft palate (Sippel, 1907; Barge, 1937; Pasteels, 1950; 

Shah and Crawford, 1980; Koch and Smiley, 1981). 

The differing characteristics of the medial edge epithelial cells in the 

alligator (cobblestones, villous, migrating), chicken (keratinized stratified 

squamous) and mouse (cell death) suggest that palatal differentiation could 

be regulated by epithelial-mesenchymal interactions. To test this hypothesis, 

palatal shelves of alligators, chicks, and mice were separated into 

epithelia and mesenchyme, recombined in various heterotypic, homotypic, 

isochronic, and heterochronic combinations, and cultured. Additionally, 

palatal epithelia and mesenchyme were cultured in isolation and in 

combination with mandibular epithelia and mesenchyme, respectively 

(Ferguson and Honig, 1984). 

If palatal epithelium from alligator is cultured in isolation, it disintegrates 

after approximately three days, whereas palatal mesenchyme cultured 

in isolation differentiates into bony, cartilaginous, and muscular anlagen; 

control recombinations differentiate normally (Fig. 29E). Palatal 

shelf epithelium recombined with mandibular mesenchyme differentiates 

into typical stratified squamous mandibular epithelium. Conversely, mandibular 

epithelium recombined with palatal shelf mesenchyme differentiates 

into a typical palatal epithelium (Fig. 29F). These results suggest 

that, in the alligator, differentiation of palatal epithelium is regulated by 

an instructive epithelial-mesenchymal interaction. This interpretation is 

confirmed in recombinations between vertebrate species. Thus, alligator 

palatal epithelium recombined with a chick palatal mesenchyme exhibits 

the typical avian stratified squamous pattern of medial edge epithelial differentiation. 

Equally, alligator palatal epithelium recombined with mouse 

palatal mesenchyme exhibits massive medial edge epithelial cell death (Fig. 

29H). Contrariwise, either chick or mouse palatal epithelium recombined 

with alligator palatal mesenchyme shows differentiation characteristic of 

the alligator (Fig. 29G). These results show that the pattern of epithelial 

differentiation is regulated by the source of the mesenchyme (Ferguson 

and Honig, 1984). 

The alligator oronasal cavity is not as restricted as that of mammals, 

which may explain why alligator palatal shelves grow horizontally (Ferguson, 

1981a, b, 1982b). Further development of the palate involves the appearance 

and growth of osseous, muscular, and cartilaginous blastemata, 

the expansion of the palatal plexus of blood vessels, and the development 

of numerous domed tactile receptors (Figs. 3A and B) from subepithelial 

condensations of mesenchymal (Merkel) cells (Fig. 24]). Small outpouchings 

of the nasal cavity expand to form the maxillary sinuses, which 

enlarge laterally and medially to invade the palatal processes of the maxillae 

after hatching. 

The fibrous superior flap of the basihyal valve arises by a posteroinferior 

extension of palatal shelf closure (Figs. 25D-F). Crocodilians possess no 

true soft palate; the superior flap of the basihyal valve descends from 

ORGANOGENESIS 

427 

the palate to seal behind a lower, more rigid flap, that lies posterior to the 

tongue and is supported by the hyoid cartilage (Wood Jones, 1940). The 

superior valve flap is attached anteriorly to the posterior nasal choanae (in 

the pterygoid bone), runs parallel to the pterygoid palate for a short distance, 

and so forms a small area of nasopharynx between the pterygOid 

bony palate and the upper mucosa of the superior basihyal valve flap 

(Ferguson, 1981a). A failure to recognize the structure of the basihyal valve 

led Muller (1967) to misinterpret this space as a division of the nasopharyngeal 

duct (by a process of the pterygoid bone) into a "cavum 

ventrale" and a "cavum dorsale" (see her Figs. 9 and 10). Muller (1967) also 

confused the near simultaneous closure of the tectoseptal processes and 

secondary palatal shelves in crocodilians (see Ferguson, 1981a, 1984a). 

Very little is known about the development of maxillary, palatal and 

salivary glands (Rose, 1893b; Woerdeman, 1920; Reese, 1925; Barge, 1937; 

Fahrenholz, 1937; Kochva, 1978). The maxillary glands may arise from 

invaginations of the oral epithelium, closely related to the invaginating 

dental epithelium (Woerdeman, 1920). A review of the development of oral 

glands in Reptilia includes some data on crocodilians (Kochva, 1978). 

D. Tongue 

The tongue develops from three principal anlagen on the pharyngeal aspect 

of the branchial arches. The paired lingual swellings of the first arch 

form the anterior two-thirds of the adult tongue, whereas the midline 

tuberculum impar of the second and the paired hypobranchial eminences 

of the third form the posterior one-third (Ferguson, 1982b, 1984a; Figs. 25H 

and I, 27F). The epithelia of these anlagen come together and merge (Fig. 

27F). Behind the tongue lie the developing epiglottis, larynx, and pharynx 

(Fig. 27F). The inferior flap of the basihyal valve arises from a superior 

outgrowth of the paired hypobranchial eminences (Figs. 25H and I, 27F). 

Crocodilian lingual development resembles that of mammals and birds 

(portmann, 1950; Scott and Symons, 1974; Sperber, 1981). 

During palatogenesis, as in the adult, the tongue lies low in the oronasal 

cavity (Ferguson, 1981a,b, 1982b). The body of the tongue contains fibrous 

tissue anteriorly and lipid posteriorly, but intrinsic lingual musculature is 

absent (Ferguson, 1981a, b, 1982b, 1984a). Anlagen for the Mm. genioglossus, 

hyoglossus, geniohyoid, and intermandibularis develop, but their origins 

are poorly known. It has been suggested that all the lingual musculature 

develops from the geniohyoid anlage, which itself has split off from 

the ventral longitudinal muscle anlage formed by the 4th to 8th trunk 

myotomes (Edgeworth, 1907). Later stages of development of the lingual 

musculature have been illustrated by Humboldt (1807), Rathke (1866), 

Voeltzkow (1899), Gappert (1903), Taguchi (1920), Sewertzoff (1929), 

Wettstein (1954), and Ferguson (1981a, b, 1982b, 1984a). 

The lingual glands arise from thickened epithelia on the dorsum of the 

tongue. These glands secrete salt in the estuarine crocodile, Crocodylus

42B 

REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCOOILIANS 

ORGANOGENESIS 

429 

porosus (Taplin and Grigg, 1981), and in C. acutus and C. johnsoni but apparently 

not in Alligator mississippiensis or Caiman crocodilus (Taplin et al., 1982). 

Numerous taste buds develop on the tongue and in the floor of the mouth 

(Ferguson, 1981a, b, 1982b). As taste buds do not form in tongues which 

develop from excised mandibular arches, cultured in vitro, their induction 

and maintenance may be dependent on neural factors (Ferguson et al., 

1982, 1983a). 

E. Ear 

Bilateral auditory pits arise in the usual vertebrate fashion from ectodermal 

thickenings which invaginate to form the auditory vesicles (Rathke, 1866; 

Reese, 1908, 1915a). These thick-walled cavities lie close to the lateral walls 

of the hindbrain and open to the exterior. The development of parts of the 

middle ear has been studied in Alligator mississippiensis and compared with 

that of other reptiles and birds (Simonetta, 1956). The intertympanic canal 

is apparently a remnant of the vault of the embryonic pharynx, which is cut 

off from the main part of the pharynx by the backward growth of the 

posterior flange of the parasphenoid. A small posterior diverticulum (apparently 

homologous with the pharyngeal pouches of many mammals) is 

incorporated as a small appendix to the intertympanic canal (Simonetta, 

1956). The structure and development of the derivatives of the tubotympanic 

cavity, that is, the Eustachian tubes, the cavum tympani, the mandibular 

recess, and the epitympanic recess have been described by 

Simonetta (1956). The histogenesis of the crocodilian columella auris and 

other aspects of otic morphogenesis have been mentioned by Rathke 

(1866), Peter (1868), Hasse (1873), Van Beneden (1882), Parker (1883), Retzius 

(1884), Gadow (1888), Versluys (1903), Gaupp (1906), Shiino (1914), 

Goldby (1925), Kalin (1933), De Beer (1937), Wettstein (1937, 1954), Cordier 

and Dalcq (1954), Guibe (1970c), and Frank and Smit (1974). The cartilages 

form part of the chondrocranium and their structure and development 

have been reviewed previously in this series (Iordansky, 1973; A. d'A. 

Bellairs and Kamal, 1981). The structure and function of the crocodilian ear 

have been reviewed by Baird (1970), A. d'A. Bellairs (1971), and Wever 

(1978). 

F. EYB 

During early development the eye is a conspicuous feature of the crocodilian 

face (Figs. 18-20). Its development has been described by Rathke 

(1866), Voeltzkow (1899), Reese (1908, 1912, 1915a), Jolk (1923), and Wettstein 

(1954). The reptilian eye was reviewed by Underwood (1970), 

whereas Walls (1942), Rochon-Duvigneaud (1954, 1970), and Crescitelli 

(1977) reviewed vertebrate visual systems in general. All the available data 

suggest that the early morphogenesis of the crocodilian eye is essentially 

similar to that of other vertebrates, and several features are illustrated in 

Figs. 18, 19,20, 27A and F. 

The structure and development of the premandibular, mandibular and 

hyoid head cavities associated with the early embryology of the extrinsic 

ocular muscles in Alligator mississippiensis have been described by Wedin 

(1949, 1953, 1955). These cavities are interpreted as the myocoeles of somites. 

The origin, migration, and changing disposition of the ocular muscle 

anlagen are also described (see Table VII, Figs. 30A-D). The anlagen of 

TABLE VII 

The Origin of the Extrinsic Ocular Muscles in Alligator mississippiensis" 

Muscle Nerve Supply Origin 

The premandibular myocoele 

The superior rectus Oculomotor From the dorso-medial 

wall 

The inferior rectus 

The medial rectus 

} 

Oculomotor 

From a common anlage of 

the caudo-ventra-medial 

wall, above and slightly 

medial to the inferior 

oblique 

The inferior oblique Oculomotor From the caudo-ventra-lateral 

wall, below and a little 

lateral to the common 

anlage of the inferior and 

medial recti 

The mandibular myocoele 

The superior oblique Trochlear Rostro-dorsal to the premandibular 

myocmere 

The hyoid myocoele 

Caudo-dorsal to the pre 

mandibular myocoele, the 

common anlage being in a 

horizontal plane at right 

angles to the median plane 

The abducens complex 

comprising: 

The lateral rectus 1 

The r~tr~ctor palpebrae 

From a postero lateral portion 

of the common anlage 

Abducens 

supenons 

The retractor bulbi 1 

From an antero-medial 

The retractor membranae Abducens portion of the common annictitantis 

lage 

"Data from Wedin (1949, 1953); see Fig. 35.

430 

.' ,~i 

A 

REPRODUCTIVE SIOLOGY AND EMBRYOLOGY OF CROCODILIANS 

NIV 

! 

!. 

I b 

l_~ 

T 

B 

ORGANOGENESIS 

paper, the embryology of the crocodilian eye glands is completely unknown, 

although Rathke (1866) and Reese (1925) comment briefly on their 

structure in late embryos. 

G. Chondrocranium and Osteocranium 

The development and structure of the crocodilian chondrocranium and 

osteocranium have been reviewed respectively by Bellairs and Kamal 

(1981) and lordansky (1973) in this series. No experimental manipulations 

(d. Toerien, 1965, 1967; Toerien and Rossouw, 1977) have been performed 

on the developing crocodilian skull. 

H. Teeth 

431 

C 

Fig. 30. Alligator mississippiensis. (A) Diagrammatic figure of a 9-mm embryo illustrating the 

spatial disposition of the eye muscle anlagen relative to the eye, the premandibular myocoele 

(P.c., dotted) and various other cephalic structures. (After Wedin 1949, 1953.) (B) Schematic 

figure of the eye muscle anlagen in a 13.5-mm embryo. (After Wedin, 1953.) (C) Schematic 

figure of the eye muscle anlagen in a 20-mm embryo. (After Wedin, 1953.) (D) Schematic 

figure of the eye muscle anlagen in a 34-mm embryo. (After Wedin, 1953.) ABM, Abducens 

musculature; DP!, depressor palpebrae inferioris; G. CIL, ciliary ganglion; GV, trigeminal 

ganglion; 10, inferior oblique; IR, inferior rectus; LR, lateral rectus; MR, medial rectus; NIl, 

optic nerve; NIII, oculomotor nerve; NIV, trochlear nerve; NV!, abducens nerve; NO, 

notochord; OTO, otic vesicle; pc, premandibular myocoel; RB, retractor bulbi; RMN, retractor 

membrane nictitantis; RPS, retractor palpebae superioris; SO, superior oblique; SR, superior 

rectus. 

the muscles appear in the following sequence: (1) abducens, (2) superior 

oblique, (3) inferior oblique, (4) superior rectus, (5) medial and inferior 

recti, and (6) retractor membranae nictitantis (which arises much later than 

the others). Except for the fact that the inferior oblique arises before the 

superior rectus, this sequence coincides with that seen in the chick (Adelmann, 

1927). The first muscle anlage to appear is not the first to reach its 

final position (see Fig. 30; Wedin, 1953). Despite the title of Meek's (1893) 

D 

The first dental elements to form are rudimentary papillae called denticles 

(Rose, 1892a, b, 1894; Green, 1930; Deraniyagala, 1936, 1939; Edmund, 

1962, 1969; Figs. 25A-I, 280 and F). Green (1930) described them as resembling 

a selachian denticle and drew attention to the structural similarity 

between them and the egg tooth of Ornithorhynchus. The denticles arise as 

outgrowths from the oral epithelium and later differentiate into very small 

teeth, which make dentine but little or no enamel. They never become 

functional but sink into the mesenchyme and are resorbed. The dental 

lamina grows down near the posteromedial aspect of the trough of denticle 

epithelium, as the latter sinks beneath the oral epithelium (Westergaard 

and Ferguson, 1984 and in preparation). The sequence of appearance of the 

early tooth germs does not correspond to Zahnreihen sensu Edmund, 1962, 

1969 (Westergaard and Ferguson, 1984 and in preparation; also review in 

Westergaard, 1980). 

Subsequent development of the dentition was reviewed by Edmund 

(1962, 1969), and the few papers that have appeared since (Soule, 1967, 

1970; Miller, 1968; Miller and Radnor, 1970; Berkovitz and Sloan, 1979; 

Slavkin et al., 1982, 1984; Owens and Ferguson, 1982) emphasize the 

similarities between crocodilians and mammals. The stages of dental development 

are similar (Slavkin et al., 1984) and the enamel protein of Alligator 

mississippiensis is immunologically cross reactive with mammalian enamel 

protein (Slavkin et al., 1982). Unlike the teeth of other reptiles, those of 

crocodilians are attached in the jaws by a periodontal ligament, and both 

its development and that of root cementum are similar to those in mammals 

(Soule, 1967, 1970; Berkovitz and Sloan, 1979; Owens and Ferguson, 

1.982). Alligator tooth germs, explanted at the bell stage and cultured in 

vitro on chemically defined media, develop satisfactorily (Ferguson and 

Honig, unpublished data). Moreover, if these tooth germs are separated 

into the outer dental epithelium and inner dental papilla mesenchyme and 

then recombined with the equivalent structures from developing mouse 

molars, the recombinants produce enamel and dentine (Ferguson and


Honig, unpublished data). The harmonious development of chimaeric alligator/mouse 

teeth indicates that the reciprocal epithelial-mesenchymal 

interactions involved in amelogenesis and dentinogenesis can operate 

across vertebrate classes. These results reinforce the contention that alligator 

embryonic material is promising for the study of general problems 

in developmental biology. 

ORGANOGENESIS 

ec 

433 

I. Central Nervous System 

The development of the crocodilian central nervous system has been described 

by Rathke (1866), Parker (1883), Voeltzkow (1899, 1903a), Neumayer 

(1906), Reese (1908, 191Oa, 1915a), Wettstein (1937), and Dalcq and 

Pasteels (1954). Pasteels (1970) and Senn (1979) review the literature on 

Reptilia in general, whereas Wettstein (1937), Cordier (1954), and Anthony 

(1970) describe the structure of the adult central nervous system and comment 

on its development. 

The early development of the central nervous system, including the 

closure of the neural folds and the fate of the notochord, was described in 

Section V. The anterior end of the neural canal enlarges sequentially to 

form the vesicles of the forebrain, midbrain and hindbrain (Figs. 18-19). 

The cranial flexure develops so that the huge midbrain forms a conspicuous 

"bulge" at the back of the head (Figs. 18-21, 31A and B). The developing 

brain cavity is large and retort-shaped with comparatively thick 

walls of compactly arranged cells (Reese, 1908, 1915a; Fig. 31A). The walls 

of the forebrain are quite thick, especially anteriorly; the wall of the midbrain 

is somewhat thinner, especially in the region of the cranial flexure, 

and it is still thinner posteriorly over the hindbrain (Fig. 31A; Reese, 1908, 

1915a). Thus the dorsal wall of the hindbrain is reduced to a mere membrane, 

whereas the ventral wall is also thin, but the lateral walls are thicker 

(Reese, 1908, 1915a). The hindbrain is wider but not as deep as the forebrain 

(Fig. 31A). In the acute angle caused by cranial flexure is the anterior 

end of the notochord (Fig. 31A), which is distinctly vacuolated by Stages 2 

to 3. Commencing around Stage 5, the walls of the forebrain thicken bilaterally, 

later forming the cerebral hemispheres (Parker, 1883; Reese, 1908, 

191Oa, 1915a; Figs. 31A and B). 

Differentiation in the walls of the brain begins with the formation of an 

inner granular and an outer clear zone, and cranial nerve fibers develop 

from the base of the brain (Reese, 1908, 1915a; Van Campenhout, 1952). 

The median third ventricle has a thick ventral wall and a thin dorsal wall, 

extended to form a large paraphysis. The two lateral ventricles, the cavities 

of the cerebral hemispheres, have quite thick walls except on the side next 

to the third ventricle (Reese, 1908, 1915a). The forming sense organs are 

associated with various regions of the developing brain in a typical vertebrate 

fashion. Crocodilians apparently develop a white fibrillar stratum 

below the grainy layer of the cerebellum (Cordier, 1954), which itself arises 

j~~'PI1 

1.:.. 

.ti~ m~ 

B 

i'-~. 

Fig. 31. Alligator mississippiensis. (A) Sagittal section through the head of a 7-mm embryo 

illustrating the early appearance of the paraphysis (P), velum (V), cerebellum (C), hypophysis 

(H) and pharyngeal sac (PS). (After Neumayer, 1906, and Reese, 191Oa.) (B) Sagittal section 

through the head of a 13-mm embryo. Insert 1 shows a parasagittal section of the same 

embryo. Note the vesicular appearance of the paraphysis (which lies like a tube between the 

forebrain and superficial ectoderm) and the forward growth of the velum into the lateral 

ventricle, where it eventually forms the choroid plexus. Insert 2 shows the hypophysis at this 

stage of development. (After Reese, 191Oa.) C Cerebellum; CH, cerebral hemisphere; CP, 

choroid plexus; FB, forebrain; H, hypophysis; HB, hind brain; HS, hypophyseal stalk; I, 

infundibulum; IP, infundibular pit; LV, lateral ventricle; M, mandible; MB, midbrain; NA, 

neuroporus; NT, notochord; 01, 02, 03, lateral diverticula of hypophysis; OC, optic chiasma; 

P, paraphysis; PC, posterior commissure; PS, pharyngeal sac; PV, post velar arch; TP, tuberculum 

posterius; TTR, torus transversus; V, velum. 

from a fold between mid and hindbrains (Figs. 31A and B). Migration of 

nerve cells to the outer wall of the neopallium results in the formation of a 

cerebral cortex. 

The pineal (epiphysis) is absent (Voeltzkow, 1903a; Reese, 191Oa; 

Krabbe, 1939; Hamasaki and Eder, 1977), so that the circulating melatonin 

is presumably synthesized elsewhere, perhaps in the Harderian glands 

(Roth et al., 1980). The large paraphysis was mistaken for an epiphysis 

(Fig. 31) by earlier workers. The development of the paraphysis has been 

described by Voeltzkow (1903a) and Reese (191Oa). In alligator embryos 7 

mm in length (stage uncertain), it is first seen as a wide evagination of the 

roof of the forebrain, just cephalad to the velum, forming a transverse 

ventral depression of the dorsal wall of the forebrain (Fig. 31A). Posterior 

to the velum, the roof of the forebrain is slightly arched to form the begin

434 REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANB 

ning of the postvelar arch, which is limited posteriorly by a thickening of 

the dorsal wall, the future posterior commissure (Fig. 31A). Shortly after its 

appearance, the paraphyseal evagination becomes partially constricted 

from the brain and forms a rounded, hollow diverticulum, connected with 

the forebrain cavity by a wide opening (Fig. 31B). Later, the paraphysis 

elongates so that in embryos 7-cm long (and older), it is seen as a tubular 

structure, with thin smooth walls, slightly curved away from the cerebral 

hemispheres over the top of the diencephalon (Fig. 31B). Simultaneously, 

the velum increases in size and projects into the forebrain cavity as a 

transverse ridge, which ends in an acute posteriorly directed angle and a 

thicker obtuse angle projecting forward under the paraphysis (Fig. 31B). 

Subsequently, vesicles lined by cuboidal epithelium appear in the velum 

and form the choroid plexus which projects into the lateral ventricle (Fig. 

31B). In crocodilians, the fate of the cells that form the epiphysis in other 

animals is unknown. 

The hypophysis (pituitary) originates as an invagination of stomodaeal 

ectoderm (Rathke's pouch) beneath the floor of the forebrain (see Section 

VIlA and Figs. 27B, 31A and B). The pharyngeal sac is a smaller, less 

definite invagination of thickened epithelium (Figs. 31A, -8; Reese, 1910a), 

that develops caudal to the hypophysis and apparently disappears without 

trace in older embryos (Reese, 1910a), although it may be associated with 

the development of the pharyngeal tonsil (Killian, 1888). The original hypophyseal 

invagination becomes the stalk of a branched hollow structure, 

which, by lengthening of the stalk, recedes to some distance from the roof 

of the mouth (Fig. 31B) and loses its connection with the stomoda~al ectoderm 

and eventually loses its stalk (Reese, 1910a). During its development, 

the hypophyseal cavity consists of a central region and three outgrowths; 

the largest extending back until it nearly reaches the notochord, 

the second extending in the same direction from the base of the hypophyseal 

stalk, and the third extending toward the floor of the infundibulum 

(Fig. 31B; Reese, 1910a). These outgrowths become more numerous and 

the body of the hypophysis becomes completely solid as development 

progresses (Reese, 1910a). 

The body of the hypophysis is associated with a downward growth from 

the floor of the infundibulum (Fig. 31B). The adult crocodilian pituitary has 

three lobes, two of which originate from the ectodermal hypophyseal invagination 

and one from the infundibular downgrowth (Guibe, 1970e). 

Baumgartner (1916), Saint Girons (1970a, b), and Pasteels (1970) have described 

the embryology, morphology, and cytology of the reptilian pituitary, 

and these topics are considered elsewhere (Pearson, Chapter 9, this 

volume). The most comprehensive study of the development of the cranial 

nerves is that of Van Campenhout (1952) on Crocodlflus niloticus. Others 

(Shiino, 1914; Wettstein, 1937, 1954; Guibe, 1970d; Pasteels, 1970) provide 

occasional data on the development of the peripheral nervous system. 

The embryonic crocodilian central nervous system becomes functional 

ORGANOGENEBIB 

435 

about Stage 14. At this stage, whole body contralateral withdrawal reflexes 

can be elicited by gently touching the lateral flank or jaw margins or limbs 

of the embryo. As development progresses, such reflexes become more 

marked, more complex, and can be elicited for longer periods of time. At 

Stage 20, the body of the embryo may wriggle both away from and toward 

the stimulus, also the limbs may move. Jaw opening and closing reflexes 

can be elicited and the embryo exhibits spontaneous movements including 

swallowing. In general, the first reflexes or movements to appear embryologically 

can be elicited for progressively longer periods of time as development 

progresses. Moreover, when eggs are opened the most recent 

reflexes or movements to appear developmentally disappear first as the 

embryo dies, whereas the developmentally earliest movements are the last 

ones to disappear. What effects these activities have on the embryology of 

certain structures (such as joints and the gastrointestinal system) is unknown 

for crocodilians, but has been described in other animals (Humphrey, 

1968, 1969a, b, 1971a, b; Gottlieb, 1973; Oppenheim, 1973; Vince, 

1973; Freeman and Vince, 1974). 

oJ. 

VertebrBe Bnd Ribs 

Several accounts of the morphology and development of the crocodilian 

vertebral column and ribs are available (Voeltzkow and D6derlein, 1901; 

Shipley and McBride, 1904; Shiino, 1914; Higgins, 1923; Goodrich, 1930; 

Emelianov, 1937; Wettstein, 1937; Mookerjee and Bhattacharya, 1948a, b; 

Devillers, 1954a, b; Chiasson, 1962; Guibe, 1970f, g; Seidel, 1978). In the 

light of current knOWledge, some of these (e.g., Voeltzkow and D6derlein, 

1901; Higgins, 1923) may be misleading, but the most relevant recent reviews 

are those by Williams (1959) and Hoffstetter and Gasc (1969). 

The ribs arise between two adjoining vertebral segments shortly after 

the part-sclerotomes have formed a series of paired sclerotomes along the 

body axis; each rib is said to derive from the scleroblastic cells of both the 

original cranial and caudal part-sclerotomes (Higgins, 1923). The devel 

oping ribs become associated with the neural arches (Higgins, 1923; 

Emelianov, 1937). As development progresses, centers of chondrification 

appear within the membranous ribs and extend throughout their length. 

Whereas the developing ribs are associated with the vertebral column at 

o~e end, their main body lies between the epaXial and hypaxial muscles, 

~lth the exception of the first pair (or ribs of the atlas vertebra), which lie 

Internal to the hypaxial muscles and external to the peritoneal lining of the 

coelom (Mookerjee and Bhattacharya, 1948a, b). 

The gastralia or abdominal ribs are said to originate by direct ossification 

of membranous centers situated in the subcutaneous tissue outside the 

three abdominal muscles (Voeltzkow and D6derlein, 1901). They insinuate 

t?emselves between these muscles secondarily and apparently do not denve 

from ossification of the abdominal muscle tendons.


K. Respiratory System 

ORGANOGENESIS 

437 

The embryology of the upper respiratory tract is little known. Edgeworth 

(1907) suggests that the crocodilian laryngeal muscles form from an anlage 

in the second branchial arch, but gives no details. Soller (1931) described 

the hyoid apparatus, larynx and its musculature in late embryos of Alligator 

mississippiensis, Caiman crocodilus, and Crocodylus niloticus. The pharynx and 

trachea in late embryos of A. mississippiensis have been described by Moser 

(1902), Hesser (1905), Reese (191Oc, 1915a, b, 1926), Hilber (1932), Broman 

(1939), and Guibe (1970h). 

In 30-somite (Stage 4) alligator embryos, the lung primordia arise from 

the ventral pharynx just caudad to the branchial clefts (Reese, 1915b). Later 

(circa Stage 14), the trachea separates from the esophagus and divides into 

two bronchial primordia, both surrounded by mesoderm (Reese, 1915b; 

Broman, 1939). When the branchial clefts close, there is a temporary closure 

of the anterior end of the esophagus and trachea (Reese, 191Oc, 1915a, 

b, 1926); both subsequently regain their lumina, the trachea earlier than the 

esophagus. Lung rudiments each consist of an endothelial component divided 

into three main lobes (each subdivided into numerous lobules) and a 

mesodermal component with a smooth outline (Reese, 1915b). The lobules 

usually consist of a single layer of endoderm surrounded by a thin layer of 

flattened mesoderm in which lie the pulmonary vascular anlagen (Reese, 

1915b). Condensations of mesoderm around the trachea and bronchi differentiate 

into the cartilaginous rings and associated structures (Reese, 

1910c, 1915b). 

L. Cardiovascular System 

The primary organogenesis of this system is described by Reese (1908, 

1915a). The heart, lined by a distinct endothelium, appears first as a slight 

bulge to the right of the neural canal just anterior to the first somite. 

Vitelline vessels also occur at this time. The initially thin mesodermal wall 

increases in thickness as the muscle columns differentiate; they are particularly 

conspicuous ventrally in the presumptive ventricles. Further development 

of the heart has been described by Rathke (1866). Greil (1903), Shipley 

and McBride (1904), Felix (1906a), Hochstetter (1906a, c). Kerr (1919), 

Goodrich (1930), Stephan (1954), Wettstein (1954), White (1968, 1970), 

Springer (1970), and Guibe (1970i). Hochstetter's (1906a) account is the 

most comprehensive and Goodrich (1930) described crocodilian cardiac 

development in a phylogenetic context. 

Although the heart of most reptiles is three chambered, that of crocodilians 

is four chambered, due to the development of a complete interventricular 

septum (Beddard and Mitchell, 1895; Shaner, 1924; Goodrich, 1930; 

Wettstein, 1954; Foxon, 1955; Bellairs and Attridge, 1975; Webb, 1979). The 

evolution of the crocodilian and avian interventricular septum have been 

FP 

VI 

v 

J I . 

DA 

Fig. 32. Heart and aortic arches of a 

crocodilian embryo, diagrammed from the 

ventral aspect. I, II, III. IV. V, VI. First to 

sixth aortic arches; C, coeliac artery; CC. 

common carotid artery; DA, dorsal aorta; 

DC, dorsal carotid artery (right side nearly 

atrophied); FP, Foramen of Panizza; LA, left 

aortic arch; P, pulmonary artery; RA, right 

aortic arch; VC, ventral carotid artery; VS, 

ventral subclavian artery (Modified from 

Shipley and McBride. 1904; Shindo, 1914; 

Goodrich, 1930; De Beer. 1959; Pasteels. 

1970.) 

considered in detail (Webb, 1979). The septum develops by the inward 

growth of the edge of the interventricular anlage from behind and above, 

from the septum aorticum (between the right and left systemic trunks), 

from the front, and also from the endocardial rudiment of the right septal 

valve (Hochstetter, 1906a; Goodrich, 1930). Although manifesting this "advanced" 

cardiac condition (similar to that in birds) in its early development, 

the crocodilian heart still shows the sharp, double flexure of the 

conus seen in dipnoans (Hochstetter, 1906a; Kerr, 1919). Likewise, the 

lateral valve of the right atrioventricular orifice becomes muscular; this 

feature is absent in mammals, but exaggerated in birds (Shaner, 1924; 

GOodrich, 1930). Crocodilians retain the left aortic arch (Fig. 32), which 

only OCCurs as an abnormality in birds (Goodrich, 1930). Thus, in crocodilians, 

the right aortic arch (carrying oxygenated blood) arises from the left 

Ventricle, whereas the left aortic arch and pulmonary trunk arise from the 

right ventricle (Fig. 32). The right and left aortic arches communicate via 

the foramen of Panizza (Fig. 32), which is located near the base of the 

Systemic trunks, just anterior to the semilunar valves (Shaner, 1924; Kerr, 

1919; Goodrich, 1930; Webb, 1979). This foramen apparently arises as a 

secondary perforation of the aortic septum comparatively late in development, 

just before the closing of the interventricular foramen (Grei!, 1903). 

The development of the arterial and venous systems has been described 

by Balfour (1881), Zuckerkandl (1895a, b), Voeltzkow (1901), Shipley and

143S 

REPRODUCTIVE BIOLOGY AND EMSRYOLOGY OF CROCODILIANS 

cBride (1904), Hochstetter (1906a, c), Reese (1908, 1915a), Shiino (1914), 

Shindo (1914), Kerr (1919), Goodrich (1930), Stephan (1954), Wettstein 

(1954), DeBeer (1959), Guibe (1970i), and Bellairs and Attridge (1975), The 

fates of the embryonic aortic arches are illustrated in Fig. 32. The right 

dorsal carotids atrophy almost completely, and frequently the left common 

carotid artery gives rise to the dorsal and ventral carotids of both sides. The 

right and left 4th aortic arches fuse to form the dorsal aorta, whereas the 

6th aortic arches become the pulmonary arteries (Fig, 32). The anterior 

cardinal veins lie adjacent to the notochord and unite with the posterior 

cardinal veins to form the Ducti Cuvieri, which in turn connects with the 

meatus venosus of the liver (Reese, 1908, 1910c, 1915a). The adult lymphatic 

system was described by Ottaviani and Tazzi (1977), but its development 

is poorly understood. 

M. Diaphragm 

Crocodilians are unique among reptiles in having thoracic and abdominal 

cavities separated by a structure analogous to the mammalian diaphragm, 

but which has been little studied (Butler, 1889; Hochstetter, 1906b; Goodrich, 

1930). The diaphragm apparently develops from the secondary fusion 

of many parts: the posthepatic septum (including the ventral oblique hepatic 

ligament), the ventral pulmonary fold, and parts of the gastric mesentery. 

Along its edge, the diaphragm is associated with many of the back 

muscles. The large diaphragmaticus muscle runs from the liver to the 

pelvis and assists the costal breathing mechanism by retracting the liver 

and expanding the pleural cavity by stretching the diaphragm thus powering 

inspiration (Gans and Clark, 1976). The diaphragm also separates off 

eight coelomic recesses and cavities (Hochstetter, 1906b). 

ORGANOGENESIS 

_............ - .......\ 

--- 

,/ 

\ 

,,/ , 

, , " « 

, I 

I I 

P 

°n/Nf))';:' 

'--"', .: 

: "~ ('.~ \ )' 

: " . "'"J .••' ./ 

I 

, •.•••' \ D 

PG 

5 : ..... ,..::..;';, ....•. 

, . 

LU~.!.e.;..,.\ \ 

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T 

439 

Fig. 33. Alligator mississippicllsis. Diagram 

of a reconstruction of the embryonic gastrointestinal 

system approXimately 3 weeks after 

egg deposition. Digestive canal shown 

in solid black, glands in solid lines and em. 

bryo outline in dotted lines (after Reese 

1910). A, Allantois; C, cloaca; D, 

duodenum; E, eye; H, hindgut; L, liver; 

LU, lung; M, mouth, O. oesophagus; P, 

pharynx; PA, pancreas; PC, post-anal gut 

of Reese (1910); PL, posterior limb bud; S, 

stomach; T, trachea; Y, yolk sac. 

N. Gastrointestinal System 

The most extensive accounts of the development of the gastrointestinal 

system are those of Reese (191Oc, 1913c) but some comments are given by 

Rathke (1866), Clarke (1891), Voeltzkow (1899), Reese (1908, 1912, 1915a, 

1926), Grunwald (1931), Wettstein (1954), Elias (1955), Arvy and Bonichon 

(1958), Guibe (1970b), Gabe (1970b), Miller and Lagios (1970), and Skozylas 

(1978). 

After early development (Section V), the foregut is a shallow enclosure 

of the anterior endoderm (Figs. l3C and D) and the blastopore connects the 

hindgut region with the exterior. Subsequently (Figs. 14 and 15), the foregut 

becomes a cavity wider laterally than dorsoventrally and extends only 

to about the cranial third of the trunk, opening onto the yolk sac (Reese, 

191Oc). With the appearance of the definitive hindgut, the anterior and 

posterior intestinal portals are identifiable where the unenclosed midgut 

joins the foregut and hindgut (Reese, 191Oc).

440 REPRODUCTIVE BIOLOGY AND EMBRYOLOGY OF CROCODILIANS ORGANOGENESIS 

The liver first appears as an endodermal diverticulum from the ventral 

foregut surrounded by splanchnic mesoderm (Fig. 33; Reese, 1910c; Elias, 

1955). A small projection from the wall of the duodenum differentiates into 

the bile duct, which ends blindly on the anteroventral edge of the liver and 

is closely related to the pancreatic duct. The pancreas derives from one 

dorsal and two ventral diverticula of the foregut, which evaginate just 

anterior to the hepatic rudiments (Reese, 1910c; Gabe, 1970b; Miller and 

Lagios, 1970). Their subsequent fate is unknown, but in the late embryo 

the pancreas lies between the posterior end of the stomach and the anterior 

end of the duodenum and opens into the latter by two or more short ducts 

(Reese, 1910c). 

o. UrogenitBI System and Sex Determination 

The literature on the embryology and structure of crocodilian urogenital 

systems is large (Remak, 1845, 1855; Rathke, 1866; Wiedersheim, 1890a, b, 

1891, 1897; VoeItzkow, 1892, 1899; Wilson, 1896, 1900; Szakall, 1899; Felix, 

1906b; Loyez, 1905, 1906; Reese, 1908, 1912, 1915c, 1924; Moens, 1912; 

Taguchi, 1920; Forbes, 1934, 1937a, b, 1938a, b, c, 1939, 1940a, b, 1961; 

Gerrard, 1954; Wettstein, 1954; Godet, 1961; Forsburg and Olivecrona, 

1963; Ramaswami and Jacobi, 1965; Guibe, 1970j; Pasteels, 1970; Fox, 1977; 

Ferguson and ]oanen, 1982, 1983; Webb and Smith, 1984) and has been 

reviewed by Fox (1977). Here follows a more detailed account of events in 

crocodilians with particular emphasis on data related to temperature dependent 

sex determination. It is based on a number of publications and on 

my observations on Alligator mississippiensis, Crocodylus johnsoni, and C. 

porosus. Good descriptions and illustrations of the developing gonads and 

Mullerian ducts in A. mississippiensis are available (Forbes, 1940a) as are 

data on the histology of hatchling gonads and reproductive ducts (Ferguson 

and ]oanen, 1983). Histological data are also available for hatchling C. 

johnsoni (Webb and Smith, 1984). 

The first indication (at approximately Stage 1) of the development of the 

excretory system is the appearance of approximately six ciliated tubules 

opening into the anterior coelom (Wiedersheim, 1890a, b, 1891). On either 

side of this pronephros a large vascular coil covered by coelomic epithelium 

appears and forms the muItilobate glomus. Later, the pronephros 

degenerates as the mesonephros originates from nephrotomes 11-23 

(which begin posterior to the pronephros and extend to the anterior cloacal 

border). There is no clear demarcation between pronephros and 

mesonephros (Wiedersheim, 1890a, b, 1891). The mesonephros develops 

branched tubules (with internal Malpighian corpuscles) which do not open 

into the coelom (Wiedersheim, 1890a, b, 1891; Reese, 1908, 1912, 1915a). 

They function as collecting ducts through embryogeny and for some time 

posthatching (Forbes, 1940a), but they ultimately become transformed into 

epididymal structures in the male or disappear in the female. The mesonephric 

duct arises from a mesodermal blastema closely related to the 

pronephros (Reese, 1908, 1912, 1915a) and grows posteriorly receiving 

mesonephric tubules as they form. The same coelomic epithelium, that 

gave rise to the mesonephros, also forms the metanephros. The metanephros 

develops convoluted tubules, glomeruli, and gives origin to the 

ureters, which join the mesonephric ducts before the latter open into 

the cloaca (Forbes, 1940a; Fox, 1977). 

Forbes (1940a) described three phases in gonadal development in Alligator 

mississippiensis: the period of genital ridge formation, the period of 

bisexuality, and the period of visible sex differentiation. The following 

summary derives primarily from that study. No information is available on 

the origin and migration of the germ cells in crocodilians. The genital ridge 

is seen first as a thickening of the coelomic epithelium on the ventral and 

ventromedial surface of the mesonephros, continuous medially with the 

primordium of the adrenal cortex. This epithelium becomes organized into 

primary sex cords, rete cords, and germ cells. Embryos of 75-88-mm headtrunk 

length (exact stages uncertain) show a period of bisexuality when 

both male and female primitive gonadal components are present (Forbes, 

1940a, 1964). The gonad has a well-defined outer layer of simple germinal 

epithelium and a subjacent zone of medullary cords. Subsequently, visible 

sex differentiation takes place. In ovaries, a well-defined germinal epithelium 

develops in which the germ cells are fairly numerous, whereas the 

medullary cords regress and are distended by large irregular cavities. The 

testes retain limited cortical areas external to the tunica albuginea, whereas 

the medullary cords differentiate as seminiferous tubules containing germ 

cells. Both males and females exhibit a degree of embryonic bisexuality; 

immature females retain Wolffian ducts and a small mass of testis-like 

medullary tissue (the medullary rest) in the ovary; immature males may 

retain variable testicular cortical areas and occasional Mullerian duct segments 

(Forbes, 1934, 1937a, b, 1938a, b, c, 1939, 1940a, b, 1961, 1964; 

Ferguson and ]oanen, 1983). 

Controversy regarding the origin of the Mullerian ducts was reviewed 

by Fox (1977: 52); the generally accepted view is that of Forbes (1940a) and 

Forsburg and Olivecrona (1963). In the female, the Mullerian ducts become 

the OViducts, whereas in the male they remain until around the time of 

hatching when they disintegrate except for short rudimentary segments, 

which may persist throughout life (Forbes, 1940a, Ferguson and ]oanen, 

1983). Vestiges at the anterior poles of the testes are common and apparently 

homologous with the mammalian appendix testis (Forbes, 1940a). 

There is no information available on the development of oviducal glands 

(Such as albumen secreting and shell glands). 

With degeneration of most of the embryonic mesonephros, the Wolffian 

duct has little role in excretion, but it persists in both sexes (Ferguson and 

Joanen, 1983). In males, the ends of the Wolffian ducts closest to the testes 

are mUch coiled and form the epididymis in conjunction with persistent 

44'1


mesonephric tubules. The posterior Wolffian ducts form the ducti deferens, 

which open into the cloaca near the base of a groove that runs along 

the upper surface of the single penis (Moens, 1912; Reese, 1924; Neal and 

Rand, 1936; Bellairs and Attridge, 1975). The development of the crocodilian 

clitero-penis has been described by Rathke (1866), Voeltzkow (1892, 

1899), Moens (1912), and Reese (1924). It grows out from the ventral abdominal 

wall, anterior to and distinct from the cloaca. As the cloacal lips 

develop, the clitero-penis withdraws between them and differentiates into 

its adult component parts (Figs. 34A-H). Anal musk glands and abdominal 

pores arise by epithelial invagination (Reese, 1921, 1924). 

Although sex is fully determined and the gonads are differentiated at 

the time of hatching in all crocodilian species investigated (Ferguson and 

Joanen, 1982, 1983), the degree of differentiation of the external genitalia is 

variable. Sexing by cloacal examination is impossible in Alligator mississippiensis 

(Joanen and McNease, 1978) or in Cavialis gangeticus (Singh, personal 

communication) less than 0.6 m in length. However, male and female 

Crocodylus porosus, C. johnsoni, and C. niloticus can be distinguished at 

hatching (Webb et aI., 1983c; Hutton, personal communication). These 

differences may be the result of the shorter incubation times of some 

species, for example, A. mississippiensis (see Table III) as suggested by 

Webb et a1. (1983c), but this would not explain the etiology in C. gangeticus 

and possibly other crocodilians. Striking differences in the degree of differentiation 

of the external genitalia and in the size of the latter are present 

from Stage 7 and marked after Stage 14 (Figs. 34A-H) between A. mississippiensis, 

C. johnsoni, and C. porosus. Thus, development of the external 

genitalia may be different from the onset in alligators and crocodiles. The 

influence of incubation temperatures on the development of the external 

genitalia is unknown, as is the influence of the degree of differentiation of 

the external genitalia at hatching (and their subsequent rate of development) 

on the age or size at which sexual maturity is attained in various 

crocodilians Cfable I). Perhaps the onset of gametogenesis is only one of 

the factors that determine sexual maturation. 

In Alligator mississippiensis, both laboratory and field experiments have 

shown that incubation temperature determines the sex of the hatchling 

and hence of the adult (Ferguson and Joanen, 1982, 1983), and this is 

confirmed by preliminary data for Crocodylus johnsoni (Webb et aI., 1983e; 

Webb and Smith, 1984), C. porosus (Webb, personal communication) and C. 

niloticus (Hutton, personal communication). Temperature dependent sex 

determination is Widespread among turtles and lizards (Bull, 1980). That 

heteromorphic sex chromosomes appear to be absent from all living 

crocodilians studied to date (21 out of the 29 species) (Cohen and Gans, 

1970) makes it likely that all crocodilian species have a temperature dependent 

mechanism. Female alligators are HY pOSitive, similar to snakes and 

turtles (V. Lance, personal communication). 

In Alligator mississippiensis, incubation of eggs at 340C produces only 

Fig. 34. Crocodylus niloticus. Drawings of the development of the external genitalia (after 

Voeltzkow, 1899, 1901). (A, B) Stage 10. Ventral and lateral views. (C, D) Stage 13. Ventral and 

lateral views. (E, F) Stage 19. Ventral and lateral views. (G, H) Stage 25. Ventral and lateral 

views. (I) Higher power view of the cross-section of the umbilical stalk seen in E. A, Allantois; 

G, gut loops; GA, genital anlage; GB, genital bulb (glans in male); GF, genital fold; GS, shaft of 

genital primordium; L, limb buds; 0, omphalo-mesenteric artery; U, umbilicus; UA, umbilical 

artery; UV, umbilical vein. 

I


ORGANOGENESIS 

445 

males, whereas incubation at 30°C or below produces only females (Ferguson 

and Joanen, 1982, 1983). At 32°C, the sex ratio varies from 87% females: 

13% males to 100% males, whereas the sex ratio is approximately 

50% females:50% males at 31°C (Ferguson and Joanen, 1982, 1983, unpublished 

observations). Similarly, in Crocodylus niloticus (Hutton, personal 

communication) incubation at 34°C produces males, and at 30°C, it produces 

females. In C. porosus (Webb and Smith, 1984; Webb, personal communication) 

incubation at 32°C produces 100% males and at 30°C 100% 

females; 34°C incubation causes a high mortality of C. porosus eggs. Despite 

this apparent uniformity amongst species, there is at least one exception. 

In C. johnsoni, incubation at 34°C produces 100% females, at 31°-33°C 

males (in various ratios), and at 26°-30°C 100% females (Webb and Smith, 

1984). This bimodal pattern of production of females at both high and low 

temperatures and males at intermediate temperatures is similar to the situation 

in Chelydra serpentina (Yntema, 1976, 1979), which itself contrasts to 

the normal turtle pattern in which high temperatures produce females and 

low temperatures produce males (Bull and Vogt, 1979, 1981; Bull, 1980). In 

all experiments differential embryonic mortality has been eliminated as a 

factor producing the skewed sex ratios, which have also been recorded in 

natural nests in different environments (Ferguson and Joanen, 1982, 1983; 

Webb and Smith, 1984) and in adult populations (see Table II). 

In an attempt to define the temperature sensitive period (TSP) for sex 

determination, experiments have been performed involving the shift of 

eggs from an all male to an all female producing temperature and vice 

versa at various times of development. If eggs of Alligator mississippiensis 

are shifted from 30° to 34°C, the TSP is between days 14 and 21 (Stages 13 

16), but if eggs are shifted from 34° to 30°C, the TSP is between days 28 and 

35 (Stages 20-21); (Ferguson and Joanen, 1983). Thus, the temperature of 

egg incubation prior to the TSP (run-in temperature) does influence sex 

determination but not irreversibly. The relationship is opposite to what 

one might expect in that the higher run-in temperature (which accelerates 

macroscopic external development) delays the TSP and embryos remain 

sexually labile for much longer. 

In Crocodylus porosus (Webb, personal communication) shifts from 30° to 

32°C give a TSP between 20 and 25 days (Stages 15-18), whereas shifts 

from 32° to 30°C give a TSP between 40 and 45 days (Stage 23). The TSP of 

C. johnsoni is poorly documented, but preliminary data for shifts from 31° 

to 30°C put it between 20 and 30 days (Stages 15-20) (Webb and Smith, 

1984). The embryonic stages during the TSPs both for switches of high to 

low or low to high temperature are remarkably similar in these three 

species. Moreover, the stages are similar to the equivalent ones for Chelydra 

(Yntema, 1968); the TSP for shifts of the eggs of emydid turtles from 25°C 

(male producing temperature) to 31°C (female producing temperature) occurs 

at Stage 16 (Yntema, 1968), whereas the TSP in shifts from 31° to 25°C 

is Stage 22 (Bull and Vogt, 1981). Double shift experiments in which a pulse 

of one temperature occurs on a background of the other revealed that the 

time required to produce males (25°C) is less than to produce females 

(31°C) (Bull and Vogt, 1981). Therefore in Alligator mississippiensis, C. 

porosus, and emydid turtles, cooler temperatures act earlier and more rapidly 

in determining sex (females in crocodilians, males in turtles). 

The situation is more complex in biomodal species with two temperature 

thresholds. No data are available for Crocodylus johnstoni but the TSPs of 

Chelydra serpentina are between Yntema Stages 16 and 20 for shifts from 26° 

to 30°C, Stages 12 and 15 for shifts from 30° to 26°C, Stages 13 and 17 for 

shifts from 20° to 26°C, and Stages 12 and 18 for shifts from 26° to 20°C 

(Yntema, 1979). Thus, it appears that less incubation at 30°C (to Yntema 

Stage 16) is required to determine femaleness than is required at 26°C (to 

Yntema Stage 19) to determine maleness. Embryos of C. serpentina, incubated 

at the male producing temperatures of 22° or 24°C must be exposed 

to 30°C for at least four hours per day in order to ensure female 

development (Wilhoft et al., 1983). 

To date the data on temperature sensitive periods for sex determination 

(in both crocodilians and turtles) have remained puzzling and no one has 

satisfactorily explained them or assimilated them into any model of the 

mechanism for temperature dependent sex determination. Part of the reason 

for this may have been the tendency to assume that the TSP equates 

with the time of embryonic sex determination (which would therefore have 

to occur over at least 2 to 3 widely different stages of development). This 

assumption may be erroneous, and the concepts and nomenclature of TSPs 

and even temperature-dependent sex determination itself may be misleading, 

and the term gonadal size-dependent sex determination may reflect 

more accurately the underlying biological processes. A new theory regarding 

these phenomena has been proposed (Ferguson, submitted). 

P. Endocrine Glands 

The development of the reptilian pituitary gland is reviewed by Pearson 

(Chapter 9, this volume); that of crocodilians is discussed in Section VII.I. 

The development of the thyroid gland resembles the basic vertebrate 

pattern. With the approximation of the mandibular arches, a deep narrow 

groove is formed in the anterior floor of the pharynx; it is uncertain 

Whether this location implies a first pouch origin. The epithelium of the 

caudal part of this groove thickens ventrally and invaginates to form the 

anlage of the thyroid gland. The gland becomes cut off from the pharynx 

and migrates caudally in the form of solid cords, which break up into 

groups of cells that form the primary follicles. These then become encapsulated 

to form the adult gland (Reese, 1908, 191Oa, c, 1915a, 1931b; Lynn, 

1960. 1970; Gabe and Saint Girons, 1970a).

qq& 

REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCODILIANS 

The parathyroids, like those of reptiles in general, are derived from 

branchial pouches three and four (Gabe and Saint Girons, 1970b). Van 

Bemmelen (1886, 1888) described a single pair of parathyroids derived from 

pouch three in late embryonic and adult specimens of Alligator mississippiensis 

and Crocodylus porosus. Hammar (1937) observed two pairs of 

parathyroid glands in young embryos of C. porosus, but noted that the pair 

from pouch four disappeared in adults, a finding supported by the work of 

Clarke (1970) on Caiman crocodilus. The glands derived from pouch four are 

small, and it is probable that they fuse with those from pouch three during 

late development (Clarke, 1970). The parathyroids are not associated with 

the thyroid glands but are located in or near the thymus, in dose proximity 

to their embryonic relationship with the third and fourth aortic arches. 

In Alligator mississippiensis, the primitive adrenal cortex arises from a 

thickened mass of coelomic epithelium lying on the ventromedial surface 

of the mesonephros and extending laterally from the dorsal mesentery to 

the genital primordium (Forbes, 1940a; Gabe, 1970a; Martoja, 1970). 

Shortly after its appearance, buds arise and extend rapidly as cortical cords 

into the mesenchyme between the mesonephros and aorta. Adrenal medullary 

tissue appears as aggregations of basophilic cells in the mesenchyme 

dorsal to the cortical cords and medial to the mesonephros. It migrates 

ventrally into the cortical mass (Lawton, 1937; Forbes, 1940a). 

G. Thymus and Immune System 

The observations available on the thymus are mainly topographic descriptions 

of its size and position in late embryos and young adults, in which it 

occupies much of the total length of the neck (Rathke, 1866; Van Bemmelen, 

1888; Hammar, 1909; Reese, 1908, 191Oc, 1915a; Pischinger, 1937; 

Wettstein, 1954; Bockman, 1970). Histological descriptions of the early thymus 

report nothing about its development (Dustin, 1914; Taguchi, 1920; 

Wettstein, 1954; Bockman, 1970). The crocodilian thymus, like that in other 

reptiles, derives from the branchial pouches (Reese, 191Oc), but precisely 

which ones is unknown (Bockman, 1970), although the second has been 

suggested (Reese, 191Oc). Study of the development of the structure and 

function of the crocodilian thymus, which is known to be involved with 

adaptive immunity (Bockman, 1970), would be of great interest as a comparison 

with the avian immune response, which involves the unique Bursa 

of Fabricius. Bursa equivalents in crocodilians remain to be investigated. 

R. Limbs and Tail 

Aspects of the development of crocodilian limbs have been described by 

Rathke (1866), Kiikenthal (1893), Zuckerkandl (1895a, b), Voeltzkow 

(1899), Goeldi (1900), Braus (1906), Reese (1908, 1915a), Kerr (1919), Tornier 

(1928), Goodrich (1930), Steiner (1934), Wettstein (1937), Deraniyagala 

ORGANOGENEBIS 

(1939), Kniisel (1944), Devillers (1954c), De Beer (1959), Guibe (1970g, k), 

Pasteels (1970), Walker (1972) and Honig (1984). Rathke (1866), Voeltzkow 

(1899), Reese (1908, 1915a), Wettstein (1937, 1954), Deraniyagala (1939), 

and Ramaswami (1946) mention some features of tail development. 

In Alligator mississippiensis, the limb buds first appear about Stage 6 

(Figs. 23 and 35). The posterior buds develop faster than the anterior, at 

least during the early stages of their embryology, contrary to Reese (1915a). 

The chronology of limb development is outlined in Section VI and is diagramed 

in Fig. 23. The limb buds develop distinct apical ectodermal ridges 

(Figs. 23 and 35A-H). Grafting experiments within alligator embryos and 

between alligator and chicken embryos (Honig, 1984) have revealed that 

the apical ectodermal ridge and zones of polarizing activity (ZPA) regulate 

differentiation as has been shown in other tetrapods (see reviews in Fallon 

and Caplan, 1983). These experimental digit duplications resemble those 

seen as natural abnormalities in the wild (Fig. 36L). Normally, there are 

five digits on the forelimb, but only four on the hindlimb (Fig. 23). Claws 

differentiate on the tips of three digits of both limbs, but are absent from 

the two outer digits of the forelimb and the one outer digit of the hindlimb 

(Fig. 23). The ventral concaVity of the embryonic claw is filled by the 

neonychial pad, a soft rounded cushion formed by a thickening of the 

epidermis superficial to the sole of the claw (Kerr, 1919). These structures 

prevent the embryo from tearing its membranes as it moves about during 

late development; they detach shortly after hatching, leaving behind the 

functional claws, the morphogenesis of which has not been reported. 

Explanted limb buds that are grown either in vitro or as grafts on the 

chick chorioallantoic membrane show good mesenchymal differentiation, 

particularly in the cartilages (Honig and Ferguson, unpublished data). 

The development of skeletal elements in the limb has been described by 

Kiikenthal (1893), Voeltzkow (1899), and Steiner (1934). There is a similar 

ity between avian and crocodilian embryos as regards the lateral deflexion 

of the wrist-hand axis from that of the forearm and in the reduction of the 

two outer digits. The tarsus of the avian embryo has a distinctly crocodilian 

appearance, with a dorsolateral process of the astragalus contacting the 

lower end of the fibula above the calcaneum (Walker, 1972). In the embryo, 

the iliac, pubic, and ischiadic cartilages are in continuity and the ace 

tabulum is closed. The piercing of the acetabulum and separation of the 

pubis take place late in development (Goodrich, 1930). 

Little is known about tail development (Figs. 18-22) apart from com 

~ents on its changing relative length. Deraniyagala (1939) described a 

highly vascularized terminal hook (or kink) on the tail of embryos of 

Crocodylus porosus, which arises by fusion of the last three rings of caudal 

scales. This hook is clearly evident in Alligator mississippiensis, C. porosus, 

and C. johnstoni (Figs. 18-22). Ramaswami (1946) reported that the kink 

Contains only connective tissue cells and concluded that it was a transient 

feature of unknown function, but Deraniyagala (1939) hypothesized that 

qq7

Fig. 36. Alligator mississippiensis. Details of the caruncle. (A) Dorsal view of the snout of a 

hatchling illustrating the position of the caruncle. (B) Scanning electron micrograph of hatchling, 

note the smooth pedicle and bifid points of the caruncle, that are used to slit the eggshell 

membranes. (C) Stage 24. Coronal histological section of the caruncle. Note the vascular loose 

fibrous central core and the enlarged strateum corneum. (D) Stage 28. Coronal histological 

section. Note the denser avascular fibrous core and the greatly enlarged strateum corneum. 

Fig. 35. Alligator mississippiensis. Early limb development (compare with Fig. 23.). (A) Stage 7 

(day 7). Scanning electron micrograph (SEM) of the hindlimb bud (H). Note also the developing 

tail (T) and allantois (A). (B) Stage 11 (day 12). Histological section of the apical ectodermal 

ridge (AER) on the forelimb bud. Photograph courtesy of Dr. J. Fallon. (C) Stage 9 (day 9). 

Lateral SEM view of the hindlimb bud. Note the apical ectodermal ridge (AER). (D) Stage 9 

(day 9). Lateral SEM view of the forelimb bud. Note that it lacks an apical ectodermal ridge 

and is less well developed than the hindlimb bud. (E) Edge on SEM view of the hindlimb bud 

depicted in C. Note the apical ectodermal ridge (arrowed). (F) Edge on SEM view of the 

forelimb bud depicted in D. The apical ectodermal ridge is not apparent. (G) Stage 10 (day 10). 

SEM of a hindlimb bud. Note the apical ectodermal ridge (arrowed). (H) Stage 10 (day 10). 

SEM of a forelimb bud. Note the apical ectodermal ridge (arrowed).

450 REPRODUCTIVE BIOLOGY AND EMBRYOLDGY OF CROCODILIANS 

DEVELOPMENTAL ABNORMALITIES 

4/51 

its hypertrophy could have produced the terminal caudal fin of some extinct 

crocodiles. 

S. Integument and Its Glands 

Information on reptilian integumentary embryology is reviewed by Maderson 

(Chapter 7, this volume). The most detailed description for crocodilians 

is by Voeltzkow (1899) who also reports the development of the 

integumentary glands and bony scutes (osteoderms). The latter have been 

extensively described by Schmidt (1914), and much of their development 

occurs posthatching. According to Deraniyagala (1939), embryos of 

Crocodylus porosus form scales about 26 days after egg laying; the process is 

largely complete by 45 days when pigmentation apparently begins. The 

macroscopic appearance and chronology of scale formation are described 

in Section VI. There is substantial variation in the coloration pattern of 

hatchling alligators, the yellow stripes being variable in both form and 

position. We do not know what determines this pattern and how it 

changes with growth and development. 

Data on the development of the various types of integumentary glands 

indicate only that these arise as invaginations of ectoderm into the dermis 

(Reese, 1921); the surrounding mesenchyme forms gland capsules of connective 

tissue and muscle (Reese, 1921). 

T. Caruncle 

The small caruncle facilitates slitting of the extraembryonic and shell membranes 

during hatching. It has been described for most crocodilians 

(Voeltzkow, 1891, 1892, 1893, 1899; Pooley, 1962, 1969a; Deraniyagala, 

1936, 1939; McIlhenny, 1935; Joanen, 1969; Singh, 1975). At the time of 

hatching, it is a small hard structure (approximately 1 mm in length and 2 

mm in diameter) located at the tip of the snout (Figs. 36A, B). It is bifid and 

ends in two sharp points; as the embryo "nods" its head in ovo, these rub 

against, pierce, and slit the enclosing membranes (Figs. 26G, H, I, 36A and 

B). The chronology of its macroscopic development is described in Section 

VI and illustrated in Fig. 26. It is initiated (and present histologically) at 

Stage 15 and becomes evident at Stage 16. It persists for approximately two 

weeks posthatching, then falls off leaving a scar that later heals. Hatchlings 

show a 1-2 mm "halo" of smooth and lightly pitted integument around the 

base of the caruncle (Figs. 36A and B). 

The structure and development of the caruncle were first described in 

Crocodylus porosus (Sluiter, 1893), and this work formed the basis for subsequent 

descriptions and illustrations (Voeltzkow, 1899; De Beer, 1949; 

Fioroni, 1962; Edmund, 1969; Guibe, 1970m). The caruncle of crocodilians, 

like that of Testudines, Sphenodon and birds, is epidermal and is formed 

principally by a hyperplastic stratum corneum so that it is structurally 

~, ' 

unrelated to the dentition. In this regard, it is very different from the true 

"egg teeth" of the Squamata, which are formed as part of the premaxillary 

dentition (De Beer, 1949; Fioroni, 1962; Edmund, 1969; Guibe, 1970m). 

Some minor confusion may result when the crocodilian caruncle is called 

the "egg tooth" by field workers (e.g., McIlhenny, 1935; Deraniyagala, 

1936, 1939; Pooley, 1962, 1969a; Neill, 1971; Singh, 1975). 

In Crocodylus niloticus, C. porosus, C. johnsoni, and Alligator mississippiensis, 

the caruncle first appears at Stage 15 as a pair of epithelial outgrowths 

on the tip of the snout (Sluiter, 1893; Voeltzkow, 1899; Fig. 26A). By Stage 

20, these have approximated, and the tissue between them has grown out 

to form a single consolidated mass (Figs. 26B and C). The ensemble forms a 

lightly pigmented horny outgrowth terminating in two points (Fig. 260). It 

has been suggested that the caruncle forms entirely by a thickening of the 

strateum corneum (Rose, 1892a; Sluiter, 1893; Voeltzkow, 1899; Fioroni, 

1962; Guibe, 1970m) in which calcium salts may be deposited (Fioroni, 

1962). However, coronal sections (Figs. 36C, D) of the caruncle in A. mississippiensis 

show a previously overlooked sizable fibrous, dermal core, in 

addition to the hyperplastic stratum corneum. Initially this dermal core is 

composed of loosely arranged fibroblasts and numerous blood vessels (Fig. 

36C). 

As development proceeds the dermal core becomes less vascular and 

more fibrous; at hatching it is almost completely fibrous (Fig. 360). Posthatching, 

this fibrosis continues until the caruncle breaks off, a process 

facilitated by epithelial ingrowths at the base and the diminished blood 

supply in the dermal core. The epithelial ingrowths then establish a normal 

epidermis. In parallel with the changes of the connective tissue, the 

keratinocytes of the stratum corneum of the developing caruncle become 

more numerous and densely packed as keratinization proceeds (Figs. 36C, 

D). The thickness of the peridermal layer decreases but the distinctive 

interface between periderm and stratum corneum persists (Figs. 36C, D). 

The presence of a fibrous core seems logical because an unsupported pile 

of dead epidermal cells, 2-mm broad by 1-mm high, would be weak and 

prone to breakage, unlike the actual condition at the time of hatching. The 

crocodilian caruncle with its fibrous core is not dissimilar to that of monotremes 

in which the core contains a nodule of bone (Hill and De Beer, 1949) 

so that the two are more likely to be homologous than has hitherto been 

assumed (De Beer, 1949). 

VIII. 


Case reports garnered from the literature are listed in Table VIII. In most 

cases, the etiology and pathogenesis of such malformations are unknown. 

Comparable summaries exist for lizards, snakes, and turtles (A. d'A. Bellairs, 

1981; Ewert, 1979).

TABLE VIII 

A Systematic Catalogue of Reported Developmental Abnormalities 

in Crocodilians 

Type oi Maliormation 

Double yolks 

Twins 

Species 

A. mississippiensis 

C niloticus 

C. aculus 

C. POroSU5 

Notes 

Often laid first or 

last by young iemales 

A. mississippiensis Stage 1 and Stage 25 

o tetraspis 

C. nilolicus 

C. porosus 

C. nilolicus 

Reierence 

Ferguson and Joanen 

(1983) 

Blomberg (1979) 

Neill (1971) 

Webb et a1. (1983b) 

Type oi Maliormation 

(b) cyclopia 

(c) anophthalmia 

Species 


O. tetraspis 


C. palustris 

TABLE VIII 

(Continued) 

Notes 

Teratogen induced 

and spontaneous 

Spontaneous 

Reference 

Fig. 37c; Ferguson 

(l982b, 1984) 

Tryon (1980) 

Ferguson (unpublished 

data) 

Whitaker and Whitaker 

(1976a, b) 

Pooley (1969a) 

Singh and Tandan 

Reese (1906); Joanen 

(personal communica 

C. niloticus 

C. gangeticus Deiective develoption) 

ment oi optic (1978); Subba-Rao and 

Hatchlings Tryon (1980) 

Hutton and Loveridge 

(personal communication) 

Stage 18 

Ferguson and Webb 

(unpublished data) 

Stage 20, hatchlings Ferguson and Webb 


(d) exophthalmia 

(e) corneal and iris 

malformations 

(2) Jaw deiects 

(a) cleit lip and palate 


C gangelicus 

placodes and 

vesicle 

Bustard (1979); Singh 

and Bustard (1982) 

Ferguson (1981a. 1982b, 

1984) 

Singh and Bustard 

(1982) 

C. jo/msol1i 

A. mississippiensis Teratogen induced Figs. 37i-i; Ferguson 

Axial biiurcation and par A. mississippiensis Single head with Fig. 37b or spontaneous, (l981a, 1982b, 1984) 

tial twinning 

cleft lip and palate 

see text 

C. niloticu5 

Two vertebral col 

Spontaneous 


umns, 2 sets oi 

(personal communicalimbs 

and tails 

tion) 

(b) reduced upper or A. mississippiensis 

Stage 21 doubleheaded 

embryo munication) 

genically induced 1984); Fig. 37e 

Joanen (personal com 

Surgically or terato 

Ferguson (1981a, 1982b, 

lower jaws 

Double head, two Hutton and Loveridge 

and spontaneous, 

tails and iour hind (personal communication) 

see text 

C. porusus 

legs 

Hot and cold nests Webb and Messel 

eNS maliormations (1977); Webb et a1. 

(1) Spina biiida 

(2) Exencephaly and 

anancephaly 

(3) Encephalocele 

(4) Microcephaly 

(5) Hydrocephalus 

(6) Inner ear and cerebellar 

maliormations 

A mississippiensis 


A. mississippiclIsis 

C. nilotirus 

C. nilolic"s 

A. mississippiens;s 



Often laid by young 

iemales 


females 


iemales 

High incubation 

temperatures 

Teratogen induced 

Low incubation temperatures. 

see Section 

II 

Fig. 37a; Ferguson 

(1981a, 1982b, 1984) 


data) 


data) 



Hu tton and Loveridge 



data) 

Fig. 37j; Ferguson 

(1981a, 1982b, 1984) 

Joanen (personal communication); 

Ferguson 


Craniofacial nlalformations 

(1) Eye deiects 

(a) microphthalmia A. mississippiensis Teratogen induced Fig. 37d; Ferguson 

and spontaneous (1982b, 1984) 

(c) laterally skewed 

lower jaws 

(3) Dental deiects 

C. johnsoni 

C. niloticus 

C. palustris 

O. tetraspis 

C. gangeticus 


C. porosus 

C. ;011ll50ni 

C. gangeticus 

Basihyal valve 

absent 

C. porGsus 

Several teeth in one 

socket 

A. mississippiensis Tooth erupting into 

the nose 

(l983b) 

Webb and Manolis 

(1983) 

Pooley (1969a); Hutton 

and Loveridge (personal 

communication) 

Kalin (1936a, b) 

Tryon (1980) 


(1982) 


data) 

Webb and Messel 

(1977); Webb et a1. 

(1983b) 

Webb and Manolis 

(1983) 


(1982) 

De Jong (1928) 

Ferguson (1982b) 

(continued)

TABLE VIII 

(Continued) 

TABLE VIII 

(Continued) 

Type of Malformation Species Notes Reference 

Type of Malformation Species Notes Reference 

(3) Vertebral column 

Skeletal malformations (a) twisted back A. mississippiensis Eggs incubated up Fig. 37k; Ferguson (un 

(1) Limbs 

(scoliosis, 

right, i.e., at right published data) 

(a) extra limbs 

A. mississippiensis Extra fifth hind leg Ferguson (observation 

kyphosis or 

angles to the nest 

attached to dorsal of animal at St Augustine 

Alligator 

lordosis) 

base and spontaneous 

surface of the 

back at the pelvic Farm, Florida) 

C. porosus 

Desiccation 

Kar (1979) 

level 

High and low incubation 

tempera 

Webb et at. (1983b) 

A. mississippiensis Extra fifth leg attached 

to right 

tures 

(b) extra digits 

(c) absent digits 

(d) syndactyly 

(2) Tail 

(a) kinked tail 

(b) reduced or absent 

tail 


C. porosus 

hindleg 

Eight toes on each 

of the front limbs 

and the left hind 

foot, seven on the 

right hind foot, 

teratogenical1y induced 


Eight toes on each 

of the hind feet 

Giles (1948) 

Ferguson (1981a, 1982b) 

Deraniyagala (1936, 

1939) 

Fig. 371 

Ferguson (1982b) 

C. johnsoni 

A. mississippifllsis Teratogenically induced 


C. niloticus Hutton and Loveridge 




temperatures 


temperatures and 

eggs incubated 

upright, i.e, at 

right angles to the 



data) 


data) 

(b) extra vertebrae 

(c) missing vertebrae 

Visceral malformations 

(1) Herniation of thoracic 

and/or abdominal 

contents 

C. niloticus 

G. gangeticus 


C. porosus 

C. niloticus 

C. porosus 


G. gangetieus 

C. porosus 

C. niloticus 

(2) Hermaphroditism A. mississippiensis 

Color malformations 

C. johnson; 

Evident only on 

gonadal histology 


temperatures, evident 

only on 

gonadal histology 

Modha (1967); Pooley 

(1969a) 


(1982) 

Case (1896) 

Rheinhardt (1874); Baur 

(1886, 1889) 

Kalin (1936a, b) 

Webb et at. (1983b) 

Fig. 37a; Ferguson (unpublished 

data) 


(1982) 

Webb et al. (1983b) 



Forbes (1940a) 

Webb and Smith (1984) 

nest base (1) Albinism or partial A. mississippiensis Some genetic cases Ferguson, unpublished; 

G. gangeticus Singh and Bustard albinism and also related to McIlhenny (1935); 

(1982) incubation tem Allen and Neill (1956); 

C. novaeguineae High incubation Bustard (1969, 1971) peratures Neill (1971) 

temperatures C. porosus Kar and Bustard, 1982; 

C. niloticus Pooley (1969a); Hutton Ferguson and Webb 

and Loveridge (per 


sonal communication) C. ;ohnstoni Ferguson and Webb 

C. palustris Deraniyagala (1939); 


Whitaker and 

C. niloticus 

Blake and Loveridge 

Whitaker (1976a, b, (1975) 

1977b) C. acutus Neill (1971) 

C. novaeguineae Photograph of a 

pure white albino 

wi th red eyes 

C. crocodi/u$ Completely black 

A. mississippiensis Red and yellow 

C. porosus Kar (1979); Kar and Bustard 

(1982b); Webb et 

al. (1983b) 



(2) Melanistic 

data) 

(3) Erythristic 

C. porosus Genetic cause sus Webb et al. (1983b) colour 

pected 

C. niloticus Modha (1967); Pooley 

(1969a) 

dF;Pi, 

Whitaker (personal communication) 

Allen and Neill (1956) 

Allen and Neill (1956)


Embryos produced by young and very old females are frequently abnormal 

(Section IlD and Table IV). Defects include spina bifida (Fig. 37A), 

exencephaly, cyclopia (Fig. 37C), microphthalmia (Fig. 370), exophthalmia, 

hydrocephalus (Fig. 37J), reduced upper or lower jaw (Fig. 37E), cleft 

lip and palate (Figs. 37F-I), scoliosis (Fig. 37K), and herniation of the 

thoracic or abdominal viscera. 

During incubation, extremes of temperature, abnormalities in hydric or 

gaseous environment, and variations in the orientation of eggs (i.e., sitting 

upright as opposed to lying flat) all cause malformations (Table VIII and 

Section II). Animals incubated at high temperatures, which hatch prematurely, 

often have a bump or bulge on their cranial platform (Alligator 

mississippiensis, Ferguson, unpublished data; Crocodylus porosus, Webb 

et al., 1983b; C. johnsoni, Webb, personal communication; C. niloticus, 

Modha, 1967). This represents the midbrain bulge and is probably caused 

by enhanced ossification of the skull before the cranial flexure has straightened 

out. As with gonadal growth, this emphasizes how an alteration 

in incubation temperature may introduce asynchronies in development. 

Craniofacial malformations have been induced experimentally in embryos 

of Alligator mississippiensis by either surgical excision of various regions 

of the neural crest or by teratogens such as 5-fluoro-2-desoxyuridine 

(FUDR) or hydrocortisone (Ferguson, 1981a, 1982b, 1984a). Teratogens 

can be injected into the albumen or the yolk. Injection into the chorioallantoic 

blood vessels is technically more difficult, but preferable because uniform, 

homogeneous deformities result from eggs treated with the same 

dose of the same drug at the same developmental stage. Table IX sum- 

Fig. 37. Alligator mississippiensis. Examples of malformed embryos. (A) View of embryo from 

egg laid by a young mother. Note the meningicoel covering the thoracic spina bifida (arrowed), 

the thoraco-abdominal herniation (A) and the kyphotic spine. Such embryos tend to 

die after 10-20 days incubation. (B) Spontaneous malformation recovered from a 50-day egg 

laid by a young female. Radiograph illustrating axial bifurcation. There are two vertebral 

columns, two pairs of upper and lower limbs, two pelves, two shoulder girdles. and two tails, 

but only one head. The latter has bilateral cleft lip and palate. (C) Lateral view of a 24-day 

embryo with FUOR induced monorhiny (one stage before cyclopia). Note the single nasal 

proboscis (arrowed), absent midface and small mandible. The eyes contact each other in the 

ventral midline. (D) Right lateral view of a developmentally retarded 24-day embryo with 

FUOR induced microophthalmia (right side) and anophthalmia (left side). (E) Macroscopic 

view of a hatchling alligator treated with 0.01 mg FUOR on day 10. The otherwise normal 

embryo has almost no lower jaw. but a normal palate and upper jaw. (F) Lateral view of a 29 

day embryo with FUOR induced bilateral cleft lip and palate (arrowed). The lower jaw is 

trapped behind the cleft premaxilla. (G) Intraoral view of a hatchling alligator with FUOR 

induced bilateral cleft lip (large arrows) and palate (small arrows). The wide palatal cleft 

extends into the nasal cavities anteriorly and into the nasopharyngeal duct posteriorly. (H) 

Transverse section through the head of a 20-day embryo with FUOR induced bilateral cleft lip 

and palate. H. & E. and Alcian Blue. Note the small slender projections representing the 

palatal shelves (P). (I) Transverse section through a hatchling with FUOR induced bilateral 

cleft lip and palate. Mallory. Note the cleft maxilla, distorted tongue and floor of the mouth, 

genioglossus (G), hyoglossus (H), intermandibularis (I), palatal submucosa (S), junction between 

nasal ciliated columnar and oral stratified squamous epithelia (arrowed) and continuous 

oral and nasal cavities. (J) Lateral view of a hatchling with FUOR induced hydrocephalus 

and bilateral cleft lip and palate. (K) Dorsal view of a hatchling with kyphosis and scoliosis. 

~is type of spontaneous malformation is frequently seen in eggs incubated at an incorrect 

onentation, e.g., at 45-90° to the nest base. Such embryos are usually unable to hatch spon 

taneously and the malformations often become progressively worse after hatching. (L) 

Crocodylus johnsoni. View of a hatchling with complete duplication of the digits in its right pes. 

(M) Detail of part L. Similar malformations can be produced experimentally by grafts of 

SUpernumerary ZPA's (zones of polariZing activity). Unless otherwise indicated the scale 

markings are divided into 1 mm intervals. 

M


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Craniofacial malformations can also be induced by altering the diet of 

penned breeding animals (Ferguson 1981a, 1982b, 1984a). However, such 

regimes produce a wide spectrum of malformations and are therefore of 

less experimental value but important in farming, management, conservation, 

and population studies. 

, 

IX. 

SHELL-LESS, SEMI·SHELL-LESS, AND IN VITAO 

CULTURE TECHNIBUES 

In many surgical and teratological investigations, it is useful to be able to 

follow development throughout incubation by observing, even manipulating, 

the embryo through the shell. Numerous investigators have explanted 

chicken embryos and egg contents into petri dishes and grown them in 

shell-less culture (for review, see Ferguson, 1984a). The developing alligator 

is a particularly good animal for shell-less culture because it obtains 

little calcium from the shell as compared to turtles and birds (Section IILD). 

Alligator embryos collected within 12 hours of egg laying, before attachment 

to the shell, may be prepared for shell-less culture (Ferguson, 1981a). 

An incision made around the longitudinal axis of the shell permits removal 

of the top one-third; this is followed by sterile excision of the underlying 

shell membrane. The egg contents are then explanted into a 120 mm 

sterile, vented petri dish under a laminar flow hood. This dish is covered 

and placed inside a larger one containing filter paper saturated with sterile 

water for high humidity. The entire assembly is incubated at 30°C and 

100% humidity under sterile conditions. Development proceeds normally 

in such preparations up to approximately Stage 18, after which malformations, 

particularly in the snout, cranium, and limbs, occur. Probably these 

result from altered tensile forces impinging on the embryo. 

As the yolk and albumen of alligator eggs are more viscous than those of 

avian eggs, a technique of semi-shell-less culture has been developed; with 

slight modifications, this can be used successfully at any period of development 

(Ferguson, 1981a, 1982b, 1984a). Sterile incisions are made around 

the longitudinal axis of the shell, as described earlier, and the top one-third 

of the shell and shell membrane are removed. The yolk and albumen are 

viscous enough to remain intact inside the lower two-thirds of the shell so 

that the natural geometry of the egg contents is maintained (Fig. 38). Maintenance 

of such embryos in sterile incubators at 30°C and 100% humidity 

results in normal development (Fig. 38), which can be filmed and/or experimentally 

manipulated. The modest extrinsic calcium needs of the embryo 

are met by the intact lower two-thirds of the shell. For optimum 

results, the technique should be performed within 24 hours of egg laying, 

before the embryo has attached to the top of the shell membrane or developed 

a chorioallantois; such embryos will develop normally and hatch 

approximately 65 days later (Fig. 38). There are no statistically significant 

DAY 12 DAY 20 DAY 28 

DAY 30 DAY 35 

DAY 40 

MM 

DAY 55 

Fig. 38. Alligator mississippiensis. Montage illustrating the development of embryos in semi 

Shell-less culture. Note the chorioallantoic blood vessels overlying the embryos.

4S2 


ACKNOWLEDGMENTS 

4S3 

differences between the weights and crown-rump lengths of these embryos 

and those recovered from normally incubating eggs of the same age. 

The technique, combined with the application of 0.01 mg. FUDR to a 

Stage 10 embryo (to inhibit lower jaw formation), has permitted the direct 

observation of palatal development in normal and teratogen-treated embryos 

in situ. Initially, the treated embryos are developmentally retarded, 

but they exhibit rapid compensatory growth and apart from an extremely 

small lower jaw, they exhibit no other macroscopically detectable abnormalities. 

The treated embryos remain viable and hatch around 65 days but 

die shortly thereafter because they cannot feed. The semi-sheIl-less culture 

technique may be equally useful for studying the development of other 

structures. 

Alligator eggs can be windowed for experimental purposes, preferably 

during the first 12 hours after egg laying, before the embryo attaches to the 

shell. Candling and removal of the eggshell and shell membrane gives 

poorer results in analyzing older embryos; many embryos die because they 

drop from the inside of the shell membrane into the underlying yolk, 

tearing their chorioallantoic blood vessels. Only 5% survive this later windowing 

technique (Honig, 1984; Honig and Ferguson, unpublished data). 

Embryonic rudiments, e.g. mandibular arches, palatal shelves, limb 

buds, teeth, genital ridges and lungs have been successfully cultured in 

vitro (Ferguson et al., 1982, 1983a, b; Ferguson and Honig, 1984). Tissues 

can be cultured for long time periods in either chemically defined or serum 

supplemented media, and at the abnormally high temperature of 37°C; 

procedural details have been presented previously (Ferguson et al., 1982, 

1983, 1984; Ferguson and Honig, 1984). 

x. CONCLUSIONS 

The embryology of crocodilians is an interesting and important topic, with 

applications not just in herpetology but also in medicine, dentistry, and 

general developmental biology. Further research is required and areas in 

need of attention have been highlighted. There is a need to extend what 

little knowledge we now possess by systematic studies of development in 

various species, particularly in the more atypical such as the gavial and 

Tomistoma. One could hardly do better than to close with a quotation from 

Parker's (1883) monograph on the development of the crocodilian skull, 

which is as relevant today as it was 100 years ago: "As the crocodile is 

known to be one of the most ancient types inhabiting this terraqueous 

globe, his development is full of interest in relation to those countless 

Reptilian forms that have succumbed to secular changes of the earth, and 

have left neither son nor nephew in the regions where they once were 

dominant." 

ACKNOWLEOGMENTS 

In 1978 the alligator specimens referred to in this chapter were collected 

during the tenure of a Winston Churchill Travelling Fellowship from the 

Winston Churchill Memorial Trust, London, in 1980 during the tenure of a 

Sir Thomas and Lady Edith Dixon Research Scholarship from The Queen's 

University of Belfast, and in 1981 during the tenure of a Wellcome Trust 

Research Visiting Fellowship from the Wellcome Trust, London. I am 

grateful to all these Bodies for their support and confidence. 

I would like to thank sincerely Mr. Ted Joanen and the staff of the 

Rockefeller Wildlife Refuge, Louisiana, for their outstanding assistance in 

obtaining alligator specimens an,d for their generous hospitality and advice 

during my visits with them. The Anatomy Department, Belfast, secretaries, 

Miss Anne Richardson and Miss Janice Smith, and the Divisional 

secretaries in Ann Arbor, Ms. S. Konchal and Mrs. K! Vernon, transformed 

my scribbles into a beautifully typed manuscript. I would also like to thank 

Mr. George Bryan for photographic assistance, Dr. K. Lewis and the staff 

of the Q. U. B. Biomedical and Medical Libraries for assistance in procuring 

copies of many obscure papers, Professors A. d'A. Bellairs, C. Gans, 

P. F. A. Maderson, and Drs. F. Billett, G. Webb, and B. Westergaard for 

critical analysis of the original manuscript, and the Academic Council of 

The Queen's University of Belfast for a grant to offset the cost of translating 

some important German papers cited in this chapter. Dr. G. Webb and the 

Conservation Commission of the Northern Territory, Australia, greatly 

facilitated my study of Crocodylus porosus and C. johnsoni embryos and 

provided valuable financial assistance toward my traveling expenses. 

Many individuals volunteered copies of unpublished data or papers in 

press and participated in useful discussions: I thank them all, but especially 

Ted Joanen, Larry McNease, Grahame Webb, Jon Hutton, John 

Loveridge, Alistair Graham, Bjarne Westergaard, and Larry Honig. 

Alligator research expenses were generously provided by grants from 

the Medical Research Council of Great Britain (Grants No. G979/386/C and 

811361OCB), the Eastern Health and Social Services Board, Northern Ireland 

(Grant No. E109/74/75), the National Institutes of Health, U.S.A. 

(Grant No. DE-02848 and DE-03569), and the Louisiana Wildlife and 

F~sheries Commission. This work was carried out under my U.S. Federal 

FISh and Wildlife permit No. PRT2-2511 and under my Northern Ireland 

Wildlife permit No. B/WLl/78, B/WL2/80. 

. Finally, it is with deep regret that I record the sudden death of my good 

friend Professor J. J. Pritchard, for whose advice and encouragement I will 

always be grateful. His death, shortly after the work for this chapter had 

commenced was a great shock and loss to me, and doubtless this chapter 

would have been enriched by his critical editorial pen had he lived to see it 

materialize.

464 REPROOUCTIVE BIOLOGY ANO EMBRYOLOGY OF CROCODILIANS 

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15. 

~

•••••••••••• 

4BO 

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