The ovary, oocyte development, and egg vitelline envelope of the pipefish, Syngnathus scovelli


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The ovary, oocyte development, and egg vitelline envelope of the pipefish, Syngnathus scovelli
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xvii, 191 leaves : ill. ; 29 cm.
Begovac, Paul Christopher, 1956-
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Subjects / Keywords:
Ovary -- anatomy & histology   ( mesh )
Oogenesis   ( mesh )
Vitelline Membrane -- analysis   ( mesh )
Fishes -- growth & development   ( mesh )
Anatomical Sciences thesis Ph.D   ( mesh )
Dissertations, Academic -- Anatomical Sciences -- UF   ( mesh )
bibliography   ( marcgt )
non-fiction   ( marcgt )


Thesis (Ph.D.)--University of Florida, 1988.
Bibliography: leaves 176-190.
Statement of Responsibility:
by Paul Christopher Begovac.
General Note:
General Note:

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University of Florida
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All applicable rights reserved by the source institution and holding location.
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aleph - 000961860
oclc - 18721431
notis - AET3564
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Copyright 1988


Paul Christopher Begovac



I would first like to express my gratitude to my

mentor, Robin Wallace, for setting high professional

standards and serving as an excellent role model for

conducting scientific research. He provided unrestricted

support and freedom to pursue problems of special interest

to me. His lessons will serve me well throughout my

career. A special thanks is due Dr. Kelly Selman, who

taught me electron microscopy and allowed unlimited use of

her laboratory on my frequent trips to Gainesville. Dr.

Chris West was a constant source of critical thought and

conversations with him were instrumental in the early

phases of the vitelline envelope work. I also thank Dr.

David Evans for serving as my outside committee member and

for his review of the dissertation. The work presented in

Chapter II is reproduced by permission of Alan R. Liss,

Inc. (J. Morphol. 193:117-133).

I am indebted to Dr. Paul Linser and Margaret Perkins

for teaching me the art of monoclonal antibody production.

Mrs. Lynn Milstead helped simplify much of my research with

her outstanding drawings. The use of the collecting

facilities at Cedar Key were kindly provided by Dr. Frank

Mature of the Department of Zoology. Some of my most

enjoyable days were spent collecting near Seahorse Key

amidst the pelicans, ibises, ospreys, and frigate birds. I


thank my peers, Deni Galileo and Steve Dworetzky, for our

many discussions about each others experiments and science

in general, but particularly the friendship we shared

together in graduate school. Drs. Mark Greeley, Peter Lin,

Teresa Lin-Petrino, and Sam Edwards were good friends and

served as sounding boards for the interpretation of many

experimental results. My family continually provided

steadfast support and perspective throughout graduate


Finally, and most importantly, I'd like to thank my

dear wife, Susan, for her enduring love and constant

support throughout the ups and downs of the past five

years. Her sacrifices, hard work, encouragement, and

laughter always kept me optimistic about the future.



ACKNOWLEDGEMENTS ............. .................. .... iv

LIST OF FIGURES .............................. ...... .vii


ABSTRACT .............................................xv


I INTRODUCTION ... .... ......................... 1
Statement of Purpose...................***... 4
Terminology ....... ... .......................**7

II THE OVARIAN ANATOMY .......................... 8
Introduction .................................8
Materials and Methods ........................ 9
Results ......................................10
Discussion .................................. ** 35

Introduction *...............**.........**.........*47
Materials and Methods ........................ 48
Results ......................*.....**..**.........*****52
Discussion ......... .. ........ ....... .....***80

Introduction ...... .......... .. ......... ... 93
Materials and Methods ........................ 95
Results ......................... ........... 01
Discussion .................*.***.... .******* 121

Introduction ...**...*...........................*127
Materials and Methods ........................129
Results ......................................138
Discussion ......................... ...... 163


REFERENCES ... ......... ............... ................ 176

BIOGRAPHICAL SKETCH .................... .............. 191


Figure Page

1. Photomicrograph of fresh ovary spread.........13

2. Transverse section of entire ovary depicting
various stages of development.................... 13

3. Transverse section of germinal ridge apposed
to the ovarian wall............................. 13

4. Region comparable to dashed area outlined in
Figure 2.........................................13

5. Extreme mature edge of ovarian sheet.............13

6. Postovulatory follicle near the mature edge......13

7. Electron micrograph of a transverse section of
the ovarian wall.................................18

8. Electron micrograph of luminal epithelium........18

9. Electron micrograph of luminal epithelial cell
blebbing into ovary lumen........................ 18

10. Transverse section showing a portion of the
germinal ridge region of the ovary ...............22

11. Intracellular filaments in prefollicle cell
in the germinal ridge............................ 22

12. Separation of contiguous cell borders of meiotic
oocytes in germinal ridge by prefollicle cell....22

13. Follicle separation from the germinal ridge......24

14. Primordial follicle separating from the
germinal ridge...................................24

15. Photomicrograph of a portion of fresh ovary
illustrating the relationship of the oocyte
germinal vesicle to the ovarian lumen............28

16. Transverse section depicting mature edge
luminal epithelium...............................28

17. Unmyelinated nerve present at the mature edge....28

18. Connective tissue elements in stromata...........28

19. India ink preparations of fresh intact ovaries
demonstrating extent of lymphatic network........32


Figure Page

20. Lymphatic space extending between the lateral
borders of adjacent young oocytes................32

21. Triangular lymphatic space in basolateral region
between adjacent early developing oocytes........32

22. Follicle relations to the luminal epithelium.....34

23. Oocyte relations to the ovarian wall.............34

24. Shared theca on lateral aspects of adjacent
oocytes .. ......... ...... .* *** ******************* 34

25. Schematic summary of ovary and relations in situ.38

26. Electron micrograph of an oogonium...............55

27. Light micrograph of germinal ridge pulsed
with 3H-thymidine after 2 days in culture.........55

28. Light micrograph of germinal ridge pulsed
with 3H-thymidine after 10 days in culture.......55

29. Electron micrograph of leptotene oocyte..........55

30. Electron micrograph of pachytene oocyte..........55

31. High magnification of germ cell-specific,
dense-cored granules.....................***....*****. 55

32. Light micrograph of perinucleolar oocytes
in the primary growth phase.....................*****60

33. Light micrograph of dispersing Balbiani
vitelline body in perinucleolar phase oocyte.....60

34. Electron micrograph of a portion of the
Balbiani vitelline body................ ......... 60

35. Higher magnification of multivesicular bodies
associated with the Balbiani vitelline body......60

36. Electron micrograph of later stage
perlnucleolar oocyte...............********* *****60

37. Photomicrograph of a portion of intact ovary
stained with rhodamine 123.......... .............60

38. Photomicrograph of a portion of ovary stained
with acridine orange....................*********60


Figure Page

39. Light micrograph of cortical alveoli-stage
oocyte........................................... 64

40. Electron micrograph of oocyte cortex in early
cortical alveoli-stage oocyte.................... 64

41. Electron micrograph of vitelline envelope........64

42. Electron micrograph of lipid in cortical
alveoli stage-oocyte............ ....... .......... 64

43. Phase and fluorescence micrographs of oocytes
stained with concanavalin A...................... 64

44. Light micrograph of early vitellogenic oocyte....69

45. Light micrograph of midvitellogenic oocyte.......69

46. Light micrograph of a late vitellogenic oocyte...69

47. Electron micrograph of early vitellogenic
oocyte depicting yolk-sphere formation...........69

48. Electron micrograph of early vitellogenic
oocyte with transitional yolk spheres............69

49. Electron micrograph of mature yolk sphere........69

50. Electron micrograph of pinocytic vesicles in
early vitellogenic oocyte........................ 69

51. Photomicrograph of maturation-stage follicles....73

52. Photomicrograph of in vitro matured follicles....73

53. Photomicrograph of in vivo matured eggs..........73

54. Maturation response of follicles treated with

55. Protein content of individual follicles from
0.4-1.3 mm diameter..............................76

56. SDS-PAGE of proteins present in various sized
follicles from <0.2-1.3 mm diameter..............78

57. Schematic diagram of sequential stages of oocyte
development as viewed in an optimal section......83

58. Model of oocyte progression through ovary
during oocyte development..................... ...90

Figure Page

59. Light micrograph of vitelline envelope...........103

60. Light micrograph of developing follicles
stained with PAS ....... ...... .................... 103

61. Wheat germ agglutinin-staining of developing
oocytes.........* .... ****......********* ** **** *103

62. Electron micrograph of vitelline envelope in
early cortical alveoli-stage oocyte..............107

63. Electron micrograph of vitelline envelope in
later cortical alveoli-stage oocyte..............107

64. Electron micrograph of vitelline envelope in
a midvitellogenic stage oocyte..............*....****107

65. Electron micrograph of vitelline envelope of
mature egg......................... ** ***********107

66. Photomicrograph of manually isolated vitelline
envelopes.......... ... ..* *************.*********** 10

67. Electron micrograph of the vitelline envelope
pellet obtained by the extraction protocol.......110

68. Higher magnification of the nonextractted
vitelline envelope pellet........................110

69. High magnification of insoluble residue..........110

70. SDS-PAGE of sequentially extracted vitelline
envelope pellet proteins........... ... ......... 114

71. SDS-PAGE of limited proteolytic digestion
products of vitelline envelope proteins and egg
proteins..9.... ..... ....*.................** ..... 118

72. SDS-PAGE of vitelline envelope proteins and egg
proteins stained with Coomassie blue and PAS.....120

73. Immunoblot specificity of 5D6 antibody to pellet
and supernatant extraction-protocol fractions....140

74. Immunoblot specificity of 5D6 antibody to plasma
and liver proteins...............................142

75. Immunocytochemistry of 5D6 antibody to early
developing follicles...... ........ ...... ......... 145

76. Immunocytochemistry of 5D6 antibody to
vitellogenic oocytes .. ............ .............. 145

Figure Page

77. Immunocytochemistry of 5D6 antibody to mature
egg.............................................. 145

78. Control immunocytochemistry with 5All antibody
to ovarian follicles............................. 145

79. Immunocytochemistry of 5D6 antibody to liver..... 145

80. Immunoelectron microscopy of 5D6 antibody to
early vitellogenic stage oocyte..................147

81. Immunoelectron microscopy of 5D6 antibody to late
vitellogenic stage oocyte ....... .... ............. 147

82. Control immunoelectron microscopy of
vitellogenic oocyte..............................149

83. Two-dimensional SDS-PAGE of extracted
vitelline envelope proteins...................... 152

84. Immunoblot of two-dimensional SDS-PAGE of
vitelline envelope proteins ..... ......... ........ 152

85. Autoradiogram of 35S-methionine-labeled
proteins within intact follicle.................. 155

86. Immunoblot of 35S-methionine-labeled proteins
within intact follicle.......... ............... 155

87. Verification of cell dissociation procedures.....158

88. Higher resolution autoradiogram of
35S-methionine-labeled proteins within
intact follicle..................................160

89. Autoradiogram of 35S-methionine-labeled proteins
within denuded ooocyte........................... 160

90. Autoradiogram of 35S-methionine-labeled proteins
within isolated follicle cell/theca complexes....160

91. Immunocytochemistry of 5D6 antibody to Fundulus
heteroclitus follicle.................... .........162

92. Control immunocytochemistry of 5All antibody
to Fundulus heteroclitus follicle ....... .......162


BL Basal lamina

C Collagen

CA Cortical alveoli

CAO Cortical alveoli-stage oocyte

CE Coelomic epithelium

CON A Concanavalin A

CT Connective tissue

D Dorsal

DAB Diaminobenzidine

E Eggs

EC Endothelial cells

EDTA Ethylenediaminetetraacetic acid

F Filaments intracellularr)

FC Follicle cell

G Golgi complexes

GR Germinal ridge

GV Germinal vesicle

GVBD Germinal vesicle breakdown

3H- Tritiated

H&E Hematoxylin and eosin

IEF Isoelectric focussing

kDa Kilodaltons

L Ovarian lumen

LE Luminal epithelium

LIV Liver

LP Lipid


LY Lymphatic space

L-15 Leibovitz 15

mm Millimeter

N Unmyelinated nerve

NU Oocyte nucleus

0 Oocyte

OG Oogonia

OW Ovarian wall

PAGE Polyacrylamide gel electrophoresis

PAS Periodic acid-Schiff's

PBS Phosphate-buffered saline

PC Peritoneal cavity

PF Primordial follicle

PFC Prefollicular cells

RCA Ricinus communis agglutinin

SDS Sodium dodecyl sulfate

SEM Standard error of the mean

SM Smooth muscle

T Thecal elements

TBS Tris-buffered saline

ug Microgram

ul Microliter

um Micrometer

V Vein

VE Vitelline envelope

VN Ventral

VO Vitellogenic oocyte









Wheat germ agglutinin

Yolk sphere

Primary yolk sphere

Transitional yolk sphere

Mature yolk sphere




Abstract of Dissertation Presented to the Graduate
School of the University of Florida in Partial Fulfillment
of the Requirements for the Degree of Doctor of Philosophy




April 1988

Chairman: Robin A. Wallace
Major Department: Anatomy and Cell Biology

The ovarian anatomy, stages of oocyte development,

and biochemical properties of the vitelline envelope were

documented for the pipefish, Syngnathus scovelli. This

unusual vertebrate ovary possesses a sequential array of

developing follicles originating from an outpocketed region

of the luminal epithelium, the germinal ridge.

Folliculogenesis occurs within the germinal ridge and newly

formed follicles enter into a linear progression of

developing follicles extending to the opposite mature edge

where ovulation occurs. Later stage oocytes are polarized

with respect to the luminal epithelium.

Six stages of oocyte development were identified as

follows: I) Oogonia; II) Primary growth subdivided into a)

chromatin-nucleolus and b) perinucleolar phases; III)

Cortical alveoli formation; IV) Vitellogenesis; V) Oocyte

maturation; and VI) Mature egg. The 3H-thymidine

incorporation studies establish the germinal ridge as the

proliferative segment of the ovary. Yolk sphere formation

appears to involve a multivesicular body-like structure

that may relate to multivesicular bodies observed in

perinucleolar and cortical alveoli stage oocytes. Oocyte

of 1.1 mm diameter have the capacity to undergo in vitro

maturation in response to the steroid 17-alpha-hydroxy-


The biochemical composition of the vitelline envelope

was examined in further detail. Two major protein families

of 109 and 98kDa were extracted from egg vitelline

envelopes, with at least the 98kDa protein being a

glycoprotein. Periodic acid-Schiff's staining of ovarian

sections reveals an external layer that is strongly

PAS-positive compared to the internal layer. Monoclonal

antibodies prepared against the 109 and 98kDa vitelline

envelope proteins were specific for the vitelline envelope

by immunocytochemistry and immunoblot analyses. The

vitelline envelope proteins were localized by

immunoelectron microscopy to the internal layer of the

vitelline envelope. Synthesis of at least the 98kDa family

occurs within the ovarian follicle. The precise cellular

origin of the vitelline envelope proteins, however, was not



These studies have established the framework for

future studies pertaining to oogenesis and oocyte

development in the pipefish. The sequential pattern of

follicle development in the pipefish ovary offers a

potentially useful model system for study of especially

early events in oogenesis and oocyte development.



That fish which is called Belone, at the
season of reproduction, bursts asunder, and in this
way the ova escape; for this fish has a division
beneath the stomach and bowels like the serpents
called typhlinae. When it has produced its ova it
survives and the wound heals up again.
Book VI, Chapter 12
3rd Century B.C.

This passage, requoted from Gudger (1905, p. 449), is

the earliest reference to reproduction in the pipefish.

The peculiar brood pouch of the pipefish was, until the

mid-19th century, the subject of considerable debate

regarding which sex carried the developing embryos.

Gudger's treatise recalls the history of this early

confusion up until the late 19th century and provides a

colorful survey of the Lophobranch literature until that

time. The account by Gudger is the most complete writing

dealing with the historical aspects of pipefish


The Gulf pipefish, Syngnathus scovelli, is a teleost

of the order Gasterosteiformes and a member of the family

Syngnathidae (Fritzsche, 1984). This group of teleosts is

widely distributed throughout the world, primarily in the

tropical climates, but is also found in more temperate



latitudes (Bohlke and Chaplin, 1970). Pipefish have been

of general interest mainly as aquaria species because of

the male brood pouch, where embryos develop in a manner

similar to those of the closely related seahorse. Most of

the documented reports pertaining to pipefish deal with

phylogeny (Herald, 1942; Herald, 1959; Herald and Dawson,

1972; Dawson, 1977; Fritzsche, 1980), subtropical species

distribution (Reid, 1954; McLane, 1955; Tagatz, 1968;

Whatley, 1969; Dawson, 1970; Robins, 1971; Dawson, 1972;

Herald and Randall, 1972; Dawson and Randall, 1975; Paxton,

1975; Dawson, 1977; Gilmore, 1977; Schooley, 1980; Dawson,

1981; Dawson, 1984), and temperate species distribution

(Briggs, 1975; Bayer, 1980). Sexual dimorphism (Berglund

et al., 1986) and properties of the male brood pouch (Quast

and Howe, 1980; Haresign and Shumway, 1981) have also been

examined. Field and life history studies specific to

Syngnathus scovelli have appeared (Joseph, 1957; Whatley,

1969; Brown, 1972). In Florida, Syngnathus scovelli are

abundant along the Gulf Coast (Reid, 1954; Joseph, 1957;

Brown, 1972) as well as along the Atlantic Coast near the

Indian River and Mosquito Lagoon (Gilmore, 1977; Schooley,

1980). Freshwater populations of Syngnathus scovelli are

found in the St. John's River (McLane, 1955; Tagatz, 1968)

and other Florida locations (summarized in Brown, 1972).

The reproductive processes in the pipefish, however,

have received little study. The earliest depiction of the


pipefish ovary was provided by Cunningham in 1898, but no

subsequent studies by Cunningham followed up on the initial

description of this remarkable ovary. Gudger (1905)

examined the breeding habits and segmentation of the egg

during early embryogenesis in the pipefish, Syngnathus

floridae. Gudger was not concerned with the ovary itself,

but rather postfertilization aspects. The uniqueness of

the ovarian structure initially described by Cunningham

(1898) was then generally lost until the 1960s when two

reports appeared concerning vitelline envelope formation

(Anderson, 1967) and cortical alveoli formation and

vitellogenesis (Anderson, 1968). Subsequently, Wallace and

Selman (1981) reaffirmed the unusual aspects of the

pipefish ovary. In short, the processes of oogenesis and

oocyte development in this rather unique vertebrate ovary

have not been examined in sufficient detail.

The study of oogenesis and oocyte development has a

long history and its significance is attested by the many

volumes dealing with female gametogenesis (Wilson, 1928;

Raven, 1961; Biggers and Schuetz, 1972; Jones, 1978; Metz

and Monroy, 1985; Browder, 1985). There has been a broad

interest in how eggs develop from a comparative standpoint

between different classes of vertebrate and invertebrate

species. There has also been an interest in specific

questions such as germ cell proliferation (Tokarz, 1978),

oocyte growth (Wallace, 1985), and oocyte maturation

(Masui, 1985) common to many different classes of

organisms. Certain organisms lend themselves to the study

of particular aspects of egg development because of the

availability of large numbers of gametes (sea urchins), the

ability to maintain animals in the laboratory (Xenopus

laevis, mice), or the importance of the organism for

commercial (salmon, cows, pigs) or clinical (human)

applications. The study of specific aspects of

gametogenesis among these many varied animals have

contributed to the overall understanding of how eggs

develop. One additional reason for studying female

gametogenesis is the importance of events, which occur

during oocyte development, for the storage of developmental

information (Raven, 1961) and nutrients essential for


The reasons for examining oogenesis and oocyte

development in the pipefish stem primarily from the unique

arrangement of follicle development that is distinct among

vertebrate ovaries. The pipefish has no significant

commercial value but is most appropriate for studying

certain basic aspects of early oogenesis. There is also an

interest from a comparative perspective for understanding

female gametogenesis among teleosts in general.

Statement of Purpose

The first objective of this project is to establish

the morphological foundation for the study of oogenesis and


oocyte development in the pipefish, Syngnathus scovelli.

The unusual organization of follicle development within the

ovary offers promise in terms of expanding our

understanding of the earliest events that occur during

oogenesis and oocyte development. Previously, these

aspects of oocyte development have not attracted the

attention of investigators to the extent that other areas

of oocyte development such as vitellogenesis (Wallace,

1985) and oocyte maturation (Masui, 1985) have received.

The initial emphasis is placed on understanding the ovarian

anatomy and the relationship of developing oocytes within

this structure. These important cellular relationships

should provide the framework for understanding how the

pattern of developing follicles is elaborated and

maintained. The second level of analysis will deal

specifically with the germ cells: oogonia, oocytes and

eggs. Central to the investigation of oocyte development

in the pipefish is the establishment of a staging series

that identifies the cellular characteristics of developing

oocytes. This series will provide a framework for

designing experiments and relating experimental

observations to the developmental sequence in the ovary. A

concurrent goal of these studies will be to identify

potentially fruitful avenues for future experimentation.

The second objective of this project will be to begin

examining in more detail one process initiated early during


oocyte development, that of vitelline envelope formation.

The morphological features of vitelline envelope formation

have been addressed previously (Anderson, 1967), therefore,

this will not to be the major emphasis of the present

studies. The biochemical properties of the vitelline

envelope and the relationship of the resident proteins to

the structure of the vitelline envelope will be the focus

of this section. The biochemistry of the vitelline

envelope, particularly the protein constituents, have not

been sufficiently analyzed in teleost fishes. Thus, these

studies are intended to provide an understanding of the

principal biochemical constituents of the vitelline

envelope, a major structural element formed during oocyte


The overall objective of this research project is to

establish the pipefish ovary as a useful model system for

investigating oogenesis and oocyte development. The

biochemical properties of the vitelline envelope are a

specific area to be examined in further detail. Within the

framework these studies provide, the spatial organization

of developing oocytes should facilitate experimentation of

the earliest events in oogenesis and oocyte development.

These processes have previously been difficult to address

due to the random organization typical of vertebrate

ovaries in general.



The terminology used throughout the dissertation will

follow standard usage; however, at this point it will be

useful to clarify a few specific terms. The term oocyte

will be used to describe the female meiotic germ cell. The

term follicle will describe the oocyte, its surrounding

vitelline envelope, and investment of follicle cells and

theca components. The term theca will apply to the

connective tissue network surrounding the oocyte-follicle

cell complex. The term lymphatics will be considered as

distinct from the theca proper. Maturational will be used

to describe the oocyte which is capable of but has not

undergone germinal vesicle breakdown. Finally, the term

vitelline envelope will be used to describe the

extracellular matrix surrounding the developing oocyte and

ovulated, mature egg. The term chorion will be applied

only to the protective covering surrounding the embryo

proper. The terminology of the teleost egg envelope

covering has had a long and complicated history with many

different designations being ascribed to this particular

structure (Anderson, 1967; Dumont and Brummett, 1985). The

usage as I have defined vitelline envelope reflects the

biochemical and mechanical similarities of the vitelline

envelope prior to fertilization and distinguish this

structure from the postfertilization chorion.



In 1898, Cunningham described an ovary which was

particularly well suited for use in the study of early

events in oogenesis. He observed that the pipefish ovary

consisted of "but one germinal lamina which extends along

the ovarian tube lengthwise, and the germ cells are present

only at the extreme edge of this lamina" (1898, p. 128).

He also noted that oocytes were arranged in decreasing

order of development towards the germinal lamina edge.

Since Cunningham's early observations, the pipefish has not

been widely utilized experimentally. Published reports

concerning reproductive processes in the pipefish and/or

the closely related seahorse include breeding and embryo

segmentation (Gudger, 1905), vitelline envelope formation

(Anderson, 1967), cortical alveoli and yolk formation

(Anderson, 1968), and a short mention of the pipefish ovary

in a review (Wallace and Selman, 1981). Outside of these

reports, little is known about oogenesis in the pipefish.

The exceptional features of this ovary prompted me to

begin reexploring oogenesis in the pipefish. General

descriptions of teleost ovaries (Dodd, 1977) and a detailed



analysis for the Fundulus heteroclitus ovary (Brummett et

al., 1982) have appeared but no such study is available for

the pipefish ovary. Light and electron microscopic

analyses were utilized to provide an understanding of the

pipefish ovarian anatomy. My initial emphasis was to

provide a description of the general anatomical features of

the pipefish ovary in relation to follicle development.

Materials and Methods

Female Gulf pipefish, Syngnathus scovelli, were

collected in the vicinity of Seahorse Key near Cedar Key,

FL using a beam trawl with one-quarter inch mesh netting.

Fish were collected on a monthly basis from November 1984

to October 1985. After collection, fish were transported

back to the Whitney Laboratory, equilibrated to room

temperature, and transferred to aquaria for maintenance.

Ovaries were processed for analyses within 72 hours of


Ovaries were dissected from freshly killed fish and

immediately fixed with 4% glutaraldehyde and 2%

paraformaldehyde in 0.1 M sodium phosphate buffer, pH 7.4

(Karnovsky, 1965). After several minutes, ovaries were cut

transversely into small segments and transferred into fresh

fixative for two hours at 230 C. The ovary segments were

washed with several changes of 0.1 M sodium phosphate

buffer, pH 7.4, for two hours at 230 C, postfixed at 40 C

for 90 minutes in 1% osmium tetroxide and 0.1 M sodium


phosphate buffer, pH 7.4, dehydrated through graded alcohol

series and flat-embedded in Epon 812. One-micron sections

were cut for orientation purposes and stained with 0.1%

toluidine blue containing 1% sodium borate. Silver and

silver-gold thin sections were cut and stained with uranyl

acetate and lead citrate (Reynolds, 1963). Sections were

observed on a JEOL 100 transmission electron microscope at

60kV. Some tissue for light microscopy was processed as

described above without osmication and embedded in JB-4

(Polysciences) resin. One-micron sections were cut and

stained with Harris's hematoxylin and eosin (H&E), Masson's

trichrome, periodic-acid Schiff (PAS) (Luna, 1968), and

toluidine blue as described.

Demonstration of lymphatic spaces was accomplished by

injection of India ink using an insulin syringe and

25-gauge needle (Perez-Clavier and Harrison, 1978). The

needle was introduced parallel to and between the ovarian

wall and underlying follicles of fresh ovaries. Injection

pressure was controlled by hand. Images were photographed

with a camera mounted on a stereomicroscope.


General Morphology

The ovaries are paired structures that fuse caudally

to give rise to a short oviduct that leads to the genital

papilla. The teleost oviduct does not share embryological

homology with the oviduct of other vertebrates. However,


functionally, it does serve to transport eggs from the

ovary to the genital papilla for transfer to the male brood

pouch. Ovaries from adult females (mean length 121 mm)

reach approximately two cm in length and contain up to

approximately 200 eggs. They are supported from the dorsal

body wall by a mesovarium through which the blood supply

and nerves to the organ traverse. The ovary lies

lengthwise in the peritoneal cavity and eggs are generally

found along the entire length of the ovarian lumen. The

ovary itself is a cylindrical tube whose outer layer is

composed of a visceral coelomic epithelium overlying a

multilaminar smooth muscle wall. The inner layer of the

cylindrical tube consists of the ovarian luminal

epithelium. Between the two epithelial layers, a tubular

sheet of follicles and supporting stromata reside in a

sandwichlike configuration.

The pipefish ovary represents an unusual vertebrate

ovary due to the temporal and spatial relationship of

oocyte development. As Figures 1 and 2 demonstrate, the

germinal ridge is located along the entire length of one

edge of the ovarian sheet. It is in this germinal ridge

that proliferative germinal elements reside. There are two

variations observed with respect to the germinal ridge. In

one case, the germinal ridge and early developing follicles

form a curl protruding into the ovarian lumen (Fig. 2). In

the other variation, the germinal ridge and succeeding

Fig. 1. Photomicrograph of a portion of fresh ovary
that has been opened lengthwise and spread out to
demonstrate sequential array of developing follicles in
the ovarian sheet. Germinal ridge (GR) is present at the
upper edge of the spread with intermediate-stage follicles
present at the lower portion of micrograph. x 40.

Fig. 2. Transverse section of an entire ovary depicting
various stages of oocyte development. Note that the
follicles are sandwiched between the outer ovarian wall
and inner luminal epithelium. The germinal ridge
(arrowhead) and young developing follicles have curled to
protrude into the ovary lumen (L). The mature edge is
indicated by the dotted line overlying the largest oocytes
(0). Note also the region of the ovary lacking follicles
between the ovarian wall and the luminal epithelium
(arrow). General dorsal (D) and ventral (VN) relations
are indicated. One um Epon, toluidine blue. x 100.

Fig. 3. Transverse section of germinal ridge apposed to
the ovarian wall (OW). Germinal ridge (*) containing
several meiotic oocytes (large nuclei) present in the
outpocketed region of the luminal epithelium (arrowheads).
Note the primordial follicle (white arrow) separating from
the germinal ridge. JB-4 resin, H&E. x 440.

Fig. 4. Region comparable to dashed area outlined in
Figure 2. Stromal tissue found associated with the
regions between adjacent follicles near the mature edge.
Note arterioles (arrows) interspersed among the connective
tissue (CT). (OW) Ovarian wall. (0) Oocyte. JB-4 resin,
H&E. x 270.

Fig. 5. Extreme mature edge of ovarian sheet with vein
(V) among the connective tissue. Luminal epithelium
(arrow) diverges from ovarian wall (OW) to overlie
prematurational oocyte (0). Endothelial cells
(arrowheads) can be observed lining a lymphatic space
(LY). JB-4 resin, H&E. x 260.

Fig. 6. Postovulatory follicle (arrows) near the mature
edge with hypertrophy of follicle remains. (L) Ovary
lumen. One um Epon, toluidine blue. x 140.




DpW ~


developing follicles are immediately apposed to the ovarian

wall such that no curl protrudes into the lumen (Fig. 3).

Regardless of which configuration exists, follicles always

assume a sandwichlike configuration as follicle development

progresses. At the opposite edge of the sheet, the mature

edge (Figs. 2, 4, 5), oocytes undergo maturation and

ovulate into the ovarian lumen. Postovulatory follicles

are sometimes observed at the mature edge (Fig. 6). The

interior cavity of these structures after ovulation is

continuous with the ovarian lumen. Postovulatory follicles

are distinct from atretic follicles, which also occur in

the ovary. Atretic follicles typically contain cellular

debris from the degenerating oocyte. Follicles between the

germinal ridge and the mature edge form a sequential spiral

with respect to oocyte development such that each follicle

is followed by follicles of progressively increasing

developmental age. The sequential array is consistent in

the early follicles but becomes somewhat intermingled and

less linear near the mature edge. This is most likely due

to the pronounced expansion of oocytes during the

vitellogenic growth period. Specific aspects of oocyte

development will be dealt with in Chapter III. The

sequential spiral lies in a transverse plane with respect

to the lengthwise axis of the fish. In situ, the germinal

ridge generally lies in relation to the dorsal body wall

and the mature edge generally lies in relation to the


ventral body wall (Fig. 2). These relationships are

maintained throughout the entire length of the ovary.

Ovaries were examined during every month of the year

and follicles of all developmental stages were present in

adult fish. In addition, ovulated eggs were present within

the ovarian lumen at all times of the year. There is no

apparent regression of the ovarian structure or follicle

development with respect to season. The availability of

fish, however, does decrease during the winter months

(December-February) but nonetheless, fish remain

reproductively active. Pipefish can be maintained in

laboratory aquaria if efforts are made to secure a constant

source of live food such as brine shrimp and locally

collected small crustaceans. Fish were maintained with

such a feeding regime with the ovaries still having a

normal appearance as indicated by the presence of all

stages of developing oocytes.

Outer Ovarian Wall

The outer ovarian wall forms the outermost boundary

of the ovarian cylindrical tube and encircles the ovary in

a continuous fashion along its entire length. The ovarian

wall has follicles associated with it along most of its

circumference of the ovary but is devoid of underlying

follicles along a region beyond the mature edge as the

epithelium passes back towards the germinal ridge. This

area (Fig. 2) allows for distension of the ovarian lumen to


accommodate and store eggs after ovulation. The ovarian

wall is covered externally by squamous visceral coelomic

epithelium continuous with the epithelium of the peritoneal

cavity. Internal to the coelomic epithelium is a smooth

muscle layer which is several cell layers thick (Fig. 7).

The smooth muscle cells are typical in structure and are

predominantly arranged in a circumferential orientation

perpendicular to the long axis of the ovary. Smooth muscle

cells with a more longitudinal orientation were

occasionally observed in the innermost and outermost region

of the wall. A few fibroblasts are observed among the

muscle cells. Arterioles are present among the smooth

muscle cell layers at various levels of the lamina whereas

venules are generally confined to the inner and outer

portions of the wall boundaries. Internal to the ovarian

wall, stromal elements supporting individual follicles are

found. Unmyelinated nerves also reside between the smooth

muscle cells at various levels of the ovarian wall lamina

(Fig. 7). They appear to lie in a longitudinal orientation

throughout the ovary. Myelinated nerves were not observed.

Inner Luminal Epithelium

The luminal epithelium forms the inner boundary of

the cylindrical ovarian tube. It is a simple epithelium

consisting of squamous-to-cuboidal cells resting on a basal

lamina (Fig. 8). The apical cell surfaces possess

microvilli that are found throughout the ovarian lumen.

Fig. 7. Electron micrograph of a transverse section of
the ovarian wall. Coelomic epithelium (CE) of outer
ovarian wall facing peritoneal cavity (PC) overlies
several layers of smooth muscle (SM) cells. Note the
orientation of the outermost longitudinal and inner
circular muscle cells. Large bundles of unmyelinated
nerves (N) present in the muscle layers among connective
tissue elements. (*) Connective tissue cell. x 7,980.

Fig. 8. Luminal epithelium (LE) overlying a thin walled
vein (V). Note tight junctions (arrows) and desmosomes on
the apical and lateral borders, respectively, of the
epithelial cells. Microvillar processes are seen
extending into the ovarian lumen (L). The basal border of
these epithelial cells rests on a basal lamina
(arrowheads). x 3,240.

Fig. 9. Blebbing of luminal epithelial cell into ovary
lumen (L). Highly secretary nature of these cells is
apparent here and in Figure 8. Note the abundance of
intracellular filaments (F). x 9,760.



L .


8 g


The epithelial cells in the germinal ridge regions of the

ovarian sheet are typically more squamous than those of

other regions. Epithelial cells overlying more developed

follicles are generally more cuboidal. Certain epithelial

cells possess a columnar shape and display blebs that

protrude into the ovarian lumen (Fig. 9). The apical

borders of the epithelial cells are joined by tight

junctions. Desmosomes of variable size also occur on the

lateral cell borders. Most of the luminal cells appear to

be highly synthetic since they possess the organelles

required for the synthesis of large amounts of protein

(i.e., rough endoplasmic reticulum, Golgi, ribosomes)

(Figs. 8, 9). A striking feature of the epithelial cells

is the abundance of intracellular filaments (Fig. 9), which

are present in cells throughout the ovarian lumen. Beneath

the luminal epithelium, capillaries and venules are present

at various positions along the follicle developmental

spiral. Stromata consisting of connective tissue as well

as lymphatics are also present in regions beneath the

epithelium. Oogonia are associated with the luminal

epithelium compartment near the germinal ridge (below).

Germinal Ridge

The germinal ridge region of the ovary lies along the

entire length of one edge of the ovarian sheet (Figs. 1, 2,

3). The ridge is a consistent feature of all ovaries

examined. The germinal ridge constitutes the proliferative


stem-cell segment of the ovarian sheet from which oocytes

are derived and follicles are formed. The germinal ridge

is well defined and contains oogonia, early meiotic

prophase I oocytes, somatic "prefollicular" cells, and

occasional degenerating cells. The germinal ridge is

distinguished as an outpocketing of the luminal epithelium

as defined by its subjacent basal lamina (Fig. 10).

External to the basal lamina enclosing the germinal ridge

is a variable connective tissue layer. Meiotic germ cells

in the germinal ridge initially lie in direct apposition

with one another sharing cell borders to a variable degree

(Figs. 10, 11). Intercellular bridges between germ cells

have not been observed. The germ cells do not lie in

direct contact with a basal lamina but are apposed to

prefollicular (follicle-cell precursors) cells (Figs. 10,

11, 12). The prefollicular cells (Fig. 11) and follicle

cells of a distinct primordial follicle (Figs. 13 a,b)

contain a complement of filaments within their cytoplasm

whereas early meiotic oocytes do not. These filaments are

essentially the same as cytoplasmic filaments observed in

luminal epithelial cells (Fig. 9). Follicle cells and

their precursors also lie in direct contact with a basal

lamina on one of their surfaces (Figs. 10, 11).

Follicle formation occurs within the germinal ridge.

The prefollicular cells within the germinal ridge are

observed extending cell processes between the contiguous

Fig. 10. Transverse section showing a portion of the
germinal ridge region of the ovary. Note the continuity
of the basal lamina (arrowheads) as it lies beneath the
luminal epithelium (LE) and how it diverges from the
luminal epithelium cells to envelop the germinal ridge to
completely surround it. Groups of oocyte (0) germ cells
and prefollicle cells (PFC) can be seen in the germinal
ridge. Note the contiguous borders (arrows) of adjacent
oocytes. x 5,720.

Fig. 11. Intracellular filaments (arrows) in
prefollicle cell (PFC) of the germinal ridge. Basal
lamina surrounding germinal ridge contacts one surface of
the prefollicle cell. Meiotic oocytes (0) with shared
cell borders are indicated. x 13,380.

Fig. 12. Separation of contiguous cell borders of
meiotic oocytes (0) in germinal ridge by prefollicle cell
(PFC) process (arrows) extending between germ cells. Note
the desmosomal contact (arrowhead) with adjacent
prefollicle cell. x 16,200.

LE ,

Fig. 13. a) Follicle separation from the germinal ridge
(GR). Definitive follicle cells (*) surround the
primordial follicle (PF) as it "pinches" (arrows) away
from the ridge. Note the thin connective tissue layer and
lymphatic space (LY) surrounding the germinal ridge and
primordial follicle. The basal lamina (arrowheads) can be
followed and noted to enclose the entire germinal ridge
and primordial follicle. Oocyte (0) in definitive
follicle of early follicle progression is indicated. (LE)
Luminal epithelium. x 3,660. b) Higher magnification of
boxed area in Figure 13a demonstrating definitive follicle
cell containing abundant intracellular filaments (arrows).
These filaments compare to those seen in Figure 11. x

Fig. 14. a) Primordial follicle (PF) as it separates
from the germinal ridge (GR). Note the basal lamina
(arrows) as it surrounds germinal ridge and primordial
follicle. (FC) Follicle cell. x 6,120. b) Higher
magnification of separation site of primordial follicle
from germinal ridge depicted in Figure 14a. The basal
lamina almost completely separates the primordial follicle
from the germinal ridge (arrows). Note the maintenance of
distinct compartments as this separation occurs. x 14,200.

7 4



1 '


W '


I ..-.



meiotic oocytes, thereby completely separating the oocyte

from other germ cells (Fig. 12). Meiotic oocytes undergo

some primary growth while still within the germinal ridge.

Primordial follicles, still associated with the germinal

ridge, are observed at various stages of separation from

the ridge (Figs. 13 a,b). This separation appears to occur

as the primordial follicle "pinches" off of the germinal

ridge to become a definitive follicle unit surrounded by

its own basal lamina (Figs. 14 a,b). This basal lamina

surrounding the newly-formed follicle, therefore, is

derived originally in part from that which is overlying the

germinal ridge. The newly formed follicles in the

progression have a thin theca and often have lymphatic

spaces around their borders. The young follicles then

enter into the sequential follicle developmental spiral.

As the oocytes progress through different stages, they

assume their complement of thecal cells which appear to be

derived from mesenchymal tissue. Thus, oocytes and

associated follicle cells are originally derived from the

luminal epithelial compartment of the ovary, whereas the

cells of the theca are not.

Mature Edge

The mature edge of the ovarian sheet is defined by

the position of the largest follicles that are ready for

maturation and ovulation (Figs. 2, 5). Prematurational

oocytes are present along this edge of the ovary in one or


two longitudinal rows. The oocyte animal pole (defined by

position of the nucleus) is oriented towards the luminal

epithelium as indicated by the oocyte germinal vesicle that

can be visualized through the luminal epithelium (Fig. 15).

The germinal vesicle is generally within approximately a

20-30 degree radius whose axis runs through the oocyte from

the luminal epithelium to the ovarian wall. This

organization indicates that the oocytes possess a polarity

in relation to the general ovarian anatomy. Large veins

and venules are present beneath the luminal epithelium in

this region of the ovary, with arterioles generally absent

(Fig. 5). Stromal and lymphatic components, present to

variable degree, are found on the lateral aspects of the

follicles. Bundles of smooth muscle whose fibers are

oriented parallel to the long axis of the ovary are also

present (Fig. 16). The muscle cells appear beneath the

luminal epithelium as it moves away from the ovary wall to

lie over the large follicles. Unmyelinated nerves can also

be found among the longitudinal smooth muscle cells (Fig.


The luminal epithelial cells at the mature edge

appear to undergo changes including vacuolation that may

precede ovulation. This degeneration is confined only to

the region of the luminal epithelium that apposes the

follicle. Subsequent to ovulation, a postovulatory

follicle remains and the follicle cells undergo marked

Fig. 15. Photomicrograph of a portion of fresh ovary
illustrating the relationship of the late-stage oocyte
germinal vesicle to the ovarian lumen. This view is
looking directly down onto the luminal epithelial surface
of an ovary that has been opened up lengthwise. Oocyte
germinal vesicles (arrows) are clearly visible just
beneath and oriented towards the luminal epithelium. Note
also the intermingling of intermediate-size follicles in
this region of the ovary. x 20.

Fig. 16. Transverse section depicting mature edge
luminal epithelium (LE) with bundles of smooth muscle (SM)
cells having a longitudinal orientation. x 6,300.

Fig. 17. Unmyelinated nerve (N) present amongst the
longitudinally oriented smooth muscle (SM) cells at the
mature edge. x 6,900.

Fig. 18. Connective tissue consisting of collagen (C)
and connective tissue cells (*). x 8,960.

* *.
'*t a i I

I ~*

r. *
St. ..


hypertrophy (Fig. 6). The follicular hypertrophy includes

the presence of rough endoplasmic reticulum and Golgi

elements in the now-columnar follicle cells.

Stromal and Lymphatic Compartments

The stromal elements of the ovarian sandwich consist

of fibroblasts, connective tissue fibers, and blood vessels

in addition to a substantial lymphatic compartment. The

sandwichlike nature of the ovary gives rise to triangular

spaces in the apicolateral (associated with the ovarian

lumen) or basolateral (associated with the ovarian wall)

portions between follicles. Stromal elements are present

in these spaces (Fig. 4). Collagen fibers are

multidirectionally oriented throughout the stromal network.

They are interposed by stromata consisting of fibroblasts

and blood vessels that form the connective tissue (Figs. 4,

18). The connective tissue is found in the thecal region

of each follicle as well as in the bordering spaces between

adjacent follicles, and is particularly pronounced between

the largest prematurational follicles of the mature edge.

The vascular supply can also be found amongst the stromal

elements interior to the ovarian wall (Fig. 4).

Additionally, capillaries are abundant beneath the luminal

epithelium at all levels of the follicle developmental

spiral. At the mature edge, venules and collecting veins

are localized just beneath the luminal epithelium and can

occasionally be seen to be continuous with small vessels

overlying the largest prematurational follicles.

A significant portion of the stromal network contains

a well developed lymphatic compartment that is lined by

attenuated endothelial cells. Spaces between adjacent

endothelial-cell processes can be observed as well as

desmosomal contacts between the cells. India ink

injections were used to demonstrate the extent of the

lymphatic compartment (Fig. 19). Upon introduction of the

syringe needle between the ovarian wall and the underlying

follicles, ink is observed flowing between follicles within

several seconds. With a slight positive pressure, the ink

disperses to fill much of the interstitial space among

follicles throughout the ovary within minutes. Thus, the

lymphatic network is extensive throughout the ovary. In

the early spiral portion of the follicle progression, the

lymphatic space occupies much of the space between

follicles (Fig. 20). The lymphatics can sometimes be seen

to extend from the inner aspect of the ovarian wall all the

way to the luminal epithelium between adjacent follicles.

The lymphatics, however, are most frequently observed in

the apico- and basolateral regions between adjacent

follicles (Fig. 21). Large follicles tend to have greater

amounts of connective tissue in addition to variable

lymphatic regions on their lateral borders.

Fig. 19. India ink preparations of fresh intact ovaries
demonstrating extent of lymphatic network in basolateral
spaces between follicles. Note that the lymphatics end
abruptly at the extreme mature edge (arrows) as eggs (E)
are visible in ovarian lumen. The ovary on the right
demonstrates extent of lymphatics in the early follicle
progression. x 27.

Fig. 20. Lymphatic space (LY) extending between the
lateral borders of adjacent young oocytes (0). x 3,300.

Fig. 21. Triangular lymphatic space (LY) in basolateral
region between adjacent early developing oocytes (0) in
relation to the ovarian wall (OW). Distinct endothelial
cell (EC) lines the lymphatic space. x 7,850.


Fig. 22. Follicle relations to the luminal epithelium
(LE). Note the somewhat cuboldal follicle cells (FC) with
thick basal lamina (arrows) enclosing the oocyte (0) and
follicle cells. Note that the thecal elements (T) are
becoming more well developed. x 7,650.

Fig. 23. Oocyte (0) relations to the ovarian wall (OW).
Note the presence of a lymphatic space (LY) between the
theca (T) and the ovarian wall. x 3,450.

Fig. 24. Shared theca (T) on lateral aspects of
adjacent oocytes (0). The theca contains primarily
connective tissue elements, and note that individual
follicles are not enclosed by a distinct epithelial cell
layer. x 5,550.

a -4F vu



The sandwichlike arrangement of follicles between the

luminal epithelium and ovarian wall results in a connective

tissue sheath which does not just circumscribe individual

follicles. The arrangement of connective tissue is such

that follicles in fact have shared rather than individual

thecal elements on their lateral aspects. On the apical

aspect of the follicles, the theca is situated just beneath

the luminal epithelium (Fig. 22), while on the basal

aspect, the theca is apposed to the ovarian wall (Fig. 23).

On the lateral portion of the follicles, the theca is

shared with adjacent follicles (Fig. 24). Lymphatics may

lie outside the theca all along the follicle spiral but are

not considered part of the theca proper. The theca is

initially sparse near newly formed follicles but becomes

better developed as oocyte development progresses. Thus,

the combination of connective tissue elements and lymphatic

network form the supporting structure for developing



According to Dodd (1977), the pipefish ovary would be

classified developmentally as a cystovarian type and thus

bears similarity to those of many other bony fishes in

having a coelom-derived ovarian lumen continuous with the

oviduct. The pipefish ovary has also been classified as an

asynchronous type (Wallace and Selman, 1981), possessing a

heterogeneous population of follicles of all developmental


stages. The pipefish is reproductively active throughout

the year instead of cycling annually as is the case for

many other fishes (Matthews, 1938; Barr, 1963; Braekevelt

and McMillan, 1967; Moser, 1967). The yearlong

reproductivity of the pipefish is corroborated by

observations that males carrying embryos can be found

during all seasons (Brown, 1972). Unlike other teleost

ovaries, the pipefish ovary lacks ovigerous lamellae common

to fish such as F. heteroclitus (Brummett et al., 1982) and

Sebastodes paucispinis (Moser, 1967). Rather, there exists

a cylindrical tubular structure in which the follicles are

sandwiched between an outer ovarian wall and an inner

luminal epithelium. The follicles are arranged in a spiral

of sequential development originating from the germinal

ridge and extending to the mature edge. The processes of

oocyte growth and development in this ovary parallel those

in many other fishes. A schematic drawing (Fig. 25)

summarizes the distinguishing features of the pipefish

ovary and follicle relationships in relation to the

environment in situ. It emphasizes follicle progressions,

germinal ridge and lumen epithelium, lymphatics, and shared

thecal components.

The arrangement of predominantly circular smooth

muscle cells and longitudinal nerve fibers in the ovarian

wall is probably important in egg release from the ovary.

In the pipefish, egg laying occurs as the male and female

Fig. 25. Schematic summary of ovary and relations in
situ. The upper diagram indicates the paired ovaries
which join to give rise to the oviduct. Note that the
ovaries have a bilateral symmetry with respect to the
early follicle progressions that lie on the lateral aspect
of each ovary. The dashed line indicates a segment of the
ovary that is enlarged in the lower diagram maintaining in
situ relations. The middle section has been unfurled to
allow visualization of the follicles and ovarian lumen.
Note that a strip of luminal epithelium has been removed
except near the germinal ridge along with the underlying
connective tissue elements to visualize the follicle
progression in the ovarian sheet. The association of the
luminal epithelium with the germinal ridge is indicated.
The germinal ridge in the reflected segment of the luminal
epithelium depicts primordial follicles in various stages
of separation. The sandwichlike nature of the ovary is
apparent in the unfurled region with follicles lying
between the ovarian wall and the luminal epithelium.
Arterioles, veins, and lymphatic spaces are indicated in
typical locations. Smooth muscle bundles near the mature
edge and smooth muscle fibers in the ovarian wall are
apparent. A theca encloses each follicle and can be seen
to be shared on lateral borders. This schematic is
prepared to highlight features of the ovary and is not
drawn fully to scale.



bundles -

Ovari an


swim in unison during which time the female deposits eggs

into the male brood pouch (Gudger, 1905). Therefore, the

expulsion of eggs from the female into the brood pouch

requires a precisely coordinated timing of egg release from

the ovary so that egg transfer can be effected.

Consequently, it is probable that the spawning response is

controlled in part by neural input. Therefore, I suggest

that egg release may be controlled with a contraction wave

of the ovarian wall beginning cranially and proceeding in a

caudal direction, in effect squeezing the eggs out of the

ovarian lumen. The eggs would then be transferred through

the oviduct and deposited in the male brood pouch.

The presence of longitudinally oriented smooth muscle

cells beneath the luminal epithelium at the mature edge is

a bit perplexing. In this region of the ovary, a

transition from circular to longitudinal fibers is noted

where the inner luminal epithelium dissociates from the

ovarian wall to overlie the prematurational and

preovulatory follicles. I intend here primarily to note

this observation. Contractile mechanisms involving

microfilaments or contractile cells have been implicated as

participating in ovulation, but the evidence is not

conclusive (reviewed by Goetz, 1983). Spontaneous

contractions of the F. heteroclitus ovary have also been

reported (Winner and Taylor, 1985). It is clear that no

specific conclusion can be drawn from the current study.

However, the pipefish ovary may provide an opportunity to

address the question of contractile involvement in

ovulation in a more detailed manner because the position of

preovulatory follicles is precisely arranged.

The germinal ridge, which runs the entire length of

the ovary, serves as a continuing source of primordial

follicles for future egg production. The distinguishing

feature of this germinal ridge is its existence as an

outpocketing of the luminal epithelium, demonstrating that

the germinal ridge and luminal epithelium are co-occupants

in the same compartment of the ovary. This observation

represents the first clear evidence in teleosts with

respect to the association of the luminal epithelium with

folliculogenesis. There is a report for the guinea pig

that indicates that germ cells and prefollicular cells are

compartmentalized in the surface epithelium (Jeppesen,

1975). In teleosts, however, there has been much

speculation about the compartment in which the oogonia and

early oocytes reside and from which source the follicle

cells are derived. Suggestions include oocytes being

derived from the luminal epithelium (Bullough, 1942; Lehri,

1968) as well as being formed from amongst the stromata

beneath the lumen epithelium (Brummett et al., 1982).

Also, follicle cells have been described as having a

mesenchymal (Mendoza, 1943; Braekevelt and McMillan, 1967)

or an epithelial origin (Bullough, 1942; Moser, 1967).

Observations on the viviparous cyprinodont, Neatoca

bilineata, indicate that many of the gonial nests are

derived from invaginations of the ovigerous fold epithelium

(Mendoza, 1943). In this report, the suggestion was made

that the gonial nests pinch off as a complete unit from the

epithelium and, therefore, arise from the ovarian

epithelium. Mendoza (1943, p. 90) described follicle cells

as being "modified fibrocytes of subepithelial connective

tissue." The data presented here provide additional

understanding of these observations and clarify the process

of folliculogenesis.

My observations on the pipefish ovary indicate that

both the germ cells and the prefollicle cells originally

reside within the ovarian luminal epithelial compartment.

This study has also defined a mechanism by which the

primordial follicle becomes enclosed by a basal lamina sac,

a process which has not been addressed previously. My

interpretation of folliculogenesis from static images is as

follows: Follicle formation occurs within the germinal

ridge. An early event in the process is the formation by

oogonial division of primary oocytes that are in the early

stages of meiosis prophase I. These early oocytes

initially share cell borders but subsequently become

completely separated from one another as prefollicle cells

extend processes between the contiguous meiotic oocytes,

thereby completely separating a newly formed oocyte from


other meiotic germ cells. The enclosed oocyte then

undergoes some initial growth while still within the ridge.

Ultimately, the primordial follicle becomes distinct from

the germinal ridge as it "pinches" off to become a newly

formed definitive follicle surrounded by its own basal

lamina and thin connective tissue sheath. The basal lamina

of the follicle thus originates from that of the germinal

ridge. The germinal ridge remains contiguous with the

luminal epithelial compartment, with the continuity of the

basal lamina surrounding the germinal ridge being

maintained. The new follicle then enters into the

sequential follicle developmental spiral. Subsequently, as

follicle growth proceeds, mesenchymally derived cells come

into association with the definitive follicle to form the

theca of the developing follicle. The theca becomes

increasingly well developed as oocyte growth proceeds. The

process repeats itself as each primordial follicle leaves

the germinal ridge to enter into the follicle progression.

A fundamental conclusion is that both the early germ

cells and the prefollicle cells originate from within the

luminal epithelial compartment. This is indicated by the

outpocketing of the luminal epithelium to form the germinal

ridge. The oocytes in the germinal ridge are a product of

oogonial division, which also occurs in the germinal ridge

of the luminal epithelium. Filaments in pipefish follicle

cells were first noted by Anderson (1967). The


demonstration in this study of cytoplasmic filaments in the

prefollicle cells and early definitive follicle cells

provides a convenient cytoplasmic marker and suggests that

follicle cells may be derived from a source similar to the

luminal epithelial cells, which also possess a substantial

number of filaments in their cytoplasm.

The orientation of the oocyte germinal vesicle

relative to the luminal epithelium indicates an organized

polarity of the late-stage oocytes with respect to the

ovarian anatomy. In the frog, Xenopus laevis, the

orientation of the oocyte to the ovarian structure has been

determined to be random (Van Gansen and Weber, 1972).

However, there does not seem to be any documentation of

these relationships for teleost ovaries. In the pipefish,

oocyte polarity in relation to the ovary is not apparent in

early oocytes but instead becomes notable in the later

stages of oocyte development. Although the orientation of

the oocyte to the luminal epithelium is not precise, the

germinal vesicle is observable quite readily within

approximately a 20-30 degree radius with respect to a

perpendicular axis between the luminal epithelium to the

ovarian wall passing through the oocyte. The establishment

of this relationship during follicle development remains an

interesting problem.

The shared theca, particularly between adjacent

oocytes, is different from the situation in other fish in


which a theca and surface epithelium appears to enclose

individual developing follicles. The developing follicles

of the pipefish thus share to a large extent the same

general external environment. Therefore, recruitment of

oocytes into the various stages may well be a result of not

only external cues but internal regulation as well.

Specialized thecal cells (Nagahama, 1983) have not been

observed in the theca but cannot be excluded. Thus, the

theca appears to form a general supportive network within

the ovary and coexists with the extensive lymphatic

portions to form the complete ovarian stromata.

The pipefish ovary possesses an extensive lymphatic

network in the interstitial regions between follicles.

This, in effect, forms a "sea" in which the developing

follicles reside bounded by the luminal epithelium apically

and the ovarian wall basally. Relevant to the lymphatic

compartment is the vascular supply to the ovary. The

distribution of arterioles near the ovarian wall and

capillaries and veins near the luminal epithelium suggests

a directional vascular flow from the ovarian wall towards

the luminal epithelium. Such a flow would bathe the

follicles with blood nutrients and plasma proteins, most

notably vitellogenin, the yolk protein precursor (Wallace,

1985). Plasma protein bathing would be expected due to the

high capillary permeability of vessels in the ovary, as

indicated by several studies using exogenous tracers


(Dumont, 1978; Ellinwood et al., 1978; Abraham et al.,

1982; Selman and Wallace, 1982, 1983). The exact function

of the lymphatics in the ovary remains unclear. It has

been suggested that the lymphatic compartment helps to

regulate plasma flow and interstitial fluid volume

(Ellinwood et al., 1978). It may also serve to transport

ovarian secretions such as hormones back into the systemic

circulation. The lymphatic compartment in the pipefish

ovary is thought to have a dynamic structure in order to

accommodate changes in follicle size during oocyte growth.

Compliance of the lymphatic network would allow follicle

growth to proceed in a stable fashion within the ovary.

The unusual features of the sequential follicle

development and germinal ridge make the pipefish ovary

suited for experimental analyses of particularly early

events in oogenesis. In addition to sharing many general

features and processes with other teleost ovaries, the

pipefish ovary possesses several intriguing variations.

The sequential pattern of follicle development, germinal

ridge participation in folliculogenesis, shared theca and

lymphatics among follicles, and the apparent polarization

of later stage oocytes with respect to the luminal

epithelium distinguish the pipefish ovary from other

vertebrate ovaries. With the exception of the closely

related seahorse (Anderson, 1967), the sequential

arrangement of follicles has not been described for other


vertebrate ovaries. This sequential arrangement is found,

however, in a variety of insects (Gutzeit and Sander,




Oocyte development is spatially and temporally

arranged in the ovary of the pipefish. The unusual pattern

of oocyte development in the pipefish was first described

by Cunningham in 1898. More recent studies have dealt

specifically with vitelline envelope formation (Anderson,

1967), cortical alveoli formation and vitellogenesis

(Anderson 1968), and ovarian anatomy (Begovac and Wallace,

1987). However, an adequate staging series for oogenesis

and oocyte development has not been established for this

experimentally useful animal.

Several staging series for teleost oogenesis have

been described (Wourms, 1976; Shackley and King, 1977;

Selman and Wallace, 1986). These staging series are

helpful for understanding cellular events during oogenesis

and serving as the basis for experimentation and comparison

to other species. In two previous reports dealing with

pipefish oogenesis (Anderson, 1967, 1968), oocyte stages

were defined by convenience according to size. Together,

these studies examined three different species of fish and

consequently are confusing because of inconsistencies in

the use of various stage designations and the application



of these stages to different fish in the same study. The

current investigations concerning pipefish oogenesis have

indicated the need for a well defined staging series for

oocyte development. Therefore, this report provides such a

series for the pipefish, Syngnathus scovelli. The stages

of oogenesis and oocyte development described are

identified according to the cytological characteristics and

physiological parameters of the developing oocytes.

Materials and Methods


Adult female Syngnathus scovelli, the Gulf pipefish,

were collected near Cedar Key, FL, transported back to the

Whitney Laboratory, and maintained in sea water aquaria.

Tissues were processed for microscopy within 72 hours of

collection. Ovaries were dissected from freshly killed

fish, rinsed briefly in solution FO (Wallace and Selman,

1978), and immediately fixed with 4% glutaraldehyde and 2%

paraformaldehyde in 0.1 M sodium phosphate buffer, pH 7.4

(Karnovsky, 1965), for 2 hours at 230 C. Individual

follicles of various sizes were also dissected from ovaries

using watchmaker forceps, measured with an ocular

micrometer, and fixed as described. Tissues were washed in

0.1 M sodium phosphate buffer, postfixed with 1% osmium

tetroxide, and embedded in Epon 812 as detailed previously

(Begovac and Wallace, 1987). Tissues were also prepared as

described above for light microscopy, with and without


osmication, and embedded in JB-4 resin (Polysciences) or

Historesin (LKB). Thick (1-3 um) sections were cut and

stained with Harris's hematoxylin and eosin, Masson's

trichrome, periodic acid-Schiff's reagent (PAS), alcian

blue, pH 2.5 (Luna, 1968), and 0.1% toluidine blue in 1%

sodium borate. For electron microscopy, silver and

silver-gold thin sections were cut and stained with uranyl

acetate and lead citrate (Reynolds, 1963). Sections were

observed with a JEOL 100 transmission electron microscope

operated at 60 kV.

Lectin binding was observed on tissues fixed with 4%

paraformaldeyde in 0.1 M sodium phosphate, pH 7.4, and

embedded in Historesin. Sections were incubated with

phosphate-buffered saline containing hemoglobin (2 mg/ml)

to block nonspecific staining. Slides were then incubated

with fluorescein-conjugated concanavalin A (5 mg/ml; 1:50

dilution), washed, and viewed with an epifluorescence


Tissue Culture

Ovaries and follicles were cultured in a medium

consisting of 85% Leibovitz L-15 (Sigma) containing 1 mM

freshly prepared glutamine and 10 mM HEPES buffer, pH 7.6,

at 200 C (hereafter referred to as 85% L-15). This culture

media was empirically determined to be optimal with respect

to cell viability over extended culture periods.

Tritiated-thymidine incorporation was examined in

isolated longitudinal ovary segments containing the

germinal ridge. These ovary segments were cultured in 85%

L-15 medium and incubated at 200 C for periods up to 14

days. The incubation media was replaced every 2-3 days

with fresh media. On days 0, 2, 5, 7, 10, the ovary

fragments were pulsed with 20 uCi/ml methyl-3H-thymidine

(ICN; specific activity 6.7= Ci/mmol) for 24 hours and then

processed as described above for light microscopy. Thick

sections of these ovaries were cut and the slides coated

with NTB-2 autoradiographic emulsion (Kodak). Slides were

exposed for various times at 40 C and the emulsion

developed for 90 seconds at 680 C with 1 part Dektol

(Kodak) and 2 parts water. Sections were lightly stained

with toluidine blue after development.

The fluorescent dyes rhodamine 123 (Johnson et al.,

1980) and acridine orange (Allison and Young, 1969;

Albertini, 1984) were used to assess the intracellular

localizations of mitochondria and lysosomes, respectively.

Intact ovary segments were incubated in 85% L-15 containing

either rhodamine 123 (10 ug/ml) or acridine orange (1-10

ug/ml). Ovaries were incubated for 30-60 minutes at 200 C

and then rinsed three times for 10 minutes each in media

lacking the dye. The ovaries were pinned out on a

microscope slide having silicon borders to allow

visualization of the entire ovarian sheet. These


preparations were observed with phase optics and

epifluorescence microscopy. Rhodamine 123 was visualized

using a rhodamine bypass filter. Acridine orange

localization was determined with fluorescein and rhodamine

bypass filters in combination to discriminate between

ribonucleoprotein- and lysosome-specific staining. [The

rhodamine bypass filter eliminates the background green

fluorescence that is due to the ribonucleoprotein and

allows visualization of the lysosomal network (Albertini,


Oocyte competence to undergo oocyte maturation in

vitro was determined after culture media optimization. In

preliminary experiments, various steroids were tested for

the ability to stimulate oocyte maturation in vitro. The

steroid 17-alpha-hydroxy-20-beta-dihydroprogesterone

(17-alpha,20-beta-DHP) was found to be the most effective

and therefore was utilized for further experiments.

Individual follicles were dissected free of ovarian stroma

in solution FO using watchmaker forceps. Oocytes were

separated into various size groups (0.7, 0.8, 0.9, 1.0, 1.1

mm diameter) and checked for the presence of a germinal

vesicle (oocyte nucleus). Follicles from several animals

were pooled for each experiment. For each size group,

follicles were placed in 10x35 mm petri dishes containing 3

ml of 85% L-15 media. Experimental treatment groups

received 17-alpha,20-beta-DHP (1 ug/ml final concentration)


in 10 ul ethanol carrier. Control treatment groups

received 10 ul of ethanol alone added to the media.

Follicles were incubated at 200 C and scored for germinal

vesicle breakdown after 40 hours of incubation using a

dissecting microscope. Follicles that underwent atresia

during the culture period were discounted from the results.

Protein content for various size follicles was

determined according to the Pederson modification of the

Lowry method (Pederson, 1977). Follicles were dissected

free of stromal tissue and homogenized in 100 ul of

solution FO. Follicles larger than 0.9 mm diameter were

homogenized as described and the sample divided into two

aliquots with the sum of these aliquots giving the total

protein content per follicle value. Homogenates were then

processed exactly as per the Peterson procedure. Bovine

serum albumin was used to generate the protein standard



Stage I: Oogonia (approximately 10 um diameter)

The oogonia constitute the stem cells of the germ

cell lineage. Oogonia are found in the luminal epithelium

near the germinal ridge and within the germinal ridge

itself. The germinal ridge is an outpocketed region of the

ovarian luminal epithelium (Begovac and Wallace, 1987).

Oogonia are oval to somewhat spherical in shape and are

conspicuous by their smaller size and large

nuclear-to-cytoplasmic ratio (Fig. 26). Oogonial nuclei

are oval in shape and contain one to three nucleoli.

Dense-cored granules of unknown composition are also

observed within the nucleus (Fig. 26). These dense-cored

granules are specific to nuclei of the germ cell lineage

and are also present in larger oocytes. The oogonial

chromatin is somewhat electron dense compared to the more

electron lucent nature of chromatin found in other meiotic

oocytes. The oogonial cytoplasm contains mitochondria,

ribosomes, and scant endoplasmic reticulum, but appears to

lack Golgi complexes. Aggregates of nuage material are

frequently associated with the mitochondria. Nuage

provides a good, reliable marker for germ cells (Eddy,

1975). Oogonia are contiguous with other germ cells and

somatic cells but are not in immediate contact with the

basal lamina of the luminal epithelium or that surrounding

the germinal ridge.

To identify proliferating germ cells, ovary segments

were pulsed with 3H-thymidine. These studies demonstrated

that only germ cells within the germinal ridge incorporated

thymidine (Figs. 27, 28); oocyte nuclei at later stages in

the follicle progression did not incorporate 3-thymidine.

Germ cells were observed to incorporate 3H-thymidine even

after being cultured in vitro for 2, 5, 7, and 10 days

(Fig. 28). Somatic cells, including follicle cells and

Fig. 26. Electron micrograph of an oogonium (OG)
present within the germinal ridge. Note the oval shape of
the nucleus and the single nucleolus. Mitochondria, nuage
material (arrows), and endoplasmic reticulum are present
within the cytoplasm. Some dense-cored granules are seen
above the nucleolus (arrowhead). x 14,000.

Fig. 27. Light micrograph of germinal ridge region of
ovary that had been pulsed with 3H-thymidine after 2 days
in culture. Note the heavy concentration of silver grains
(arrows) that completely cover the nuclei of germ cells.
Note the unlabeled germ cell nucleus (arrowhead) in the
germinal ridge and the larger oocyte (0) with an unlabeled
nucleus. JB-4 resin, toluidine blue. x 470.

Fig. 28. Light micrograph of germinal ridge segment of
ovary pulsed with 3H-thymidine after 10 days in culture.
Germ-cell labeling (arrows) still occurs after this
culture period. JB-4 resin, toluidine blue. x 510.

Fig. 29. Electron micrograph of leptotene
chromatin-nucleolus oocyte (0). Cellular organelles are
located in a juxtanuclear position around the centrioles
(arrow). Note that Golgi complexes (arrowheads) have
appeared. The nucleus is spherical and the chromatin
remains electron dense. x 12,640.

Fig. 30. Electron micrograph of chromatin-nucleolus
stage oocyte (0) in pachytene. Long synaptonemal
complexes (arrows) and micronucleolar fragments are
present in the nucleus. Note that the nucleus is more
electron lucent than those shown in Figures 26 and 29.
Dense-cored granules (arrowhead) can also be observed in
the nucleus. x 8,080.

Fig. 31. Higher magnification electron micrograph of
dense-cored granules specific for germ cell nuclei. x



26 2.28

Ai .



29 3


connective tissue cells, incorporated thymidine during

these time periods as well.

Stage IIa: Primary Growth, Chromatin-Nucleolus Phase
(approximately 10-20 um diameter)

The primary growth stage is initiated by true

oogenesis, or oocyte formation. It is the period of

follicle formation and an increase in the amount of

cytoplasm and cellular organelles within the oocyte.

Chromatin-nucleolus oocytes are found within the

germinal ridge and have not yet become incorporated into

definitive follicles. They are formed by the

transformation of oogonial daughter cells into prophase I

meiotic oocytes. It is unclear as to how this process

occurs and whether the oogonial division that produces the

oocytes is symmetric or asymmetric. It is difficult to

positively identify leptotene oocytes from the transforming

oogonia, but loss of the perinuclear ring of mitochondria

and the appearance of Golgi elements (Fig. 29) appear to be

noticeable changes that distinguish an early oocyte from an

oogonium. The organelles are localized in a polarized

juxtanuclear aggregation that seems to form around the

centrioles (Fig. 29). The nucleus of leptotene oocytes

become more spherical in shape as compared to oogonial

nuclei. The nucleus of leptotene oocytes also contains

dense-cored granules and the chromatin remains somewhat

electron dense. Oocytes in zygotene and pachytene can more

easily be distinguished by the appearance of synaptonemal

complexes within the nucleus (Fig. 30). During zygotene,

the synaptonemal complexes are short, as initial

chromosomal pairing occurs, and nucleoli can still be

observed in the nucleus. By pachytene, the synaptonemal

complexes are long and the nucleoli have become more

dispersed and fragmented (Fig. 30). Zygotene and pachytene

nuclei also become noticeably less electron-dense when

compared to oogonia and leptotene nuclei (Figs. 26, 29,

30). The nuclei of later chromatin-nucleolus oocytes also

contain dense-cored granules (Figs. 30, 31). Oocytes in

zygotene and pachytene grow slightly, with an increase in

cell organelles including mitochondria, Golgi complexes,

ribosomes, and endoplasmic reticulum. These organelles are

typically located in a juxtanuclear aggregate within the

oocyte. Nuage continues to be present in the cytoplasm,

often associated with mitochondria. Leptotene, zygotene,

and pachytene oocytes generally share at least part of

their cell borders with contiguous oocytes, and the

remaining cell surfaces are in contact with prefollicular

cells within the germinal ridge.

The later chromatin-nucleolus oocytes become arrested

in early diplotene of meiotic prophase I as synaptonemal

complexes dissociate within the oocyte nucleus. During

this phase, the ooplasmic matrix increases along with the

numbers of cell organelles. The organelles tend to remain

polarized in a juxtanuclear location. Nucleoli of typical


structure reappear adjacent to the inner layer of the

nuclear envelope. The appearance of multiple nucleoli

seems to coincide with the formation of the definitive

follicle as it separates from the germinal ridge. Final

follicle formation occurs with these diplotene-arrested

oocytes. The definitive follicle now consists of an oocyte

surrounded by a monolayer of flattened follicle cells with

the entire complex enclosed by a basal lamina. Sparse

thecal elements associate with the follicle external to the

basal lamina at this stage. The details of

folliculogenesis have been presented elsewhere (Begovac and

Wallace, 1987).

Stage lib: Primary Growth, Perinucleolar Phase (20-140 um

This phase of primary growth typically begins as the

definitive follicle separates from the germinal ridge. In

the light microscope, the perinucleolar oocytes are

transparent with the nucleus clearly visible. The nucleus

is generally spherical and contains multiple nucleoli in a

perinuclear position adjacent to the inner layer of the

nuclear envelope (Figs. 32, 33, 34). Dense-cored granules

can still be observed in oocyte nuclei. During this stage,

the nucleus increases in size and the nucleoli increase in


Perinucleolar oocytes also increase in size due to

the continued elaboration of cytoplasm, mitochondria,

Golgi, ribosomes, endoplasmic reticulum, and multivesicular

Fig. 32. Light micrograph of perinucleolar oocytes in
the primary growth phase. Note the multiple nucleoli in a
perinuclear position within the germinal vesicle (GV).
The Balbiani vitelline body is also clearly visible in two
of the oocytes, first as a juxtanuclear aggregate (arrow)
and later assuming a crescentlike shape (arrowheads).
JB-4 resin, H&E. x 330.

Fig. 33. Light micrograph of perinucleolar phase oocyte
illustrating perinuclear ring organization of the Balbiani
vitelline body (arrows). (GV) Germinal vesicle. JB-4
resin, H&E. x 310.

Fig. 34. Electron micrograph of a perinucleolar oocyte
with the Balbiani vitelline body containing mitochondria
and an aggregate of multivesicular bodies (arrows).
Oocyte nucleus (NU) contains perinuclear nucleoli
(arrowhead). (BL) Basal lamina. x 7,500.

Fig. 35. Higher magnification of multivesicular bodies
[one of which is labeled with an (*)] associated with the
Balbiani vitelline body. x 27,300.

Fig. 36. Later stage perinucleolar oocyte (0) with
mitochondria and Golgi complexes (G) in the peripheral
ooplasm. A basal lamina (arrowhead) overlies the squamous
follicle cells (FC). Note that the oocyte has started to
elaborate numerous microvillar processes (arrow). x 7,800.

Fig. 37. Photomicrograph of a portion of intact ovary
that has been stained with rhodamine 123 to follow
movements of mitochondria in developing oocytes. Note the
regions of juxtanuclear fluorescence in the smallest
oocytes (arrow). As oocytes increase in size, the
staining becomes perinuclear and finally disperses
throughout the oocyte (arrowhead). x 85.

Fig. 38. Photomicrograph illustrating the lysosomal
localizations in developing oocytes using acridine orange
staining. Note that the staining pattern is similar to
that of rhodamine 123 (Fig. 37). Initially a juxtanuclear
staining (arrow) is observed, followed by a
crescent-shaped staining (arrowhead), becoming
perinuclear, and finally dispersing throughout the oocyte
(*). x 82.

M. F.


bodies (Figs. 34, 35, 36). In early perinucleolar oocytes,

most of these cellular organelles are typically present as

a juxtanuclear mass (Fig. 32, 34) commonly known as the

Balbiani vitelline body (Guraya, 1979) or mitochondrial

cloud (Heasman et al., 1984). Mitochondria and

multivesicular bodies comprise the major components of the

Balbian! vitelline body (Fig. 34). Multivesicular bodies

initially appear during the middle of the perinucleolar

stage, are heterogeneous with respect to size, shape, and

contents (Fig. 35), and remain into early vitellogenesis.

As stage lib progresses, the juxtanuclear Balbiani

vitelline body changes to form first a crescent shape (Fig.

32), then progresses to a perinuclear ring (Fig. 33) that

finally disperses throughout the oocyte by late

perinucleolar growth. This progression can be also

visualized in living oocytes using the vital dyes rhodamine

123 for mitochondria (Fig. 37) or acridine orange for

acidic, lysosomal elements (Fig. 38).

In the early perinucleolar phase, the peripheral

ooplasm contains cytoplasmic matrix, ribosomes, and smooth

endoplasmic reticulum, while the oocyte surface is

typically smooth. After the Balbiani vitelline body

disperses in late perinucleolar growth, mitochondria and

multivesicular bodies are found throughout the oocyte

cytoplasm, whereas the abundant Golgi complexes become

mostly confined to the peripheral cytoplasm (Fig. 36). The


oocyte has also begun to elaborate numerous microvillar

processes towards the overlying follicle cells (Fig. 36).

The follicle cells also extend microvilli, but do so

later than the oocyte. Follicular investments surrounding

perinucleolar oocytes include squamous follicle cells,

connective tissue elements, and blood vessels. The

follicle cells possess a large nucleus and primarily

mitochondria and rough endoplasmic reticulum in the

cytoplasm, with only scant Golgi complexes.

Stage III: Cortical Alveoli Formation (140-260 um diameter)

This stage is characterized by the initial appearance

of three components: 1) cortical alveoli; 2) vitelline

envelope; and 3) lipid (Fig. 39). The cortical alveoli and

vitelline envelope initially appear at approximately the

same time (Fig. 40a). Cortical alveoli, indicated by

PAS-positive reactivity, are first observed in the light

microscope throughout the peripheral cytoplasm of oocytes.

This is confirmed by ultrastructural observations

indicating that the cortical alveoli are membrane-limited

structures having a homogeneous appearance (Fig. 40a). The

smaller cortical alveoli sometimes contain a denser

inclusion that is not membrane bound (Fig. 40b). As

oocytes increase in size, the cortical alveoli increase in

number and become heterogeneous in size as they fill much

of the oocyte cytoplasm. The larger cortical alveoli stain

less intensely with PAS, while in the electron microscope,

Fig. 39. Light micrograph of cortical alveoli-stage
oocyte with much of the oocyte filled with larger sized
cortical alveoli (CA). The vitelline envelope (VE) is
present between the oocyte and overlying follicle cells.
Lipid (LP) is present in a perinuclear position. Multiple
nucleoli are still present in the germinal vesicle (*).
JB-4 resin, H&E. x 140.

Fig. 40. a) Electron micrograph of the cortex in an
early cortical alveoli-stage oocyte. The vitelline
envelope (VE) has appeared between the microvillar
processes and cortical alveoli (CA) are present. Note the
Golgi complexes (G) intermingled with the mitochondria.
(FC) Follicle cell. x 10,700. b) Cortical alveolus with
electron dense inclusion (arrow). x 19,200.

Fig. 41. Vitelline envelope (VE) of later cortical
alveoli-stage oocyte (0) now having trilaminar appearance
due to continued formation. (FC) Follicle cell. x 12,240.

Fig. 42. Electron micrograph of lipid (LP) in cortical
alveoli-stage oocyte. The void in the center of the lipid
spheres represents an artifact caused by partial
extraction of the lipid during fixation and processing.
x 13,920.

Fig. 43. a) Phase micrograph of cortical alveoli-stage
oocyte (CAO) and vitellogenic oocyte (VO). x 325. b)
Fluorescence photomicrograph of same section treated with
the fluorescent lectin, Con A. Note the positive staining
of the different size cortical alveoli populations
(arrows). The smaller cortical alveoli generally react
most intensely. Note also that the vitelline envelope
stains (arrowheads) with this lectin. x 325. c) Control
micrograph containing sequentially cut section of same
block treated with 0.5 M glucose and the fluorescent
lectin. No reactivity is observed. x 325.



they are more electron lucent and appear to contain a

filamentous network. Additional details regarding cortical

alveoli may be found in Anderson (1968).

The vitelline envelope appears coincident with

cortical alveoli in follicles of approximately 140 um

diameter. The formation of this structure has been

previously described (Anderson, 1967). In the light

microscope, the vitelline envelope is first observed as a

thin band between the oocyte and overlying follicle cells

(Fig. 39); it also stains positively with PAS.

Ultrastructurally, the vitelline envelope first becomes

apparent with the appearance of a homogeneous material

between the microvillar processes of the oocyte (Fig. 40a).

As the follicle reaches approximately 200 um diameter, a

trilaminar appearance of the vitelline envelope is attained

(Fig. 41), with architecturally complex ZI, Z2, and Z3

layers (terminology of Anderson, 1967). In the later

cortical alveoli-stage oocytes, the Z3 layer becomes

thickened relative to Zl and Z2.

A third process that occurs in cortical alveoli-stage

oocytes is the formation and accumulation of lipid

droplets. Lipid accumulation is first apparent in

follicles approximately 200 um in diameter. The lipid

droplets at this size are small and aggregated in

perinuclear positions (Figs. 39, 42). The lipid mass

continues to enlarge in a perinuclear position in later

cortical alveoli-stage oocytes. In freshly isolated

oocytes, the lipid has a dark appearance while later on

during oocyte development, the lipid droplets increase in

size and appear brilliant orange in the living oocyte. The

processes of lipogenesis and lipid accumulation during

oocyte development are poorly understood.

During the cortical alveoli stage, the oocyte surface

begins to display pinocytic vesicles at the base of the

microvillar processes. Microvilli continue to lengthen as

the vitelline envelope thickens. The follicle cells also

elaborate microvilli that begin to interdigitate with the

oocyte processes. Follicle cells remain somewhat squamous

in shape and contain mitochondria and rough endoplasmic

reticulum, while Golgi complexes become more abundant. The

specialized micropylar cell also appears during this stage

and is involved in the formation of the micropyle.

Concanavalin A treatment of oocytes demonstrated that

the cortical alveoli and the vitelline envelope bind this

lectin (Figs. 43a, b). Staining is abolished in the

presence of 0.5 M glucose (Fig. 43c) and/or mannose.

Cortical alveoli bind concanavalin A throughout oocyte

growth although the intensity of staining is lower in the

larger cortical alveoli, presumably due to a loss of

vesicular contents during tissue preparation. The

vitelline envelope binds this lectin stains throughout

oocyte growth as well.


Stage IV: Vitellogenesis (260-1,100 um diameter)

Vitellogenesis involves the accumulation of

exogenously derived yolk protein precursors into the oocyte

and is responsible for the majority of oocyte growth

(Wallace, 1985). Oocytes in early vitellogenesis are

characterized in the light microscope by the appearance of

small yolk spheres in the oocyte periphery and interior

(Fig. 44). As oocyte growth proceeds, the interior yolk

spheres become heterogeneous in size (Fig. 45), while

smaller yolk spheres still are present in the peripheral

ooplasm. The accumulation of yolk spheres in the oocyte

interior results in the progressive displacement of

cortical alveoli and lipid to the peripheral cytoplasm.

The larger yolk spheres apparently form by fusion of the

smaller yolk spheres. In late vitellogenic oocytes, most

of the individual yolk spheres disappear and a large

central fluid yolk mass is formed (Fig. 46). The cytoplasm

containing small yolk spheres, cortical alveoli and lipid

is displaced to a thin rim at the oocyte periphery.

At the ultrastructural level, yolk spheres are

somewhat electron-dense, membrane-limited structures (Figs.

47-49). They also contain highly electron-dense inclusions

(Figs. 47-49). The ultrastructural appearance of this

inclusion remains unchanged in osmicated and nonosmicated

tissues, but the inclusion occasionally has a fine granular

appearance. Although, the biochemical nature of the dense

Fig. 44. Light micrograph of early vitellogenic oocyte
containing numerous small yolk spheres (Y) within its
interior. Note the displacement of cortical alveoli (CA)
to the oocyte periphery. Lipid (LP) is still present
internally near the germinal vesicle and amongst the yolk
spheres. x 120.

Fig. 45. Light micrograph of midvitellogenic oocyte in
which the yolk spheres (Y) are becoming larger in size.
Cortical alveoli (CA) and lipid become more progressively
displaced to the peripheral ooplasm. x 255.

Fig. 46. Light micrograph of a late vitellogenic stage
oocyte in which most of the yolk spheres (Y) have
coalesced to form a large central fluid yolk mass. Lipid
droplets (*) remain in the peripheral part of the yolk
mass along with cortical alveoli (CA). Vitelline envelope
(arrowhead). x 150.

Fig. 47. Transmission electron micrograph depicting
early vitellogenic oocyte with yolk spheres in the process
of being formed. The primary yolk spheres (YI) containing
condensing yolk and vesicular structures are illustrated.
Fully formed yolk spheres (YIII) containing the highly
electron dense, yolk-specific inclusions (arrow) are also
present. x 17,400.

Fig. 48. Electron micrograph of early vitellogenic
oocyte with transitional yolk-spheres (YII). Note the
condensing yolk material and the presence of vesicular
structures in these transitional yolk spheres. Note in
particular the transitional yolk sphere (YII) containing
the electron-dense, yolk-specific marker (arrow) in
addition to vesicular elements. (LP) Lipid. x 20,300.

Fig. 49. Large mature yolk sphere (YIII) of
midvitellogenic oocyte containing a heterogeneous
population of the yolk-specific inclusions (arrowheads).
Note that the yplk spheres are membrane bound (arrows).
Other smaller yolk spheres are also present. x 12,800.

Fig. 50. Electron micrograph of oocyte periphery
illustrating the pinocytic nature of vitellogenic oocytes.
Note that these pinocytic vesicles are clathrin-coated
(arrows). (VE) Vitelline envelope. x 36,600.


-- 4 CA

45 4


47 4'

49 -50

inclusion is unclear, it does provide a specific, reliable

marker for following yolk formation.

Early vitellogenic oocytes possess many small,

somewhat electron-dense structures at the oocyte periphery

and interior among the cortical alveoli (Fig. 47). There

are several populations of these membrane-bound structures

including primary, transitional and mature yolk spheres.

The primary spheres appear to contain proteinaceous

material interspersed among multivesicular elements (Fig.

47). The intermediate, transitional spheres contain

proteinaceous material, variable vesicular elements, and

the yolk-specific, electron-dense inclusion (Fig. 48). The

mature spheres contain homogeneous condensed yolk and a

heterogeneous size and distribution of the yolk-specific

marker (Fig. 49). Yolk sphere formation is first apparent

in the oocyte periphery, with accumulation occurring in the

internal regions of the oocyte as the mature yolk spheres

fuse with one another. Oocytes continue to display

pinocytic vesicles during vitellogensis (Fig. 50). These

vesicles are clathrin-coated and most likely are involved

in the sequestration of yolk protein precursors as has been

documented in other animals (Wallace, 1985).

Several other events occur during the vitellogenic

stage. By midvitellogenesis, the vitelline envelope has

lost its trilaminar appearance and has undergone

condensation, particularly of the Z2 and Z3 layers. Lipid


droplets continue to accumulate and increase in size. The

lipid becomes brilliantly colored and consequently

contributes the major coloration of later stage oocytes.

Cortical alveoli probably continue to be formed during

vitellogenesis as well (Selman et al., 1986).

Follicle cells surrounding vitellogenic oocytes

become somewhat cuboidal in shape. The follicle cells

possess abundant rough endoplasmic reticulum, mitochondria,

and Golgi complexes. Follicle-cell microvillar processes

extend deep into the vitelline envelope pore canals

approaching the oocyte surface. Both follicle-cell and

oocyte microvillar processes are present within a single

pore canal.

Stage V: Oocyte Maturation (1,100-1,300 um diameter)

When oocytes reach this stage, they become capable of

resuming first meiotic division by undergoing germinal

vesicle breakdown. Structurally, the postvitellogenic

oocyte contains a large central fluid yolk mass and has at

its periphery a thin rim of cytoplasm containing mostly

cortical alveoli and lipid (Fig. 46). The vitelline

envelope is highly compacted and possesses pore canals

prior to maturation. However, at some undefined point

during maturation and prior to ovulation, the pore canals

disappear as the oocyte and follicle-cell microvillar

processes are retracted or lost.

Figs. 51-53. Photomicrographs of maturation-stage
follicles and mature eggs demonstrating both the animal
pole (left) and vegetal pole (right) in each pair.

Fig. 51. Prematuration follicles with disperse lipid
droplets throughout the oocyte and the germinal vesicle
(arrow) clearly visible beneath the animal pole.

Fig. 52. Postmaturation follicles that have been
matured in vitro by treatment with the steroid
17-alpha,20-beta-DHP. Note the loss of the germinal
vesicle over the animal pole (left follicle). x 18.

Fig 53. Normal appearance of in vivo matured eggs.
Note the loss of the germinal vesicle (arrow) and the
lipid redistribution towards the animal pole. x 18.

Fig. 54. Maturation response of various size follicles
to treatment with the steroid 17-alpha,20-beta-DHP.
Values represent the means and SEM of the percentage of
follicles within each size pool that underwent germinal
vesicle breakdown (GVBD). Experimental and control values
are indicated and represent three separate experiments.
Follicles (10-40) were incubated in each follicle size
pool. There in an increase in competence to undergo in
vitro maturation with respect to follicle diameter.
Follicles that are 1.1,mm in diameter are fully competent
to undergo 100% germinal vesicle breakdown. Control
groups receiving ethanol vehicle (EtOH) showed no response
except for a single case in the 1.1 mm diameter group.

M 17a, 20A -DHP
80 M EtOH

am 60


0.7 0.8 0.9 1.0 I.I
Follicle diameter (mm)


The germinal vesicle is located near the animal pole

just beneath the oocyte surface (Fig. 51). The

identification of the maturational stage has been

established by examining fresh ovaries and using in vitro

follicle culture to define the size at which oocytes become

fully competent to respond to steroid stimulation by

undergoing germinal vesicle breakdown. The results of the

culture experiments (Fig. 54) indicate that follicles of

1.1 mm diameter undergo 100% germinal vesicle breakdown in

response to 17-alpha,20-beta-DHP. Oocytes matured in vitro

have a similar appearance to in vivo matured oocytes (Figs.

52, 53) except for being smaller in size. Lipid droplets

in prematurational oocytes are generally distributed around

the entire oocyte periphery prior to maturation. After

maturation, in vivo or in vitro, most lipid droplets

aggregate towards the animal pole region where the oocyte

nucleus was previously located. The vegetal pole therefore

becomes cleared with only scattered lipid droplets still

remaining. There is an increased intensity of the orange

coloration of the oocyte that is a result of the clearing

process. After germinal vesicle breakdown, the oocytes

ovulate into the ovarian lumen and become mature eggs.

A determination of protein content in different sized

follicles indicated that a slight change in the rate of

protein accumulation occurs as oocytes reach the

maturational stage and become eggs (Fig. 55). No change in


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Fig. 56. SDS-PAGE profiles of predominant proteins in
follicles and eggs <0.2-1.3 mm diameter. Follicle
diameters are listed above their respective lanes.
Molecular weight standards are listed to the left of the
gel. Note that no change in protein patterns occurs as
oocytes undergo maturation (1.1-1.2 mm diameter) and
become mature egg (1.3 mm diameter). Note also that the
major protein constituents of all follicles above 0.4 mm
diameter are the various yolk proteins, which are distinct
from the the cytoplasmic proteins indicated for the
smallest (yolk-free) oocytes. Approximate protein content
of samples are: < 0.2 mm, no value; 0.3 mm, no value; 0.4
mm, 7.9 ug; 0.5 mm, 16.0 ug; 0.6 mm, 27.1 ug; 0.7 mm, 20.5
ug, 0.8 mm, 27.8 ug, 0.9 mm, 25.1 ug; 1.0 mm, 35.0 ug; 1.1
mm, 39.3 ug; 1.2 mm, 44.1 ug; 1.3 mm, 50.8 ug.

Follicle Diameter Immi

ddd d d d d M

200- :i




22- .



the major yolk proteins takes place during maturation (Fig.

56), unlike the situation recently found for the oocytes of

several other teleosts (Greeley et al., 1986b).

Stage VI: Mature egg (approximately 1,300 um)

This stage is the culmination of oocyte development

and represents a cell capable of being fertilized.

Macroscopically, the egg is oval to pear-shaped and

brilliantly colored orange because of the lipid droplets.

The nucleus.has undergone dissolution as a result of final

maturation and the egg and the animal pole has an opaque

appearance (Fig. 53). The mature egg retains the large

central fluid yolk mass. Cortical alveoli are uniformly

distributed around the oocyte periphery beneath the

oolemma. The lipid droplets, however, become more

aggregated towards the animal pole while the vegetal pole

has fewer lipid droplets and is more clear (Fig. 53). The

vitelline envelope consists of a thin Zl layer and highly

compacted Z2, Z3 layers and lacks the pore canals seen in

the envelope surrounding oocytes. The micropyle is visible

on the outer egg surface since there are no cellular

investments surrounding the egg. Eggs are stored within

the ovarian lumen until the time of fertilization and are

somewhat fragile. After fertilization, they become turgid

when the vitelline envelope elevates to form the chorion.


Oocyte development in the pipefish is considered to

be asynchronous (Wallace and Selman, 1981), with a

heterogeneous population of different-stage oocytes. Many

of the cellular processes that occur during oocyte

development in the pipefish parallel those found in other

nonmammalian vertebrates. These include oogonial

proliferation (Tokarz, 1978), formation and dispersal of

the Balbiani vitelline body (Guraya, 1979; Heasman et al.,

1984), formation of cortical alveoli and a vitelline

envelope (Anderson, 1967, 1968; Selman et al., 1986),

vitellogenesis (Anderson, 1968; Selman and Wallace, 1983;

Wallace, 1985) and oocyte maturation (Masui, 1985; Greeley

et al., 1986a). The unique features of oogenesis in the

pipefish, however, relate to the ovarian anatomy whereby

oocytes are sequentially arranged according to

developmental age (Begovac and Wallace, 1987). This

arrangement is particularly useful when examining specific

cellular events such as oogonial dynamics, vitelline

envelope formation, and vitellogenesis. One is able to

study such processes with respect to a developmental time

frame on a single cross section of the ovary in which all

oocyte stages have been treated in the same manner.

Additionally, the very earliest events of oogonial

proliferation and early follicle formation, because of

their specific location within the ovary, become accessible


to experimental manipulation. Experiments of this nature

are much more difficult in other vertebrate ovaries because

of the random organization of developing oocytes.

Therefore, many of the advantageous features for studying

oogenesis in the pipefish stem from this temporal and

spatial pattern of oocyte development. The schematic

diagram shown in Figure 57 highlights oocyte development in

the pipefish and summarizes the cytological and

physiological changes seen during specific stages.

The initial phase of oocyte development involves the

true genesis of an oocyte as a result of oogonial

proliferation and differentiation. The dynamics of

cellular divisions that result in this transformation are

unclear. The period of oogonial proliferation varies from

species to species and may be continuous or cyclical in

teleosts (reviewed by Tokarz, 1978). In the pipefish,

oogonial proliferation is probably continuous because fully

developed ovaries are present throughout the year (Begovac

and Wallace, 1987). The present 3H-thymidine experiments

indicate the qualitative labeling of proliferating germ

cells. Complete nuclear labeling with 3H-thymidine will

occur in mitotically active cells and preleptotene meiotic

germ cells, whereas localized labeling will be observed

over nucleoli in the amplification state (Coggins and Gall,

1972; Coggins, 1973). Consequently, proliferating oogonia

and preleptotene oocytes are the two populations of germ

Fig. 57. Schematic diagram illustrating sequential
stages of oocyte development as viewed in an optimal
section. This schematic, in conjunction with a schematic
of the ovarian anatomy (Chapter II, Figure 25) provides a
reconstruction of oocyte development within the ovarian
structure. The orientation of this drawing reflects the
general relationships that oocytes possess in vivo. Stage
designations are indicated as Roman numerals on the inner
layer. The positions of the oocytes up through the first
vitellogenic oocyte accurately reflect the approximate
position of these stages in the developmental spiral.
Later vitellogenic stages are drawn in for completeness of
stages observed. Nuclei are depicted in most of the
oocytes. The vitelline envelope and follicular
investments have not been illustrated. The label (VI) is
positioned over the animal pole of the mature egg. This
schematic was prepared to highlight the reorganization of
the major cytological features within the oocytes as they
progress through the oocyte developmental spiral. It is
not fully drawn to scale.

Balbiani vitelline
body / \

I. Oogonia
U. Primary Growth
a. Chromatin-Nucleolus Phase
b. Perinucleolar Phase

MT. Cortical Alveoli Formation
3. Vitellogenesis
3. Oocyte Maturation
ME. Mature Egg

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