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Distribution of actin in isolated seminiferous epithelia and denuded tubule walls of the rat.

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THE ANATOMICAL RECORD 213:63-71 (1985)
Distribution of Actin in Isolated Seminiferous
Epithelia and Denuded Tubule Walls of the Rat
Department ofdnatomy, Faculty of Medicine, University of British Columbia, Vancouver,
British Columbia, Canada V6T 1 W5
We have studied the distribution of actin, using NBD-phallacidin a s
a probe, in isolated sheets of seminiferous epithelia and denuded tubule walls of the
rat. Sheets of intact seminiferous epithelia were separated from tubule walls using
EDTA in PBS. The isolated epithelia and denuded tubule walls were fixed, mounted
on slides, made permeable with cold acetone ( -2OoC), and then treated with NBDphallacidin.
Actin was observed in myoid cells, in ectoplasmic specializations of Sertoli cells,
and in Sertoli cell regions adjacent to tubulobulbar processes of late spermatids. In
myoid cells, filament bundles course in circular and longitudinal directions relative
to the tubule wall. In Sertoli cells viewed at an angle perpendicular to the epithelial
base, actin filaments in ectoplasmic specializations adjacent to junctional complexes
circumscribe the bases of the cells. Filament bundles in ectoplasmic specializations
adjacent to germ cells closely follow the contour of and are arranged parallel to the
long axis of the developing acrosome. Sertoli cell regions adjacent to tubulobulbar
processes of late spermatids stain intensely with NBD-phallacidin.
Isolated seminiferous epithelia, combined with NBD-phallacidin as a probe for
actin, provide a n ideal model system in which to study further the contractile
properties of Sertoli cell ectoplasmic specializations and the possible involvement of
these structures in events that occur during spermatogenesis.
Sertoli cells are thought to be involved with many of
the morphogenetic events that occur in the mammalian
seminiferous epithelium during spermatogenesis (Fawcett, 1975; Russell, 1980a; Vogl et al., 1983). Among
these events are the movement of spermatocytes through
the blood testis barrier, the translocation of spermatids
to the luminal surface of the tubule, and the release of
spermatozoa from the epithelium. Although the underlying mechanisms of these events are unknown, the
elaborate cytoskeletal network of Sertoli cells is likely
involved (Fawcett, 1975; Means et al., 1980; Russell,
1980a; Vogl et al., 1983).
Ectoplasmic specializations (Russell, 1977) are a particularly interesting group of Sertoli cell structures that
are partly composed of cytoskeletal elements and that
may be involved with some of the events described above.
These structures occur adjacent to the basally situated
junctions between Sertoli cells and in regions of adhesion to germ cells. They are characterized by a dense
layer of filaments sandwiched between the plasma membrane and a cistern of endoplasmic reticulum (Brokelmann, 1963; Flickinger and Fawcett, 1967; Nicander,
1967; Dym and Fawcett, 1970). Because these regions
contain actin (Toyama, 1976; Franke et al., 1978) and
possess a magnesium-dependent ATPase activity (Gravis
et al., 1976),they are generally thought to be contractile.
Using techniques similar to those developed for studying intestinal epithelia (Bjerknes and Cheng, 19811, we
have recently succeeded in isolating intact sheets of
(i, 1985 ALAN
seminiferous epithelia from the testes of ground squirrels (Vogl and Soucy, 1985). We are using this isolation
procedure, together with NBD-phallacidin as a probe for
filamentous actin (Barak et al., 19801, to study further
the structure and function of Sertoli cell ectoplasmic
specializations. This methodology has allowed us to visualize the three-dimensional arrangement of actin filament bundles in Sertoli cells and in myoid cells of
seminiferous tubules.
We initiated this study to determine the arrangement
of actin filament bundles in seminiferous tubules of the
rat. We chose t,he rat because it is used by most investigators as the model system in which to study mammalian spermatogenesis.
Three adult male Sprague-Dawley rats (392, 396, 375)
were used for these studies.
Seminiferous epithelia and denuded tubule walls were
obtained in the following manner. The testes were excised from animals anesthetized with sodium pentobarbitone administered intraperitoneally and perfused, via
the spermatic artery on the posterior aspect of each
testis, with phosphate-buffered saline (PBS) (150 mM
NaC1; 5 mM KC1; 0.8 mM KHzPO,; 3.2 mM NazHPO,;
Received August 27, 1984; accepted March 28, 1985
adjusted to pH 7.3 with 0.1 N NaOH) containing 20 mM
EDTA (ethylenediamine-tetraacetic acid). After 5 minutes, perfusion was stopped and the testes were transferred to a petri dish containing 5 mM EDTA in PBS.
The organs were then decapsulated and the seminiferous tubule masses cut, using two scalpels in a scissorlike
fashion, into small segments. During this procedure,
seminiferous tubules separated from interstitial tissue,
and sheets of seminiferous epithelium separated from
tubule walls. Identification of the various components
was faciliated by using a Zeiss Stereomicroscope SR
fitted with a darkfield condensor. Interstitial tissue was
teased away from tubular elements, and the latter, containing epithelial sheets, denuded tubule walls, and intact tubules, were collected with a polyethylene pipette
and transferred to a 15-ml centrifuge tube. Total time
from decapsulating a testis to completing the harvesting
of tubular elements was 10 minutes. All solutions used
in the protocol described above were at room
Samples to be used for fluorescence microscopy were
centrifuged at a low setting and the supernatants replaced with PBS containing 5 mM EDTA and 3.7% paraformaldehyde. After 10 minutes in fixative, the tissue
was washed three times with PBS. To help visualize
filaments associated with spermatogenic cells, some of
the material was mechanically fragmented by aspiration with a syringe fitted with first an 18 gauge then a
25 gauge needle. Isolated epithelia, denuded tubule
walls, and mechanically fragmented samples were
mounted together on polylysine-covered slides, treated
with cold acetone (-2O"C), and air-dried. Samples
mounted on slides were rehydrated for 10 minutes with
PBS, then exposed for 20 minutes, a t room temperature,
to one of the following: 1)PBS (control for autofluorescence); 2) PBS + 1.65 x
M NBD-phallacidin (fluorescent probe for filamentous actin); 3 ) PBS + 1.65 x
lop6 M NBD-phallacidin + 1.04 x lo-* M phalloidin
(competitive specificity control); 4) PBS + 1.04 x lop4
M phalloidin (control for phalloidin in treatment 3). The
slides were washed twice with PBS then mounted with
1:l (by volume) glycero1:PBS containing 0.02% sodium
Fluorescence was recorded on Tri-X film using Zeiss
Photomicroscope I11 fitted with filters used for detecting
fluorescein. Spermatids were staged using criteria established by Leblond and Clermont (1952).
Tissue for electron microscopy was processed as follows. A sample of tubular elements containing a mixture of epithelial sheets, denuded tubule walls, and
intact tubules was centrifuged a t a low setting and the
supernatant replaced with a fixative containing 1.5%
glutaraldehyde, 1.5% paraformaldehyde, and 0.1 M sodium cacodylate (pH 7.3). After 2 hours, the sample was
washed with buffer then some of the material was aspirated through first an 18 gauge then a 25 gauge needle.
All tissue was postfixed on ice for 1 hour with 1%Os04
in 0.1 M sodium cacodylate (pH 7.3).The material was
processed further using standard techniques for electron
microscopy. Thick sections (1 pm) were photographed
with a Zeiss Photomicroscope 111. Thin sections were
studied with a Phillips 300 operated a t 60 kV.
of intact seminiferous tubules, sheets of seminiferous
epithelium, and denuded tubule walls. These different
elements were visible with darkfield optics during the
isolation treatments and their presence was confirmed
in thick sections of fixed material (Figs. 1,2).
Two major features were evident in tissue viewed with
the electron microscope. First, the isolated sheets of
seminiferous epithelium lacked a basal lamina (data not
shown). Second, ectoplasmic specializations adjacent to
germ cells and Sertoli cell junctional complexes remained intact. In fact, ectoplasmic specializations of the
Sertoli cell often remained attached to spermatids that
had been mechanically dissociated from the epithelium
(Figs. 3,4).
Using NBD-phallacidin as a probe for actin, we observed strong fluorescence in myoid cells and in Sertoli
cells. In Sertoli cells, ectoplasmic specializations were
labeled, as were regions adjacent to tubulobulbar
Although the fluorescence emitted from labeled actin
in myoid cells was most clearly evident in denuded tubule walls, such as the one shown in Figure 5, it was also
visible in intact seminiferous tubules (not shown). In the
latter structures, the fluorescence pattern of labeled actin in myoid cells was visualized simultaneously with
that in Sertoli cell junctional complexes.
Within each flat and somewhat polygonally shaped
myoid cell, linear tracts of actin were observed extending across the cell from one border to another as shown
in Figure 5 . These tracts were oriented predominantly
in two directions, with those adjacent to one cell surface
organized in a circular manner relative to the seminiferous tubule wall and those adjacent to the other surface
arranged longitudinally.
The pattern of actin we observed in Sertoli cells was
very different from that in myoid cells. A strong fluorescence was emitted by actin in ectoplasmic specializations of junctional complexes and appeared to outline
the base of each Sertoli cell. When an epithelial sheet
was viewed perpendicular to the epithelial base, as in
Figure 6, a honeycomb pattern was observed. By adjusting the plane of focus, or by looking a t the edges of an
epithelial sheet where the elongate Sertoli cells were
attached by their lateral surfaces to the slide (Fig. 71,
the distribution of NBD-phallacidin in more apical epithelial regions could be observed. In apical regions, fluorescence was associated almost exclusively with
developing spermatids. Although certainly evident in
intact epithelial sheets, the fluorescence emitted from
actin in these regions was more easily studied in fragmented epithelia.
Unlike the fluorescence in regions adjacent to Sertoli
cell tight junctions, the pattern of fluorescence in apical
epithelial regions associated with spermatids was different a t different stages of spermiogenesis. Because 1)we
had demonstrated that ectoplasmic specializations of
Sertoli cells often remained attached to spermatids that
were mechanically dissociated from the epithelium, and
2) the fluorescence we observed appeared to closely follow the outer contours of the germ cells, we interpreted
most of the fluorescence as coming from actin in ectoplasmic specializations of Sertoli cells and not from the
germ cells themselves. The earliest stage of spermatid
with which staining was associated was approximately
Using the isolation procedure described above, we were stage 8 (Fig. 8a, a'). At this stage, ectoplasmic specialiable to obtain testicular samples containing a mixture zations occurred adjacent to the region of the spermatid
Fig. 1. Thick (1pm) section of isolated seminiferous epithelia (asterisk) and denuded tubule walls (arrows) obtained using techniques
described in the text. Bar = 100 pm. x 347.
Flg. 2. Thick (1 pm) section of an isolated sheet of seminiferous
epithelium. Notice that a tubule wall is absent. A group of three
spermatogonia is indicated by the arrow. Bar = 150 pm. x 198.
Fig. 3. Electron micrograph of a spermatid that has separated from
the seminiferous epithelium. An ectoplasmic specialization (arrow-
heads) of a Sertoli cell is attached to the region juxtaposed to the
acrosome. Bar = 2.5 pm. ~ 9 , 7 8 9 .
Fig. 4. A magnified view of the region indicated by the rectangle in
Figure 3. Filament bundles within the attached Sertoli cell ectoplasmic specialization are indicated by the arrowhead. Also indicated
are 1)the nuclear envelope of the spermatid, 2) inner acrosomal membrane, 3) outer acrosomal membrane, 4) plasma membrane of the
spermatid, 5) plasma membrane of the Sertoli cell, and 6) “outer”
membrane of a Sertoli cell cistern of endoplasmic reticulum. Bar =
0.50 pm. ~ 6 4 , 0 0 0 .
Fig. 5. Distribution of NBD-phallacidin in myoid cells of a denuded
seminiferous tubule wall. A single myoid cell is indicated by the
arrows. The fluorescence occurs as linear tracts that course across the
cells in predominantly two directions. Bar = 50 pm. x590.
Fig. 6. Actin distribution, as indicated by NBD-phallacidin, at the
base of an isolated sheet of seminiferous epithelium. The base of a
single Sertoli cell is indicated by the arrows. The fluorescence corre-
sponds to the position of ectoplasmic specializations adjacent to junctions between Sertoli cells. Bar = 40 pm. X890.
Fig. 7. Actin distribution predominantly in Sertoli cell ectoplasmic
specializations adjacent to spermatids. Shown in this figure are clusters of spermatids that occur in apical recesses of Sertoli cells. Notice
that the spermatid heads are clearly marked by the fluorescence emitted by ectoplasmic specializations in the adjacent Sertoli cells. Bar =
40 pm. X730.
Fig. 8. Shown here is the distribution of actin, a s indicated by the
probe NBD-phallacidin, i n ectoplasmic specializations adjacent to sequential stages of germ cells. The germ cells present in these micrographs were mechanically separated, together with adjacent Sertoli
cell regions, from epithelia isolated a s described in the text. A, acrosome; N, nucleus; RC, residual cytoplasm. Shown in panels a (fluorescence) and a ’ (phase) is a spermatid a t approximately stage 8 of
spermiogenesis. Bundles of actin filaments are obvious in the discshaped ectoplasmic specialization t h a t lies adjacent to the acrosome.
In panels h and b’ (stage 10 spermatid) and c and c’ (stage 11spermatid)
the fluorescence pattern closely follow the contours of the acrosomes.
Filament bundles are aligned parallel to the long axis of the acrosomes. In panels d and d’ (stage 15-17 spermatids) and e and e’ (stage
19 spermatids) ectoplasmic specializations outline the shapc of the
underlying spermatid heads. Also shown in panel e is a n intense
fluorescence emitted from clusters of columnar structures present along
the concave surface of the spermatids. In the region marked by the
arrows, notice t h a t the columnar units occur in two rows. We believe
t h a t signals such a s these are from the filament~richSertoli cell regions surrounding tubulobulbar processes of the germ cells. Bar = 10
pm. ~ 1 , 5 4 0 .
plasma membrane overlying the developing acrosome
and consisted of linear tracts of fluorescence that together formed a disc-shaped unit. At later stages, the
fluorescence closely followed the contours of the developing acrosome (Fig. 8b,b’,c,c’)and eventually the entire
spermatid head (Fig. 8d,d’,e,e‘). At these later stages of
spermiogenesis, linear tracts of actin were observed that
generally aligned parallel to the long axis of the acrosome andor spermatid head (Figs. 8, 9). In Sertoli cell
cytoplasm adjacent to stage 19 spermatids, linear tracts
of fluorescence occasionally were observed extending
from the tip of the spermatid head to more basal head
regions, as shown in Figure 10a.
The most intense fluorescence we observed was not
emitted by actin in myoid cells or in ectoplasmic specializations of Sertoli cells, but rather by actin in regions
adjacent to the concave surface of stage 19 spermatids
(Fig. 8e, 9a,b). Often the fluorescence pattern appeared
to consist of two rows of columnar units all aligned
perpendicular to the rim of the spermatid head. This is
particularly evident in Figure 8e and is less visible in
Figures 9a and b. This region of Sertoli cells is closely
associated with the tubulobulbar processes of late-stage
No specific fluorescence was observed in any of the
control slides (Fig. 10).
In seminiferous tubules of the rat, actin concentrations, as visualized with NBD-phallacidin, occur in three
major locations: 1)myoid cells; 2) ectoplasmic specializations of Sertoli cells; 3) regions of the Sertoli cell around
tubulobulbar processes of spermatids.
In myoid cells, actin filament bundles course predominantly in two directs. Those lying adjacent to one flat
surface of the cells course in a circular direction relative
Fig. 9. Fluorescence (a,b) and phase (a’,b’)micrographs of stage 19
spermatids, and adjacent Sertoli cell regions, labeled with NBD-phallacidin. The arrows indicate the orientations of actin filament bundles
in ectoplasmic specializations. An intense fluorescence is evident ad
jacent to the concave surface of the spermatid heads. That this signal
is composed of columnar units, similar to those shown in Figure 8e, is
indicated in regions marked by the arrowheads. Bar = 5 pm. X4,OOO.
to the tubule wall while those on the other course longitudinally. This arrangment probably enables the single
layer of myoid cells to generate contractile forces in
more than one direction. That myoid cells contain actin
and are contractile has been appreciated for some time
(Clermont, 1958; Toyama, 1977). However, the three dimensional arrangement of filaments within these cells
has not been previously been described.
Our observations of the arrangement of actin filaments in Sertoli cell ectoplasmic specializations are consistent with the actin patterns visible in published
electron micrographs of these regions. At basally situated junctions between Sertoli cells, filaments lie sandwiched between the plasma membrane and a cistern of
endoplasmic reticulum (Brokelmann, 1963; Dym and
Fawcett, 1970; Flickinger and Fawcett, 1967; Fawcett,
1975; Nicander, 1967). In the present study, we have
shown that actin filaments at these junctional sites circumscribe the bases of Sertoli cells. When intact epithe-
lial sheets are viewed from a n angle perpendicular to
the epithelial base, and numerous Sertoli cell junctional
complexes are seen simultaneously, a “honeycomb” arrangement is evident. These results are consistent with
predictions based on three-dimensional reconstructions
from electron micrographs of Sertoli cell junctional complexes (Weber et al., 1983).
In ectoplasmic specializations associated with germ
cells, actin filaments occur in linear bundles that are
generally arranged parallel to the long axis and conform
to the shape of the developing acrosome. Although Russell et al. (1980) have demonstrated in electron micrographs that ectoplasmic specializations occur adjacent
to spermatogenic cells a t stages as early as spermatocytes and round spermatids, in the present study, using
fluorescence, we observed these structures as distinct
entities only in association with approximately stage 8
and later spermatids. It is unclear why there is this
discrepancy between the two sets of data. One possible
Fig. 10. Control series for NED-phallacidin staining. Shown in panels
a (fluorescence) and a‘ (phase) is a spermatid stained with NBD-phallacidin. Actin bundles in the ectoplasmic specialization are labeled as
are filaments within the concavity of the spermatid head. Also notice
the linear signal that courses across the Sertoli cell apical process from
the tip of the spermatid head to more caudal regions of the head. In
panels b and b‘, c and c’, and d and d’ are spermatids treated with 1)
NBD-phallacidinin the presence of phalloidin, 2) phalloidin alone, and
3) buffer alone, respectively. Bar = 10 pm. X 2 , l O O .
reason is that specializations associated with spermatocytes and round spermatids may be remnants of Sertoli
cell junctional complexes and hence not recognized as
distinct entities by fluorescence. When they first appear,
ectoplasmic specializations have a disc shape and are
composed of linear bundles of filaments. As spermatogenesis continues the bundles become less distinct, but
are still visible, and the entire structure undergoes shape
changes that coincide with the shape changes in the
developing acrosome. Bundles of filaments that we detected in those cytoplasmic regions of Sertoli cells associated with stage 19 spermatids generally correspond to
the orientation of filament bundles visible in electron
micrographs published, for example, by Lalli and Clermont (1981); that is, the bundles are generally oriented
along the long axis of the spermatid head and occasionally are visible in Sertoli cell regions not directly apposed to the germ cell.
Although we have interpreted the fluorescence associated with germ cells as originating from actin in attached Sertoli cell ectoplasmic specializations, we cannot
rule out the possibility that some of the signal, particularly that associated with stage 10 and 11 spermatids,
may actually have been generated by filamentous actin
within the germ cells themselves. However, three pieces
of information are consistent with our conclusion that
most of the observed fluorescence was due to actin in
ectoplasmic specializations. First, ultrastructural data
indicate that ectoplasmic specializations often remain
attached to germ cells that are mechanically dissociated
from the epithelium (this study; Romrell and Ross, 1979).
Second, most of the fluorescence associated with germ
cells appears to originate from a region external to the
germ cell and not from the subacrosomal space-a location where filamentous actin is known to occur (Campanella et al., 1979). Third, the fluorescence pattern is
consistent with the arrangement of actin filament bun-
dles in electron micrographs of ectoplasmic specializations. The fluorescence pattern originating from actin in
germ cells that are devoid of Sertoli cell fragments has
yet to be determined.
The function of ectoplasmic specializations is not
known. The close relationship of these structures to Sertoli cell junctions and to regions of adhesion to germ
cells indicates that they may be involved with intercellular attachment and hence with spermiation (Gravis,
1978a, 1979, 1980; Romrell and Ross, 1979; Ross, 1976,
1977; Ross and Dobler, 19751, as well as with the movement of germ cells through the blood testis barrier (Dym
and Fawcett, 1970). Ectoplasmic specializations associated with germ cells have also been suggested to actively “grip” elongate spermatids (Franke et al., 1978;
Toyama, 1976)and to “rigidify” Sertoli cell apical crypts
in which spermatids mature (Russell, 1977, 1980a). Although ectoplasmic specializations are generally considered contractile (Gravis, 1978b, 1979; Toyama, 1976) we
have been unable to demonstrate myosin in these structures or to induce their contraction in ground squirrel
Sertoli cells (Vogl and Soucy, 1985).
Our observation that ectoplasmic specializations adjacent to germ cells generally conform to the contour of
the developing acrosome is consistent with, but does not
prove, the hypothesis that the structures may facilitate
the development of acrosome shape (Fawcett, 1979) or
stabilize junctional sites in some way (Vogl and Soucy,
1985). It is also interesting that filament bundles are
generally oriented parallel to the long axis of the spermatid head and not in a circular fashion as one might
expect if ectoplasmic specializations “grip” the germ
cells using contractile force.
During the late stages of germ cell differentiation in
the rat, two rows of tubular projections extend from the
concave surface of the sickle-shaped spermatid heads
into the adjacent Sertoli cell. These projections, together
with modified regions of the Sertoli cell, are termed
tubulobulbar complexes (Russell and Clermont, 1976).
These complexes consist, in part, of a filamentous network in the Sertoli cell. This filamentous network may
develop from a rearrangement of actin in ectoplasmic
specializations. This argument is supported by the observation that tubulobulbar processes appear in regions
associated with ectoplasmic specializations (Russell and
Clermont, 1976).
The function of tubulobulbar complexes is not known.
The most popular working hypotheses are that they
assist in attaching spermatids to the epithelium (Russell
and Clermont, 1976; Russell, 1979a) or facilitate the
removal of excess cytoplasm from maturing germ cells
(Russell, 1979b, 1980b, see review by Russell, 1984). It
has also been suggested that they transfer a chemical
signal to Sertoli cells that initiates sperm release
(Gravis, 1980).
Our results indicate that filament networks in regions
of the Sertoli cell adjacent to tubulobulbar processes
contain a n abundance of actin. Actin in these regions
may be contractile, or may be purely skeletal without
being contractile. Although the functional significance
of these possibilities has yet to be demonstrated, the
actin networks may play a role in anchoring the tubulobulbar processes to the Sertoli cell, facilitating the
development of spermatid head shape, or internalizing
germ cell cytoplasm.
In summary, we have been able to visualize the threedimensional arrangement of actin filament bundles in
myoid cells and in ectoplasmic specializations of Sertoli
cells in the rat. We have also presented evidence indicating that the filament networks in Sertoli cell regions
surrounding tubulobulbar processes of spermatids contain actin. Isolated sheets of seminiferous epithelia may
provide a n ideal system in which to study further the
involvement of Sertoli cell cytoskeletal elements in
events that occur during spermatogenesis.
We would like to thank Marilyn Stuart for typing the
manuscript and Bryon Grove for many helpful criticisms. This work was supported by BCHCRF grant #67
(83-1)and MRC grant #MA-8020 to A.W. Vogl.
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Suarez-Quian and Dym (1984, Annals of the New York
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NBD-phallacidin, in fixed and frozen sections of rat testis. Our results confirm and extend their original findings. These authors also reported that they could not
demonstrate myosin in Sertoli cell junctional complexes,
and suggested that microfilaments a t these sites stabilize the blood-testis barrier.
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epithelium, distributions, wall, denuded, seminiferous, activ, isolated, tubules, rat
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