3521
Journal of Cell Science 107, 3521-3534 (1994)
Printed in Great Britain © The Company of Biologists Limited 1994
Chromatin and microtubule organization during premeiotic, meiotic and early
postmeiotic stages of Drosophila melanogaster spermatogenesis*
Giovanni Cenci1, Silvia Bonaccorsi1,2, Claudio Pisano1,2, Fiammetta Verni1 and Maurizio Gatti1,2
1Istituto
Pasteur, Fondazione Cenci Bolognetti, and 2Centro di Genetica Evoluzionistica del CNR, Dipartimento di Genetica e
Biologia Molecolare, Universita’ di Roma ‘La Sapienza’, P. A. Moro, 5. 00185 Roma, Italy
*This paper is dedicated to the memory of Fritz Sobels, in recognition of his studies on Drosophila spermatogenesis
SUMMARY
Larval and pupal testes of Drosophila melanogaster were
fixed with a methanol/acetone fixation procedure that
results in good preservation of cell morphology; fixed cells
viewed by phase-contrast optics exhibit most of the structural details that can be seen in live material. Fixed testis
preparations were treated with anti-tubulin antibodies and
Hoechst 33258 to selectively stain microtubules and DNA.
The combined analysis of cell morphology, chromatin and
microtubule organization allowed a fine cytological dissection of gonial cell multiplication, spermatocyte develop-
ment, meiosis and the early stages of spermatid differentiation. We placed special emphasis on the spermatocyte
growth phase and the meiotic divisions, providing a
description of these processes that is much more detailed
than those previously reported. In addition, by means of
bromo-deoxyuridine incorporation experiments, we were
able to demonstrate that premeiotic DNA synthesis occurs
very early during spermatocyte growth.
INTRODUCTION
level, and our current conception of the mechanisms that
control the formation of male gametes is still very poor.
Recently, however, many new genes involved in Drosophila
spermatogenesis have been identified by single P element
mutagenesis (Castrillon et al., 1993; Fuller, 1993; unpublished
work of our laboratory). The molecular analysis of some of
these genes is thus expected to be completed in the next few
years, providing substantial insights into the problem of sperm
development.
A fundamental step in the mutational dissection of spermatogenesis is the precise definition of the phenotypic consequences of specific mutations at the cellular level. The
accuracy of such definition must rely on a careful description
of the cellular transformations that occur during spermatogenesis in wild-type males. D. melanogaster spermatogenesis has
been extensively studied at the ultrastructural level, and
beautiful descriptions of the premeiotic (Tates, 1971), meiotic
(Tates, 1971; Lin et al., 1981; Church and Lin, 1982, 1985)
and postmeiotic (for review see Lindsley and Tokuyasu, 1980)
stages are currently available. Light microscope analyses,
though more widely accessible, are unfortunately not as
detailed and complete. The classic cytological procedure for
light microscope analysis of Drosophila spermatogenesis is the
observation of living preparations in phase contrast (see for
example Lifschytz and Hareven, 1977; Castrillon et al., 1993;
Fuller, 1993). The morphological differentiation of living cells
is remarkably good and most cell types and stages, including
the meiotic divisions, can be readily recognized. However, the
fact that the structures observed in living cells cannot be discriminated by immuno- and histochemical staining techniques,
The differentiation of sperm from the relatively undifferentiated gonial cells is one of the most complex and elaborate of
developmental processes. This process encompasses two types
of cell divisions: the mitotic divisions needed for the multiplication of gonial cells, and the meiotic divisions leading to the
formation of haploid spermatids. In addition, both premeiotic
and postmeiotic cells undergo a series of dramatic changes in
cellular morphology that culminate in the differentiation of the
sperm tail. All these events, collectively defined as spermatogenesis, are characterized by a remarkable degree of evolutionary conservation, so that insects and mammals exhibit only
a few differences in the basic morphogenetic features involved
in sperm development. Thus, an understanding of spermatogenesis in a model system should provide a conceptual
framework of wide utility.
A model organism particularly suitable for the elucidation
of complex developmental processes is Drosophila
melanogaster. Mutational dissection of Drosophila embryogenesis, followed by the molecular characterization of the
relevant genes, has led to an increasing understanding of the
fundamental mechanisms underlying embryonic development.
Genetic analysis has also been applied to Drosophila spermatogenesis and many mutants have been isolated that disrupt
specific aspects of this process (see for example Lifschytz and
Hareven, 1977; Lifschytz and Meyer,1987; Hackstein, 1991).
Although these studies have clearly shown that the genetic dissection of spermatogenesis is feasible, very few genes involved
in sperm development have been characterized at the molecular
Key words: Drosophila, spermatogenesis, meiosis
3522 G. Cenci and others
limits the utility of this material. On the other hand, classical
fixation techniques, such as those described by Cooper (1965),
allow a clear visualization of chromosomal material but not of
other cellular structures, such as the cytoskeleton and the
spindle.
An ideal fixation technique would be one that preserves the
cellular morphology observed in vivo yet simultaneously
allowing staining with chemical and immunological reagents.
We have developed such a technique during the course of
studies on the Y chromosome loops (Pisano et al., 1993). In
this report we have exploited this technique to describe the
arrangement and the organization of microtubules, chromatin
and mitochondria in the various cell types arising during D.
melanogaster spermatogenesis.
MATERIALS AND METHODS
Stocks
All the observations were made on larval or pupal testes of an OregonR stock that has been maintained in our laboratory for about 30 years.
The flies were reared on standard Drosophila medium at 25±1°C; dissections and cytological procedures were performed at room temperature.
Fixation and staining procedures
Pupal and larval testes were dissected and fixed according to Pisano
et al. (1993). Fixed preparations were washed three times (5 minutes
each) in PBS, before incubation with the primary antibody for indirect
immunofluorescence or Giemsa staining. Three different anti-tubulin
antibodies were used for immunostaining: (1) an anti-α-tubulin monoclonal antibody (Ab) raised against chick brain microtubules
(Amersham; Blose et al., 1982) designated as A-Am; the ascites fluid
containing this antibody was diluted 1:50 in PBS. (2) An anti-β
tubulin monoclonal Ab also raised against chick brain microtubules
(Amersham; Blose et al., 1982) designated as B-Am; the ascites fluid
with this antibody was diluted 1:30 in PBS. (3) The anti-α tubulin
monoclonal Ab 3A5 raised against Drosophila embryonic tubulin
(Piperno and Fuller, 1985), which was diluted 1:2 in PBS. In every
case the slides were incubated for 45 minutes with 30 µl of diluted
primary antibody in a humid chamber at room temperature. They were
then washed three times in PBS (5 minutes each) and incubated for
45 minutes with the secondary antibody (sheep anti-mouse IgG,
F(ab′)2 fragment, conjugated with 5(6)-carboxy-fluorescein-Nhydroxysuccinimide ester (FLUOS) from Boehringer, Mannheim, cat.
no. 1214616) diluted 1:15 in PBS.
After three washes in PBS (5 minutes each) the immunostained
slides were stained with Hoechst 33258 according to Bonaccorsi et
al. (1988). Giemsa staining was performed as previously described
with 4% Giemsa (Bonaccorsi et al., 1988).
The slides were analyzed with a Zeiss III photomicroscope
equipped with a mercury light source for incident illumination.
FLUOS and Hoechst 33258 fluorescence were detected using the 0.9
(BP 450-490, FT 510, LP420) and the 0.1 (BP 365/11, FT 395, LP
397) Zeiss filter combinations, respectively. Microphotographs were
taken with Kodak T-Max 100 film.
Bromo-deoxyuridine (BrdU) labeling
Dissected testes were incubated in 10 µg/ml BrdU (Sigma) dissolved
in PBS. They were then washed for at least 3 minutes in testis buffer
(Kremer et al., 1986), squashed and fixed as described above. Testis
preparations were incubated for 1 hour with the ready-to-use
Amersham solution (code RPN 202) containing an anti-BrdU monoclonal antibody, nuclease and 1% BSA. After washing in PBS for 5
minutes the slides were incubated with the secondary antibody (sheep
anti-mouse IgG, F(ab′)2 fragment, conjugated with FLUOS from
Boehringer, Mannheim) diluted 1:15 in PBS, washed and mounted in
PBS for observation.
RESULTS AND DISCUSSION
Our methanol-acetone fixation technique (Pisano et al., 1993;
see Materials and Methods) results in good preservation of the
cell morphology observed in live material by phase-contrast
optics. As shown in Fig. 1, most of the morphological details
that can be seen in living cells are also visible in fixed preparations, suggesting that fixation artifacts are minimal. The only
intracellular structures that are substantially affected by
fixation are the Y chromosome loops. These filamentous structures are usually faint and labile in living preparations but
become clearly apparent after fixation (Bonaccorsi et al., 1988;
Pisano et al., 1993).
We have immunostained fixed preparations from both larval
and pupal testes with three different anti-tubulin antibodies (AAm, B-Am and 3A5; see Materials and Methods). It has been
shown that A-Am recognizes most of the Drosophila α-tubulin
isoforms (Matthews et al., 1989), while B-Am reacts with both
the β1- and the testis-specific β2-tubulin (Kimble et al., 1989);
the monoclonal antibody 3A5 reacts with α-tubulin (Piperno
and Fuller, 1985). These antibodies gave identical results with
all the cell types of D. melanogaster spermatogenesis. Thus,
we will not distinguish between them in describing the various
immunostaining patterns.
Following tubulin immunostaining, testis preparations were
treated with Hoechst 33258 and viewed under a fluorescence
microscope with filter combinations that detect either fluorescein or Hoechst fluorescence. This enabled us to examine
microtubule and chromatin organization in the same cell and
to correlate the behavior of these cellular components with the
other structures that can be seen under phase-contrast
microscopy. Based on these observations we have proposed a
stage subdivision of spermatocyte growth, meiotic divisions
and early spermatid differentiation. Each stage is defined only
by morphological criteria and corresponds to a facet of the
process that can be unambiguously distinguished from both the
preceding and the successive one.
Spermatogonia
Cytological analysis of the cells located at the apex of the testis
is difficult because in squashed preparations these cells are
usually very close to each other and very similar in morphology. Thus, we have been unable to discriminate between germline stem cells, primary spermatogonia and the progenitors of
the cyst cells. However, cysts containing 2, 4, or 8 secondary
spermatogonia can be easily recognized and analyzed.
Secondary spermatogonia of these cysts appear identical when
in the same phase of cell cycle, with nuclei almost entirely
occupied by chromatin (Fig. 2). Hoechst staining permits an
easy discrimination between G1 nuclei and those that have
completed DNA synthesis, which have roughly doubled the
chromatin content (Fig. 2). In most cases, the cyst cell nuclei
can be readily identified because of their elongated appearance
with the chromatin often divided into a main larger mass and
a smaller, more or less separated element. In addition, the
Drosophila spermatogenesis 3523
Fig. 1. Comparison between living and fixed
cells of Drosophila spermatogenesis. (a,b)
Live (a) and fixed (b) mature primary
spermatocytes; note the Y chromosome loops.
(c,d) Live (c) and fixed (d) primary
spermatocytes in late meiotic prophase I; note
the intranuclear granules produced by the
disintegration of the Y loops. (e,f) Live (e) and
fixed (f) prometaphases, metaphases or early
anaphases I; observation of unstained material
does not allow unambiguous identification of
the meiotic stage. (g,h) Live (g) and fixed (h)
telophases I. (i,j) Live (i) and fixed (j)
secondary spermatocytes. (k,l) Live (k) and
fixed (l) prometaphases, metaphases or early
anaphases II; also in this case examination of
unstained material does not allow precise
identification of the meiotic stage. (m,n) Live
(m) and fixed (n) telophases II. (o,p) Live (o)
and fixed (p) onion-stage spermatids; note the
clear nuclei (n) and the phase-dense
nebenkerns (nk). Bar, 10 µm.
chromatin content of these nuclei is comparable to that of postsynthetic spermatogonia, indicating that cyst cells are in an
extended G2 phase of the cell cycle (Fig. 2).
Interphase spermatogonia exhibit a dense cytoplasmic
network of microtubules that undergoes a dramatic reorganization in preparation for mitosis, culminating in spindle
formation. Spermatogonial mitoses are relatively rare, suggesting that mitotic division of these cells is very rapid. The
metaphase spindle consists of two relatively narrow and
elongated hemispindles that shorten progressively during
anaphase and telophase. At telophase the hemispindles become
broader as they flatten against the reforming nuclei; a midbody
connecting the two daughter nuclei is often visible at this time.
These structures progressively disappear as the next interphase
begins and a new cytoskeleton takes shape in the cytoplasm
surrounding each nucleus (Fig. 2).
Premeiotic DNA synthesis
The four gonial divisions generate 16 primary spermatocytes
that are morphologically indistinguishable from the parental
secondary spermatogonia in the G1 phase (we designate this
stage as S0; an early S0 stage is shown in Fig. 2). These
primary spermatocytes soon enter a program of growth accompanied by a series of characteristic changes in nuclear morphology that result in a 25-fold increase in nuclear volume
(Tates, 1971; Lindsley and Tokuyasu, 1980; Fuller, 1993).
An important question about spermatocyte growth concerns
the timing of premeiotic DNA synthesis (cf. Olivieri and
Olivieri, 1965; Lindsley and Tokuyasu, 1980). Olivieri and
Olivieri (1965) injected one-day-old males with tritiated
thymidine and examined their testes by autoradiography for
thymidine incorporation at several post injection (p.i.) time
intervals. They detected DNA synthesis in spermatogonial
cysts as early as 30 minutes p.i. In contrast, thymidine incorporation into 16-cell primary spermatocyte cysts was first
observed at 8 hours p.i. The same results were obtained by
Sato (quoted by Lindsley and Tokuyasu, 1980), using a higher
specific activity tritiated thymidine. Because only the
youngest spermatocytes were labelled at 8 hours p.i., the
labelling of these cells could result from thymidine incorporation occurring during the S phase of the 8-cell-cyst spermatogonia. Thus, these observations indicate either that premeiotic DNA synthesis occurs throughout the growth phase
of D. melanogaster spermatocytes, or that these cells do not
incorporate tritiated thymidine for some unknown reason.
To examine the occurrence of the premeiotic DNA
synthesis dissected larval testes were incubated at 25°C in
PBS containing bromo-deoxyuridine (BrdU). At different
times after the beginning of the incubation, testes were fixed
with our usual procedure, and treated with an anti-BrdU
antibody to detect BrdU incorporation. The first clearly
labelled cysts were observed after 3 hours incubation.
3524 G. Cenci and others
Labelled nuclei were observed in 2-, 4- and 8-cell spermatogonial cysts, as well as in 16-cell cysts containing very young
spermatocytes morphologically indistinguishable from
secondary spermatogonia in the G2 stage (Fig. 3). Cysts con-
taining spermatocytes even slightly larger than gonial cells
were invariably unlabelled.
The spermatocyte labeling in 16-cell cysts cannot be due to
an immunostaining artifact because the cyst cell nuclei, which
are in an extended G2 phase, do not react with the anti BrdU
antibody (Fig. 3). Moreover, the fact that three hours are the
minimum time required for labeling of both spermatogonia and
spermatocytes strongly suggests that spermatocyte labeling is
the direct consequence of BrdU incorporation into these cells,
and not the result of BrdU uptake during the last spermatogonial S phase. Thus, although we cannot explain why the
tritiated thymidine experiments failed to detect premeiotic S
phase, we conclude that premeiotic DNA synthesis occurs in
very young primary spermatocytes. The early occurrence of
this synthesis may facilitate the elevated transcriptional
activity of primary spermatocytes, which have embarked on
the production of most, if not all, mRNA needed for spermiogenesis (for review see Fuller, 1993).
Spermatocyte growth
Primary spermatocytes that have just completed DNA synthesis
(stage S1, Fig. 4a) are similar in size and morphology to the G2
secondary spermatogonia. As a primary spermatocyte begins its
growth, the nucleus progressively assumes an eccentric position
within the cytoplasm (stage S2, Fig. 4b,c). This so-called polar
spermatocyte stage (Tates, 1971) is characterized by an asymmetrical distribution of the mitochondria within the cytoplasm,
with these organelles clustered at the pole of the cell opposite
to the nucleus (Cooper, 1965; Tates, 1971). In our cytological
preparations polar spermatocytes exhibit a dense network of
cytoskeletal microtubules and a cloud of Hoechst-fluorescent,
dotted material clustered in a cytoplasmic region opposite to the
nucleus (Fig. 4b,c). These fluorescent dots most likely correspond to the Hoechst-stained mitochondrial DNA, thus identifying the cytoplasmic location of these organelles.
During the polar spermatocyte stage there is a substantial
increase in nuclear diameter accompanied by changes in
chromatin organization within the nucleus. In young polar
spermatocytes (stage S2a, Fig. 4b), the chromatin appears as a
compact mass occupying the center of the nucleus. As the polar
spermatocytes grow, chromatin subdivides into three masses
Fig. 2. Spermatogonial cysts in different phases of the cell cycle.
(a) Phase-contrast; (b) tubulin immunostaining; (c) Hoechst 33258
staining. A, an 8-cell cyst in the G1 phase. B, a 4-cell cyst at
metaphase; note the metaphase spindles in b. C, a 16-cell cyst in
early G1. Tubulin immunostaining in b clearly shows that the cells of
this cyst have just completed telophase; in one case a central spindle
is still present between the two daughter nuclei. D, an 8-cell cyst in
the G2 phase. The phases of the cell cycle can be easily deduced
taking as a standard the metaphase nuclei of cyst B and the telophase
nuclei of cyst C. It is evident that the nuclei of cyst A contain about
one half the DNA of those of cyst B, whereas the nuclei of cyst D
contain more or less the same DNA as cyst B nuclei. In cysts B and
C the cyst cell nuclei (arrows) are clearly identifiable in that they are
not involved in cell division. The DNA content of these nuclei
appears to be the same in both cysts, suggesting that the cyst cells are
in an extended G2 phase. Only one of the two cyst cell nuclei of cyst
D is recognizable (large arrow), the other could be any of the nine
additional fluorescent elements of this cyst. Cyst A contains only 9
nuclei and it is probably lacking one of the cyst cell nuclei; a large
arrow points to the identifiable cyst cell nucleus. Bar, 10 µm.
Drosophila spermatogenesis 3525
Fig. 3. Bromo-deoxyuridine labelling of 16-cell cysts of young
primary spermatocytes. (a) Hoechst 33258 staining; the arrows point
to the cyst cells. (b) Bromo-deoxyuridine incorporation detected by
antibody staining; note that the cyst cells are not labelled.
or clumps that remain closely apposed to the inner nuclear
envelope (stage S2b, Fig. 4c; see also Cooper, 1965). The
space between these clumps increases with nuclear growth, so
that in mature spermatocytes most of the karyoplasm is not
occupied by the chromatin. Two of these chromatin clumps are
larger and denser than the other and probably correspond to the
somatically-paired large metacentric autosomes (Cooper,
1965). The third chromatin mass appears to be composed by
3-5 fluorescent elements that are likely to correspond to the X
chromosome, the portions of the Y chromosome not involved
in loop formation, and the tiny fourth chromosomes (Cooper,
1965).
As nuclear growth continues, the nucleus progressively
resumes a central position within the cell and the mitochondria
disperse uniformly into the cytoplasm (stage S3). This stage is
also characterized by the first appearance within the karyoplasm of the Y chromosome loops. In D. melanogaster there
are three lampbrush-like loops, formed by the kl-5, kl-3 and ks1 fertility factors (Bonaccorsi et al., 1988). We have previously
shown that these structures develop asynchronously during
spermatocyte growth, with the kl-5 and ks-1 loops appearing
earlier than the kl-3 loop (Bonaccorsi et al., 1988). The
youngest apolar spermatocytes (stage S3, Fig. 4d) have a
centrally located nucleus surrounded by a symmetric network
of microtubules, and exhibit two phase-dark bodies that correspond to the primordia of the kl-5 and ks-1 loops (Bonaccorsi
et al., 1988).
Maturation of the apolar spermatocytes is characterized by
a further increase in nuclear size and by the elaboration of the
Y loops. The kl-5 and ks-1 loops continue to expand and soon
lose their initial compactness, revealing their underlying filamentous structure. At this stage (stage S4, Fig. 4e), a third
cluster of filamentous material, the kl-3 loop, also becomes
apparent. The three loops enlarge steadily and reach their
maximum size in mature spermatocytes (stage S5, Fig. 4f).
These cells are the largest cells produced during Drosophila
spermatogenesis. Their nuclei exhibit three fully grown Y
chromosome loops that differ both in the type of thread they
contain and in their relative degree of compactness: the kl-5
and ks-1 loops are more compact and contain a coarse thick
thread, while the kl-3 loop consists of a thinner, less twisted
and folded filament (Bonaccorsi et al., 1988). The Y loops of
mature spermatocytes occupy most of the nucleus and often
overlap, so that in many cases they cannot be distinguished
from each other. However, they can be easily identified by
Giemsa staining at pH 10 or by immunostaining with various
antibodies that specifically decorate each of these structures
(Bonaccorsi et al., 1988; Pisano et al., 1993).
The disintegration of the Y loops marks the end of the
growth phase of primary spermatocytes and the beginning of
the first meiotic division. When the loops begin to fall apart
into pieces (stage S6, Fig. 4g), the nuclei remain at more or
less the same size as in the mature spermatocytes. However, in
S6 nuclei the nucleolus has already disappeared; the chromatin
has begun to condense in preparation for meiosis but is still
close to the inner nuclear envelope. The microtubular system
of these cells has also changed from that of mature spermatocytes in that the microtubules are now more concentrated
around the nucleus than in the rest of the cytoplasm (Fig. 4g).
Traditionally, the spermatocyte growth phase is considered
as the meiotic prophase. However, as already observed by
Cooper (1965) in D. melanogaster and by Kremer and
coworkers (1986) in D. hydei, spermatocyte growth is not
accompanied by the typical progressive condensation and
synapsis of meiotic chromosomes that can be seen in the
meiotic prophase of most organisms. After premeiotic DNA
synthesis the chromosomes are dispersed within the nucleus
and become organized into three clumps as primary spermatocytes enlarge. Although there is no direct evidence that each
clump contains a couple of paired homologs, this conclusion
is generally accepted by students of Drosophila spermatogenesis (for reviews see Cooper, 1965; Fuller, 1993). In addition,
it has been suggested that homologous chromosome association in the clumps may be the consequence of the persistence
of ‘somatic pairing’ during gonial mitoses and throughout spermatocyte development (Metz, 1926; Cooper, 1965; reviewed
by Fuller, 1993).
Although each clump is likely to consist of a bivalent, no
leptotene- or zygotene-like chromosomes can be discerned
within these structures by aceto-orcein or Hoechst staining
(Cooper, 1965; present results). Thus, the clumps represent a
diffuse state of meiotic chromosomes, analogous to the state
of interphase chromatin. This decondensed state of spermatocyte chromosomes persists until the M1 stage (see below),
when chromatin condenses very rapidly, just prior to the onset
of the first meiotic metaphase. According to Cooper (1965),
when the bivalents become visible by orcein staining, they
have already attained a degree of condensation equivalent to
that of diakinetic chromosomes.
The absence of the leptotene-pachytene stages in primary
spermatocytes is probably related to the lack of synaptonemal
complex (Meyer, 1964) and meiotic crossing over in D.
melanogaster males. Interestingly, D. melanogaster females,
that exhibit a normal synaptonemal complex and undergo
meiotic recombination, also exhibit a canonical meiotic
prophase that lasts about 80 hours and comprises chromosome
condensation stages equivalent to leptotene/zygotene and
pachytene (King, 1970; Carpenter, 1975; Davring and Sunner,
1976, 1977). Thus, D. melanogaster possesses the genetic
information necessary for progressive meiotic chromosome
condensation, but this information is exploited only during
female meiosis. Meiotic chromosome condensation in males is
very rapid and is brought about in a fashion similar to that
observed in mitotic cells. This observation suggests the intrigu-
3526 G. Cenci and others
Phase-contrast
Tubulin
Hoechst
Stage
S1
S2a
S2b
S3
S4
S5
S6
Fig. 4. Primary spermatocyte
growth. (Row a) A complete cyst of
young primary spermatocytes in the
S1 stage; the arrows point to the cyst
cells. (Row b) Polar spermatocytes
in the S2a stage; the arrows indicate
the asymmetric cap of mitochondria.
(Row c) Polar spermatocytes in the
S2b stage; the arrows point to the
clusters of mitochondria. (Row d) A
young apolar spermatocyte in the S3
stage; note the primordia of the kl-5
(A) and ks-1 (C) Y chromosome
loops, and the three chromatin
clumps. (Row e) A primary
spermatocyte in the S4 stage in
which some filaments of the kl-3
loop (B) have become apparent.
(Row f) A mature primary
spermatocyte in the S5 stage
showing fully developed Y
chromosome loops; note that the
nucleolus (arrow) is surrounded by
the sex chromosome chromatin.
(Row g) A primary spermatocyte in
the S6 stage with disintegrating Y
loops. Bar, 10 µm.
Drosophila spermatogenesis 3527
ing possibility that meiotic chromosome contraction in males
exploits the same genetic program that governs chromosome
condensation during mitosis.
Meiotic divisions
As the Y loops disintegrate the nucleus shrinks and its diameter
eventually becomes about 20% shorter than that of mature
spermatocytes (stage M1, Fig. 5a,b). A distinguishing feature
of the M1 stage is the presence within the nucleus of a few
compact granules derived from Y-loop disintegration. The
microtubular system of these M1 cells has already completed
its reorganization: the network of microtubules around the
nucleus has disappeared and two prominent asters have
become apparent. During the M1a stage (Fig. 5a), the asters,
initially located on the same side of the nucleus, migrate to
opposite poles, while remaining closely apposed to the outer
nuclear envelope. By the M1a stage, the chromatin has already
attained a high degree of condensation and the three major
bivalents are clearly visible. The fourth chromosome bivalent
is not always clearly identifiable. However, we often observed
a dotted element associated with the smallest bivalent, suggesting that the fourth chromosomes lie close to the sex chromosomes, retaining their apparent vicinity during interphase.
In the M1a stage the bivalents tend to be peripherally located
near the inner of the nuclear envelope, while in the subsequent
M1b (Fig. 5b) stage they appear to move freely in the nucleoplasm.
The M1a nuclei are round or ovoid and the cytoplasm
around them has a rather homogeneous appearance under
phase-contrast microscopy. In the later M1b stage (Fig. 5b),
the nuclei become irregularly shaped and are surrounded by
phase-contrast dense, elongated elements. Most likely these
structures correspond to the system of parafusorial and astral
membranes that surround the spindle and the poles during male
meiosis (Tates, 1971; Church and Lin, 1982). During the M1b
stage the asters are more prominent than in the M1a stage, and
a few spindle fibers radiating from the asters appear to reach
the bivalents, which occupy variable positions within the
nucleoplasm. The demarcation between the nucleus and the
cytoplasm progressively disappears as meiosis proceeds, while
additional spindle fibers become connected to the bivalents,
which can be seen at different distances from the spindle poles
(stage M2; Fig. 5c). The M1b and M2 stages almost certainly
correspond to the prometaphase of the first meiotic division, a
phase in which the bivalents, connected to the spindle poles by
kinetochore microtubules, undergo complex motions necessary
for proper metaphase orientation (Church and Lin, 1982,
1985).
Prometaphase of the first meiotic division is completed
when the bivalents congress to a metaphase plate (stage M3;
Fig. 5d). During metaphase, the bivalents cluster in a single
Hoechst-bright structure equidistant from the poles and are
connected to the spindle poles by two symmetrical sets of
microtubules. In addition to these pole-to-chromosome microtubules the metaphase spindle exhibits bundles of microtubules
that do not encounter the chromosomes. These microtubules
appear to originate from the poles, run outside the congregated
bivalents and end either in the opposite pole or distally to the
metaphase plate (Fig. 5d).
Nearly all the cysts with cells in the M3 stage also contain
early anaphase I cells (stage M4a; Fig. 5e). Interestingly, in
most of these early anaphases the segregating bivalents have
already migrated a considerable distance towards the poles,
suggesting that initial anaphase chromosome movement is very
rapid. The spindle organization of early anaphase I cells is very
similar to that of metaphase I cells in the M3 stage, consisting
of two closely apposed umbrella-shaped half spindles.
However, as anaphase proceeds the spindle microtubules reorganize: the density of microtubules in the region between the
segregating sets of chromosomes (central spindle) progressively increases, while the number of microtubules located in
the proximity of the chromosomes decreases. By mid-anaphase
(stage M4b, Fig. 5f) the microtubules emanating from each
pole overlap in the central region of the spindle, so that the two
opposed umbrella-shaped half spindles of early anaphase are
now fused in a unique oval structure. By late anaphase/early
telophase (stage M4c, Fig. 5g,h) there is a further reduction in
the number of microtubules emanating from the centrosomes,
accompanied by an apparent increase of the density of central
spindle microtubules. Concomitant with this spindle reorganization, there is a further increase in the distance between the
two daughter nuclei, suggesting that the M4a and M4b-c stages
correspond to anaphase A and anaphase B, respectively.
The dense bundle of microtubules in the central spindle
formed during stage M4b-c is progressively squeezed and
eventually attains an hourglass shape (telophase I, stage M5,
Fig. 5i). In many cases, and more frequently in broken cysts,
these hourglass-shaped spindles are bent at the mid-body.
During telophase I (stage M5, Fig. 5i), the two daughter nuclei
become clearly demarcated from the cytoplasm and often
exhibit a dark dot, which stands out from the phase-contrast
clear nucleoplasm. In favourable preparations these ‘round
bodies’ (Grond, 1984) can also be seen in spermatid nuclei, but
they disappear soon after the initiation of spermatid elongation
(Lindsley and Tokuyasu, 1980). The nature and the biological
role of these intranuclear structures, also called the protein
bodies (Kremer et al., 1986) or the nucleolus-like bodies
(Lindsley and Tokuyasu, 1980), are still largely unknown
(Tates, 1971; Lindsley and Tokuyasu, 1980; Grond, 1984;
Kremer et al., 1986).
As meiosis proceeds, telophase I nuclei enlarge and the
second division asters become apparent (stages M6a and M6b;
Fig. 6). Actually centrosome separation occurs earlier and
sometimes two distinct centrosomes are already visible in late
anaphase I (stage M4c; see Fig. 5h). These centrosomes, which
contain a single centriole (Tates, 1971), progressively nucleate
the aster microtubules and migrate to the cell poles. In
telophases with hourglass-shaped spindles, two close foci of
microtubule nucleation can often be seen. As the central spindle
progressively disappears, these initial asters become more
prominent and move to opposite poles, while remaining closely
apposed to the outer nuclear envelope (stage M6a, not shown).
When the asters have completed their migration to the poles,
secondary spermatocyte nuclei have considerably enlarged and
their chromatin appears to be relatively decondensed,
occupying the center of the nucleus (stage M6b, Fig. 6a). The
M6b cells are relatively rare, indicating that this stage, which
probably corresponds to interphase II, has a short duration.
As the chromosomes condense, in preparation for meiosis
II, three Hoechst-bright elements become apparent (stage M7,
Fig. 6b). These elements (the two major autosomes and the X
or the Y chromosome) in many cases lie very close to each
3528 G. Cenci and others
Phase-contrast
Tubulin
Hoechst
Stage
M1a
M1b
M2
M3
Fig. 5(a-d). The first meiotic division in D. melanogaster males. (Row a) Late meiotic prophase nuclei in the M1a stage; note the prominent
asters and the intranuclear granules resulting from disintegration of the Y loops. (Row b) Prometaphase nuclei in the M1b stage; note the
parafusorial and astral membranes surrounding these nuclei. (Row c) A prometaphase nucleus in the M2 stage in which the nuclearcytoplasmic demarcation is no longer visible. (Row d) A metaphase I cell with congregated bivalents (M3 stage).
other due to the small size of secondary spermatocyte nuclei,
so that only one or two Hoechst-bright spots can be observed.
Soon after the chromosomes have condensed, the cytoplasmic
nuclear demarcation disappears and additional spindle fibers
penetrate the nucleus, while the chromosomes are found at
variable distances from the spindle poles (stage M8; Fig. 6b).
Drosophila spermatogenesis 3529
Phase-contrast
Tubulin
Hoechst
Stage
M3
M4a
M4b
M4c
M4c
M5
Fig. 5(e-i). (Row e) A metaphase I cell in the M3 stage (top) plus an early anaphase (bottom) in the M4a stage. (Row f) A mid-anaphase in the
M4b stage. (Row g) Two late anaphases in the M4c stage; note the prominent central spindle. (Row h) A late anaphase in the M4c stage
showing two distinct centrosomes at each pole. (Row i) A telophase in the M5 stage showing a bent central spindle and the forming secondary
spermatocyte asters. In all the panels, Hoechst staining detects both nuclear and mitochondrial DNA; note the characteristic arrangement of
mitochondria along the central spindle. Bar, 10 µm.
These features suggest that during the M7 and M8 stages chromosomes undergo the prometaphase motions necessary for
proper orientation and congression.
The subsequent stages of the second meiotic division are
analogous to those of the first meiotic division. Thus, chromo-
somes eventually congregate into a metaphase plate (stage M9,
Fig. 6c), while the density of both kinetochore and polar microtubules increases. During early anaphase II (stage M10a, Fig.
6d), the chromosomes move rapidly towards the poles and the
spindle retains a metaphase-like configuration. Later in
3530 G. Cenci and others
Phase-contrast
Tubulin
Hoechst
Stage
M6b
M7
M8
M9
M10a
M10b
M10c
M11
M11
Fig. 6. The second
meiotic division in D.
melanogaster males.
(Row a) A secondary
spermatocyte nucleus in
the M6b stage. (Row b)
Two prometaphase
secondary spermatocyte
nuclei in the M7 stage
(top) plus a
prometaphase nucleus in
the M8 stage (bottom).
(Row c) A metaphase II
in the M9 stage. (Row d)
An early anaphase in the
M10a stage. (Row e) A
mid-anaphase in the
M10b stage. (Row f) A
late anaphase in the
M10c stage. (Row g)
Two telophases in the
M11 stage. (Row h) A
telophase in the M11
stage with a bent spindle.
Note that in c-h Hoechst
staining reveals the
arrangement of
mitochondria. Bar,
10 µm.
Drosophila spermatogenesis 3531
anaphase, the daughter nuclei continue to move apart, while
the density of fibers in the central spindle progressively
increases (stages M10b, Fig. 6e; M10c, Fig. 6f). As for meiosis
I, we propose that stages M10a and M10b-c correspond to
anaphase A and B, respectively.
During telophase II (stage M11, Fig. 6g,h), the central
spindle is progressively squeezed, attaining an hourglass
shape, while the two daughter nuclei become demarcated from
the cytoplasm. Many telophase II spindles are bent at an angle
just as telophase I spindles. However, while most anaphase
spindles of broken cysts are bent, spindle bending does not
occur, or is much less pronounced, in complete cysts.
Our observations have shown that meiotic nuclei undergo
cyclic changes in the nuclear-cytoplasmic demarcation. Late
prophase/early prometaphase spermatocytes in the M1 stage
exhibit a clear demarcation between the nucleus and the
cytoplasm. This demarcation disappears as the cells enter the
M2 stage; the absence of a nuclear-cytoplasmic boundary
persists through metaphase and anaphase I (stages M3+M4)
until telophase I (stage M5), when a sharply outlined nucleus
becomes visible again. Nuclear demarcation is maintained
during interphase, prophase and early prometaphase II (stages
M6 and M7), and disappears again when secondary spermatocytes enter late prophase II (stage M8), to reappear at telophase
II (stage M11). These cyclic changes in nuclear appearance
cannot be due to the breakdown of the nuclear membrane,
which remains largely intact during male meiosis (Tates,
1971). Instead, they most likely reflect transformations of other
components of the nuclear envelope such as the nuclear
lamina. This suggestion is supported by recent data of WhiteCooper and coworkers (1993), who analyzed lamin immunostaining during Drosophila spermatogenesis, and showed that
premeiotic primary spermatocytes have a prominent nuclear
lamina that breaks down in cells undergoing meiosis.
Figs 5 and 6 clearly show that tubulin immunostaining
detects a prominent fibrous structure that connects the two
daughter nuclei during both the first and the second meiotic
ana-telophase. This structure is recognized by three different
anti-tubulin antibodies, which also react with the metaphase
spindle and the cytoskeleton. Thus, this bundle of fibrous
material is undoubtely a bundle of spindle microtubules.
However, in a recent cytological analysis of the first meiotic
division of Drosophila males, Casal and coworkers (1990)
failed to detect a central spindle by tubulin immunostaining. In
these studies dissected testes were treated with taxol and fixed
with formaldehyde, after an ethanol prefixation; the preparations were then stained by indirect immunofluorescence with
either an anti-α tubulin antibody (Sera Lab, Accurate
Chemicals, Westbury/NY, YL1/2; Kilmartin et al., 1982) or the
same anti-β tubulin antibody employed here (Amersham;
Blose et al., 1982; see Materials and Methods). Using this
method they were able to detect metaphase and early anaphase
spindles but not ana-telophase central spindles. The reason for
these results is currently unclear. Their fixation procedure
might have selectively disrupted the central spindle of anatelophases. Alternatively, and more likely, the taxol treatment
might have interfered with spindle reorganization during
anaphase, preventing the formation of the central spindle.
The central spindles of the first and the second meiotic
divisions exhibit a remarkably similar behavior during anatelophase. At mid-anaphase I (stage M4b) or II (stage M10b)
the central spindle shows an extended zone of microtubule
overlap at its equator. However, while the anaphase spindle
elongates, this apparent overlap zone does not shorten as seen
in organisms such as diatoms, where spindle elongation occurs
by microtubule sliding in the overlap zone (for review see
Cande, 1989). This observation suggests that anaphase B
spindle elongation in Drosophila male meiosis is brought
about by mechanisms that do not involve microtubule sliding
in the central spindle. Alternatively, if such a sliding does
occur, it must be accompanied by microtubule growth, so that
the extension of the overlap zone would not decrease as the
spindle poles move apart.
An interesting feature of male meiosis is the precise partition
of mitochondria between the daughter cells at each meiotic
division. Electron microscopy studies have shown that at
prometaphase mitochondria aggregate around the nucleus,
lining up parallel to the spindle axis. This arrangement persists
through telophase until the occurrence of cytokinesis, thus
mediating precise partition of these organelles (Tates, 1971;
reviewed by Fuller, 1993). Due to their DNA content, mitochondria can be also identified by light microscopy in
favourable Hoechst-stained preparations (Figs 5 and 6). In late
meiotic prophase and early prometaphase I (stage MI) mitochondria appear to be uniformly distributed within the
cytoplasm. However, as cells enter late prometaphase I (stage
M2) mitochondria aggregate around the equator of the nucleus.
This arrangement persists through metaphase, early and mid
anaphase up until the formation of the central spindle. Then
mitochondria line up along this structure and remain associated with it until the occurrence of cytokinesis, which divides
them into two equal groups (Fig. 5). The same mitochondrial
behavior is observed during meiosis II (Fig. 6), so that the four
meiotic products receive the same amount of mitochondria.
The timing of meiotic division
Previous cinematographic studies on live cells have shown that
about 72 minutes elapse from the time that primary spermatocytes enter the M1 stage to the time when anaphase begins
(Church and Lin, 1985). M1 cells, selected on the basis of their
characteristic morphology, were filmed for 1.5 hours, allowing
precise determination of the timing of the initial meiotic events
(Church and Lin, 1985). Within 30 minutes from their identification M1 cells undergo several morphological changes that
culminate in the appearance of chromosomes; this indicates
that a maximum of 30 minutes elapses between the formation
of M1 cells and the appearance of the bivalents.
From the time that chromosomes become visible to the
beginning of anaphase I there are about 42 minutes. During the
first 20 minutes of this interval the bivalents undergo a series
of complex saltatory prometaphase motions; in the subsequent
22 minutes they exhibit a persistent bipolar orientation that is
maintained until the onset of anaphase (Church and Lin, 1985).
We have determined the frequencies of the various meiotic
stages in fixed testis preparations. The frequency of each stage
should be proportional to its duration in live material. Thus, if
one ascertains the frequency and the duration in vivo of a given
meiotic stage, the duration of all the other stages can be easily
calculated from their relative frequencies. Using these criteria
we have estimated the lengths of the meiotic phases (Table 1);
calculations were made by grouping together stages that are
easily distinguishable from both the preceding and the
3532 G. Cenci and others
Table 1. The timing of meiotic cell division in D. melanogaster males
Phases and stages of meiosis
Number of cells
Relative frequencies
Estimated duration
(minutes)
L. Prophase I
E. Prometa. I
(M1a,b)
Prometa. I
Metaphase I
( M2; M3)
Anaphase I
Telophase I
(M4a,b,c; M5)
Interphase II
E. Prometa. II
(M6a,b; M7)
Prometa. II
Metaphase II
(M8; M9)
Anaphase II
Telophase II
(M10a,b,c; M11)
Total
202
16.6
44
128
10.5
28
280
23.1
61
214
17.6
47
117
9.6
26
274
22.6
60
1,215
100
266
The relative frequencies of the various stages were obtained by scoring 40 late larval/early pupal testes.
E, early; L, late; prometa, prometaphase.
Phase-contrast
Tubulin
Hoechst
Fig. 7. Spermatid differentiation in D. melanogaster. (Row a) Partial cyst with spermatids at various differentiation stages. 1, stage T1; 2, stage
T2; 3, stage T3 with nuclei smaller than nebenkerns. (Row b) Onion stage spermatids (stage T4); note the characteristically dense cytoskeleton.
Bar, 10 µm.
following ones, and by assuming that stages M1-M3 last
approximately 72 minutes (Church and Lin, 1985). For
example, we have grouped stages M1a and M1b, which include
late prophase and early prometaphase I, and are characterized
by the presence of a clear nuclear-cytoplasmic demarcation;
this demarcation is absent in the next group of stages (M2 and
M3), that correspond to late prometaphase and metaphase I.
Similarly, stages M6-M7, that comprise interphase II and early
prometaphase II, exhibit a clear nuclear-cytoplasmic demarca-
tion that disappears in late prometaphase and metaphase II
(stages M8 and M9). Since the relative frequency of stages M1M3 is 27.1%, and their duration in vivo is 72 minutes, the total
duration of meiosis is expected to be 266 minutes (72×100/27).
Thus, the lengths of stages M1a+b, M2+M3, M4+M5,
M6+M7, M8+M9 and M10+M11 should be 44, 28, 61, 47, 26
and 60 minutes, respectively (Table 1). Although these times
are only rough estimations, they clearly indicate that the first
and the second meiotic divisions have approximately the same
Drosophila spermatogenesis 3533
Fig. 8. Chromatin decondensation and
recondensation during early stages of
spermatid differentiation. Spermatid
preparations are stained with Giemsa at
pH 10; note the intense staining of the
chromatin and the pale staining of the
nebenkerns. (a) Onion-stage spermatids
(stage T4) showing a very compact
chromatin mass. (b) Spermatids with oval
nebenkerns (stage T5) and relatively
diffuse chromatin. (c) Spermatids with elongated mitochondrial derivatives showing a dense chromatin mass flattened against the side of the
nucleus underlying the growing axoneme. Bar, 10 µm.
durations (M1-M5=44+28+61=133minutes; M6-M11=47+
26+60=133 minutes). The two meiotic divisions also appear to
have very similar durations in certain plant species (reviewed
by Van’t Hof, 1974).
The timing of D. melanogaster male meiosis has been also
estimated by time-lapse cinematography of living pupal testes
(Cross and Shellenbarger, 1979). This analysis showed that
between the stage of ‘elongated nuclei’ during the first meiotic
division and the stage defined as ‘toward the end of the second
division’ there is a time interval of about 130 minutes. The
‘elongated nuclei’ probably correspond to cells in midanaphase I (stage M4b-c), while the ‘end of the second
division’ is likely to correspond to telophase II (stage M11).
We have calculated that the time elapsing between the
beginning of anaphase I (stage M4a) and the end of telophase
II (stage M11) is 194 minutes. Since the interval considered by
Cross and Shellenbarger is obviously more restricted than ours,
the two estimations appear to be comparable.
Spermatid differentiation
In telophase II cells, the mitochondria associated with the
central spindle move towards the nuclei, forming an irregular
mass on one side of the nucleus (stage T1; Fig. 7a). This mass
quickly assumes a crescent-like shape (stage T2; Fig. 7a),
which then results in a spherical shape (stage T3; Fig. 7a).
Spermatids in the T3 stage consist of a spherical mitochondrial aggregate associated with a smaller, spherical nucleus. As
spermatid differentiation proceeds, the nucleus enlarges and
mitochondria progressively fuse to form a complex organelle,
called the nebenkern, containing multiple layers of interleaved
mitochondrial membranes (Tates, 1971; Tokuyasu, 1975).
Because cross sections of this mitochondrial derivative
resemble an onion, this stage of spermatid development has
been named the onion stage (Bowen, 1922; Tates, 1971;
Tokuyasu, 1975). Light microscopy and Hoechst staining do
not allow discrimination between T3 mitochondrial aggregates
and onion-stage nebenkerns. They both appear as phase-dense,
round structures that exhibit a dotted Hoechst fluorescence, due
to the presence of mitochondrial DNA. However, onion stage
spermatids (stage T4; Fig. 7b) can be easily distinguished from
those in the T3 stage because they have a nucleus of the same
size or slightly larger than the nebenkern, and a dense
cytoskeleton with characteristic bundles of microtubules.
The onion-stage spermatid nuclei contain a very compact
chromatin mass that can be easily visualized after both Hoechst
and Giemsa staining (Figs 7b and 8a). When the elongation
process begins, the nucleus remains spherical, while the
nebenkern assumes an oval shape (stage T5, Fig. 8b). At stage
T5 the nuclear chromatin loses its previous compactness and
becomes more diffuse within the nucleoplasm. However, as
elongation proceeds, chromatin condenses again, forming a
dense mass flattened against the fenestrated side of the nucleus
underlying the elongating axoneme (Fig. 8c). This extra cycle
of chromatin decondensation and recondensation has been
already described by light microscopy in D. hydei (Kremer et
al., 1986). Although the precise biological meaning of this phenomenon is currently unclear, it has been correlated with the
molecular reorganization of the chromatin that occurs in
spermatid nuclei (Das et al., 1964; Hauschtek-Jungen and
Hartl, 1982).
We are grateful to Minx Fuller for her generous gift of the 3A5
antibody, and to Mike Goldberg and Byron Williams for critical
readings of the manuscript. This work has been in part supported by
a grant from Progetto Finalizzato Ingegneria Genetica.
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(Received 4 July 1994 - Accepted 26 August 1994)