Department of Biology, Washington University in St Louis, Campus Box 1229, 1 Brookings Drive, St Louis, MO 63130, USA
* Author for correspondence (e-mail: noguchi{at}biology2.wustl.edu)
Accepted 23 January 2003
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SUMMARY |
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Key words: Individualization, Spermatogenesis, Drosophila, Actin cone, Microtubule, Actin dynamics, Membrane dynamics
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INTRODUCTION |
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To provide information on the mechanism, we wanted to be able to manipulate
the process and to visualize the events in the cystic bulge at high
resolution. For this reason, we developed an in vitro culture system.
Previously, it has been demonstrated that isolated spermatogenic cysts of
Drosophila melanogaster and Drosophila hydei undergo
differentiation in vitro to some extent
(Cross and Shellenbarger,
1979; Fowler,
1973
; Liebrich,
1981
; Liebrich,
1982
). Cross and Shellenbarger
(Cross and Shellenbarger,
1979
) described meiosis and reported that all subsequent steps of
spermatogenesis take place in isolated and cultured cysts of D.
melanogaster. However, these studies did not investigate the
individualization step in detail. This previous work encouraged us to develop
the culture system further to permit observation and manipulation of
individualizing cysts. We have adapted the culture system so that we can
employ advanced live imaging techniques.
The main questions we have asked are: what is the driving force for cystic bulge movement? Are the actin cones responsible for movement? How is the syncytial membrane reorganized into a thin tubular structure? For these studies, we have used immunofluorescence localization of microtubules and F-actin during formation and movement of the actin cones, pharmacological manipulation of cytoskeletal elements and fluorescence recovery after photobleaching (FRAP) techniques to demonstrate that actin dynamics play a key role in the process. In addition, membrane dynamics in the cystic bulge were visualized using FM1-43. Based on these observations, we propose a model for the role of actin in individualization. This is the first detailed description of individualization in vitro and dissection of the cell biological processes involved.
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MATERIAL AND METHODS |
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Cell culture and live recordings
In vitro culture of isolated cysts was performed according to Cross and
Shellenbarger (Cross and Shellenbarger,
1979) with the following modifications. Testes were dissected from
newly eclosed adult males and transferred into culture media (modified M3
medium without bicarbonate; Sigma-Aldrich, St Louis, MO), containing 10% fetal
calf serum (Sigma-Aldrich) and standard penicilin/streptomycin cocktail
(Cross and Sang, 1978
;
Cross and Shellenbarger,
1979
). This formulation supported the best viability and
differentiation of isolated cysts, although for observation of
individualization, there were no significant differences among the several
media we tested (which were based on those used for culturing
Drosophila cells). The basal end of the testis proximal to the
seminal vesicle was cut off using a sharp glass needle, while the apical end
of the testis was held using a forceps. Then, the fully elongated cysts were
gently squeezed out from the muscle layer of the testis using the side of the
glass needle. Isolated cysts were transferred to culture media and cultured at
room temperature. This procedure does not damage the cysts and viability tests
revealed that 80-90% of the cysts were alive after 24 hours of culture in
vitro (data not shown).
In pharmacological experiments, cysts were cultured in 24-well plates (Falcon, Franklin Lakes, NJ) with 1 ml of culture media. The actin depolymerizing drug latrunculin A (LTA, Molecular Probes, Eugene, OR); the actin stabilizing drug jasplakinolide (Molecular Probes); the microtubule depolymerizing drugs colchicine (SIGMA-Aldrich) and nocodazole (SIGMA-Aldrich); the inhibitor of kinesin motor ATPase monastrol (SIGMA-Aldrich); and the myosin ATPase inhibitor 2-3-butanedion monoxime (BDM, SIGMA-Aldrich) were separately dissolved in dimethylsulfoxide (DMSO) for stock solutions. An equal volume of culture media containing twice as much as the final concentration of each drug was added directly to culture media during a recording of the movement of the cystic bulge. Serial differential interference contrasted (DIC) microscope images of the cystic bulge were collected using an inverted microscope equipped with a 10x plan fluor lens (Nikon, Japan) and CCD camera (SPOT, Diagnostic instruments inc. Sterling Heights, MI) every 10 or 12 minutes and processed and plotted using NIH image 1.62 and Microsoft Excel.
Immunofluorescence microscopy and image acquisition
Anti--tubulin antibody (DM1A, SIGMA-Aldrich) was used for tubulin
staining. Isolated cysts were fixed with 4% paraformaldehyde (EM grade,
Electron Microscope Science, PA) in phosphate-buffered saline, pH 7.2 (PBS)
for 7 minutes at room temperature. Then cell membranes were extracted by 0.1%
Triton X-100 in PBS for 15 minutes. After three washes with PBS, cysts were
incubated in blocking solution (2% Bovine serum albumin in PBS) for 1 hour at
room temperature. Cysts were kept in 1/300 dilution of primary antibody in
blocking solution for 2 hour and washed three times, followed by incubation
with secondary antibody (Alexa-488 conjugated goat anti-mouse IgG; Molecular
Probes), Alexa 568-phalloidin (Molecular Probes) and DAPI (SIGMA-Aldrich) for
2 hours. After washing with PBS, cysts were mounted on glass slides with
Mowiol (Calbiochem, San Diego, CA). Specimens were examined with an inverted
epifluorecscence microscope (ECLIPSE TE200, Nikon, Japan) with a cooled CCD
camera (Quantix, Photometrics, Tucson, AZ). The images were deconvolved
digitally using Slide book software (Intelligent Imaging Innovation, Denver,
CO).
FRAP analysis
GFP-actin was expressed in the testis by crossing the three driver
Drosophila stock pCOG-GAL4; NGT40; nanos-GAL4 with a pUASpGFP-actin
Drosophila stock. Similar to the female germline, UAS transgenes are
poorly expressed in the male germline. We tried numerous combinations of
drivers and targets that were not expressed at high enough levels, especially
late in spermatogenesis, for our purpose. Sufficient GFP-actin expression for
these experiments was achieved using the three drivers pCOG-GAL4; NGT40;
nanos-GAL-4, and a pUASp-GFP-actin transgenic line. The pUASp-vector was
originally engineered for enhanced gene expression in female germline
(Rørth, 1998). In this
combination, a large amount of GFP-actin expression was observed in germline
cells at early stages at the apical end of testis. Lower levels were observed
at later stages, including during individualization. We compared three
combinations of driver and targets: (1) pCOG-GAL4; NGT40;
nanos-GAL-4>pUASp-GFP-actin; (2) NGT40; nanos-GAL-4>pUASp-GFP-actin; (3)
nanos-GAL-4>pUASp-GFP-actin, and found that the fluorescence of GFP-actin
is dependent on the number of drivers. The pUASp-GFP-actin transgene provided
more expression when compared with a pUASt-GFP-actin line with the same
drivers.
The cysts from male progeny were collected as described above, and transferred to glass bottom dishes (World Precision Instruments inc., Sarasota, FL) coated with poly-L-lysine (Sigma diagnostics, St Louis, MO). Photobleach and subsequent time-lapse imaging were performed using a confocal microscope (TCS SP2, Leica, Iena, Germany) equipped with a 488 nm laser line and 40x and 60x HCX-plan-apo oil immersion lenses. Recovery of fluorescent intensity in the region of interest was measured using Photoshop and plotted using Microsoft Excel. In each experiment FRAP of 5-10 samples was performed.
Membrane labeling in vitro
Membrane dynamics in cystic bulges were examined by labeling cell membrane
with 0.5 µM FM 1-43 (Molecular Probes)
(Kuromi and Kidokoro, 1998)
for 10 minutes. After two washes with culture media, cysts were transferred to
a glass bottom dish and examined using confocal microscopy. In the case of
double imaging of membrane and GFP-actin, both fluorochromes were excited by
488 nm laser line and detected separately by two barrier filters (500-580 nm
for GFP-actin and 600-700 nm for FM1-43). Because FM1-43 has a wide range of
emission, barrier filter 500-580 nm passes some of the FM1-43 signal;
therefore, membrane is visible in the GFP-actin image. Membrane staining is
represented as yellow and GFP-actin is green in the merged image (see
Results).
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RESULTS |
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In elongated cysts, prior to the onset of individualization, sperm nuclei
were elongated but the DNA was not initially condensed. At this stage,
cytoplasmic microtubules were observed running along the length of the
spermatids and surrounding the nuclei (Fig.
3A). Actin was not yet accumulated around the apical side of sperm
nuclei. After nuclear condensation initiated, F-actin began to accumulate at
the basal apical side of nuclei slightly overlapping with them
(Fig. 3B). At this stage, a
number of cytoplasmic microtubules were still apparent. As the actin continued
to accumulate, it became thicker and more triangular in shape, appearing more
polarized (Fig. 3C). At this
time point, sperm nuclei had completed condensation and microtubules were
absent from the region surrounding the nuclei and in the actin accumulations.
At the onset of actin cone movement, suddenly microtubule staining disappeared
completely throughout the whole cyst (Fig.
3D,E). There is an obvious correlation between the disappearance
of cytoplasmic microtubules and onset of actin cone movement (88 out of 89
cysts with actin cones that had not yet moved showed microtubule staining; 61
out of 61 cysts with moving actin cones showed no microtubule staining).
Nocodazole treatment disrupted fibrous cytoplasmic microtubule structures, but
strong diffuse staining of tubulin remained in the cytoplasm (not shown).
However, the fluorescence intensity of diffuse cytoplasmic tubulin staining
dropped significantly after the onset of actin cone movement, suggesting that
the epitope itself has disappeared. EM observations also failed to detect
cytoplasmic microtubules during individualization
(Tokuyasu, 1974;
Tokuyasu et al., 1972
). These
results suggest that loss of tubulin staining is not a result of
depolymerization, but of destruction of tubulin or modification of C-terminal
end of depolymerized
-tubulin. In this stage axoneme tubulin cannot to
be seen because the antibody epitope is blocked by modification of the
C-terminal end of
-tubulin
(Bré et al., 1996
).
After the onset of movement, actin cones became more triangular. There were no
other major actin or microtubule structures visible.
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Although the above result suggests that microtubules are not required for
movement, it remains possible that a motor, like kinesin, might attach to the
actin cone and slide along the axoneme in the plus-end direction
(Gibbons, 1981). Axonemal
microtubules are not likely to be affected by these drug incubations because
of their stability. This idea is supported by the fact that there was no
shortening of the cyst when treated with microtubule depolymerization drugs
(not shown). The other type of microtubule motor, dynein, is unlikely to be
involved because it is minus-end directed motor. Therefore, we applied
monastrol, a kinesin inhibitor (Fig.
4C) that affects mitotic kinesins. Monastrol did not inhibit
movement even at a concentration of 100 µM, a concentration that completely
inhibits spindle formation in dividing cells
(Mayer et al., 1999
). Taken
together, our results provide no support for microtubule-based motility
processes being important for cystic bulge movement.
Although we have already shown that actin dynamics are important for
movement, we wondered whether myosin-based movement might also be important.
To test for actin-based motor activity, we used a general myosin ATPase
inhibitor, BDM (Cramer and Mitchison,
1995). This drug also did not inhibit movement even at an
extremely high concentration (Fig.
4C). These results suggest that some classes of myosins (at least
myosin I, II and V) are not directly involved in force generation.
Actin dynamics in actin cone
The above results suggest that actin polymerization is the driving force of
cystic bulge movement, as is the case for leading edge protrusion in
locomoting cells (Small et al.,
2002; Welch and Mullins,
2002
). We investigated the similarity of actin dynamics in actin
cones to actin structures at the leading edge of moving cells. We asked three
main questions:
We investigated actin dynamics in the cones by FRAP using GFP-actin. The maximum expression level of GFP-actin we were able to induce in late cysts, was relatively low (see Materials and Methods). However, the amount of GFP-actin was sufficient to permit us to perform FRAP experiments. The expression of GFP-actin did not affect cystic bulge movement (2.8±0.2 µm/minute, n=6).
Two different types of FRAP experiments were performed (Figs 6, 7; see Movies 3, 4 at http://dev.biologists.org/supplemental/). First, fluorescence of GFP-actin in moving actin cones was bleached along a line across actin cones, and fluorescent recovery was monitored. A bleached region at any point along the cone completely recovered, indicating that actin turns over everywhere in the actin cone. The bleached line moved forward with the actin cones, suggesting net movement of actin filaments as the actin cone moved (n=7; Fig. 6A,B; Fig. 7A,B). We did not detect a flow of actin from front to back of the actin cone (i.e. treadmilling). This was somewhat surprising and different from leading edge protrusion. Next, fluorescence of a whole single actin cone was bleached (Fig. 6D). We plotted the recovery of fluorescence intensities at the front and the back of the actin cone separately (Fig. 7C,D). Although recovery occurred everywhere, the recovery was quicker in the front. Time for recovery to one half relative intensity at the front was 4 minutes, while at the back, it was 6 minutes (n=6). This suggests that the front of the cone has a slightly more rapid rate of polymerization. In moving actin cones, the actin turned over completely within about 12 minutes.
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Membrane dynamics of cystic bulge
The cystic bulge is the place where membrane remodeling occurs. In previous
ultrastructural work, Tokuyasu and colleagues
(Tokuyasu et al., 1972)
observed a large amount of membranous material in the cystic bulge. In order
to reveal membrane structures in the cystic bulge and membrane surrounding the
actin cone, we labeled cell membranes using the fluorescent membrane dye, FM
1-43 (Fig. 8), which has been
extensively used for probing membrane recycling activity in various cell types
(Kuromi and Kidokoro, 1998
).
This labeling worked best on spermatids where the cyst cells were removed, so
that the dye could interact with membrane of the spermatids directly. A
solution of 0.5 µM of FM1-43 labels the plasma membrane of the cystic bulge
completely within 10 minutes. The membrane forms complex architecture inside
the bulge. Optical sectioning demonstrates that these structures are
invaginations of plasma membrane with open connections to the outside. These
invaginations were present before individualization began. The individualized
part of sperm tails directly protruded from the cystic bulge and had no
connection with inner membrane structures within the bulge. After incubation
with FM 1-43 for 20 minutes and washing, cystic bulge movement was observed
over a period of 10 minutes. Because the magnification we used was high enough
to identify single membrane vesicles, we would expect to see membrane vesicle
uptake if there is endocytic activity. However, we could not see any membrane
uptake around site of membrane remodeling. Moreover, very few stained membrane
vesicles accumulate inside the cystic bulge after incubation with dye (see
Movies 5-7 at
http://dev.biologists.org/supplemental/).
In addition, if exocytosis contributes in a major way to membrane remodeling
at this site, we would expect to see appearance of unlabeled membrane patches
in the region of the actin cones during the observations made after washing
out the dye. These unlabeled regions would be derived from fusion of unlabeled
internal membrane vesicles. However, for the entire length of the observation
(at 15 second intervals, over 7 minutes), the membrane around the neck looked
very flat and homogenous (see Movie 7 at
http://dev.biologists.org/supplemental/).
Although we did not detect exocytic or endocytic activity in the region of the
cystic bulge, we were able to easily detect membrane uptake in the cyst cells
surrounding other cysts (see Movie 8 at
http://dev.biologists.org/supplemental/)
in the same culture. Double imaging of GFP-actin and FM 1-43 revealed that the
actin cones localized to the neck of the individualized sperm tail, at the
position where the membrane is deformed into thin tubes.
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DISCUSSION |
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The driving force of movement
Our data are most consistent with the idea that actin cone movement is
driven by actin polymerization, similar to lamellipodia extension and
Listeria motility. The speed of the cystic bulge movement is similar
to the speed of movement of the leading edge of lamellipodia
(Bear et al., 2002;
Watanabe and Mitchison, 2002
).
Cystic bulge movement is altered very quickly after inhibiting either assembly
or disassembly of actin, consistent with the requirement for active actin
assembly and disassembly for movement. In FRAP experiments, we detected a
slightly faster rate of turnover at the front than in the rear of the cone. It
is likely that the faster dynamics of actin at the front is important for
movement. In addition, the observed acceleration of actin dynamics after the
onset of movement supports this idea. Arp 2/3 complex
(Mullins et al., 1998
;
Welch et al., 1997b
), which is
the key factor involved in promoting actin polymerization at the leading edge
(Welch et al., 1997a
), is
enriched at the front of actin cones, suggesting that this site is important
for force generation. All these data support the idea that the driving force
is actin polymerization.
However, some puzzling differences in actin behavior in this structure when
compared with leading edge protrusion make it difficult to explain how
assembly drives movement in this case. First, we had expected that actin would
treadmill through the actin cone from front to back, because of assembly at
the front, i.e. in the direction of movement. This has been observed in other
actin motility processes. However, this is not the case in actin cones. The
filaments in the cone move forward relative to the substrate. Second, actin
turns over at a rate that is much slower than that of actin in lamellipodia
and Listeria comet tails. In both of these structures, filaments turn
over in 1-2 minutes (Theriot and
Mitchison, 1991; Theriot et
al., 1992
), but in case of actin cones, turnover takes 12 minutes.
Another puzzle is the stability of actin cones to depolymerization by LTA.
Actin completely turns over in 12 minutes in moving cones, so we might expect
that LTA would cause depolymerization in that time frame. However, even after
2 hours of LTA treatment, cones remain. It is likely that actin in cones is
stabilized by binding of cross linkers or other proteins, but the mechanism
that regulates stability to permit turnover as the cones move, but prevents
depolymerization when assembly stops is not clear.
Despite these differences from other motility processes, our favored model involves only actin assembly as the driving force for motility. In order to explain the dynamics of actin in the cone and results of pharmacological experiments, we would suggest that there are two actin structural components in a moving actin cone. The first actin structure is the actin cone itself. The three characteristics discussed above (stability, filament translocation and slow turnover) suggest that the actin cone is a highly organized and stable structure compared to the actin network in lamellipodia. It is likely that each actin cone moves forward as one unit. The second actin structure component is an actin network near the membrane that pushes the actin cone forward by force of polymerization. Actin filaments elongate near the membrane, similar to the leading edge, but the membrane is held rigid, rather than protruding as it does at the leading edge (Fig. 9). In this case, a photobleached GFP-actin in a filament would be pushed away from the membrane, i.e. `forward' relative to the membrane. Eventually, this filament would be crosslinked into the actin cone as new actin filaments assembled. This model is consistent with our data, but we do not have information about the orientation of actin filaments and the sites of actin monomer incorporation that would provide additional support for such a model. In addition, the molecules that might be important to keep the membrane rigid and prevent its protrusion are as yet unknown.
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Microtubule-based motility is not likely to be involved in cone movement. There are no cytoplasmic microtubules, which might participate in generating force in cooperation with microtubule motors and inhibitors of microtubule dynamics, and motors do not stop movement. In addition, when actin dynamics are altered, the cystic bulge stops immediately. If the movement was microtubule based, it is not clear why actin dynamics would be important.
Further studies are required to provide support for our model of actin cone motility. Additional studies examining membrane dynamics, effect of disruption of actin polymerization regulators and ultrastructure of the actin cones will be needed for more insight into the similarities and differences in the mechanism of actin cone movement and lamellipodia extension.
The role of actin cones in individualization
Although we do not yet understand fully the mechanism of actin cone
movement, we can speculate about the role of actin cones during
individualization. We suggest that the actin cones have three roles
(Fig. 9). First, the actin
cones have the ability to push the cystic bulge forward, using actin
polymerization. Second, the actin cones sweep the cytoplasm and organelles out
of the sperm flagella, acting as a sieve. Finally, the actin cones must bind
the cell membrane around them and shape it into the observed thin tubular
structure. Eventually, as the actin depolymerizes at the cone tip, the
membrane must attach to the axoneme.
Microtubule structure and formation of actin cones: signal for a
dramatic transition?
There is an interesting transition that occurs as individualization begins.
Microtubule staining disappears during a very short period around the onset of
actin cone movement. Our data suggests that this disappearance is due to
tubulin degradation as movement begins. This idea is supported by previous
observations that the amount of tubulin present in individualized spermatids
was much less than in cysts prior to individualization
(Kemphues et al., 1980), and
that cytoplasmic microtubules disappear during individualization
(Tokuyasu et al., 1972
).
This transition temporally coincides with the onset of actin cone movement, rather than sperm nucleus DNA condensation. FRAP experiments demonstrated that actin dynamics also accelerated after the onset of movement. Therefore, we suspect that a global signal orchestrates these events to trigger the onset of individualization.
Membrane remodeling does not require endocytosis or exocytosis.
Conventional endocytosis may not be important for movement of the cystic
bulge, because FM1-43 staining of cell membrane demonstrated that membrane
uptake did not take place around the actin cones and blocking endocytosis
using temperature shift of the shibire (dynamin) mutant did not
affect cystic bulge movement. In addition, no concentration of -adaptin
has been observed in the region around the actin cones, suggesting that no
coated pit formation occurs there (Rogat
and Miller, 2002
). Conversely, clathrin mutants have defects in
individualization (Bazinet et al.,
1993
; Fabrizio et al.,
1998
), but the reason that individualization fails has not been
well studied. The discrepancies in these data will only be resolved by further
analysis of the clathrin mutant phenotype and studies of the effects of loss
of function in other proteins in the endocytosis pathway.
Likewise, exocytosis may not play major role in the membrane remodeling
process, because FM1-43 membrane staining suggests that there is not a
significant amount of membrane insertion at the sites around the actin cones,
and BFA treatment did not affect the movement of cystic bulge. Our data do not
completely exclude the possibility that exocytic events participate in
remodeling, as we could not directly measure the exocytosis. However, it seems
more likely that the large number of membrane invaginations that are present
in the cystic bulge is a sufficient source of membrane to accomplish
remodeling. As seen in the movie of FM1-43 labeled spermatids (Movies 6, 7 at
http://dev.biologists.org/supplemental/,
the plasma membrane seems to be smoothly reorganized into thin tubular
structures around the actin cones. Furthermore, ultrastructural observations
have shown that the membrane around actin cones (investment cone in their
terminology) is flat, without any invaginating or docking membrane vesicles
(Tokuyasu et al., 1972) (T.N.,
unpublished). These data support the idea that the cell membrane in the cystic
bulge is directly deformed into a thin tubular structure.
Culturing cysts in vitro
The whole process of spermatogenesis from germ stem cell differentiation to
individualization into motile sperm occurs continuously during adult life.
There are many processes of interest to cell and developmental biologists that
occur during the different stages of spermatid differentiation. These
processes include meiosis, cell elongation, membrane remodeling, discarding of
cytoplasm and flagella formation. The culture system we have used is suitable
for studies in vitro of many aspects of spermatogenesis, as most of the
processes are evident in these cultures. Through culturing of testes from
various male sterile mutants and ectopically expressing a variety of genes,
including GFP-tagged proteins, it should be possible to learn much about these
events. Our studies using these cultures have successfully demonstrated that
actin polymerization is important for individualization. Studies are in
progress to fully understand how actin cones move and how they associate with
the membrane.
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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Abe, S. and Uno, S. (1984). Nuclear elongation of dissociated newt spermatids in vitro and their nuclear shortening by antimicrotubule agents. Exp Cell Res 154,243 -255.[Medline]
Bazinet, C., Katzen, A. L., Morgan, M., Mahowald, A. P. and
Lemmon, S. K. (1993). The Drosophila clathrin heavy chain
gene: clathrin function is essential in a multicellular organism.
Genetics 134,1119
-1134.
Bear, J. E., Svitkina, T. M., Krause, M., Schafer, D. A., Loureiro, J. J., Strasser, G. A., Maly, I. V., Chaga, O. Y., Cooper, J. A., Borisy, G. G. et al. (2002). Antagonism between Ena/VASP proteins and actin filament capping regulates fibroblast motility. Cell 109,509 -521.[Medline]
Bré, M. H., Redeker, V., Quibell, M., Darmanaden-Delorme,
J., Bressac, C., Cosson, J., Huitorel, P., Schmitter, J. M., Rossler, J.,
Johnson, T. et al. (1996). Axonemal tubulin polyglycylation
probed with two monoclonal antibodies: widespread evolutionary distribution,
appearance during spermatozoan maturation and possible function in motility.
J. Cell Sci. 109,727
-738.
Bubb, M. R., Spector, I., Beyer, B. B. and Fosen, K. M.
(2000). Effects of jasplakinolide on the kinetics of actin
polymerization. An explanation for certain in vivo observations. J.
Biol. Chem. 275,5163
-5170.
Castrillon, D. H., Gonczy, P., Alexander, S., Rawson, R.,
Eberhart, C. G., Viswanathan, S., DiNardo, S. and Wasserman, S. A.
(1993). Toward a molecular genetic analysis of spermatogenesis in
Drosophila melanogaster: characterization of male-sterile mutants generated by
single P element mutagenesis. Genetics
135,489
-505.
Cramer, L. P. (1999). Role of actin-filament disassembly in lamellipodium protrusion in motile cells revealed using the drug jasplakinolide. Curr. Biol. 9,1095 -1105.[CrossRef][Medline]
Cramer, L. P. and Mitchison, T. J. (1995). Myosin is involved in postmitotic cell spreading. J. Cell Biol. 131,179 -189.[Abstract]
Cross, D. P. and Sang, J. H. (1978). Cell culture of individual Drosophila embryos. I. Development of wild-type cultures. J. Embryol. Exp. Morphol. 45,161 -172.[Medline]
Cross, D. P. and Shellenbarger, D. L. (1979). The dynamics of Drosophila melanogaster spermatogenesis in in vitro cultures. J. Embryol. Exp. Morphol. 53,345 -351.[Medline]
Fabrizio, J. J., Hime, G., Lemmon, S. K. and Bazinet, C.
(1998). Genetic dissection of sperm individualization in
Drosophila melanogaster. Development
125,1833
-1843.
Fowler, G. L. (1973). In vitro cell differentiation in the testes of Drosophila hydei. Cell Differ. 2,33 -42.[CrossRef][Medline]
Fuller, M. T. (1993). Spermatogenesis. InThe Development of Drosophila melanogaster (ed. M. Bate and A. M. Arias), pp. 71-147. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.
Fuller, M. T. (1998). Genetic control of cell proliferation and differentiation in Drosophila spermatogenesis. Semin. Cell Dev. Biol. 9, 433-444.[CrossRef][Medline]
Gibbons, I. R. (1981). Cilia and flagella of eukaryotes. J. Cell Biol. 91,S107 -S124.
Gonczy, P., Viswanathan, S. and DiNardo, S. (1992). Probing spermatogenesis in Drosophila with P-element enhancer detectors. Development 114, 89-98.[Abstract]
Hendricks, L. C., McClanahan, S. L., Palade, G. E. and Farquhar, M. G. (1992). Brefeldin A affects early events but does not affect late events along the exocytic pathway in pancreatic acinar cells. Proc. Natl. Acad. Sci. USA 89,7242 -7246.[Abstract]
Hicks, J. L., Deng, W. M., Rogat, A. D., Miller, K. G. and
Bownes, M. (1999). Class VI unconventional myosin is required
for spermatogenesis in Drosophila. Mol. Biol. Cell
10,4341
-4353.
Hoyle, H. D. and Raff, E. C. (1990). Two Drosophila beta tubulin isoforms are not functionally equivalent. J. Cell Biol. 111,1009 -1026.[Abstract]
Hudson, A. M. and Cooley, L. (2002). A subset
of dynamic actin rearrangements in Drosophila requires the Arp2/3 complex.
J. Cell Biol. 156,677
-687.
Keller, H. U. (2000). Redundancy of lamellipodia in locomoting Walker carcinosarcoma cells. Cell Motil. Cytoskel. 46,247 -256.[CrossRef][Medline]
Kemphues, K. J., Kaufman, T. C., Raff, R. A. and Raff, E. C. (1982). The testis-specific beta-tubulin subunit in Drosophila melanogaster has multiple functions in spermatogenesis. Cell 31,655 -670.[Medline]
Kemphues, K. J., Raff, E. C., Raff, R. A. and Kaufman, T. C. (1980). Mutation in a testis-specific beta-tubulin in Drosophila: analysis of its effects on meiosis and map location of the gene. Cell 21,445 -451.[Medline]
Kuromi, H. and Kidokoro, Y. (1998). Two distinct pools of synaptic vesicles in single presynaptic boutons in a temperature-sensitive Drosophila mutant, shibire. Neuron 20,917 -925.[Medline]
Laughran, L. J., Stanley, H. P. and Bowman, J. T. (1976). Electron microscopic study of postcytokinetic cell fusion in an autosomal male sterile mutant (ms(2)3R) of Drosophila melanogaster. J. Ultrastruct. Res. 56,21 -30.[Medline]
Liebrich, W. (1981). In vitro spermatogenesis in Drosophila. I. Development of isolated spermatocyte cysts from wild-type D. hydei. Cell Tissue Res. 220,251 -262.[Medline]
Liebrich, W. (1982). The effects of cytochalasin B and colchicine on the morphogenesis of mitochondria in Drosophila hydei during meiosis and early spermiogenesis. An in vitro study. Cell Tissue Res. 224,161 -168.[Medline]
Lifschytz, E. and Hareven, D. (1977). Gene expression and the control of spermatid morphogenesis in Drosophila melanogaster. Dev. Biol. 58,276 -294.[Medline]
Lindsley, D. I. and Lifschytz, E. (1972). The genetic control of spermatogenesis in Drosophila. In The Genetics of Spermatozoon (ed. R. A. Beatty and S. Gluecksohn-Waelsch), pp.203 -222. Copenhagen: Bogtrykkeriet Forum.
Lindsley, D. I. and Tokuyasu, K. T. (1980). Spermatogenesis. In Genetics and Biology of Drosophila, 2nd edn (ed. M. Ashburner and T. R. Wright), pp.225 -294. New York: Academic Press.
Mayer, T. U., Kapoor, T. M., Haggarty, S. J., King, R. W.,
Schreiber, S. L. and Mitchison, T. J. (1999). Small molecule
inhibitor of mitotic spindle bipolarity identified in a phenotype-based
screen. Science 286,971
-974.
Misumi, Y., Miki, K., Takatsuki, A., Tamura, G. and Ikehara,
Y. (1986). Novel blockade by brefeldin A of intracellular
transport of secretory proteins in cultured rat hepatocytes. J.
Biol. Chem. 261,11398
-11403.
Mullins, R. D., Heuser, J. A. and Pollard, T. D.
(1998). The interaction of Arp2/3 complex with actin: nucleation,
high affinity pointed end capping, and formation of branching networks of
filaments. Proc. Natl. Acad. Sci. USA
95,6181
-6186.
Robinson, D. N. and Cooley, L. (1997).
Examination of the function of two kelch proteins generated by stop codon
suppression. Development
124,1405
-1417.
Rogat, A. D. and Miller, K. G. (2002). A role for myosin VI in actin dynamics at sites of membrane remodeling during Drosophila spermatogenesis. J. Cell Sci. 115,4855 -4865.[CrossRef][Medline]
Romrell, L. J., Stanley, H. P. and Bowman, J. T. (1972). Genetic control of spermiogenesis in Drosophila melanogaster: an autosomal mutant (ms(2)10R) demonstrating disruption of the axonemal complex. J. Ultrastruct. Res. 38,578 -590.[Medline]
Rørth, P. (1998). Gal4 in the Drosophila female germline. Mech. Dev. 78,113 -118.[CrossRef][Medline]
Small, J. V., Stradal, T., Vignal, E. and Rottner, K. (2002). The lamellipodium: where motility begins. Trends Cell Biol. 12,112 -120.[CrossRef][Medline]
Theriot, J. A. and Mitchison, T. J. (1991). Actin microfilament dynamics in locomoting cells. Nature 352,126 -131.[CrossRef][Medline]
Theriot, J. A., Mitchison, T. J., Tilney, L. G. and Portnoy, D. A. (1992). The rate of actin-based motility of intracellular Listeria monocytogenes equals the rate of actin polymerization. Nature 357,257 -260.[CrossRef][Medline]
Tilney, L. G., Connelly, P. S., Vranich, K. A., Shaw, M. K. and
Guild, G. M. (2000). Actin filaments and microtubules play
different roles during bristle elongation in Drosophila. J. Cell
Sci. 113,1255
-1265.
Tokuyasu, K. T. (1974). Dynamics of spermiogenesis in Drosophila melanogaster. 3. Relation between axoneme and mitochondrial derivatives. Exp. Cell Res. 84,239 -250.[Medline]
Tokuyasu, K. T., Peacock, W. J. and Hardy, R. W. (1972). Dynamics of spermiogenesis in Drosophila melanogaster. I. Individualization process. Z Zellforsch. Mikrosk. Anat. 124,479 -506.[Medline]
Townsley, F. M. and Bienz, M. (2000). Actin-dependent membrane association of a Drosophila epithelial APC protein and its effect on junctional Armadillo. Curr. Biol. 10,1339 -1348.[CrossRef][Medline]
Tracey, W. D., Jr, Ning, X., Klingler, M., Kramer, S. G. and
Gergen, J. P. (2000). Quantitative analysis of gene function
in the Drosophila embryo. Genetics
154,273
-284.
van der Bliek, A. M. and Meyerowitz, E. M. (1991). Dynamin-like protein encoded by the Drosophila shibire gene associated with vesicular traffic. Nature 351,411 -414.[CrossRef][Medline]
Watanabe, N. and Mitchison, T. J. (2002).
Single-molecule speckle analysis of actin filament turnover in lamellipodia.
Science 295,1083
-1086.
Welch, M. D., DePace, A. H., Verma, S., Iwamatsu, A. and
Mitchison, T. J. (1997a). The human Arp2/3 complex is
composed of evolutionarily conserved subunits and is localized to cellular
regions of dynamic actin filament assembly. J. Cell
Biol. 138,375
-384.
Welch, M. D., Iwamatsu, A. and Mitchison, T. J. (1997b). Actin polymerization is induced by Arp2/3 protein complex at the surface of Listeria monocytogenes. Nature 385,265 -269.[CrossRef][Medline]
Welch, M. D. and Mullins, R. D. (2002). Cellular control of actin nucleation. Annu. Rev. Cell Dev Biol. 18,247 -288.[CrossRef][Medline]
Wilkinson, R. F., Stanley, H. P. and Bowman, J. T. (1974). Genetic control of spermiogenesis in Drosophila melanogaster: the effects of abnormal cytoplasmic microtubule populations in mutant ms(3)10R and its colcemid-induced phenocopy. J. Ultrastruct. Res. 48,242 -258.[Medline]