We thank P. Verrijzer, J. Johansen H., Okhura for antibodies and A. Debec, the Bloomington and Vienna stock centers for flies. Thanks to C. Jackson for suggestions and stimulating discussions.

1. Drosophila testes collection
NOTE: Young males (0-15 h post-eclosion) are necessary for examining diploid germ cells (i.e., spermatogonia and spermatocytes).
<ol>
	<li>Select young flies as much as possible to minimize interference from developing spermatids.</li>
	<li>Anesthetize flies by brief exposure to carbon dioxide and transfer to a fly pad14,15.</li>
	<li>Under a dissecting microscope (10x magnification), use forceps to remove the head.</li>
	<li>Transfer 5 to 10 decapitated flies to 100 &#181;L of Ringer&#8217;s buffer (183 mM KCI, 47 mM NaCl, 10 mM Tris-HCI, 1 mM EDTA, complete protein inhibitor, pH 6.8) on a silicone-coated glass slide on a black background support.</li>
	<li>With dissecting microscope at a 40x magnification, use one pair of number 5 forceps to hold a&#160;fly between the thorax and the abdomen, and, with other forceps, pull the abdomen away from the thorax. Rapidly isolate the testes and adjacent tissues from the fly carcass15 and place in a new drop of Ringer&#8217;s buffer on the same slide.</li>
	<li>Once all the testes are ready, clean the testes from the remaining tissues (accessory gland and external genitalia).</li>
	<li>Remove excess buffer from the drop containing the dissected testes. Typically, use a p200 pipette with a 10 &#956;L tip fitted on top of a 200 &#956;L tip. This allows the removal of the liquid excess through a small aperture. Place a drop of 4% paraformaldehyde fixative on top of the drop containing the dissected testes, and then rapidly transfer the testes to a tube containing fresh fixative solution.</li>
</ol>
2. Paraformaldehyde fixation
<ol>
	<li>Prepare 2 mL of fixative solution just before use: 600 &#181;L of 4% paraformaldehyde diluted in phosphate buffered saline (PBS) with 30 &#181;L of NP40 20% plus 1200 &#181;L of heptane in a 2 mL siliconized microcentrifuge tube. Use 4% paraformaldehyde in PBS prepared in 600 &#956;L aliquots and stored at -20&#176;C. 
	Caution: Heptane and paraformaldehyde are toxic and should be handled in a fume hood. Dispose of them according to the host institute&#8217;s regulations.</li>
	<li>Transfer the testes using a p200 micropipette with a wide aperture tip to the PFA/NP40/heptane solution. Shake the tube up and down for 30 s by hand. Continue fixing the testes on a rotary mixer at room temperature for 30 min.</li>
	<li>Stop the rotary mixer and place the tube in a tube rack to allow phase separation and for the testes to settle to the bottom of the tube.</li>
	<li>Remove all the heptane and the fixative solution and rinse the testes quickly three times with 1x PBS.</li>
	<li>Transfer to a clean 200 &#181;L individual tube. Perform all incubations in these small tubes to minimize the amounts of antibodies employed.</li>
	<li>Remove excess buffer using a 200 &#181;L micropipette with both a P200 and a P10 tip attached. Take care not to aspirate testes.</li>
	<li>Keep testes in 1x PBS on ice until all samples are ready.</li>
</ol>
3. Antibody staining in solution
<ol>
	<li>Discard the PBS and leave testes in 0.3% PBS-T for 30 min at room temperature to permeabilize cell membranes.</li>
	<li>Pre-incubate in the same buffer plus 5% normal goat serum (NGS) for 1 h.</li>
	<li>Add 200 &#181;L of the primary antibody diluted in 0.3% PBS-T plus 5% NGS (Table of Materials).</li>
	<li>Incubate the primary antibody overnight at 4 &#176;C (or room temperature) with gentle agitation on a rocker.</li>
	<li>Wash 3 times in 0.3% PBS-T for 15 min on a rocker at room temperature. Allow the testes to settle to the bottom of the tube by gravity.</li>
	<li>Add 200 &#181;L of secondary antibody diluted 1:500 from the DyLight conjugated series.</li>
	<li>Incubate the secondary antibody in a dark chamber for 4 h at room temperature.</li>
	<li>Wash in 0.3% PBS-T for 15 min on a rocker at room temperature. Then repeat with 0.2% PBS-T and 0.1% PBS-T, each time for 30 min on a rocker at room temperature. Do a final wash in 1x PBS for 30 min to remove the detergent.</li>
	<li>Add 180 &#181;L of DAPI solution at 1 &#181;g/mL in 1x PBS for 15 min. The stock solution is 10 mg/mL in H2O. Avoid making stock solution in PBS, which can reduce the DAPI staining signal on diffuse chromatin in larger spermatocytes.</li>
	<li>Wash 3x for 5 min each.</li>
	<li>Use a hydrophobic barrier pen to draw a circle on a clean slide.</li>
	<li>Aspirate testes and put them on the slide. Pre-rinsing the wide aperture tip with 0.1% PBS-T prevents the testes from sticking to the tip. Remove excess PBS.</li>
	<li>Add a drop (approximately 100 &#181;L) of Citifluor AF1.</li>
	<li>Gently place a glass coverslip over the tissue, taking care to avoid trapping air bubbles.</li>
	<li>Seal the coverslip to the slide using clear nail polish.</li>
	<li>Observe slides on a microscope.</li>
</ol>
4. Colocalization analyses
<ol>
	<li>Acquire confocal images. We used a Zeiss LSM 710 confocal microscope (63x Plan Apochromatic oil) with a z-resolution of 0.5 or 1 &#181;m.</li>
	<li>Select different cells from independent testes.</li>
	<li>Import images to processing software (e.g., Imaris).</li>
	<li>Define the nucleus by manual segmentation in order to exclude the neighboring nuclei. Use the Coloc software; it collects the Pearson&#8217;s correlation coefficients (PCCs) inside the entire volume between 2 fluorophores.</li>
</ol>

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G., Cenci, G., Gatti, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Immunostaining of Drosophila testes. 2011 (10), 1273-1275 (2011).</li><li>Bonaccorsi, S., Giansanti, M. G., Cenci, G., Gatti, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Paraformaldehyde fixation of Drosophila testes. 2012 (1), 102-104 (2012).</li><li>White-Cooper, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spermatogenesis:+analysis+of+meiosis+and+morphogenesis.">Spermatogenesis: analysis of meiosis and morphogenesis.</a> Methods in Molecular Biology. 247, 45-75 (2004).</li><li>Kibanov, M. V., Kotov, A. A., Olenina, L. 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The authors have nothing to disclose.

This protocol provides a method for successful immunolabeling of whole-mount adult Drosophila testes. As reported in other procedures young males must be used (we even shortened the age to less than 20 hours after eclosion) to minimize the presence of developing sperm, which risks obstructing the spermatocytes. The dissection should be rapid and the antibody dilution must be adjusted to optimize the signal-to-noise ratio21,22,23. Because using other protocols, we did not observe staining throughout the depth of the testis, we developed a fixation method combining PFA together with NP40 and heptane. We now routinely use 600 &#181;L of 4% paraformaldehyde in PBS with 30 &#181;L of 20% NP40 plus two volumes of heptane. This is critical to allow for sufficient permeabilization. Reducing the amount of NP40 to 10 &#181;L visibly reduces the labeling. All the subsequent steps were carried out in suspension and the excess buffer was removed by allowing the testes to descend gently to the bottom of the tube by gravity. For the last step transferring samples to a slide, we rinse the tip in 0.1% PBS-T to prevent the testes sticking to the cone. According to observations, the immunohistological staining protocol works well with a variety of nuclear proteins, including the markers used here, as well as probes of epigenetic histone modifications, lamin and nucleoporins. We note that the protocol can also be used for larval testes, in which case the heptane should be omitted, as it makes the testes more difficult to recover intact.
The protocol is especially designed to maintain undamaged stacks of the spermatocytes together for an efficient and uniform labeling of the testis. We illustrated the protocol with the large late stage spermatocytes, whose misshapen nuclei make them among the most difficult cells to preserve. Furthermore, in these spermatocytes the nuclear labeling often becomes dilute probably because of the size of the nuclei. For example, it is commonly observed that DAPI or lamin labeling is stronger in the cells at the testis apex and becomes progressively weaker and more disperse as the spermatocytes are growing3,24. This also could be one of the reasons that the distribution of nucleoporins has only been described in earlier stage cells (spermatogonia and early spermatocytes)17.
Excellent images of immunostaining of Drosophila S5 spermatocytes have been published, particularly for the testis-specific t-BRD and dany proteins25,26 using either 0.3% sodium deoxycholate protocol or following squashing of the tissue21,22. In our hands, squashes lead to a significant non-uniformity of staining in the depths of the organ and as well sacrificing the 3D structure. Even if the use of 0.3% sodium deoxycholate gave better results, we still observed non-uniformity. Thus, we prefer NP40 for its reproducibility and quality of labeling. The confocal images permit reconstruction of 3D images for nuclear proteins. For example, we have successfully applied this method using many different probes, illustrated here by the intranuclear MINT domain, intimately associated with the autosomal chromatin masses and the nuclear pore network. Intriguing 3D distribution of nuclear pores of these spermatocyte nuclei is now available for further studies. The images permit reproducible quantification of colocalization from different spermatocytes from the same testis and from different testes. To our knowledge this is the first time that accurate quantification by Pearson correlation coefficient has been measured on such samples. It gives the confidence required for accurate immunolabelled protein levels comparison of different testes.

Drosophila testes are a valuable model for the study of nuclear architecture. In mitotic spermatogonial cells, the nuclear volume is largely occupied by chromatin. These cells undergo four rounds of division, producing a cyst of 16 primary spermatocytes. Over the next several days, the spermatocytes pass through 6 developmental stages (S1-S6), greatly increasing in volume, approximatively 20-fold1,2. They also change their nuclear morphology. The outline of the nucleus, revealed by the lamin Dm0, becomes irregular and shows deep invaginations1,3. This is accompanied by extensive gene expression changes and modifications in chromatin organization1,4,5,6,7. The interphase chromosomes are well-defined in stage S4-S5, forming three distinct masses corresponding to the 2 major bivalent autosomes and the X-Y pair docked with the nucleolus.
Mad1 was originally isolated as a key component of the mitotic checkpoint (reviewed in8). In interphase, it is described as located primarily at the nuclear envelope but also in the nucleoplasm9,10,11. In Drosophila testis, we reported that Mad1 is prominent in the nucleoplasm, intimately associated with the two major autosomal chromatin masses and to a lesser extent with the XY chromatin. We defined this chromatin-associated structure as MINT (Mad1-containing IntraNuclear Territory)12. Four other proteins were associated with MINTs in spermatocytes: the nucleoporin Mtor/Tpr, the SUMO peptidase Ulp1, the mitotic checkpoint protein Mad2 and a subunit of a Polycomb-like complex Raf212.
To complete the description of MINTs, we needed to use antibodies and were confronted with the technical problems generally encountered with immunostaining in testis: a poor permeability resulting in a significant non-uniformity of staining in the depth of the testis, particularly in the large spermatocytes. Squashed preparations increased antibody access but led to compressed images, restricting the 3D analyses, and preventing measurement of quantitative fluorescence, including colocalization. The problem of antibody penetration could also be addressed by permeabilizing agents such as 0.3% Na deoxycholate13, but this procedure did not in our hands yield reproducible results. Because we performed many co-labeling experiments, we looked to improve the permeabilization protocol. The combination of 1% NP40 plus heptane resulted in more reproducibility, particularly in studying the large spermatocytes.
This protocol performs fixation and immunostaining of testes in suspension, improving the sample quality as well as enhancing reproducibility, and rendering it suitable for confocal microscopy. We used the protocol to better define the spatial relationships of certain nuclear envelope proteins with the nucleoplasmic structures of large spermatocytes. This method will permit better investigations of the changes that accompany spermatocyte development, for example the dynamic structural changes of the nucleus including chromatin, nucleoplasm and the nuclear pores.

<table><tbody><tr><td>&lt;em&gt;&lt;strong&gt;Drosophila srains&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>ketelGFP y; ketelGFP/y+CyO</td><td></td><td></td><td>Gift from A. Debec (Institut Jacques Monod, UMR 7592, Paris, France)</td></tr><tr><td>Mad1-GFP</td><td></td><td></td><td>J Cell Sci. 2011 May 15;124(Pt 10):1664-71.</td></tr><tr><td>Mtor RNAi&amp;nbsp;</td><td>VDRC</td><td>ID 110218</td><td>Vienna Drosophila RNAi Center</td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DAPI&amp;nbsp;</td><td>Thermo</td><td>D1306</td><td>Stock solution must be prepared in H2O</td></tr><tr><td>Dumont # 5 Fine Forseps&amp;nbsp;</td><td>Fine Science Tools</td><td></td><td></td></tr><tr><td>MBP Wide Bore Pipette Tips</td><td>Fisherbrand</td><td>11917744</td><td>Wide aperture tip</td></tr><tr><td>Thermo Scientific&amp;nbsp;0.2 mL Individual Tube</td><td>Thermo</td><td>AB-0620</td><td></td></tr><tr><td>2 ml micro centrifuge tubes&amp;nbsp;</td><td>Sigma</td><td>T3531-200EA&amp;nbsp;</td><td>Siliconized 2ml tube</td></tr><tr><td>Citifluor AF1</td><td>Citifluor</td><td>AF1</td><td></td></tr><tr><td>Dakopen&amp;nbsp;</td><td>Sigma</td><td>Z377821-1EA&amp;nbsp;</td><td></td></tr><tr><td>Normal Goat Serum (NGS)</td><td>Sigma</td><td>NS02L-1ML</td><td></td></tr><tr><td>Paraformaldehyde 4%</td><td>Sigma</td><td>P6148-500G&amp;nbsp;</td><td>Paraformaldehyde (4%) dissolved in PBS 1X&amp;nbsp;; aliquot by 600 ml&amp;nbsp;</td></tr><tr><td>PBS 1X</td><td>Thermo</td><td>14190094</td><td>DPBS, no calcium, no magnesium</td></tr><tr><td>0.1% (0.2 or 0.3%) PBS-T&amp;nbsp;</td><td></td><td></td><td>PBS 1X&amp;nbsp; containing 0.1% (0.2 % or 0.3%) Triton X-100</td></tr><tr><td>Triton X-100, for molecular biology,&amp;nbsp;</td><td>Sigma</td><td>T8787</td><td></td></tr><tr><td>&lt;strong&gt;Antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>GFP booster</td><td>Chromotek</td><td>gba488</td><td>1/200</td></tr><tr><td>Mouse anti-Mtor</td><td></td><td></td><td>1/40 gift from J. Johansen, Iowa State University</td></tr><tr><td>Rat anti-Nup62</td><td></td><td></td><td>1/200 gift from H. Ohkura (University of Edinburgh, UK</td></tr><tr><td>Guinea-pig anti-Ulp1</td><td></td><td></td><td>1/200 gift from C. P. Verrijzer, Erasmus University, Rotterdam</td></tr><tr><td>Rabbit anti-Raf2</td><td></td><td></td><td>1/200 gift from C. P. Verrijzer, Erasmus University, Rotterdam</td></tr><tr><td>DyLight conjugated series&amp;nbsp;</td><td>Thermo</td><td></td><td>1/500 Donkey anti-(guinea-pig, mouse,&amp;nbsp; rabbit or rat) as second antibodies</td></tr></tbody></table>

immunostaining of whole-mount drosophila testes for 3d confocal analysis of large spermatocytes

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm development. However, immunolabeling of the whole Drosophila testis is often associated with significant non-uniformity of staining due to antibody penetration. Squashed preparations only partially overcome the problem since it decreases the 3D quality of the analyses. Herein, we describe a whole-mount protocol using NP40 and heptane during fixation together with immunolabeling in liquid media. It preserves the volume suitable for confocal microscopy together with reproducible and reliable labeling. We show different examples of 3D reconstitution of spermatocyte nuclei from confocal sections. The intra- and inter-testes reproducibility allows 3D quantification and comparison of fluorescence between single cells from different genotypes. We used different components of the intranuclear MINT structure (Mad1-containing Intra Nuclear Territory) as well as two components associated with the nuclear pore complex to illustrate this protocol and its applications on the largest cells of the testis, the S4-S5 spermatocytes.

The main obstacle to obtaining adequate staining of Drosophila testes is the limited penetrability of antibodies, which has been partially resolved by using squashed preparations but at the cost of inducing a restriction of the 3D analyses (for example 3D representation or measure of colocalization). The procedure permits uniform labeling and preserves the volume of all the cells of the testis. Here we have focused the illustration of the method on the 3D representation of S4-S5 spermatocytes, as they are the largest cells and thus more sensitive to deformation from squashing. We used different nuclear markers, including nucleoporins and the labeling of the intranuclear MINT domain (Mad1-containing Intra Nuclear Territory) described by the mitotic protein Mad112. As MINTs could be revealed by several different antibodies, we used them to test the robustness of the protocol for quantitative 3D colocalization, a method that necessitates a correct spatial resolution.
The protocol described above permits the acquisition of confocal images of sufficient quality to then be treated for 3D reconstitution. Four adjacent serial sections of a stage S5 spermatocyte are depicted in Figure 1A, showing a representative immunostaining of MINT (Mad1 immunolabeling using GFP booster in red) together with DAPI (in cyan). The chromosomes are organized into the characteristic 3 major distinct clusters and the MINT structure is shown in fine detail around the DNA, intimately associated with--but distinct from--the autosomal chromatin masses. The resolution also permits detection of the presence of Mad1 at the nuclear periphery as well as between the autosomal masses of chromosomes 2 and 3. We used Imaris software for the 3D representation. The nuclear surface was defined by the low level presence of Mad1 at the nuclear envelope (in grey) and MINT was defined by high level accumulation of Mad1 inside the nucleus (in red) together with DAPI staining (in blue) (Figure 1A). The 3D reconstitution movie well illustrates the volume and the relationship of Mad1 (at the nuclear envelope and in the MINT) together with the chromosomes in stage S4&#8211;S5 spermatocytes.
The distribution of nucleoporins has been only poorly described in these cells, probably due to the difficulty detecting them in these large misshapen nuclei. The distribution of Nup154 has only been reported once previously16 and Nup153 and Nup98 were only studied in the spermatogonia and small, early spermatocytes17. The 3D reconstitutions of large S5 spermatocytes stained by the nucleoporin Nup62 and by importin &#946; (product of the ketel gene) were feasible and are represented in Figure 1C and Figure 1D, respectively. These nuclear pore-associated components appear non-uniformly distributed over the nuclear rim as dots of irregular size. For example, the importin &#946; gathers around the autosomal chromatin similarly to where a lamin network has been already described3. This kind of distribution with pore-rich areas and large pore-free areas has been described by electron microscopy in rodent spermatocytes18. Understanding the changes in lamin and nucleoporin distribution would help to understand the nuclear envelope structure in these G2 cells just prior to male meiosis.
While conserving the 3D volume of the cells this protocol also permits antibodies to penetrate uniformly into the depth of every cell, allowing quantitative analysis such as colocalization. The Pearson correlation coefficient (PCC) is a well-established metric. It can range from +1 (denoting perfect positive correlation) to &#8722;1 (perfect negative correlation)19. To evaluate the localization of MINT partner proteins relative to Mad1 in spermatocytes, we have previously estimated the PCC on a single section of 0.5 &#181;m using ImageJ in S5 spermatocytes12 (Figure 2A,B). We tested the protocol more deeply by using 3D colocalization on the whole nuclei instead of just a surface. The volume of the nuclei was defined by manual segmentation and the PCC was measured using the Colocation tool. We analyzed eight S5 spermatocyte nuclei, from the same testis or different testes. Figure 2B shows a 3D representation of a nucleus that served to measure the PCC. Figure 2D shows the measurement of the PCC on eight different S5 spermatocyte nuclei from three different testes. PCC values are not different, varying from 0.6 and 0.75, indicating a high value for colocalization. These are indistinguishable from the PCC defined on a single section. Thus, the procedure permits a reproducible uniform labeling and preserves a good volume of all the cells of the testis.
Because the protocol allows reliable measurement of colocalization from independent testes, it permitted comparison of staining from different fly genotypes. This was illustrated here by comparison of WT flies and flies RNAi depleted for Mtor (generated by expression of dsRNA UAS lines and the Bam-Gal4 driver that achieve substantial depletion in late spermatogonia and early spermatocytes20). As described for the colocalization experiments, Mtor-RNAi-depleted testes and wild types testes were dissected and fixed together and the confocal images were acquired under constant settings. Respecting these conditions, the comparison of individual cells becomes possible. We observed that the MINT is more affected than the nuclear envelope by the loss of Mtor as shown by representative Mtor RNAi-depleted (upper) and wt (lower) cells (Figure 3)12.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61061/61061fig01.jpg" /> 
Figure 1: Illustration of immunostaining of Drosophila S4-S5 spermatocytes nuclei using different probes.&#160;(A) Images of a single stage 5 spermatocyte nucleus immunostained with &#945;-GFP antibody (GFP booster) detecting the MINT structure by Mad1-GFP (red) together to DAPI staining (blue). It shows with precision the relationship between Mad1 and the two autosomal chromatin masses (represented as asterisks). Mad1 is also faintly detected at the periphery of the nucleus. All images are serial Z-projections of a stack of 3 (0.5 &#181;m) successive confocal slices. Confocal images were acquired on a confocal microscope (63x, Plan Apochromatic oil DIC objective lens). Scale bars are 5 &#181;m. (B) Frame from a movie rendering of the same spermatocyte nucleus. The nuclear surface was defined by low expression of Mad1 at the nuclear envelope (in greys) and MINT was defined by high expression of Mad1 inside the nucleus (in red). (A, B) Reprinted with permission from previously published figure (from12). (C) 3D image of a stage 5 spermatocyte nucleus with simultaneous immunostaining of Nup62 (yellow) and MINT (GFP booster in red). Scale bar is 4 &#181;m. (D) 3D image of a stage 5 spermatocyte nucleus with simultaneous immunostaining of the importin-&#946; Ketel (yellow) together with DNA (DAPI in blue). Scale bars are 5 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61061/61061fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61061/61061fig02.jpg" /> 
Figure 2: 2D and 3D Colocalization quantification of Mad1 together with Ulp1.&#160;(A) Merged images of a stage 5 spermatocyte nucleus immunostained with GFP booster antibody detecting Mad1 (green) and with anti-Ulp1 antibody (red) represented as a 1 &#181;m thick stack of a spermatocyte nucleus. On the right is indicated the Pearson intensity correlation coefficient calculated using the Coloc-2 ImageJ plugin. The PCC was estimated to 0.73 (Reprinted with permission from Raich12). (B) Merged image of stage 5 spermatocyte nucleus (corresponding to nucleus G in the adjacent table) immunostained with GFP booster antibody detecting Mad1 (green) and with anti-Ulp1 antibody (red) represented in a 3D projection. The volume of the nucleus was defined by manual segmentation. On the right, the table lists the Pearson intensity correlation coefficients calculated on 8 different spermatocytes (A to H) from 3 different testes (1 to 3). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61061/61061fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61061/61061fig03.jpg" /> 
Figure 3: Immunofluorescence analysis of Mtor RNAi-treated spermatocytes compared to wild type.&#160;Cells depleted of Mtor using MtorRNAi (ID 110218) cells (upper panels) and wild type cells (lower panels) were stained for Mad1, Raf2 and Mtor proteins. The testes were fixed and stained together, mounted on the same slide and images acquired with the same parameters. The RNAi cells could be distinguished from WT by the absence or presence respectively of Mad1-GFP. The figure is representative of maximum projections of 4 x 0.5 &#181;m stacks. Scale bar is 5 &#181;m (Reprinted with permission from previously published figure in12). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61061/61061fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes at JoVE.com

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

We present here an improved protocol for the whole-mount preparation and immunostaining of Drosophila testes, suitable for confocal microscopy and also allowing reproducible and reliable labeling. We illustrate this protocol by 3D representation and quantification of colocalization experiments on the large S4-S5 spermatocytes.

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm ...

immunostaining-whole-mount-drosophila-testes-for-3d-confocal-analysis

Nanyang Technological University

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JoVE Journal

Biology

4.1K Views.  Université de Paris, F 75006, Paris, France. We present here an improved protocol for the whole-mount preparation and immunostaining of Drosophila testes, suitable for confocal microscopy and also allowing reproducible and reliable labeling. We illustrate this protocol by 3D representation and quantification of colocalization experiments on the large S4-S5 spermatocytes.

Video: Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Developmental Biology

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Transillumination-Assisted Dissection of Specific Stages of the Mouse Seminiferous Epithelial Cycle for Downstream Immunostaining Analyses

Spermatogenesis is a unique differentiation process that ultimately gives rise to one of the most distinct cell types of the body, the sperm. Differentiation of germ cells takes place in the cytoplasmic pockets of somatic Sertoli cells that host 4 to 5 generations of germ cells simultaneously and coordinate and synchronize their development. Therefore, the composition of germ cell types within a cross-section is constant, and these cell associations are also known as stages (I&#8211;XII) of the seminiferous epithelial cycle. Importantly, stages can also be identified from intact seminiferous tubules based on their differential light absorption/scatter characteristics revealed by transillumination, and the fact that the stages follow each other along the tubule in a numerical order. This article describes a transillumination-assisted microdissection method for the isolation of seminiferous tubule segments representing specific stages of mouse seminiferous epithelial cycle. The light absorption pattern of seminiferous tubules is first inspected under a dissection microscope, and then tubule segments representing specific stages are cut and used for downstream applications. Here we describe immunostaining protocols for stage-specific squash preparations and for intact tubule segments. This method allows a researcher to focus on biological events taking place at specific phases of spermatogenesis, thus providing a unique tool for developmental, toxicological, and cytological studies of spermatogenesis and underlying molecular mechanisms.

This protocol describes transillumination-assisted microdissection of segments of adult mouse seminiferous tubules representing specific stages of seminiferous epithelial cycle, and cell types therein, and subsequent immunostaining of squash preparations and intact tubule segments.

Spermatogenesis is a unique differentiation process that ultimately gives rise to one of the most distinct cell types of the body, the sperm. ...

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Imaging Centrosomes in Fly Testes

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell differentiation. Centrosomes are essential to determine the cell division axis during mitosis and to nucleate cilia during interphase. The identity of the proteins that mediate these dynamic changes remains only partially known, and the function of many of the proteins that have been implicated in these processes is still rudimentary. Recent work has shown that Drosophila spermatogenesis provides a powerful system to identify new proteins critical for centrosome function and formation as well as to gain insight into the particular function of known players in centrosome-related processes. Drosophila is an established genetic model organism where mutants in centrosomal genes can be readily obtained and easily analyzed. Furthermore, recent advances in the sensitivity and resolution of light microscopy and the development of robust genetically tagged centrosomal markers have transformed the ability to use Drosophila testes as a simple and accessible model system to study centrosomes. This paper describes the use of genetically-tagged centrosomal markers to perform genetic screens for new centrosomal mutants and to gain insight into the specific function of newly identified genes.

Imaging of centrosomal proteins during Drosophila spermatogenesis is a powerful method to identify new proteins critical for centrosome biology as well as to elucidate the particular function of known players in this process.

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and organogenesis. Immunostaining allows for the visualization of developing tissues and organs. However, a protective cuticle that forms at the end of embryogenesis prevents permeation of antibodies into late-stage embryos and larvae. While dissection prior to immunostaining is regularly used to analyze Drosophila larval tissues, it proves inefficient for some analyses because small tissues may be difficult to locate and isolate. Sonication provides an alternative to dissection in larval Drosophila immunostaining protocols. It allows for quick, simultaneous processing of large numbers of late-stage embryos and larvae and maintains in situ morphology. After fixation in formaldehyde, a sample is sonicated. Sample is then subjected to immunostaining with antigen-specific primary antibodies and fluorescently labeled secondary antibodies to visualize target cell types and specific proteins via fluorescence microscopy. During the process of sonication, proper placement of a sonicating probe above the sample, as well as the duration and intensity of sonication, is critical. Additonal minor modifications to standard immunostaining protocols may be required for high quality stains. For antibodies with low signal to noise ratio, longer incubation times are typically necessary. As a proof of concept for this sonication-facilitated protocol, we show immunostains of three tissue types (testes, ovaries, and neural tissues) at a range of developmental stages.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and ...

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes differentiation from a stem cell population, mitotic amplification, and meiosis. In addition, post-meiotic germ cells undergo a dramatic morphological reshaping process as well as a global epigenetic reconfiguration of the germ line chromatin&#8212;the histone-to-protamine switch.
Studying the role of a protein in post-meiotic spermatogenesis using mutagenesis or other genetic tools is often impeded by essential embryonic, pre-meiotic, or meiotic functions of the protein under investigation. The post-meiotic phenotype of a mutant of such a protein could be obscured through an earlier developmental block, or the interpretation of the phenotype could be complicated. The model organism Drosophila melanogaster offers a bypass to this problem: intact testes and even cysts of germ cells dissected from early pupae are able to develop ex vivo in culture medium. Making use of such cultures allows microscopic imaging of living germ cells in testes and of germ-line cysts. Importantly, the cultivated testes and germ cells also become accessible to pharmacological inhibitors, thereby permitting manipulation of enzymatic functions during spermatogenesis, including post-meiotic stages.
The protocol presented describes how to dissect and cultivate pupal testes and germ-line cysts. Information on the development of pupal testes and culture conditions are provided alongside microscope imaging data of live testes and germ-line cysts in culture. We also describe a pharmacological assay to study post-meiotic spermatogenesis, exemplified by an assay targeting the histone-to-protamine switch using the histone acetyltransferase inhibitor anacardic acid. In principle, this cultivation method could be adapted to address many other research questions in pre- and post-meiotic spermatogenesis.

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

This protocol describes the dissection and cultivation of intact testes and germ-line cysts from Drosophila melanogaster pupae. This method allows microscopic observation of spermatogenesis ex vivo. Furthermore, we describe a pharmacological assay of the effect of inhibitors on specific stages of germ-cell development in pupal testes.

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

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Upright Imaging of Drosophila Egg Chambers

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Transillumination-Assisted Dissection of Specific Stages of the Mouse Seminiferous Epithelial Cycle for Downstream Immunostaining Analyses

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Dissection and Staining of Drosophila Larval Ovaries

Imaging Centrosomes in Fly Testes

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

Super-Resolution Live Cell Imaging of Subcellular Structures