This work was supported by a grant (R01GM098394) from NIH and National Institute of General Medical Sciences as well as grant 1121176 from National Science Foundation.

1. Track A. Preparation of Live Testes
<ol>
 <li>Prepare PBS by dissolving 8.0 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4 in 800 ml of distilled water and adjust the pH to 7.4. Bring the volume to 1 L and sterilize by autoclaving.</li>
 <li>Isolate testes as previously described 27.</li>
 <li>Prepare DAPI staining buffer by diluting 1 mg/ml stock to 1 &mu;g/ml in PBS. Use aluminum foil to protect the tube from light and store at -20 &deg;C.</li>
 <li>After isolating the testes, immerse the specimen in 6 &mu;l of DAPI staining buffer on top of a positively charged glass microscope slide for 10 min. Testes adhere well to commercially available positively charged slides, allowing for easy manipulation of the specimen. Positively charged slides are covalently modified to impart a static positive charge to the glass surface. Similar to polylysine coating, this promotes the interaction of the testes with the surface of the slide.</li>
 <li> Wash excess DAPI by replacing the staining buffer twice with 6 &mu;l of PBS. Use a piece of filter paper to wick away the buffer after each washing step. Use caution as to not accidentally remove the testes along with the buffer during each wash.</li>
 <li>Add 6 &mu;l of PBS to the specimen. The amount of buffer used should be adjusted for the size of the coverslip. The volumes provided in this protocol are for an 18 x 18 mm coverslip. Optional: It is advisable to pierce the testes near the relative location of the cell type of interest. This will ensure that there is minimal pressure on these cells during the squashing step, thereby maintaining their structural integrity. Piercing of the testes can be performed using a sharp, clean scalpel.</li>
 <li>Carefully place a coverslip on top of the specimen.</li>
 <li>Seal the edges of the coverslip using clear nail polish to ensure that the buffer does not evaporate from the live specimen while imaging.</li>
 <li>Use a marker to designate the position of the testes for ease in locating the specimen on the slide and proceed to imaging.</li>
	<li>Wash excess DAPI staining buffer by replacing the buffer twice with 6 &mu;l of PBS. Use a piece of filter paper to wick away the buffer after each washing step. Be careful to not accidentally remove the specimen along with the buffer during each wash.</li>
</ol>
2. Track B. Preparation of Fixed Testes
<ol>
 <li>Prepare Fix buffer by diluting 37% formaldehyde stock solution in PBS to a final concentration of 3.7% formaldehyde in 1x PBS</li>
 <li>Prepare DAPI staining buffer by diluting 1 mg/ml DAPI stock to 1 &mu;g/ml in PBS.</li>
 <li>Immerse the testes in a 6 &mu;l drop of PBS on top of a positively charged glass microscope slide.</li>
 <li>Manually orient the testes in a linear fashion or as otherwise preferred.</li>
 <li>Use a piece of filter paper to wick away the PBS and replace it with 6 &mu;l of Fix buffer for 5 min.</li>
 <li>Use a piece of filter paper to wick away the Fix buffer and immerse the specimen in 6 &mu;l of DAPI staining buffer for 10 min.</li>
 
 <li>Carefully place an 18 x 18 mm coverslip on top of the specimen.</li>
 <li>Seal the edges of the coverslip using clear nail polish to ensure that the buffer does not evaporate from the specimen while imaging.</li>
 <li>Use a marker to designate the position of the testes for ease in locating the specimen on the slide and proceed to imaging.</li>
</ol>
3. Track C. Preparation of Immunostained Testes
Coverslip siliconization: Immerse multiple 18 x 18 mm coverslips in a small tray containing siliconizing solution and incubate for 1 min at room temperature under a fume hood. Make sure that the coverslips are fully exposed and are not stacked on top of each other.
<ol>
 <li>Place the specimen in a 5 &mu;l drop of PBS on the siliconized coverslip. Orient the testes as desired and pierce the testes using a sharp scalpel. Gently place a positively charged glass microscope slide over the coverslip, allowing the PBS to become evenly dispersed between the coverslip and the slide. Use a small piece of filter paper to wick away the excess buffer between the coverslip and slide. As the buffer is removed by the filter paper, the increase in pressure will squash the testes. Many specimens should be prepared simultaneously, as some may be lost or damaged over the course of the protocol. A glass engraver can be used to label the slides if different types of specimens will be prepared simultaneously.
	 <ol>
		 <li>Wash the coverslips 3x for 1 min each in water, followed by 3x for 1 min each in 70% ethanol, and a final wash for 1 min in water. Allow the siliconized coverslips to air-dry under a fume hood.</li>
		</ol>
	</li>
	<li>Drop the slides into liquid nitrogen and allow the specimen to freeze for 5-10 min.</li>
 <li>Remove the slides from the liquid nitrogen using a large forceps. Use a scalpel to quickly remove the coverslip. The specimen should remain on the slide. Be careful not to smear the specimen during this process by sliding the coverslip along the slide.</li>
 <li>Incubate the slides in prechilled methanol in a glass Coplin staining jar at -20 &deg;C for 15 min. </li>
 <li>Transfer the slides to a glass Coplin staining jar containing prechilled acetone at -20 &deg;C for 30 sec.</li>
 <li>Wash the slides for 1 min in PBS at room temperature using a glass Coplin staining jar.</li>
 <li>Prepare PBST-B by supplementing PBS with 0.1% Triton-X100 and 1% Bovine Serum Albumen (BSA). 5% normal serum from the same host species as the secondary antibody (from step 3.14) may be used instead of BSA to reduce the background noise produced from nonspecific secondary antibody binding.</li>
 <li>Incubate the slides for 10 min in PBST-B in a glass Coplin staining jar for to block nonspecific sites.</li>
 <li>Fill the wells of the moisture chamber with water. Humidity inside the chamber will minimize the evaporation of antibody solutions from the specimen during the incubation steps.</li>
 <li>Prepare PBST-BR by supplementing PBST-B with 100 &mu;g/ml RNase A. RNase A degrades RNA in the specimen and also serves as an additional blocking agent by absorbing strongly to the glass surface.
	 <ol>
		 <li>Place an approximately 1x1 cm piece of Parafilm on top of the specimen to spread the antibody solution evenly and protect the antibody solution from evaporating. Close the moisture chamber and incubate the specimen in the primary antibody for 1 hr at room temperature.</li>
		</ol>
	</li>
 <li>Remove the slides from PBST-B. Use a piece of filter paper to dry the area of the slide around the specimen, using caution not to dry the specimen itself. Finally, place the slide in the moisture chamber with the specimen facing up</li>
 <li>Gently add 100 &mu;l of primary antibody diluted in PBST-BR on top of the specimen (1:200 is generally a good starting concentration for uncharacterized antibodies).
	 <ol>
		 <li>Cover the specimen with an approximately 1 x 1 cm piece of Parafilm and close the moisture chamber. Incubate the specimen in secondary antibody for 1 hr at room temperature.</li>
		 </ol>
	</li>
 <li>Open the moisture chamber and use forceps to gently remove the Parafilm from the slide. Incubate the slides for 5 min in PBST in a glass Coplin staining jar at room temperature to wash. Repeat this process for a total of three washes.</li>
 <li>Remove the slides from PBST-B and place them in the moisture chamber with the specimen facing up. Gently add 100 &mu;l of secondary antibody diluted in PBST-BR on top of the specimen.</li>
 <li>Open the moisture chamber and use forceps to gently remove the Parafilm from the slide. Again, wash the specimen 3x for 5 min each in PBST in a glass Coplin staining jar followed by 3x for 5 min each in PBS.</li>
 <li>Use a Kimwipe and filter paper to carefully dry the surface of the slide, being careful to avoid touching or drying the specimen.</li>
 <li>Add 6 &mu;l of Mounting Media to specimen, cover with a clean coverslip (not coated with silicone), and the seal edges with nail polish.</li>
 <li>Use a marker to designate the position of the testes for ease in locating the specimen on the slide and proceed to imaging.</li>
</ol>
4. Imaging Notes
Imaging can be achieved using an upright or inverted regular light microscope or confocal microscope. It is important that the microscope be equipped with phase-contrast, especially for the imaging of live testes specimens (Track A). This feature has recently become available on confocal microscopes.
When the testes break spontaneously (Track A), the tip (stem cell region) is usually kept intact and is easily identifiable, providing a good marker to initially locate and use for orientation.
Centrioles are fairly small structures and images should therefore be taken using 63X or 100X objectives, a zoom factor of 4-6X, and a resolution of at least 512 x 512 pixels when possible.

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Free access to this publication was supported by Leica Microsystems.

Studying centrosome biology in fly testes using genetically-tagged centrosomal markers is a useful method for assessing centrosome function and activity in both a wild-type and mutant context. In particular, Track A is suitable for rapid screening of centrosomal abnormalities such as malformation, misegregation, instability, or abnormal length in an effort to identify new mutants. Furthermore, spermatid cilia in live preparations remain motile for approximately 15 min after dissection and the use of live testes in Track A also allows for sperm motility to be easily addressed. Since the activity of sperm motile cilia is directly related to centrosome function, assays can be performed to determine the effects of various centrosomal mutations on ciliary function. Track B can be used for more specific observations especially when statistical data is required such as for counting the number of centrioles per cell and or number of cells per cyst. Track C is most useful for detailed observations that require staining with antibodies. Examples include labeling of a particular cell type such as stem cells, staining of a protein that doesn't have an available tag such as acetylated tubulin, or to verify the absence or mislocalization of a protein in a mutant.
When imaging centrosomes and centrosome-related structures, using genetically-tagged markers rather than antibodies is not only experimentally easier, but also provides more robust and reproducible results. Therefore, the use of genetically-tagged centrosomal proteins is a reliable approach for mechanistic and quantitative analyses that require a large data set. For example, the use of genetically-tagged centrosomal markers has been particularly valuable for the quantification of centriole length. Such analysis has revealed that various mutations can be classified into two categories based on the variability in centriole length. One category includes mutations that change centriole length but do not affect the standard deviation 1 and the other category includes mutations that affect both centriole length and the standard deviation in length 11. Such quantitative data can provide useful insight into the function of particular centrosomal genes. Centrosomal mutants that exhibit defective centriole length with an increase in standard deviation may be due to a destabilization of centriolar structure. However, defective centriole length with a normal standard error may indicate that the mutation does not structurally destabilize the centriole and is more likely to be due to a change in regulatory mechanism controlling centriole length. Due to inconsistencies in immunostaining, quantitative analyses are difficult with the use of antibody markers alone.
The centrosome is a large, complex proteinaceous structure and many of its proteins are only found within its interior. Using genetically-tagged centrosomal markers rather the antibodies allows one to consistently label internal components of the centrosome whose epitopes may be otherwise inaccessible to antibody markers. For example, localization studies of Bld10 using antibodies finds the protein to be enriched at the distal and proximal ends of the centriole 13, while Bld10-GFP shows a more uniform distribution 1. However, it is also important to consider the level of expression of a particular genetically-tagged protein, as this can affect protein distribution. Localization of Sas-4-GFP and SAS-6-GFP expressed under their endogenous is restricted to the proximal ends of the centriole 1,16,35 On the other hand, Sas-4-GFP and SAS-6-GFP expressed under the strong ubiquitin promoter are localized along the entire length of the centriole 12,14 . Another important consideration is the effect of the genetic tag on protein function. Analyzing if genetically-tagged proteins are functional can be tested by introducing the transgenic protein into a mutant background and examining if the transgenic protein rescues the mutant phenotype.
Fixation of Drosophila testes can be performed using a variety of chemical fixatives. Here, we describe fixation with both formaldehyde (Track B) and methanol-acetone (Track C). However, either fixative can be used interchangeably and since various fixatives may chemically disrupt native antibody epitopes, the selection of the appropriate fixative for immunostaining must be determined experimentally. The following fixatives and incubation conditions are commonly employed: 3.7% formaldehyde, 5 min at room temperature; methanol, 15 min at -20 &deg;C; acetone, 10 min at -20 &deg;C; methanol, 15 min at -20 &deg;C. followed by acetone, 30 sec at -20 &deg;C; ethanol, 20 min at -20 &deg;C. Although fixation with acetone, methanol, and ethanol do not require an extended permeabilization step for immunostaining, fixation with formaldehyde should be followed by a 1 hr incubation in PBST-B at room temperature to permeabilize cell membranes and allow antibody access to intracellular epitopes. Furthermore, certain fixatives are more suitable for particular antigens. For example, formaldehyde functions well for fixation of small proteins, whereas methanol and acetone are well-suited for the fixation of large molecular complexes 36.
Immunostaining of Drosophila testes has been previously described for observing chromatin structures and the microtubule cytoskeleton 29,37. Here (Track C), the procedure has been optimized for the analysis of centrosomal structures containing genetically-tagged proteins. We provide a detailed description of this procedure to guide individuals who are inexperienced in working in Drosophila testes. This procedure also includes modifications to improve the preservation and morphology of the testes, such as by using siliconized coverslips and positively charged glass microscope slides.

Drosophila testes are a suitable organ system to study a variety of cellular and developmental processes and have been reviewed extensively over the years 1-9. This manuscript focuses on the use of Drosophila testes to study the centrosome, a conserved cellular organelle. As in other systems, the centrosome of the Drosophila testes function in mitosis, meiosis, and ciliogenesis 10. Centrosomes are composed of a pair of microtubule-based structures known as centrioles surrounded by a complex protein network referred to as pericentriolar material (PCM). The centriole pair is comprised of an older mother centriole and a younger daughter centriole. As the cell progresses towards mitosis, both centrioles separate, duplicate, and acquire a large amount of PCM to ultimately form two distinct centrosomes. The centrosome containing the original mother centriole is referred to as the mother centrosome and the centrosome containing the original daughter centriole is referred to as the daughter centrosome.
Drosophila testes are ideal for studying the molecular basis of centrosome biology by fluorescent microscopy for a variety of reasons.
<ol>
 <li>Most of the Drosophila proteins that are required for centrosome biology in the testes are conserved among eukaryotes, suggesting that insight relevant to centrosome biology in humans and other species can be gained by studying the centrosome in Drosophila testes 1,11-14 .</li>
 <li>Performing mutant analysis in Drosophila provides a significant advantage since, unlike many other models, centrosomal mutations in Drosophila are not embryonic lethal, thus allowing for classical genetic analysis of centrosome function. This unique feature of Drosophila is due to the presence of maternal contribution that persists during the critical stages of embryonic development. 1,11-14 . Thus, one can a) study mutations that completely eliminate centrosome formation and b) study the fate of a normal centrosome that was formed in the early embryo by maternal contribution in a mutant context after the maternal contribution has been depleted (the principle of the method is described in 11).</li>
 <li>Functional transgenes with genetically encoded fluorescent tags that label the centrosome are available. Many of these lines use the protein's own promoter to drive transcription of the transgene in order to prevent strong overexpression. This is particularly important because overexpression of proteins often results in artifacts that interfere with analysis of centrosome function 1,11 .</li>
 <li>The centriole and centrosome are uniquely long throughout Drosophila spermatogenesis, thus allowing for rapid and easy analysis of the centrosome by imaging.</li>
 <li>Consecutive steps in spermatogenesis and centrosome biology are organized chronologically along the testes, starting at the testis tip with the centrosomes of sperm stem cells and ending at the bottom of the testes with reduced centrosome size and activity in mature sperm cells (Figures 1 and 2). This allows for easy identification and analysis of centrosome function during different stages of sperm development.</li>
 <li>Testes are easily dissected from male larvae, pupae, and adults 27.</li>
 <li>During spermatogenesis, the centrosome and its centrioles proceed through multiple compositional, structural and functional states to function in mitosis, meiosis, and cilium formation. During these processes, the centrosome assembles, duplicates, migrates, anchors to specific parts of the cell, matures, divides, and creates a cilium. Furthermore, the centriole of the mature spermatid gives rise to a centriolar precursor named the PCL 1. Also in the mature spermatid, the centrosome undergoes a process called centrosome reduction whereby it loses many components of the PCM and the centriole (Figures 1 and 2). Thus, studies in the testes allow one to address multiple aspects of centrosome life such as centrosome retention in the stem cells, centriole duplication, centriole stability, centriole elongation, centriole separation and segregation, PCM recruitment, ciliogenesis, PCL formation, centrosome reduction, and astral microtubule nucleation among others.</li>
 <li>Finally, studies of centrosome biology are aided by the other well-known characteristics of Drosophila that have made it a preferred model organism for biological studies. These include a short generation time, ease of genetics, as well as random and site-directed mutagenesis.</li>
</ol>
Together, the above stated characteristics of Drosophila testes provides a model where the centrosome can be studied by easy, rapid, and detailed imaging. The techniques described in this paper have been applied to investigate many aspects of the centrosome biology including centriole formation 11, centriole duplication 15, PCM recruitment 16, centrosome regulation 17, and ciliogenesis 18. These techniques have also been applied to study the centrosome in other areas of biology such as meiotic regulation 19, spindle assembly 20, and centrosome activity in asymmetrical stem cell division 21 among many others.
Imaging of testes in the centrosome starts with obtaining flies that express genetically-tagged centrosomal proteins and isolating the testes from male larvae, pupae, or adult flies. These flies are available from several research groups 1,11,15,22-25 . The larval testes contain all stages of spermatogenesis prior to meiosis and are useful when analyzing mutations that are lethal in the pupa or adult. However, Late pupal or young adult testes are the most robust and contain all pre- and post-meiotic stages of spermatogenesis, thereby making them preferable for analysis. Since the number of sperm cells decreases as the fly ages, the use of adult testes is also appropriate for studying the centrosome in the context of aging. 26. A method of testis isolation from adult flies has been previously described 27.
Imaging of centrosomes and their function in Drosophila testes can be achieved via three related tracks that are presented here (Figure 3). Selection of which track is most appropriate is dependent on the nature of the question the investigator is addressing.
Track A involves imaging of live testes. It is the most rapid of the three tracks but can only be applied when specimens do not need to be preserved and when immunostaining is not required. In Track A, testes are mounted (intact, pierced or cut) on slides and carefully squashed under a coverslip to form a single layer of easily identifiable cells. The cells are then visualized using both phase-contrast microscopy and fluorescent microscopy. The use of phase-contrast is particularly important for the analysis of Drosophila testes because it reveals cellular information that is not visible with other forms of transmitted illumination and thus allows for quick identification of various stages of sperm development 28,29 . However, Track A has two main disadvantages. First, the morphological integrity of cells is often compromised once the testes are ruptured. Second, unfixed cells sometimes move within the specimen, making the imaging of multiple confocal layers within a specified region difficult. To promote the analysis of specific cell types, testes may be pierced or cut in order to direct the way the testis ruptures such that squashing under the coverslip minimally affects the morphology of the cell type of interest.
Track B involves chemical fixation of the testes. This track requires an intermediate amount of time for specimen preparation and has the advantage that fixed specimens can be saved for later analysis. Furthermore, fixation serves to make cellular structures more rigid, minimizing movement of the specimen during imaging. However, phase-contrast microscopy becomes much less informative after chemical fixation, making some stages of sperm development difficult to identify.
Track C is the most time-intensive, but has the added benefit that cellular structures are fixed and immunostained, allowing for the visualization of proteins that are not available with the appropriate genetics tags. There are many antibodies available both commercially and from various research groups for immunostaining centrosomes and centrosome-related structures in Drosophila testes.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/ Equipment</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments/Description</td>
 </tr>
 <tr>
 <td>DAPI</td>
 <td>Invitrogen</td>
 <td>D1306</td>
 <td></td>
 </tr>
 <tr>
 <td>Positively Charged Slides</td>
 <td>AZER Scientific</td>
 <td>EMS200A+</td>
 <td></td>
 </tr>
 <tr>
 <td>Feather Microscalpel</td>
 <td>Electron Microscopy Sciences</td>
 <td>72045-30</td>
 <td></td>
 </tr>
 <tr>
 <td>37% Formaldehyde or Paraformaldehyde</td>
 <td>Fisher Scientific</td>
 <td>BP531-500</td>
 <td></td>
 </tr>
 <tr>
 <td>Phosphate Buffered Saline (PBS)</td>
 <td>Boston Bioproducts</td>
 <td>BM-220</td>
 <td></td>
 </tr>
 <tr>
 <td>18x18 mm coverslips number 1.5</td>
 <td>VWR</td>
 <td>48366 205</td>
 <td></td>
 </tr>
 <tr>
 <td>Nail Polish</td>
 <td>Electron Microscopy Sciences</td>
 <td>72180</td>
 <td></td>
 </tr>
 <tr>
 <td>Sigmacote</td>
 <td>Sigma-Aldrich</td>
 <td>SL2-100 ml</td>
 <td></td>
 </tr>
 <tr>
 <td>Methanol</td>
 <td>Fisher Scientific</td>
 <td>A412-4</td>
 <td></td>
 </tr>
 <tr>
 <td>Acetone</td>
 <td>Fisher Scientific</td>
 <td>A949-1</td>
 <td></td>
 </tr>
 <tr>
 <td>Triton-X100</td>
 <td>Sigma-Aldrich</td>
 <td>T9284-100ML</td>
 <td></td>
 </tr>
 <tr>
 <td>BSA</td>
 <td>Jackson ImmunoResearch</td>
 <td>001-000-162</td>
 <td></td>
 </tr>
 <tr>
 <td>RNAse A</td>
 <td>5 prime</td>
 <td>2900142</td>
 <td></td>
 </tr>
 <tr>
 <td>Filter paper</td>
 <td>Whatman</td>
 <td>1001-055</td>
 <td></td>
 </tr>
 <tr>
 <td>Glass engraver</td>
 <td>Dremel</td>
 <td>290-01</td>
 <td></td>
 </tr>
 <tr>
 <td>TCS SP5 confocal microscope</td>
 <td>Leica </td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Mounting Media</td>
 <td>Electron Microscopy Sciences</td>
 <td>17985-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Immuno Stain Moisture Chamber, Black</td>
 <td>Electron Microscopy Sciences</td>
 <td>62010-37</td>
 <td></td>
 </tr>
 <tr>
 <td>Glass Coplin Staining Jar, Screw Cap</td>
 <td>Electron Microscopy Sciences</td>
 <td>70315</td>
 <td></td>
 </tr>
 </tbody>
</table>

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.

Centrosomes undergo multiple morphological and functional transformations over the course of spermatogenesis. This characteristic of spermatogenesis makes the testes a useful system to study various aspects of centrosome biology. One such readily observed process is centrosome elongation. In spermatogonia, the centriolar marker Ana1-GFP marks the 0.6 &mu;m long centriole (Figure 4a). This centriole elongates during spermatogenesis and reaches a length of 2.5 &mu;m in nearly mature spermatids. Since centrioles in Drosophila sperm are uniquely long, imaging can be used to make quantitative statements regarding centrosome elongation (Figure 4b). Analysis of centriole length can also be performed in a mutant background and various mutants have been identified that alter centriole growth (Figure 5). Examples are always early (aly) and cannonball (can), mutations that arrest spermatogenesis before the onset of meiosis 30 but do not block centriole elongation. In these mutants, centrioles of the mature spermatocyte grow to about ~2.4 &mu;m in comparison to centrioles of control cells which reach a maximum of 1.8 &mu;m.
<img src="/files/ftp_upload/50938/50938fig1.jpg" alt="Figure 1" fo:content-width="6in" fo:src="/files/ftp_upload/50938/50938fig1highres.jpg"/> 
Figure 1. Spermatogenesis and Centrosome Biology. A) Development of Stem Cells and Spermatogonia. All sperm cells originate from Stem Cells found at the apical tip of the testes. Each Stem Cell divides asymmetrically to form a Stem Cell that inherits the mother centrosome and a progenitor spermatogonium that inherits the daughter centrosome 31. As Spermatogonia form, they become surrounded by 2 cyst cells, which continue to surround the Spermatocytes and Spermatids throughout spermatogenesis. Spermatogonia divide 4 times to produce 16 Spermatogonia, each containing two centrioles. B) Development of Spermatocytes and Meiosis. During Spermatocyte development, centrioles duplicate once more to generate four centrioles organized into two pairs per cell and 64 centrioles per cyst. Early during Spermatocyte development, each centriole moves to the plasma membrane, docks to it, and elongates to form a structure that resembles a primary cilium18,32-34 . In mature Spermatocytes, centriole length reaches ~1.8 &mu;m. The centrosomes play an essential role in meiosis. During meiosis I, the centriole pairs that are still attached to the plasma membrane move towards the center of the cell and create cell membrane invaginations at their distal tip. The PCM around the centrioles grows, nucleates astral microtubules, and colocalizes with the spindle pole. During the transition to meiosis II, the centriole pair separates so that only a single centriole is present at each spindle pole. C) Spermiogenesis. At the end of meiosis II, the Early Round Spermatid has a single centriole attached to the invaginated plasma membrane. A large round mitochondrial derivative is found near the nucleus and later starts to elongate along the growing axoneme in Later Round Spermatids. During spermatogenesis, the axoneme forms and elongates inside the cytoplasm. In addition a new centriolar structure known as the PCL (Proximal Centriole Like) appears near the preexisting centriole1. At the end of spermatogenesis, cells sharing a common cyst separate from each other to become fully mature motile sperm. The diagrams of Spermatocyte development (B) and Spermiogenesis (C) depict only one cell out of the 16 Spermatocytes and 64 Spermatids, respectively per cyst, and do not depict the cyst cells. <a href="https://www.jove.com/files/ftp_upload/50938/50938fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img src="/files/ftp_upload/50938/50938fig2.jpg" alt="Figure 2" fo:content-width="4in" fo:src="/files/ftp_upload/50938/50938fig2highres.jpg"/> 
Figure 2. The Drosophila Testes. An overlay of a phase-contrast and GFP fluorescence micrograph of a whole-mount testis expressing the pan-centriolar marker Ana-1-GFP. Distinct area corresponding to panels 1-3 of Figure 1 are separated by a dashed red line A) Stem Cells and Spermatogonia, B) Spermatocytes and Meiosis, C) Spermiogenesis.
<img src="/files/ftp_upload/50938/50938fig3.jpg" alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/50938/50938fig3highres.jpg"/> 
Figure 3. Tracks Towards Imaging Centrosomes in Drosophila Testes.
<img src="/files/ftp_upload/50938/50938fig4.jpg" alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/50938/50938fig4highres.jpg"/> 
Figure 4. Centrosome Elongation During Spermatogenesis. A) Each stage shows a phase-contrast image (left), fluorescence image (middle), and magnified image of the centrosome (right). The phase-contrast image demonstrates the particular cell stage based on the position and morphology of the cell, nucleus, nucleolus, and the mitochondria. The fluorescence image shows DNA stained with DAPI (blue). The centrosome image shows centrosomes labeled by Asterless-GFP (Asl, top images) and Ana1-GFP (Ana1, bottom image). A white circle highlights the cell membrane in both phase-contrast and fluorescence images. The dashed white circles and the gray circle in the fluorescence image highlight the position of the nucleus and nucleolus, respectively. Yellow arrow points to the Y-chromosome loops. Red dashed circles mark the mitochondria. Stages 1-6 refers to 29 and stages 13-17 refers to description of cell stages in 32. Stage 1: Primary Spermatocyte. Identified as a small cell a tip of the testis. Stage 2a: Polar Spermatocyte. Identified by the presence of a mitochondria cap on one side of the nucleus. Stage 3: Apolar Spermatocyte. Identified by its relative larger size than the primary spermatocyte and lack of a mitochondria cap. Stage 4: Primary Spermatocyte. Identified by the appearance of all: All 3 Y-chromosome loops. Stage 5: Mature Primary Spermatocyte. Largest Spermatocyte produced in Spermatogenesis, identified by a further increase in nuclear size. Stage 6: Premeiotic Primary Spermatocyte. Y-chromosome loops disintegrate and the nucleus Disappears. Stage 13: Onion Stage Spermatid. Round cell containing equally sized nucleus and mitochondria. Stage 17: Late spermatid. Identified as an elongated spermatid that remains part of a bundle of 64 spermatids with nuclei found near the base of the testes., Nuclei are slightly wider than mature spermatids found in the seminal vesicle. Scale bars, 10 &mu;m and 1 &mu;m. B) Graph depicts average and standard deviation for each stage as measured by Asterless-GFP (blue) and Ana1-GFP (green). In Stage 6, Asterless-GFP (blue) and Ana1-GFP (green) averages and standard deviations overlap and only Asterless-GFP (blue) is apparent. Starting at Stage 13, Asterless-GFP does not decorate the whole centriole and measurements were not determined (ND).
<img src="/files/ftp_upload/50938/50938fig5.jpg" alt="Figure 5" fo:content-width="5in" fo:src="/files/ftp_upload/50938/50938fig5highres.jpg"/> 
Figure 5. Abnormally Long Centrioles in Stage 6 Spermatocytes of aly and can Mutants. A-C) Light micrograph of whole testes and fluorescent panel of representative centrosome labeled by Ana1-GFP. Note the abnormal shape of the testes in aly and can mutants, which results from a lack of spermatids as a consequence of meiotic arrest. D) Graph depicts average, standard deviation, and data distribution of centriole length in wild-type, aly and can. Sample number for each data point is in parentheses. Scale bar, 1 &mu;m.

Watch this Scientific Journal Video about Imaging Centrosomes in Fly Testes at JoVE.com

Imaging Centrosomes in Fly Testes

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 ...

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15.9K Views.  University of Toledo. 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.

Video: Imaging Centrosomes in Fly Testes

Implantation of Ferumoxides Labeled Human Mesenchymal Stem Cells in Cartilage Defects

The field of tissue engineering integrates the principles of engineering, cell biology and medicine towards the regeneration of specific cells and functional tissue. Matrix associated stem cell implants (MASI) aim to regenerate cartilage defects due to arthritic or traumatic joint injuries. Adult mesenchymal stem cells (MSCs) have the ability to differentiate into cells of the chondrogenic lineage and have shown promising results for cell-based articular cartilage repair technologies. Autologous MSCs can be isolated from a variety of tissues, can be expanded in cell cultures without losing their differentiation potential, and have demonstrated chondrogenic differentiation in vitro and in vivo1, 2.
In order to provide local retention and viability of transplanted MSCs in cartilage defects, a scaffold is needed, which also supports subsequent differentiation and proliferation. The architecture of the scaffold guides tissue formation and permits the extracellular matrix, produced by the stem cells, to expand. Previous investigations have shown that a 2% agarose scaffold may support the development of stable hyaline cartilage and does not induce immune responses3.
Long term retention of transplanted stem cells in MASI is critical for cartilage regeneration. Labeling of MSCs with iron oxide nanoparticles allows for long-term in vivo tracking with non-invasive MR imaging techniques4.
This presentation will demonstrate techniques for labeling MSCs with iron oxide nanoparticles, the generation of cell-agarose constructs and implantation of these constructs into cartilage defects. The labeled constructs can be tracked non-invasively with MR-Imaging.

Goal of the presentation is to demonstrate a highly reproducible method to generate matrix associated stem cell implants in cartilage defects, which can be visualized with MR imaging. Stem cells are labeled with FDA-approved Ferumoxides, mixed with agarose, implanted into cartilage defects and imaged with a 7T MR scanner.

The field of tissue engineering integrates the principles of engineering, cell biology and medicine towards the regeneration of specific cells and ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

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 ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to facilitate sensory functions in cells. The level of a protein may be regulated differently at centrosomes than at other .cellular locations, and the variation in the centrosomal level of several proteins at different points of the cell cycle appears to be crucial for the proper regulation of centriole assembly. We developed a quantitative fluorescence microscopy assay that measures relative changes in the level of a protein at centrosomes in fixed cells from different samples, such as at different phases of the cell cycle or after treatment with various reagents. The principle of this assay lies in measuring the background corrected fluorescent intensity corresponding to a protein at a small region, and normalize that measurement against the same for another protein that does not vary under the chosen experimental condition. Utilizing this assay in combination with BrdU pulse and chase strategy to study unperturbed cell cycles, we have quantitatively validated our recent observation that the centrosomal pool of VDAC3 is regulated at centrosomes during the cell cycle, likely by proteasome-mediated degradation specifically at centrosomes.

Here, a novel quantitative fluorescence assay is developed to measure changes in the level of a protein specifically at centrosomes by normalizing that protein&#8217;s fluorescence intensity to that of an appropriate internal standard.

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Cryopreservation of Zebrafish Spermatogonia by Whole Testes Needle Immersed Ultra-Rapid Cooling

Current trends in science and biotechnology lead to creation of thousands of new lines in model organisms thereby leading to the necessity for new methods for safe storage of genetic resources beyond the common practices of keeping breeding colonies. The main purpose of this study was to adapt the needle immersed vitrification (NIV) procedure to cryopreserve whole zebrafish testes. Cryopreservation of early-stage germ cells by whole testes NIV offers possibilities for the storage of zebrafish genetic resources, especially since after transplantation they can mature into both male and female gametes. Testes were excised, pinned on an acupuncture needle, equilibrated in two cryoprotective media (equilibration solution containing 1.5 M methanol and 1.5 M propylene glycol; and vitrification solution containing 3 M dimethyl sulfoxide and 3 M propylene glycol) and plunged into liquid nitrogen. Samples were warmed in a series of three consequent warming solutions. The main advantages of this technique are (1) the lack of spermatozoa after digestion of warmed testes thus facilitating downstream manipulations; (2) ultra-rapid cooling enabling the optimal exposure of tissues to liquid nitrogen therefore maximizing the cooling and reducing the required concentration of cryoprotectants, thereby reducing their toxicity; (3) synchronous exposure of several testes to cryoprotectants and liquid nitrogen; and (4) repeatability demonstrated by obtaining viability of above 50% in five different zebrafish strains.

The main purpose of this study was to adapt the needle immersed vitrification (NIV) procedure to cryopreserve whole zebrafish testes. Additionally, the repeatability of the method in five different zebrafish strains was tested.

Current trends in science and biotechnology lead to creation of thousands of new lines in model organisms thereby leading to the necessity for new methods ...

Oligomerization Dynamics of Cell Surface Receptors in Living Cells by Total Internal Reflection Fluorescence Microscopy Combined with Number and Brightness Analysis

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of clustering. Here, we describe an imaging approach to determine the average oligomeric state of mEGFP-tagged-receptor homocomplexes in the membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis. N&#38;B is a method similar to fluorescence-correlation spectroscopy (FCS) and photon counting histogram (PCH), which are based on the statistical analysis of the fluctuations of the fluorescence intensity of fluorophores diffusing in and out of an illumination volume during an observation time. In particular, N&#38;B is a simplification of PCH to obtain information on the average number of proteins in oligomeric mixtures. The intensity fluctuation amplitudes are described by the molecular brightness of the fluorophore and the average number of fluorophores within the illumination volume. Thus, N&#38;B considers only the first and second moments of the amplitude distribution, namely, the mean intensity and the variance. This is, at the same time, the strength and the weakness of the method. Because only two moments are considered, N&#38;B cannot determine the molar fraction of unknown oligomers in a mixture, but it only estimates the average oligomerization state of the mixture. Nevertheless, it can be applied to relatively small time series (compared to other moment methods) of images of live cells on a pixel-by-pixel basis, simply by monitoring the time fluctuations of the fluorescence intensity. It reduces the effective time-per-pixel to a few microseconds, allowing acquisition in the time range of seconds to milliseconds, which is necessary for fast oligomerization kinetics. Finally, large cell areas as well as sub-cellular compartments can be explored.

We describe an imaging approach for the determination of the average oligomeric state of mEGFP-tagged-receptor oligomers induced by ligand binding in the plasma membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis.

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of ...

Imaging Centrosomes in Fly Testes

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Chapters in this video

Implantation of Ferumoxides Labeled Human Mesenchymal Stem Cells in Cartilage Defects

Imaging Cell Shape Change in Living Drosophila Embryos

Isolation of Drosophila melanogaster Testes

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

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

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

Cryopreservation of Zebrafish Spermatogonia by Whole Testes Needle Immersed Ultra-Rapid Cooling

Oligomerization Dynamics of Cell Surface Receptors in Living Cells by Total Internal Reflection Fluorescence Microscopy Combined with Number and Brightness Analysis