Name: ex vivo culture of drosophila pupal testis and single male germ-line cysts: dissection, imaging, and pharmacological treatment
Uploaded: 2014-09-11T14:00:00
Duration: 8 min 35 s
Description: Watch this Scientific Journal Video about Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment at JoVE.com

The authors wish to thank T. Noguchi for helpful advice on the initial establishment of the culture technique in our laboratory. Research in the lab of R. R.-P. was funded by the DFG within the international collaborative research center TRR81.

Ethics statement: 
The experimental procedures described in this protocol involve exclusively work with Drosophila melanogaster and are not subject to animal welfare laws in Germany.
1. Preparation of Media, Tools, and Flies
<ol>
	<li>Prepare modified commercially available powdered M3 growth medium without potassium bicarbonate17. CAUTION: Irritating to eye and skin. Supplement the medium with 10% fetal bovine serum, 100&#160;U/ml penicillin, and 100&#160;&#181;g/ml streptomycin. Use heat-inactivated and insect-culture-tested fetal bovine serum for best results. Filter the complete medium (referred to hereafter as medium) through a 0.2&#160;&#181;m bottle-top filter unit and store in aliquots at -20&#160;&#176;C. Before use, filter (0.2&#160;&#181;m syringe filter tip) the medium again to remove precipitates.</li>
	<li>Prepare inhibitor stock solutions for the pharmacological assays. Dissolve the chemicals in the appropriate solvent and store in aliquots. For example, prepare a stock solution of 28.69&#160;mM anacardic acid (CAUTION: Irritating to eye and skin) in dimethylsulfoxide.</li>
	<li>Prepare tools for the disruption of dissected pupal testes and dissection of single cysts. Use two sharp and stiff needles, rather than a pair of forceps. The capillary force that develops between the two arms of a pair of forceps makes the dissection of single cysts in small amounts of medium very difficult. For example, use the tip of a stainless steel insect pin inserted into an inoculation loop holder.</li>
	<li>In order to obtain appropriately aged pupae, seed ca. 10 large culture vials (tubes with 4&#160;cm diameter) with a sufficient number of adult flies (ca. 40&#8211;60 flies). To analyze the H-P switch, use flies expressing H2AvD-RFP and ProtamineB-eGFP in their endogenous pattern12,16,22. Incubate the vials under standard conditions at 25&#160;&#176;C for ca. 4&#8211;5 days until third-instar larvae crawl up and start pupation on the walls of the vials25.</li>
</ol>
2. Marking or Collecting White Pre-pupae to Obtain 24-hr-old Pupae for Dissection
<ol>
	<li>To test the influence of certain chemicals on the H-P switch, dissect pupal testes at ca. 24 hr APF12. To obtain staged pupae, take the prepared fly culture vials with pupating larvae (see section 1.4) and identify as many white pre-pupae as possible. Pre-pupae during the very first step in pupation appear as motionless, non-feeding, rather shortened larvae with everted spiracles and white-colored puparium. The puparium becomes tan-brown within 30 to 60 min, which indicates the next stage of pupal development26.</li>
	<li>Mark the positions of the pre-pupae with a permanent marker on the outside of the vials. Incubate the vials at 25&#160;&#176;C for ca. 24 hr.
	<ol>
		<li>Alternatively, pick the pre-pupae from the side of the vials with a small brush moistened with PBS buffer. Then transfer the pre-pupae to the side of a fresh 50&#160;ml reaction tube and incubate at 25&#160;&#176;C for ca. 24 hr.</li>
	</ol>
	</li>
</ol>
3. Dissection of Pupal Testes at ca. 24 hr after Puparium Formation
<ol>
	<li>Distribute several drops (ca. 80&#160;&#181;l each) of medium on a 10&#160;cm plastic cell culture or Petri dish. Using a broad pair of forceps or a brush moistened with medium, pick the pupae at the 24&#160;hr APF stage from the incubated vials (see section 2). Transfer one pupa to each drop of medium on the dish.</li>
	<li>For the dissection, use a stereo-microscope equipped with a fiber-optic illuminator. Do not heat the testes above RT during the dissection as this might prevent proper development in culture.
	<ol>
		<li>Dissect pupae with two pairs of No. 5 forceps. With one pair, grab the pupa at its most posterior part (the posterior spiracles) and hold it in position.</li>
		<li>Grab the anterior spiracles with the other pair of forceps and open the pupal operculum at its anterior end. Peel off the pupal case until at least part of the thorax is exposed.</li>
		<li>Continue to hold the pupa in position by its most posterior end. To release the internal pressure of the pupa, insert both arms of a pair of forceps into the head region of the pupa.</li>
		<li>Rip the pupa open by opening the forceps and moving the forceps in the anterior direction. Ensure that the opening generated with this movement is as large as possible in the first attempt. Avoid damaging the pupa at its posterior end before the pressure has been released as this might result in the testes being pushed directly through the small opening and most often being disrupted.</li>
		<li>Observe that once the pupa is opened, the released internal pressure of the pupa presses out agglomerates of fat body cells and tissue through the large opening in the head region. Check whether the pupal testes have already been pushed out of the pupa with this material; this occurs in some cases. The testes are not connected to any other tissue at 24 hr APF and therefore might be expelled easily from the pupa23.</li>
		<li>Avoid squeezing the pupa with the forceps as this might damage the testes if they remain in the pupa. Increase the size of the opening using one pair of forceps.</li>
		<li>Then hold the pupa at its most posterior end. Close the other pair of forceps and gently streak out the contents of the pupa from posterior to anterior to release the testes from the pupa.</li>
	</ol>
	</li>
	<li>Collect the testes by pulling them into a pre-cut 200&#160;&#181;l pipette tip. Try to transfer only a very small amount of medium to reduce the number of fat body cells transferred with the testes. Place the testes into medium in a fresh culture dish. Wash the testes several times with medium until only few fat body cells remain.</li>
	<li>For live imaging, transfer testes to a glass-bottom culture dish with medium and use an inverted imaging microscope. For pharmacological assays, continue with step 5.</li>
</ol>
4. Dissection of Male Germ-line Cysts and Live Imaging
<ol>
	<li>Transfer one or more pupal testis to a glass-bottom culture dish.</li>
	<li>Use a stereo-microscope with fiber-optic illumination and two stainless steel needles (see section 1.3) for the dissection of male germ-line cysts.
	<ol>
		<li>With one needle, carefully try to pin down one testis at its anterior or posterior end. Hold it in position. Insert the other needle into the testis and disrupt the testis sheath by moving the needle sideways. Many of the cysts will be released from the testis by this movement.</li>
		<li>Continue to hold the testis in position with one needle. Gently streak out the remaining cysts from the testis with the other needle.</li>
		<li>Separate aggregated cysts either by gently moving a needle sideways through the cluster of cysts or by transferring the cysts to a fresh culture dish with medium using a 200&#160;&#181;l pipette tip.</li>
	</ol>
	</li>
	<li>Move the culture dish to an inverted imaging microscope for live imaging of single cysts. Disperse cysts again with a needle if necessary.
	<ol>
		<li>Identify the stage of the cysts by using phase-contrast and fluorescence microscopy.</li>
		<li>Focus on a cyst of the desired stage. Confirm focus and position of the cyst frequently during longer imaging experiments, as the cysts do not adhere to the bottom of the culture dish and therefore tend to float out of focus. 
		NOTE: Coating the dish with poly-L-lysine for immobilization is detrimental to the development of early germ-line cysts. Instead, individual cysts can be isolated. With some practice, it is possible to single out a particular cyst by very gently pushing it with the needle. This will allow the transfer of single cysts to separate culture dishes with a glass Pasteur pipette with a drawn-out tip that fits into the field of view of the microscope. The isolated cysts then can be easily relocated in the culture dishes even after long-term incubation.</li>
	</ol>
	</li>
</ol>
5. Pharmacological Assay with Intact Pupal Testes
<ol>
	<li>Fill each well of a 24-well cell culture plate with 500&#160;&#181;l of medium for one experiment of 12 replicates. Transfer one pupal testis to each well, using a pre-cut 200&#160;&#181;l pipette tip.
	<ol>
		<li>Check for the presence of protamines in the isolated testes using an inverted fluorescence microscope. The testes should be free of ProtamineB-eGFP signal, which indicates that the H-P switch has not taken place in any of the cysts. Replace testes that already show ProtamineB-eGFP-positive cysts.</li>
		<li>To the first 12 wells, add 500&#160;&#181;l of medium containing diluted inhibitor compound (anacardic acid) to reach the desired final concentration (150&#160;&#181;M) in 1&#160;ml of medium per well. Use the remaining wells as untreated controls; add 500&#160;&#181;l of medium with the same amount of solvent used as with the inhibitor compounds. Note: If many different compounds and solvents are used, it might be more efficient to use medium alone as control and to test the influence of increasing solvent concentrations in separate experiments.</li>
	</ol>
	</li>
	<li>Incubate the plate at 25&#160;&#176;C for 24 hr in the dark.</li>
	<li>Check again for the presence of protamines in the incubated testes as described above. The control testes should now show one or more ProtamineB-eGFP-positive cysts, which indicates transition through the H-P switch.</li>
	<li>Using a pipette with a 200&#160;&#181;l tip, transfer the testes to poly-L-lysine-coated slides, make squash preparations and stain with appropriate fluorescent antibodies as described elsewhere12,27,28.</li>
</ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibition+of+histone+acetyltransferase+activity+by+anacardic+acid+sensitizes+tumor+cells+to+ionizing+radiation.">Inhibition of histone acetyltransferase activity by anacardic acid sensitizes tumor cells to ionizing radiation.</a> FEBS Lett. 580 (18), 4353-4356 (2006).</li><li>Aldaz, S., Escudero, L. M., Freeman, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+Drosophila+imaginal+disc+development.">Live imaging of Drosophila imaginal disc development.</a> PNAS. 107 (32), 14217-14222 (2010).</li><li>Niki, Y., Yamaguchi, T., Mahowald, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+stable+cell+lines+of+Drosophila+germ-line+stem+cells.">Establishment of stable cell lines of Drosophila germ-line stem cells.</a> PNAS. 103 (44), 16325-16330 (2006).</li><li>Church, K., Lin, H. P. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetochore+microtubules+and+chromosome+movement+during+prometaphase+in+Drosophila+melanogaster+spermatocytes+studied+in+life+and+with+the+electron+microscope.">Kinetochore microtubules and chromosome movement during prometaphase in Drosophila melanogaster spermatocytes studied in life and with the electron microscope.</a> Chromosoma. 92 (4), 273-282 (1985).</li><li>Rebollo, E., Gonz&#225;lez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+the+spindle+checkpoint+in+Drosophila+spermatocytes.">Visualizing the spindle checkpoint in Drosophila spermatocytes.</a> EMBO rep. 1 (1), 65-70 (2000).</li><li>Cheng, J., Hunt, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-lapse+Live+Imaging+of+Stem+Cells+in+Drosophila+Testis.">Time-lapse Live Imaging of Stem Cells in Drosophila Testis.</a> Curr. Protoc. Stem Cell Biol. 11, 2E.2.1-2E.2.8 (2009).</li></ol>

The authors have no competing interests.

The presented protocol describes two different applications based both on the ability to dissect Drosophila testes and single germ-line cysts at a given developmental stage and on the ex vivo development of these cells and tissues in a culture dish. One application is the pharmacological manipulation of vital processes during sperm development. Importantly, this allows access to post-meiotic spermatogenesis that is not easily gained using functional analyses based on fly genetics. The other application is the microscopic observation of living and developing germ cells and testes. Especially this application benefits from the ability of Drosophila cells and organs in culture to survive and develop at RT and under normal atmosphere. Hence, an inverted imaging microscope equipped with a heated stage (heating to standard breeding temperature of 25&#160;&#176;C) is sufficient to obtain meaningful results in live-imaging experiments. Furthermore, many transgenic fly strains expressing fluorescent protein-tagged proteins for live imaging exist or can be easily established.
To obtain pupal testes or germ-line cysts of a specific developmental stage, it is crucial that one is familiar with the timing of Drosophila development. The exact timing for each stage might vary among laboratories, culture conditions, and fly strains and must be established empirically. As outlined above, this is especially important for pharmacological assays that, for example, target the H-P switch. For such experiments, it is advisable to always check for spermatids with a ProtamineB-eGFP signal before the actual inhibitor test because the timing can vary slightly even within one population.
Following the protocol above should enable the experimenter to dissect testes from early pupal stages as intact organs free of attached fat body tissue. These testes and, even more so, isolated germ-line cysts are very fragile. Hence, it is crucial to develop the manual dexterity and experience necessary for handling and transferring testes and cysts using forceps, needles, and pipettes. Especially studies targeting meiotic and early post-meiotic germ-cell development would benefit from this. While it is comparatively easy to dissect cysts of late spermatids with long flagella from adult testes, it is difficult to obtain earlier stages. Dissecting these cysts from early pupal testes following this protocol is much more efficient. Keep in mind that in a usual fly strain, only ca. 50% of the pupae will be male. Therefore, prepare to dissect at least twice as many pupae as the number of testes required. Alternatively, it is possible to identify male L3 larvae using a stereo-microscope, as the testes can be identified as translucent round organs embedded in the lateral fat body tissues of the intact larva23,34, and to collect them in fresh culture vials that can be used to pick pre-pupae for staging and dissections. It is also possible to identify male pre-pupae35.
We noticed that both the isolated germ-line cysts and the intact pupal testes in culture are sensitive to light, as has also been reported previously18. This light sensitivity might also hold for some chemical compounds used in pharmacological assays. Therefore, it is best to keep the culture dishes of an assay in the dark during incubation. Likewise, when planning time-lapse imaging experiments with developing testes and, especially, isolated cysts, keep in mind that too-long exposure times and/or too-high sampling rates may inhibit ex vivo development. The appropriate sampling rate should be determined experimentally.
Bacterial and/or fungal infection and over-growth in the culture dishes poses a severe problem. Infected culture dishes with too many bacteria do not support ex vivo development of the testes and cysts. Therefore, the described culture medium contains penicillin and streptomycin (see step 1.1), but during long-term incubation, these antibiotics might be degraded and their activity might diminish. This could be one of the reasons for the limited time of survival (max. 72&#160;hr) of cultured cysts observed when using the protocol above. Yet this time frame could not be extended by regular replacement of the culture medium in the dishes. We conclude that other factors might influence the survival in culture, such as a lack of growth signals from other tissues of the genital tract. On the other hand, post-meiotic development is quicker in Drosophila than in mammals. In Drosophila, spermiogenesis takes approximately 90 hr, and the H-P switch takes place between 50 and 60 hr after meiosis9,12. Thus, the usual survival time of around 48 hr achieved with this protocol is well suited to follow pivotal developmental processes in spermatogenesis, such as meiotic divisions or the H-P switch. Although bacterial or fungal contamination can be avoided by using clean tools and media, the presented protocol does not make use of strictly sterile work conditions because flies and fly testes already contain bacteria when raised under standard conditions. Nevertheless, some applications of this culture technique might benefit from flies dissected or even raised under aseptic conditions (for protocols, see&#160;17,36,37).
Pharmacological assays depend on the use of chemical compounds acting as inhibitors or activators of various enzymes. The compounds commonly used in such experiments often differ widely in their target specificity. As an advantage, this offers the potential to target whole enzyme classes, for example, with anacardic acid inhibiting p300/CBP, PCAF, and MYST family members of HATs38. The downside of this could be potential off-target or cytotoxic effects induced by such compounds. Therefore, each compound used in an assay with pupal testes or cysts should be tested for its cytotoxicity and specific effect at different concentrations. Furthermore, slight variations in the culture conditions, compound concentration or activity, and variations in the cell permeability might lead to variability of the induced effects. Therefore, it is necessary to monitor the success of the treatment in each experiment. For example, when we used anacardic acid at 150 &#181;M, we observed slight variability in the strength of the chromatin condensation phenotype between and sometimes within some testes, while acetylation of histone H4 was consistently decreased.
Pharmacological assays with Drosophila testes in culture have been used to analyze pre- and post-meiotic mechanisms of spermatogenesis4,12,14,21. Since it is difficult to access post-meiotic spermatogenesis with genetic tools to target players with essential pre-meiotic and meiotic functions, this ex vivo approach constitutes an alternative. Furthermore, in principle, the method could be adapted to pulse-chase experiments to follow the development of treated germ cells over time. However, we have not yet tested this possibility. Making use of early pupal testes is an advantage in pharmacological assays. First, the thin outer sheath of the young pupal testes (around 24 hr APF) could allow improved permeability of chemical compounds. Second, when studying the post-meiotic H-P switch, the pupal testes dissected at 24 hr APF from the Protamine-GFP-expressing fly line enable a direct readout of whether the H-P switch was affected or not.
To date, several protocols for ex vivo cultivation of Drosophila organs and tissues including testes and germ-line cysts have been developed (e.g.,&#160;see&#160;4,14,17,20,39,40). The cultivation medium and the general conditions used in the protocol presented here were established in 197917. The culture system was later adapted to in vivo imaging of the individualization mechanism in late post-meiotic cysts isolated from adult testes4. Previously, we have reported that these conditions also support survival and differentiation of meiotic and early post-meiotic germ cells in their cysts for around 48 hr12. In contrast to other culture systems19 the observed developmental timing in these cultures was approximately consistent with the timing determined from fixed samples (for details, see12). H2AvD-RFP and Protamine-GFP fluorescence signals can be used to visualize the nuclear morphology and chromatin composition of the germ cells in culture. We found that for clear identification of developmental stages in culture, this is advantageous over other criteria, such as coiling of cysts. Importantly, the presented protocol does not involve addition of fly extract or other growth factors other than fetal bovine serum. Furthermore, we show here that the conditions are compatible with fluorescence microscopy imaging of meiotic divisions ex vivo in culture. Images can be obtained with reasonable resolution in an easy-to-use medium that supports long-term development. This is in contrast to protocols enabling short-term (ca. 3 hr) observations in Voltalef oil41,42.
The procedures described above for cultivation and pharmacological treatment of pupal testes and single male germ-line cysts are easily adaptable to the many genetic, epigenetic, cell biological, or developmental questions that can be researched in Drosophila spermatogenesis. This includes especially research focusing on germ-line stem cells. It has been shown that stem cell development can be followed by live imaging of cultured adult testes20,43. However, the peristaltic movement of the mature testis sheath interferes with image acquisition. Therefore, either imaging is done using fragmented testes43 or the number of imaging experiments performed has to be increased to account for lost datasets20. Early pupal testes (24 hr APF) are not yet enclosed with muscle cells. Even slightly older testes (36&#8211;45 hr APF) appear to show few peristaltic movements (compare Figure 3 and Supplemental video 1). Therefore, cultivation of young pupal testes as intact organs could serve as an alternative.
Furthermore, once mastered, the ability to dissect pupal testes and eventually isolated germ-line cysts will allow further applications using single cysts as a source of a homogeneous, albeit small, cell population. For example, it is thus possible to perform RT-qPCR to analyze the transcriptional regulation of specific genes during defined stages of spermatogenesis, as has already been demonstrated15.

The development of male germ cells of many species is a sequential process. It is characterized by a series of crucial events that culminates in the formation of morphologically and epigenetically highly specialized spermatozoa. In Drosophila melanogaster, germ-line stem cells at the tip of the testis divide asymmetrically to give rise to a new stem cell and the spermatogonium, i.e.,&#160;the daughter cell that is committed to differentiation (see Figure 1 for an overview)1,2. The spermatogonium undergoes four rounds of mitotic divisions and, with the onset of meiosis, a phase of impressive growth and transcriptional activity called the spermatocyte stage. The cells originating from one spermatogonium remain connected to each other and develop as a synchronized bundle wrapped by two somatic cells&#8212;a structure referred to as a cyst. After completion of meiosis, the morphology of the male germ cells is completely reorganized (schematically shown in Figure 1). Initially, round spermatids elongate to form the hydrodynamic head structure, and the flagellum develops. Eventually, the sperm cells are separated from each other by a complex, apoptosis-related mechanism called individualization3&#8211;5.
In many species, including Drosophila melanogaster and mammalian species, the morphological changes of the germ cells are accompanied by a remarkable epigenetic rearrangement of the haploid male germ-line chromatin, called the histone-to-protamine switch (H-P switch)6&#8211;9. Possibly nearly all canonical histone and many histone-variant molecules are stripped from the DNA and are replaced by small basic proteins, the protamines, resulting in the highly compact nucleoprotamine structure specific to the chromatin of mature sperm. General characteristics of the H-P switch are the replacement of canonical histones first by histone variants, then by transitional proteins, and finally by protamines. All this is accompanied by several changes to epigenetic marks, such as a surge in acetylation levels of histone H4 just prior to histone removal7,10-12.
Most, if not all, of the above-mentioned processes are essential for the development of mature, fully fertile sperm. This is true not only in Drosophila but also in humans, as mammalian spermatogenesis shares a considerable amount of similarity with the invertebrate system1,8,9. Studying male germ-cell development is greatly facilitated in the Drosophila model system. Flies are genetically highly accessible. The generation of mutants as well as the establishment of fly strains expressing fusion genes or RNA interference constructs takes little effort. However, in the study of post-meiotic spermatogenesis, the use of mutants and simple genetic tools could reach a limit. In Drosophila spermatogenesis, transcription ceases almost completely with entry into meiotic divisions. Thus, post-meiotic spermatogenesis is mainly based on translationally repressed and stored mRNAs synthesized in an extended meiotic prophase13-16. Therefore, tools like RNA interference or the use of non-conditional mutants are not suitable for studying specifically the post-meiotic roles of gene products that also fulfill essential functions in pre-meiotic or meiotic germ cell development.
One advantage of Drosophila that can be exploited in such cases is the ability of dissected intact testes and even cysts of germ cells to develop ex vivo in culture medium4,12,17&#8211;20. The dissection and cultivation of testes or germ-line cysts allows not only microscopic observation of germ-cell development in living cells, but also the use of pharmacological assays with, e.g.,&#160;inhibitors of enzyme complexes such as histone acetyltransferases (HATs), histone deacetylases (HDACs), topoisomerases, or the proteasome. Thus, it is possible to manipulate enzymatic functions during post-meiotic spermatogenesis and to monitor the developmental outcomes regardless of embryonic, pre-meiotic, or meiotic functions4,12.
In pharmacological assays, we previously analyzed the acetylation-dependent localization of a bromodomain protein during meiotic prophase as well as the relevance of histone acetylation levels for the epigenetic H-P switch in post-meiotic spermatogenesis by using specific inhibitors targeting HATs and HDACs12,21. Targeting the H-P switch in these assays was made possible by using a fly strain that expresses the fusion genes histone2AvD-RFP and protamineB-eGFP12,22 in the endogenous patterns, and the observation that the first spermatids in pupal testes pass through the H-P switch between 24 and 48 hr APF12.
The protocol presented describes the dissection of testes and cysts from Drosophila pupae, the culture conditions, and a pharmacological assay with intact pupal testes. For this purpose, pupal testes at around 24 hr after puparium formation (APF) are dissected (see Figures 1 and 2). At this time, the testes have not made connections to other tissues and are surrounded by only a thin outer sheath, which allows better penetration by the inhibitor compounds23,24. The testes are bean- or pear-shaped organs that can be easily taken into culture. These testes are dissected, cultured, and treated with specific inhibitors. Subsequently, the effects of the inhibitors are monitored using immunofluorescence analyses and fluorescence microscopy of fusion protein expression, and by comparison of the nuclear morphology of the treated germ cells to that of untreated germ cells. The conditions presented in this protocol also allow live imaging of developing testes and germ-line cysts in culture.

<table border="0" cellpadding="0" cellspacing="0" width="782">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">Anacardic acid</td>
			<td style="width:105px;">Merck Millipore</td>
			<td style="width:99px;">172050</td>
			<td style="width:247px;">brand: Calbiochem (EMD Millipore)</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Anti-acetyl-Histone H4</td>
			<td>Merck Millipore</td>
			<td>06-598</td>
			<td>brand: Upstate</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">AxioCam MRm&#160;</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">AxioObserver.Z1 inverted microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dimethyl sulfoxide</td>
			<td>Sigma</td>
			<td>D8418</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dumont 5-Inox forceps</td>
			<td>Dumont</td>
			<td></td>
			<td>multiple suppliers</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fetal Bovine Serum, Heat Inactivated</td>
			<td>Sigma</td>
			<td>F4135</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fiber optical&#160; illuminator</td>
			<td></td>
			<td></td>
			<td>multiple suppliers</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Glass Bottom Microwell Dishes</td>
			<td>MatTek</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">goat anti-rabbit IgG (H+L)-Cy5</td>
			<td>Dianova</td>
			<td>711-175-152</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Hoechst</td>
			<td>Sigma</td>
			<td>B1155</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Inoculation loop or pin holder</td>
			<td></td>
			<td></td>
			<td>multiple suppliers</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;">Austerlitz INSECT PINS stainless steel needles, size 0</td>
			<td>Plano GmbH</td>
			<td>N5018</td>
			<td rowspan="2" style="width:247px;">In our hands both these pin sizes worked well in the dissections. The minutiens are less rigid but have a smaller tip diameter.</td>
		</tr>
		<tr height="28">
			<td height="28" style="height:28px;">Austerlitz INSECT PINS stainless steel minutiens, diam. 0.1 mm</td>
			<td>Plano GmbH</td>
			<td>N5014</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Multiwell plates, 24-wells</td>
			<td>Becton Dickinson</td>
			<td>351147</td>
			<td>brand: Falcon</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pen Strep</td>
			<td>invitrogen</td>
			<td>15070</td>
			<td>brand: Gibco</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Shields and Sang M3 Insect Medium</td>
			<td>Sigma</td>
			<td>S8398</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Stemi SV6 binocular</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment at JoVE.com

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.

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 ...

ex-vivo-culture-drosophila-pupal-testis-single-male-germ-line-cysts

Research

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Biology

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

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 ...

Ex Vivo Culture of Primary Human Fallopian Tube Epithelial Cells

Epithelial ovarian cancer is a leading cause of female cancer mortality in the United States. In contrast to other women-specific cancers, like breast and uterine carcinomas, where death rates have fallen in recent years, ovarian cancer cure rates have remained relatively unchanged over the past two decades 1. This is largely due to the lack of appropriate screening tools for detection of early stage disease where surgery and chemotherapy are most effective 2, 3. As a result, most patients present with advanced stage disease and diffuse abdominal involvement. This is further complicated by the fact that ovarian cancer is a heterogeneous disease with multiple histologic subtypes 4, 5. Serous ovarian carcinoma (SOC) is the most common and aggressive subtype and the form most often associated with mutations in the BRCA genes. Current experimental models in this field involve the use of cancer cell lines and mouse models to better understand the initiating genetic events and pathogenesis of disease 6, 7. Recently, the fallopian tube has emerged as a novel site for the origin of SOC, with the fallopian tube (FT) secretory epithelial cell (FTSEC) as the proposed cell of origin 8, 9. There are currently no cell lines or culture systems available to study the FT epithelium or the FTSEC. Here we describe a novel ex vivo culture system where primary human FT epithelial cells are cultured in a manner that preserves their architecture, polarity, immunophenotype, and response to physiologic and genotoxic stressors. This ex vivo model provides a useful tool for the study of SOC, allowing a better understanding of how tumors can arise from this tissue, and the mechanisms involved in tumor initiation and progression.

Ex Vivo Culture of Primary Human Fallopian Tube Epithelial Cells

The fallopian tube (FT) is emerging as an alternative site of origin for serous ovarian carcinoma (SOC). This protocol describes a novel method for the isolation and ex vivo culture of fallopian tube epithelial cells. This system recapitulates the in vivo epithelium and allows the study of SOC pathogenesis.

Epithelial ovarian cancer is a leading cause of female cancer mortality in the United States. In contrast to other women-specific cancers, like breast and ...

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Ex vivo Culture of Mouse Embryonic Skin and Live-imaging of Melanoblast Migration

Melanoblasts are the neural crest derived precursors of melanocytes; the cells responsible for producing the pigment in skin and hair. Melanoblasts migrate through the epidermis of the embryo where they subsequently colonize the developing hair follicles1,2. Neural crest cell migration is extensively studied in vitro but in vivo methods are still not well developed, especially in mammalian systems. One alternative is to use ex vivo organotypic culture3-6. Culture of mouse embryonic skin requires the maintenance of an air-liquid interface (ALI) across the surface of the tissue3,6. High resolution live-imaging of mouse embryonic skin has been hampered by the lack of a good method that not only maintains this ALI but also allows the culture to be inverted and therefore compatible with short working distance objective lenses and most confocal microscopes. This article describes recent improvements to a method that uses a gas permeable membrane to overcome these problems and allow high-resolution confocal imaging of embryonic skin in ex vivo culture6. By using a melanoblast specific Cre-recombinase expressing mouse line combined with the R26YFPR reporter line we are able to fluorescently label the melanoblast population within these skin cultures. The technique allows live-imaging of melanoblasts and observation of their behavior and interactions with the tissue in which they develop. Representative results are included to demonstrate the capability to live-image 6&#160;cultures in parallel.

Ex vivo Culture of Mouse Embryonic Skin and Live-imaging of Melanoblast Migration

We describe the dissection and ex vivo culture of mouse embryonic skin. The culture system maintains an air-liquid interface across the tissue surface and allows imaging on an inverted microscope. Melanoblasts, a component of the developing skin, are fluorescently labeled allowing their behavior to be observed using confocal microscopy.

Melanoblasts are the neural crest derived precursors of melanocytes; the cells responsible for producing the pigment in skin and hair. Melanoblasts ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

Mouse Fetal Whole Intestine Culture System for Ex Vivo Manipulation of Signaling Pathways and Three-dimensional Live Imaging of Villus Development

Most morphogenetic processes in the fetal intestine have been inferred from thin sections of fixed tissues, providing snapshots of changes over developmental stages. Three-dimensional information from thin serial sections can be challenging to interpret because of the difficulty of reconstructing serial sections perfectly and maintaining proper orientation of the tissue over serial sections. Recent findings by Grosse et al., 2011 highlight the importance of three- dimensional information in understanding morphogenesis of the developing villi of the intestine1. Three-dimensional reconstruction of singly labeled intestinal cells demonstrated that the majority of the intestinal epithelial cells contact both the apical and basal surfaces. Furthermore, three-dimensional reconstruction of the actin cytoskeleton at the apical surface of the epithelium demonstrated that the intestinal lumen is continuous and that secondary lumens are an artifact of sectioning. Those two points, along with the demonstration of interkinetic nuclear migration in the intestinal epithelium, defined the developing intestinal epithelium as a pseudostratified epithelium and not stratified as previously thought1. The ability to observe the epithelium three-dimensionally was seminal to demonstrating this point and redefining epithelial morphogenesis in the fetal intestine. With the evolution of multi-photon imaging technology and three-dimensional reconstruction software, the ability to visualize intact, developing organs is rapidly improving. Two-photon excitation allows less damaging penetration deeper into tissues with high resolution. Two-photon imaging and 3D reconstruction of the whole fetal mouse intestines in Walton et al., 2012 helped to define the pattern of villus outgrowth2. Here we describe a whole organ culture system that allows ex vivo development of villi and extensions of that culture system to allow the intestines to be three-dimensionally imaged during their development.

Mouse Fetal Whole Intestine Culture System for Ex Vivo Manipulation of Signaling Pathways and Three-dimensional Live Imaging of Villus Development

Improved imaging technology is allowing three-dimensional imaging of organs during development. Here we describe a whole organ culture system that allows live imaging of the developing villi in the fetal mouse intestine.