We thank Bryne Ulmschneider for contributions to the protocol and Diane Barber for suggestions on the manuscript. This work was funded by a National Institute of Health grant GM116384 to T.G. Nystul and D.L. Barber.

NOTE: To measure pHi in the FSC lineage, we calculate the ratio of fluorescence intensities of pHluorin to mCherry in FSCs, pFCs, and follicle cells in physiological conditions, and convert the ratios into pHi values with standard calibration curves for each cell type7. First, live imaging experiments are performed to measure fluorescence intensities of pHluorin and mCherry in germaria dissected into a buffer containing NaHCO3, which mimics physiological conditions1</s...

<ol>
	<li>Grillo-Hill, B. K., Choi, C., Jimenez-Vidal, M., Barber, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+H++efflux+is+sufficient+to+induce+dysplasia+and+necessary+for+viability+with+oncogene+expression.">Increased H+ efflux is sufficient to induce dysplasia and necessary for viability with oncogene expression.</a> eLife. 03270, (2015).</li><li>Ulmschneider, B., Grillo-Hill, B. K., Benitez, M., Azimova, D. R., Barber, D. L., Nystul, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+intracellular+pH+is+necessary+for+adult+epithelial+and+embryonic+stem+cell+differentiation.">Increased intracellular pH is necessary for adult epithelial and embryonic stem cell differentiation.</a> J. Cell Biol. 215 (3), 345-355 (2016).</li><li>Han, J., Burgess, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+Indicators+for+Intracellular+pH.">Fluorescent Indicators for Intracellular pH.</a> Chemical reviews. 110 (5), 2709-2728 (2010).</li><li>Koivusalo, M., Welch, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amiloride+inhibits+macropinocytosis+by+lowering+submembranous+pH+and+preventing+Rac1+and+Cdc42+signaling.">Amiloride inhibits macropinocytosis by lowering submembranous pH and preventing Rac1 and Cdc42 signaling.</a> J. Cell Biol. 188 (4), 547-563 (2010).</li><li>Choi, C. -. H., Webb, B. A., Chimenti, M. S., Jacobson, M. P., Barber, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=pH+sensing+by+FAK-His58+regulates+focal+adhesion+remodeling.">pH sensing by FAK-His58 regulates focal adhesion remodeling.</a> J. Cell Biology. 202 (6), 849-859 (2013).</li><li>Rossano, A. J., Chouhan, A. K., Macleod, G. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetically+encoded+pH-indicators+reveal+activity-dependent+cytosolic+acidification+of+Drosophila+motor+nerve+termini+in+vivo.">Genetically encoded pH-indicators reveal activity-dependent cytosolic acidification of Drosophila motor nerve termini in vivo.</a> J. Phys. 591 (7), 1691-1706 (2013).</li><li>Grillo-Hill, B. K., Webb, B. A., Barber, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ratiometric+imaging+of+pH+probes.">Ratiometric imaging of pH probes.</a> Methods in Cell Biol. 123, 429-448 (2014).</li><li>Miesenb&#246;ck, G., De Angelis, D. A., Rothman, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+secretion+and+synaptic+transmission+with+pH-sensitive+green+fluorescent+proteins.">Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins.</a> Nature. 394 (6689), 192-195 (1998).</li><li>Kirilly, D., Xie, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+ovary:+an+active+stem+cell+community.">The Drosophila ovary: an active stem cell community.</a> Cell research. 17 (1), 15-25 (2007).</li><li>Sahai-Hernandez, P., Castanieto, A., Nystul, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+models+of+epithelial+stem+cells+and+their+niches.">Drosophila models of epithelial stem cells and their niches.</a> Wiley interdisciplinary reviews. Dev. Biol. 1 (3), 447-457 (2012).</li><li>Losick, V. P., Morris, L. X., Fox, D. T., Spradling, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+stem+cell+niches:+a+decade+of+discovery+suggests+a+unified+view+of+stem+cell+regulation.">Drosophila stem cell niches: a decade of discovery suggests a unified view of stem cell regulation.</a> Dev. Cell. 21 (1), 159-171 (2011).</li><li>Pocha, S. M., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+and+molecular+mechanisms+of+single+and+collective+cell+migrations+in+Drosophila:+themes+and+variations.">Cellular and molecular mechanisms of single and collective cell migrations in Drosophila: themes and variations.</a> Ann Rev. of Gen. 48, 295-318 (2014).</li><li>St Johnston, D., Ahringer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+polarity+in+eggs+and+epithelia:+parallels+and+diversity.">Cell polarity in eggs and epithelia: parallels and diversity.</a> Cell. 141 (5), 757-774 (2010).</li><li>Castanieto, A., Johnston, M. J., Nystul, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EGFR+signaling+promotes+self-renewal+through+the+establishment+of+cell+polarity+in+Drosophila+follicle+stem+cells.">EGFR signaling promotes self-renewal through the establishment of cell polarity in Drosophila follicle stem cells.</a> eLife. 3, (2014).</li><li>Margolis, J., Spradling, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+behavior+of+epithelial+stem+cells+in+the+Drosophila+ovary.">Identification and behavior of epithelial stem cells in the Drosophila ovary.</a> Development. 121 (11), 3797-3807 (1995).</li><li>Nystul, T. G., Spradling, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+epithelial+niche+in+the+Drosophila+ovary+undergoes+long-range+stem+cell+replacement.">An epithelial niche in the Drosophila ovary undergoes long-range stem cell replacement.</a> Cell stem cell. 1 (3), 277-285 (2007).</li><li>Morris, L. X., Spradling, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+live+imaging+provides+new+insight+into+stem+cell+regulation+and+germline-soma+coordination+in+the+Drosophila+ovary.">Long-term live imaging provides new insight into stem cell regulation and germline-soma coordination in the Drosophila ovary.</a> Development. 138 (11), 2207-2215 (2011).</li><li>Lee, T., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosaic+analysis+with+a+repressible+cell+marker+(MARCM)+for+Drosophila+neural+development.">Mosaic analysis with a repressible cell marker (MARCM) for Drosophila neural development.</a> Trends in neuro. 24 (5), 251-254 (2001).</li></ol>

Here, we describe a method for measuring the pHi of cells in the FSC lineage within wildtype tissue. This protocol has been developed and refined over the past five years, since we first began to study pHi in Drosophila ovarian tissue. During that time, the protocol has been used successfully by multiple investigators in our lab and on at least four different spinning disc and laser scanning microscopes. The reproducibility of our original observation, that pHi increases as cells in the FSC lineage differentiate...

Recent studies revealed a role for changes in pHi during cellular differentiation and dysplasia in vivo1,2. These studies found that pHi is remarkably consistent in cells of the same type at the same stage of differentiation, but that it changes as cells transition from one stage to another. In some cases, blocking the changes in pHi partially disrupts differentiation, suggesting that the change in pHi is not just a consequence of changes in cell fate but instead helps to promote the change in cell fate, perhaps through effects on pH-sensitive regulatory proteins or chemical reactions required for differentiation. Future studies have the potential to reveal more insight into the many different roles of pHi dynamics in vivo. However, one of the challenges of studying pHi during differentiation in vivo is obtaining accurate measurements of pHi. Unlike other features of differentiation, such as changes in cellular morphology and gene expression, pHi is a labile chemical property of the cell that is not preserved in cells that have been fixed and permeabilized with standard methods. In addition, pHi may not be stable in cells that are stressed or dying as a result of experimental manipulation. Thus, it is important to keep cells alive and as healthy as possible when measuring pHi. Several vital dyes are available that work well for measuring the pHi of cells in culture3, but in many cases they are not suitable for in vivo studies because they do not penetrate the tissue deeply or evenly enough to provide accurate measurements.
To circumvent the problem of poor dye penetration, we and others have used a genetically encoded probe, mCherry::pHluorin4,5,6,7, that can be expressed specifically in the cell types of interest and imaged in live tissue. pHluorin is a variant of GFP with a higher pKa (~ 7.0 vs. ~ 4.0) that folds more readily at higher pH; so the total fluorescence intensity emitted from a population of pHluorin molecules in the cell increases with increasing pHi8. Importantly, fluorescence is linear within the normal cytosolic range of pHi values. In contrast, the fluorescence of mCherry (pKa ~ 4.5) is insensitive to pH changes within the cytosolic range. These two reporters are covalently linked together as a single chimeric protein, encoded by a single open reading frame, so they are always present in equal amounts. Therefore, the ratio of pHluorin to mCherry fluorescence intensity provides a measurement of pHi that is normalized to the probe concentration in each cell. The ratios can then be converted to estimates of pHi values using a standard curve that is generated by obtaining the pHluorin to mCherry ratios from tissues that have been equilibrated to known pH values.
Here, we describe the methods for using mCherry::pHluorin to measure the pHi of the epithelial FSC lineage in the Drosophila ovary. This well-characterized tissue has been used to model many different aspects of epithelial biology, such as stem cell self-renewal and differentiation9,10,11, collective cell migration12, and the development and maintenance of cell polarity13,14. The follicle epithelium is produced by two FSCs that reside at the anterior edge of the tissue in a structure called the germarium15,16. These cells divide regularly during adulthood to self-renew and produce progeny, called prefollicle cells (pFCs), that can either re-enter the niche and become an FSC or differentiate into one of three different follicle cell types: polar cells, stalk cells, or main body follicle cells. We showed previously that in wildtype tissue, the pHi increases steadily during the early stages of differentiation, from a pHi of approximately 6.8 in FSCs to 7.0 in pFCs, to 7.3 in follicle cells2. Blocking this increase by RNAi knockdown of a ubiquitously expressed sodium/proton exchanger, DNhe2, severely impairs pFC differentiation, whereas increasing pHi by overexpression of DNhe2 causes a mild excess differentiation phenotype. These findings demonstrate that pHi is stably maintained in the early FSC lineage and that it can be experimentally increased or decreased in vivo. The methods described here can be used to measure pHi in either wildtype tissue or various forms of mutant tissues, including RNAi knockdown or overexpression using a Gal4 of interest, and mitotic clones.

<table>
	<tbody>
		<tr>
			<td>Fly Stocks</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>UAS-mCherry::pHluorin[1]</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>y1 w*;P{GawB}10930/CyO</td>
			<td>Bloomington Stock Center</td>
			<td>7023</td>
			<td></td>
		</tr>
		<tr>
			<td>Act-Gal4 flipout stock</td>
			<td>Bloomington Stock Center</td>
			<td>4409</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Chemicals for Buffer preparation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma Aldrich</td>
			<td>S5886</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma Aldrich</td>
			<td>P-3911</td>
			<td></td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>Mallinckrodt</td>
			<td>4912</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Thermo Fisher Scientific</td>
			<td>BP310</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>Thermo Fisher Scientific</td>
			<td>M63</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma Aldrich</td>
			<td>C-5080</td>
			<td></td>
		</tr>
		<tr>
			<td>HCO3</td>
			<td>Sigma Aldrich</td>
			<td>S-5761</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma Aldrich</td>
			<td>M-9272</td>
			<td></td>
		</tr>
		<tr>
			<td>NMDG+</td>
			<td>Sigma Aldrich</td>
			<td>M-2004</td>
			<td></td>
		</tr>
		<tr>
			<td>K2HPO4</td>
			<td>Mallinckrodt</td>
			<td>7088</td>
			<td>Use to Make KHPO4 pH 7.4</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Thermo Fisher Scientific</td>
			<td>BP362</td>
			<td>Use to Make KHPO4 pH 7.4</td>
		</tr>
		<tr>
			<td>Concanavalin A, Alexa Fluor 647 Conjugate</td>
			<td>Thermo Fisher Scientific</td>
			<td>C21421</td>
			<td>0.25 mg/ml dilution</td>
		</tr>
		<tr>
			<td>Nigericin</td>
			<td>Thermo Fisher Scientific</td>
			<td>N1495</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Dissection and mounting tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>2 Dumont Inox forceps (Size 5)</td>
			<td>Thermo Fisher Scientific</td>
			<td>NC9473431</td>
			<td></td>
		</tr>
		<tr>
			<td>2 23-gauge syringe needles</td>
			<td>Sigma Aldrich</td>
			<td>Z192457</td>
			<td></td>
		</tr>
		<tr>
			<td>9-well glass dissecting dish</td>
			<td>Thermo Fisher Scientific</td>
			<td>13-748B</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum Grease</td>
			<td>Dow Corning</td>
			<td>1018817</td>
			<td></td>
		</tr>
		<tr>
			<td>22 X 40 mM glass coverslips</td>
			<td>Thermo Fisher Scientific</td>
			<td>12545C</td>
			<td></td>
		</tr>
		<tr>
			<td>Round Glass Coverslips, 12mm diameter, 0.13-0.16mm thickness</td>
			<td>Ted Pella, Inc.</td>
			<td>26023</td>
			<td></td>
		</tr>
		<tr>
			<td>3-D mounting chamber</td>
			<td>custom manufactured</td>
			<td></td>
			<td>.stl and .ipt files for 3-D printer included as supplemental files</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Other equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Thermo Fisher Scientific</td>
			<td>13-620-183A</td>
			<td>Model: Accumet AB15</td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td>Olympus Corporation</td>
			<td>0H11436</td>
			<td>Model: SZ2-ST</td>
		</tr>
		<tr>
			<td>Confocal Microscope</td>
			<td>Leica Biosystems</td>
			<td></td>
			<td>SP5 or SP8 laser-scanning confocal microscope with a 40&#215; objective with a numerical aperture of 1.3</td>
		</tr>
	</tbody>
</table>

methods for imaging intracellular ph of the follicle stem cell lineage in live drosophila ovarian tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Here we have described the process of measuring pHi in the follicle epithelium, which involves several steps. First, the ovaries are dissected from flies of the appropriate genotype using tools for dissecting and mounting (Figure 1). The ovarioles are then imaged using quantitative fluorescence microscopy and the images are analyzed to obtain measurements of pHi. For each image, the cell types of interest are identified as described in Section 3.1 (<strong cl...

Watch this Scientific Journal Video about Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue at JoVE.com

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

The overall goal of this procedure is to measure the intracellular pH of follicle cells within live explanted Drosophila ovarian tissue. This method can help us answer key questions in stem cell and developmental biology such as, how changes to intracellular pH contribute to sulfate decisions. The main advantage of this technique is that it makes it possible to accurately quantify pH within living intact tissue.To begin the experiment, prepare mounting chambers by applying a thin coating of vacuum grease to the flatter side of a 3D printed mounting chamber. Place the flatter side on a 22 x 40 millimeter cover slip to seal the cover slip to the mounting chamber. Next, dissect the ovaries from female adult flies in 500 microliters of dissection buffer.Anesthetize three to four flies using CO2 gas and transfer the flies onto a fly pad. Use forceps to select a fly and place it into the dissection media. Pinch the thorax of the fly with one pair of forceps while tugging on the tip of its abdomen until its ovaries are exposed.Set a timer for exactly 10 minutes. After detaching the ovaries from the other organs, tease apart the ovarioles using 22 and a half gauge syringe needles. Use one syringe needle to hold the cluster of ovarioles in place at its anterior end and stroke downwards along the length of single ovarioles to carefully separate the muscle sheath from the ovaries.Incubate the dissected ovaries in dissection media for the remainder of the time on the timer. It is important the ovaries remain in the dissection media for exactly 10 minutes to provide enough time for the cells. Place a small drop of dissection media onto the glass cover slip inside the mounting chamber.Transfer the separated ovarioles using forceps. Separate later stage egg chambers from germaria associated with earlier stage egg chambers and ensure that the dissected germaria are placed at the center of the dissection media drop. Add two small drops of nail polish to opposite sides of around 12 millimeter cover slip and let it air dry for about 10 seconds.Place the round cover slip on the drop of dissection media containing separated germaria, so that the side with nail polish faces down and contacts the dissection media. Press down on the edges to flatten and secure the germaria in position. Fill mounting chamber with additional dissection media and move on to live imaging.Image ovarioles in ny dissection buffer at pH 6.5. Adjust settings such that pixel intensities of the pHluorin and mCherry images are low, but not below the limits of detection of the camera. Next, image another set of ovarioles in nygerion dissection buffer at pH 7.5 and adjust the settings such that the pixel intensities of the pHluorin and mCherry images are high but not saturated.Image ovaries and bi-carbonate dissection buffer and ensure that the pixel intensities of the pHluorin and mCherry images are not saturated with the chosen settings. If pixels are saturated, adjust the settings. After selecting appropriate live imaging settings, proceed to collecting images of at least five germaria in each of the conditions.Subtract the background by opening unprocessed images in Fiji with each channel in a separate window. Use the rectangle tool to draw a region of interest or ROI in the pHluorin channel window and a part of the image without signal. Set the lower limit of the threshold so that pixels with intensity values below the background are excluded and set the upper limit of the threshold to maximum.When the threshold is set in this way, the areas of the image without signal are mostly blue and the signal is clearly visible. If the threshold was set, navigate to the set measurements dialog box and make sure to check the limit to threshold box. Measure the mean fluorescence intensity in the ROI, go to process, then, math menu to subtract the measured background intensity from each channel and slice of the image using the subtract function.Repeat the steps for the mCherry channel window but, use the restore selection function instead of drawing a new rectangle with the rectangle tool found in the edit selection menu, to add a rectangle with the same dimensions and position on the image. Choose one or more slices in which the cell of interest has highest fluorescence intensity in the mCherry channel and make sure that the concanavalin A staining in the far red channel, is in focus within the selected slice. Next, draw an ROI around each follicle stem cell or FSC and measure the mean fluorescence intensity of pHluorin and mCherry and all of the slices chosen for measurements.Finally, divide the mean fluorescence intensity of the pHluorin channel by the mean fluorescence intensity of the mCherry channel, to calculate a ratio of pHluorin to mCherry fluorescence. Two germaria were imaged using quantitative fluorescence microscopy with UAS mCherry pHluorin and 10930 gal 4, stained with concanavalin A and ROI outlining of follicle stem cell or FSC or PFC and a follicle cell is shown in each image. The ratio of the fluorescence intensities in the GFP and mCherry channels is converted to pH I values, using a standard curve.The pH I increases with differentiation in the early FSC lineage, from 6.8 in FSCs to 7.0 in prefollicle cells to 7.3 in follicle cells. The pH I values of each cell type can be represented as a pseudocolored micrograph that shows the differences in pHluorin to mCherry ratios, display these differences in color as defined by an LUT. When attempting this procedure, it is important to remember to have all the reagents ready before hand because many steps in this protocol are time sensitive.After its development, this technique paved the way for researchers in the field of developmental biology, to explore the role of pH in sulfate decisions intracellular malano gaster.

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Developmental Biology

6.7K ビュー.  University of California, San Francisco. The overall goal of this procedure is to measure the intracellular pH of follicle cells within live explanted Drosophila ovarian tissue. This method can help us answer key questions in stem cell and developmental biology such as, how changes to intracellular pH contribute to sulfate decisions. The main advantage of this technique is that it makes it possible to accurately quantify pH within living intact tissue.To begin the experiment, prepare mounting chambers by applying a thin coati...

ビデオ: Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH...

extended live imaging of female drosophila melanogaster germline stem cell niches

Author Spotlight: Advancing Live Analysis of Drosophila Germline Stem Cell Niches

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein...

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

z-scores for assessing ovarian reserve in young patients undergoing fertility preservation

Author Spotlight: Advancing Fertility Assessment with Z-Score-Based Follicle Density Evaluation in Cryopreserved Ovarian Tissue

The present protocol describes a method for assessing ovarian reserve in patients under 25 years old who require fertility preservation through ovarian tissue cryopreservation. This method involves: (1) histological assessment of ovarian reserve in cortical samples, (2) comparison to a reference dataset, and (3) calculation of...

Assessing Ovarian Reserve in Young Patients Using Z-Scores for Follicle Density

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processing the ovarian tissue and determining ovarian reserve in young females through z-score follicle density analysis

Processing the Ovarian Tissue and Determining Ovarian Reserve in Young Females Through Z-Score Follicle Density Analysis

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

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optical quantification of intracellular ph in drosophila melanogaster malpighian tubule epithelia with a fluorescent genetically-encoded ph indicator

Optical Quantification of Intracellular pH in Drosophila melanogaster Malpighian Tubule Epithelia with a Fluorescent Genetically-encoded pH Indicator

Cellular ion transport can often be assessed by monitoring intracellular pH (pHi). Genetically Encoded pH-Indicators (GEpHIs)&#160;provide optical quantification of intracellular pH in intact cells. This protocol details the quantification of intracellular pH through cellular ex vivo live-imaging of Malpighian tubules of Drosophila melanogaster with pHerry, a pseudo-ratiometric genetically encoded...

Optical Quantification of Intracellular pH in Drosophila melanogaster Malpighian Tubule Epithelia with a Fluorescent Genetically-encoded pH Indicator

use of drosophila s2 cells for live imaging of cell division

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and...

Use of Drosophila S2 Cells for Live Imaging of Cell Division

live imaging of glial cell migration in the drosophila eye imaginal disc

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

live-imaging of the drosophila pupal eye

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging...

Live-imaging of the Drosophila Pupal Eye

a rapid, simple workflow for quantification of external adult drosophila structures

Author Spotlight: Investigating Intracellular pH Dynamics and Developing Accessible Imaging Protocols

Here, we present a rapid, low-cost, workflow for high-resolution imaging of adult Drosophila eyes to quantify patterning and growth defects. We describe our protocol for sample preparation by point-mounting, high-resolution image acquisition, and image...

High-Resolution Imaging and Analysis of Drosophila Eye Defects

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preparation of drosophila larval and pupal testes for analysis of cell division in live, intact tissue

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

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

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

preparing individual drosophila egg chambers for live imaging

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal...

Preparing Individual Drosophila Egg Chambers for Live Imaging

live imaging of drosophila larval neuroblasts

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously...

Live Imaging of Drosophila Larval Neuroblasts

live-cell imaging of drosophila melanogaster third instar larval brains

Author Spotlight: Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants 

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological...

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

a method for ovarian follicle encapsulation and culture in a proteolytically degradable 3 dimensional system

A Method for Ovarian Follicle Encapsulation and Culture in a Proteolytically Degradable 3 Dimensional System

A new method for ovarian follicle encapsulation in a 3D fibrin-alginate interpenetrating network is described. This system combines structural support with proteolytic degradation to support the development of immature follicles to produce mature oocytes. This method may be applied to culture cell aggregates to maintain cell-cell contacts without limiting...

live imaging of apoptotic cell clearance during drosophila embryogenesis

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...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

live-cell imaging of sensory organ precursor cells in intact drosophila pupae

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

the utility of stage-specific mid-to-late drosophila follicle isolation

The Utility of Stage-specific Mid-to-late Drosophila Follicle Isolation

Stage-specific isolation of mid-to-late Drosophila follicles is useful for a variety of purposes. Such follicles develop in culture, which allows for genetic and/or pharmacologic manipulations to be coupled with in vitro development assays and live imaging. Additionally, follicles can be used for molecular studies, such as isolating mRNA and...

The Utility of Stage-specific Mid-to-late Drosophila Follicle Isolation

live imaging followed by single cell tracking to monitor cell biology and the lineage progression of multiple neural populations

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations

A robust protocol to monitor neural populations by time-lapse video-microscopy followed by software-based post-processing is described. This method represents a powerful tool to identify biological events in a selected population during live imaging...

live imaging of gfp-labeled proteins in drosophila oocytes

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage.&#160;HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

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

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

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

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin rods are a hallmark of a conserved, inducible Actin Stress Response (ASR) that accompanies human pathologies, including neurodegenerative disease. Previously, we showed that the ASR contributes to morphogenesis failures and reduced viability of developing embryos. This protocol allows the continued study of mechanisms underlying actin rod assembly and the ASR in a model system that is highly amenable to imaging, genetics and biochemistry. Embryos are collected and mounted on a coverslip to prepare them for injection. Rhodamine-conjugated globular actin (G-actinRed) is diluted and loaded into a microneedle. A single injection is made into the center of each embryo. After injection, embryos are incubated at elevated temperature and intranuclear actin rods are then visualized by confocal microscopy. Fluorescence recovery after photobleaching (FRAP) experiments may be performed on the actin rods; and other actin-rich structures in the cytoplasm can also be imaged. We find that G-actinRed polymerizes like endogenous G-actin and does not, on its own, interfere with normal embryo development. One limitation of this protocol is that care must be taken during injection to avoid serious injury to the embryo. However, with practice, injecting G-actinRed into Drosophila embryos is a fast and reliable way to visualize actin rods and can easily be used with flies of any genotype or with the introduction of other cellular stresses, including hypoxia and oxidative stress.

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The goal of this protocol is to inject Rhodamine-conjugated globular actin into Drosophila embryos and image intranuclear actin rod assembly following heat stress.

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin ...

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, germ cell development, piRNA biology, and niche formation. Here, we present a dissection technique to live-image the gonad ex vivo during a period when in vivo live-imaging is highly ineffective. This protocol outlines how to transfer embryos to an imaging dish, choose appropriately-staged male embryos, and dissect the gonad from its surrounding tissue while still maintaining its structural integrity. Following dissection, gonads can be imaged using a confocal microscope to visualize dynamic cellular processes. The dissection procedure requires precise timing and dexterity, but we provide insight on how to prevent common mistakes and how to overcome these challenges. To our knowledge this is the first dissection protocol for the Drosophila embryonic gonad, and will permit live-imaging during an otherwise inaccessible window of time. This technique can be combined with pharmacological or cell-type specific transgenic manipulations to study any dynamic processes occurring within or between the&#160;cells in their natural gonadal environment.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, ...

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of the fly such as imaginal discs, the larval brain or egg chambers of adult females or the adult intestine can be extracted and kept in culture for imaging with time-lapse microscopy, providing valuable insights into cell and developmental biology. Here, we describe in detail our current protocol for the dissection of Drosophila larval brains, and then present our current approach for immobilizing and orienting larval brains and other tissues on a glass coverslip using Fibrin clots. This immobilization method only requires the addition of Fibrinogen and Thrombin to the culture medium. It is suitable for high-resolution time lapse imaging on inverted microscopes of multiple samples in the same culture dish, minimizes the lateral drifting frequently caused by movements of the microscope stage in multi-point visiting microscopy and allows for the addition and removal of reagents during the course of imaging. We also present custom-made macros that we routinely use to correct for drifting and to extract and process specific quantitative information from time-lapse analysis.

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

This protocol describes applications of sample immobilization using Fibrin clots, limiting drifting, and allowing the addition and washout of reagents during live imaging. Samples are transferred to a drop of Fibrinogen containing culture medium on the surface of a coverslip, after which polymerization is induced by adding thrombin.

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of ...

Development of Organoids from Mouse Pituitary as In Vitro Model to Explore Pituitary Stem Cell Biology

The pituitary is the master endocrine gland regulating key physiological processes, including body growth, metabolism, sexual maturation, reproduction, and stress response. More than a decade ago, stem cells were identified in the pituitary gland. However, despite the application of transgenic in vivo approaches, their phenotype, biology, and role remain unclear. To tackle this enigma, a new and innovative organoid in vitro model is developed to deeply unravel pituitary stem cell biology. Organoids represent 3D cell structures that, under defined culture conditions, self-develop from a tissue's (epithelial) stem cells and recapitulate multiple hallmarks of those stem cells and their tissue. It is shown here that mouse pituitary-derived organoids develop from the gland's stem cells and faithfully recapitulate their in vivo phenotypic and functional characteristics. Among others, they reproduce the activation state of the stem cells as in vivo occurring in response to transgenically inflicted local damage. The organoids are long-term expandable while robustly retaining their stemness phenotype. The new research model is highly valuable to decipher the stem cells' phenotype and behavior during key conditions of pituitary remodeling, ranging from neonatal maturation to aging-associated fading, and from healthy to diseased glands. Here, a detailed protocol is presented to establish mouse pituitary-derived organoids, which provide a powerful tool to dive into the yet enigmatic world of pituitary stem cells.

Development of Organoids from Mouse Pituitary as In Vitro Model to Explore Pituitary Stem Cell Biology

The pituitary gland is the key regulator of the body's endocrine system. This article describes the development of organoids from the mouse pituitary as a novel 3D in vitro model to study the gland's stem cell population of which the biology and function remain poorly understood.

The pituitary is the master endocrine gland regulating key physiological processes, including body growth, metabolism, sexual maturation, reproduction, ...