Name: methods for imaging intracellular ph of the follicle stem cell lineage in live drosophila ovarian tissue
Uploaded: 2017-09-26T14:00:00.000+00:00
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Description: 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

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 conditions1,7. Next, standard curves are generated by measuring pHluorin and mCherry fluorescence intensities in a Na+-free, K+ buffer containing the ionophore nigericin adjusted to two different pH values, 6.5 and 7.5. In the presence of nigericin, pHi equilibrates with the pH of the buffer across the plasma membrane, causing pHi to match the extracellular pH. Lastly, the standard curves are used to convert pHluorin to mCherry ratios to estimated pHi values.
1. Pre-Trial: Preparation Before Measuring pHi In Vivo
NOTE: To measure pHi in vivo, the mCherry::pHluorin transgene must be expressed in the cell type of interest. Below are some common ways to generate transgenic pHluorin flies in the FSC lineage. Generating mCherry::pHluorin clones is particularly useful for identifying FSCs, which are located on the anterior edge of an FSC clone. Tissue specific mCherry::pHluorin expression is useful for measuring pHi across the entire tissue and is also more convenient when combining with expression of an RNAi or transgene.
<ol>
	<li>Generation of transgenic pHluorin flies
	<ol>
		<li>mCherry::pHluorin-labeled FSC clones
		<ol>
			<li>To make mCherry::pHluorin FSC clones, cross UAS-mCherry::pHluorin1 to an Act-Gal4 flipout stock or other stock with a Flp/FRT inducible Gal4.</li>
			<li>To make heatshock inducible clones, heatshock adult F1s for 2 days post eclosion in empty vials for 1 h in a 37 &#176;C water bath, four times approximately every 12 h.</li>
			<li>To ensure normal growth and maximum rates of oogenesis during this period, maintain flies by providing fresh wet yeast (approximately equal parts dry baker&#39;s yeast and water) daily for at least 6 days after clone induction.</li>
		</ol>
		</li>
		<li>Tissue specific mCherry::pHluorin expression</li>
		<li>Cross UAS-mCherry::pHluorin1 to a follicle cell specific driver, y1w*;P{GawB}10930/CyO (Bloomington ID: 7023).</li>
		<li>3 days after flies start eclosing, collect and place them in vials containing wet yeast, for at least 24 h prior to dissection.</li>
	</ol>
	</li>
	<li>Making solutions 
	NOTE: It is important to prepare the following buffers on the day of the experiment because the pH and solute concentrations in the buffer can change over time.
	<ol>
		<li>Prepare bicarbonate, nigericin, and dissection buffers. Add the components in the order listed to avoid formation of precipitates (Refer to Table 1, Table 2, and Table 3).</li>
	</ol>
	</li>
</ol>
2. Trial: Measuring pHi in the FSC Lineage
NOTE: The dissection, mounting, and live imaging steps for the bicarbonate buffer and the two nigericin buffer conditions may need to be performed twice: once to determine the microscope settings and a second time to collect experimental data. See the section 2.2 below for more information.
<ol>
	<li>Dissection and mounting 
	NOTE: For materials required to perform this step please refer to Figure 1 and Table of Materials. For 3D printed mounting chambers, 3D printer files are provided as Supplemental File 1 and Supplemental File 2.
	<ol>
		<li>Dissection:
		<ol>
			<li>Prepare mounting chambers by applying a thin coating of vacuum grease to the flatter side of the 3D printed mounting chamber. Place that side on a 22 x 40 mM coverslip to seal the coverslip to the mounting chamber.</li>
		</ol>
		</li>
		<li>Dissect ovaries from female adult flies in 500 &#181;L dissection buffer (bicarbonate or nigericin buffer, Table 3)
		<ol>
			<li>Anesthetize 3 - 4 flies using CO2 gas and transfer flies onto a fly pad perfused with CO2 gas.</li>
			<li>Pick up an anesthetized fly with forceps and place into the dissection media.</li>
			<li>Pinch the thorax of the fly with one forcep, while tugging on the tip of its abdomen until its ovaries are exposed.</li>
			<li>After detaching the ovaries from the other organs, tease apart the ovarioles using 221/2 syringe needles. Carefully separate the muscle sheath from the ovaries and separate individual ovarioles from the cluster of ovarioles. An effective technique to isolate the muscle sheath is to 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.</li>
			<li>Incubate the dissected ovaries in dissection media for exactly 10 min before proceeding to the mounting step so that there is enough time for the pHi of cells to fully equilibrate with the pH of the nigericin dissection media. 
			NOTE: Ensure that the muscle sheath is removed from the ovarioles. The muscle sheath may attenuate the fluorescence signal, which makes it difficult to identify individual cells using Concanavilin-A, and may cause ovarioles to move spontaneously during live imaging.</li>
		</ol>
		</li>
		<li>Mounting
		<ol>
			<li>Place a small drop of dissection media onto the glass coverslip inside the mounting chamber.</li>
			<li>Transfer separated ovarioles using forceps.</li>
			<li>Separate later stage egg chambers from germaria associated with earlier stage egg chambers, and try to ensure that the dissected germaria are placed at the center of the dissection media drop.</li>
			<li>Add two small drops of nail polish to opposite sides of a round 12-mm coverslip and let it air dry for about 10 s.</li>
			<li>Place the round coverslip on the drop of dissection media containing separated germaria such that that side with nail polish faces down and contacts the dissection media. Press down on the parts of the edge with nail polish to flatten and secure the germaria in position. We recommend using older nail polish because it smears less over the surface of the coverslip as it is being secured in place.</li>
			<li>Fill the mounting chamber with additional dissection media. Condition specific buffer that does not contain Concanavalin-A dye can be used to fill the chamber. 
			NOTE: The total time spent dissecting and mounting should not exceed 15 min. This is important to minimize the effects of tissue damage and cell death.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Live imaging: 
	NOTE: In this step, first collect images from the bicarbonate condition and the two nigericin conditions to determine the proper microscope settings; adjust the microscope settings as necessary, and then collect images for the experimental data. Use a confocal microscope capable of imaging in 3 channels: GFP (475 excitation/509 emission), mCherry (575 excitation/610 emission), and far-red (633 excitation/647 emission). 
	NOTE: Set Microscope Settings: The pixel intensities of the images collected will be quantified to determine the estimates of pHi, so it is important that the intensity values of the signal in each image set is neither too low nor saturated. The range of intensities that can be captured in an image is defined by the image bit depth. In many cases, 8-bit images are generated by default, but we recommend acquiring the data as 16-bit images, which have a broader dynamic range. It is important to ensure that the crosstalk between fluorescent channels is minimized, so the settings should be adjusted so that the emission spectrum collected from each fluorophore does not overlap. To test these settings, take a sample image using the excitation spectrum for GFP and collect in the emission spectrum for mCherry and vice versa. If the settings have been adjusted properly, there should be no signal in either case. Set the microscope settings (i.e., voltage settings on a laser scanning confocal, laser power and pinhole size) so that the pixel intensities of the signal in all three image sets are within the dynamic range of the image file. For all our experiments, we used a white light laser scanning confocal microscope with a 40x objective to acquire 16-bit images in 1,024 &#215; 1,024 format, while we optimized voltage gain for every experiment as needed. For example, in one experiment, we set the voltage gain for GFP, mCherry, and far-red as 37.8%, 72.9%, and 260%, respectively.
	<ol>
		<li>Image ovarioles in nigericin dissection buffer at pH 6.5, and adjust settings such that pixel intensities of the pHluorin and mCherry images are low, but not below the limits of detection of the camera.</li>
		<li>Image another set of ovarioles in nigericin 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.</li>
		<li>Image ovaries in bicarbonate dissection buffer and ensure that the pixel intensities of the pHluorin and mCherry images are not saturated with the chosen settings. If the pixels are saturated, adjust the settings as necessary. 
		NOTE: Microscope setting parameters may differ based on the exact microscope setup being used, and must be optimized for each experiment. After microscope settings have been set, do not change them for the remainder of the experiment. Settings should be consistent across control and experimental conditions. In our initial pHi experiments, we generated 2 and 3 point nigericin calibration curves and found no significant difference between calculated pHi using either method. However, in general, the addition of more points in the calibration curve would be expected to improve accuracy. Therefore, we recommend generating curves with different numbers of points to determine how many are needed for a given set of experimental conditions.</li>
	</ol>
	</li>
	<li>Data collection:
	<ol>
		<li>Perform dissection, mounting, and imaging of the samples in the bicarbonate and nigericin buffer conditions. After dissection and mounting, the maximum time spent imaging one set of samples should not exceed 45 min to ensure that the fluorescence intensity measurements are made in live, healthy tissue.</li>
		<li>For each condition, acquire images for at least 5 germaria.</li>
	</ol>
	</li>
</ol>
3. Post-trial: Image Analysis
<ol>
	<li>Measure fluorescence intensities in FSCs, pFCs, and follicle cells
	<ol>
		<li>Background subtraction:
		<ol>
			<li>Open unprocessed images in FIJI with each channel in a separate window (Figure 4C).</li>
			<li>Use the rectangle tool to draw a region of interest (ROI) in the pHluorin channel window in a part of the image without signal.</li>
			<li>Optional step: Set the lower limit of the threshold so that pixels with intensity values below 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 (Figure 4A).</li>
			<li>Measure the mean fluorescence intensity in the ROI. If the threshold was set in step 3, make sure to check the &#34;Limit to threshold&#34; box in the Set Measurements dialog box.</li>
			<li>Subtract the measured background intensity from each channel and slice of the image using the Subtract function, found in the Process &#8594; Math menu.</li>
			<li>Repeat steps 3.1.1.2-6 for the mCherry channel window except that, in step 3.1.1.2, instead of drawing a new rectangle with the rectangle tool, use the Restore selection function, found in the Edit&#8594; Selection menu, to add a rectangle with the same dimensions and position on the image.</li>
		</ol>
		</li>
		<li>Identify FSCs, pFCs, and follicle cells:
		<ol>
			<li>Identify the FSCs, pFCs, and follicle cells by morphology and location using the Concanavalin-A staining to find cell boundaries (Figure 2).</li>
			<li>FSCs are thin triangular cells located on the edge of the germarium at the Region 2a/2b border. If mCherry::pHluorin is expressed in an FSC clone, FSC can also be identified as the anterior most cell of the clone.</li>
			<li>pFCs are irregular shaped cells in Region 2b, immediately adjacent to and downstream of FSCs.</li>
			<li>Follicle cells are square or columnar cells that surround the germ cell cyst in Region 3.</li>
		</ol>
		</li>
		<li>Obtaining fluorescence intensity ratios
		<ol>
			<li>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, seen in the far-red channel, is in focus within the selected slice.</li>
			<li>Draw ROI around each FSC and measure the mean fluorescence intensity of pHluorin and mCherry in all slices chosen for measurements.</li>
			<li>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.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Derive pHi values and generate pseudocolored images
	<ol>
		<li>Derive pHi values
		<ol>
			<li>In all cases, the linear regression curve should be generated using ovaries from flies of the same genotype for both experimental and control conditions. Generate a linear regression curve for each cell type using the data from two nigericin buffer conditions (Figure 3). In Excel, this can be done by plotting the data on a graph and adding a linear trendline. Alternately, see:</li>
			<li>Use the slope and the y-intercept of the linear regression curve calculated from the nigericin conditions and the equation of a line (y = mx + b) to convert the pHluorin to mCherry ratio values calculated from the samples in the bicarbonate condition to pH values. In this equation, substitute the slope form (the y-intercept) for b, put the ratio value in for x, and solve for y.</li>
		</ol>
		</li>
		<li>Generate a pseudocolored image reflecting the ratio values in FIJI.
		<ol>
			<li>Follow the procedure above for background subtraction (step 3.1.1 and Figure 4).</li>
			<li>Divide the pHluorin channel from the mCherry channel using the Image Calculator function, found in the Process menu. Be sure the &#34;Create new window&#34; and &#34;32-bit (float) result&#34; checkboxes are both checked. Change the lookup table (LUT), found under the Image menu, to Thermal or LUT of choice. See Figure 4 for other suitable options.</li>
			<li>In the adjust brightness and contrast dialog box (Image &#8594; Adjust &#8594; Brightness/Contrast), click the &#34;Set&#34; button. Set the Minimum displayed value to 0 and the Maximum displayed value to a ratio that best captures the data, usually 1.0-2.0 (Figure 5).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

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>&lt;strong&gt;Fly Stocks&lt;/strong&gt;</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>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Chemicals for Buffer preparation&lt;/strong&gt;</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>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Dissection and mounting tools&lt;/strong&gt;</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>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Other equipment&lt;/strong&gt;</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&amp;times; 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 ...

methods-for-imaging-intracellular-ph-follicle-stem-cell-lineage-live

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

6.8K Views.  University of California, San Francisco. 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.

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

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

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

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 pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Neuroscience

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

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 quantified.

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

Genetics

Visualizing the Effects of Oxidative Damage on Drosophila Egg Chambers using Live Imaging

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ribonucleoprotein particle movement, mRNA localization, organelle movement, and cytoskeletal dynamics. There are several methods for live imaging that have been developed. Due to the fact that each method involves dissecting individual ovarioles placed in media or halocarbon oil, cellular damage due to hypoxia and/or physical manipulation will inevitably occur over time. One downstream effect of hypoxia is to increase oxidative damage in the cells. The purpose of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries after induction of controlled cellular damage. Here, we use hydrogen peroxide to induce cellular oxidative damage and give examples of the effects of such damage on two subcellular structures, mitochondria and Clu bliss particles. However, this method is applicable to any subcellular structure. The limitations are that hydrogen peroxide can only be added to aqueous media and would not work for imaging that uses halocarbon oil. The advantages are that hydrogen peroxide is readily available and inexpensive, acts quickly, its concentrations can be modulated, and oxidative damage is a good approximation of damage caused by hypoxia as well as general tissue damage due to manipulation.

The objective of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries.

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ...

Nuclear Migration in the Drosophila Oocyte

Live cell imaging is particularly necessary to understand the cellular and molecular mechanisms that regulate organelle movements, cytoskeleton rearrangements, or polarity patterning within the cells. When studying oocyte nucleus positioning, live-imaging techniques are essential to capture the dynamic events of this process. The Drosophila egg chamber is a multicellular structure and an excellent model system to study this phenomenon because of its large size and availability of numerous genetic tools. During Drosophila mid-oogenesis, the nucleus migrates from a central position within the oocyte to adopt an asymmetric position mediated by microtubule-generated forces. This migration and positioning of the nucleus are necessary to determine the polarity axes of the embryo and the subsequent adult fly. One characteristic of this migration is that it occurs in three dimensions (3D), creating a necessity for live imaging. Thus, to study the mechanisms that regulate nuclear migration, we have developed a protocol to culture the dissected egg chambers and perform live imaging for 12 h by time-lapse acquisitions using spinning-disk confocal microscopy. Overall, our conditions allow us to preserve Drosophila egg chambers alive for a long period of time, thereby enabling the completion of nuclear migration to be visualized in a large number of samples in 3D.

Nuclear Migration in the Drosophila Oocyte

In Drosophila, the oocyte nucleus undergoes microtubule-dependent migration during oogenesis. Here, we describe a protocol that was developed to follow the migration by performing live imaging on egg chambers ex-vivo. Our procedure maintains egg chambers alive for 12 h to acquire multi-position 3D time-lapse movies using spinning-disk confocal microscopy.

Live cell imaging is particularly necessary to understand the cellular and molecular mechanisms that regulate organelle movements, cytoskeleton ...

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

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

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

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 Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

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 described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Biology

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 ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

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

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Rozdziały w tym wideo

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

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Visualizing the Effects of Oxidative Damage on Drosophila Egg Chambers using Live Imaging

Nuclear Migration in the Drosophila Oocyte

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Imaging Through the Pupal Case of Drosophila melanogaster

Studying Mitochondrial Structure and Function in Drosophila Ovaries