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

Summary

Cellular ion transport can often be assessed by monitoring intracellular pH (pH_i). Genetically Encoded pH-Indicators (GEpHIs) 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 pH-indicator.

Abstract

Epithelial ion transport is vital to systemic ion homeostasis as well as maintenance of essential cellular electrochemical gradients. Intracellular pH (pH_i) is influenced by many ion transporters and thus monitoring pH_i is a useful tool for assessing transporter activity. Modern Genetically Encoded pH-Indicators (GEpHIs) provide optical quantification of pH_i in intact cells on a cellular and subcellular scale. This protocol describes real-time quantification of cellular pH_i regulation in Malpighian Tubules (MTs) of Drosophila melanogaster through ex vivo live-imaging of pHerry, a pseudo-ratiometric GEpHI with a pK_a well-suited to track pH changes in the cytosol. Extracted adult fly MTs are composed of morphologically and functionally distinct sections of single-cell layer epithelia, and can serve as an accessible and genetically tractable model for investigation of epithelial transport. GEpHIs offer several advantages over conventional pH-sensitive fluorescent dyes and ion-selective electrodes. GEpHIs can label distinct cell populations provided appropriate promoter elements are available. This labeling is particularly useful in ex vivo, in vivo, and in situ preparations, which are inherently heterogeneous. GEpHIs also permit quantification of pH_i in intact tissues over time without need for repeated dye treatment or tissue externalization. The primary drawback of current GEpHIs is the tendency to aggregate in cytosolic inclusions in response to tissue damage and construct over-expression. These shortcomings, their solutions, and the inherent advantages of GEpHIs are demonstrated in this protocol through assessment of basolateral proton (H⁺) transport in functionally distinct principal and stellate cells of extracted fly MTs. The techniques and analysis described are readily adaptable to a wide variety of vertebrate and invertebrate preparations, and the sophistication of the assay can be scaled from teaching labs to intricate determination of ion flux via specific transporters.

Introduction

The goal of this protocol is to describe quantification of intracellular pH (pH_i) using a Genetically-Encoded pH-Indicator (GEpHI) and demonstrate how this method can be used to assess basolateral H⁺ transport in a model insect (D. melanogaster) renal structure, the Malpighian tubule (MT). MTs serve as the excretory organs of the fruit fly and are functionally similar to the mammalian nephron in several key respects¹. MTs are arranged as 2 pairs of tubules (anterior and posterior) in the thorax and abdomen of the fly. The single-cell epithelial tube of each MT is composed of metabolically active principal cells with distinct apical (luminal) and basolateral (hemocoel) polarity as well as intercalated stellate cells. Anterior MTs are composed of 3 morphologically, functionally, and developmentally distinct segments, notably the initial dilated segment, transitional segment, and secretory main segment, which joins to the ureter². At the cellular scale trans-epithelial ion transport into the lumen is accomplished by an apical plasma membrane V-ATPase³ and an alkali-metal/H⁺ exchanger as well as a basolateral Na⁺-K⁺-ATPase⁴, inward-rectifier K⁺ channels⁵, Na⁺-driven Cl⁻/HCO₃⁻ exchanger (NDAE1)⁶, and Na⁺-K⁺-2Cl⁻ cotransporter (NKCC; Ncc69)⁷, while stellate cells mediate Cl^- and water transport^8,9. This complex but accessible physiologic system provides excellent opportunities for investigation of endogenous ion transport mechanisms when combined with the diverse genetic and behavioral toolsets of Drosophila.

The rationale for this protocol was to describe a genetically malleable system for studying epithelial ion transport with potential for integration from cell to behavior and export of tools to other model systems. Expression of pHerry¹⁰, a GEpHI derived from a fusion of green pH-sensitive super-ecliptic pHluorin^11,12 (SEpH) and red pH-insensitive mCherry¹³, in MTs permits quantification of H⁺ transport in single MT cells through the high K⁺/nigericin calibration technique¹⁴. As many ion transporters move H⁺ equivalents, quantification of intracellular pH_i serves as a functional representation of ion movement via a variety of transporters. The Drosophila MT model system also offers powerful genetic tools in tissue-specific transgene¹⁵ and RNA interference (RNAi)¹⁶ expression, which can be combined with cellular imaging and whole-organ assays^17,18,19 of tubule function to create a robust toolset with vertical integration from molecules to behavior. This stands in contrast to many other protocols for assessing epithelial biology, as historically such measurements have relied on intricate and daunting micro-dissection, sophisticated ion-selective electrodes^20,21, and expensive pH-sensitive dyes²² with restrictive loading requirements and poor cellular specificity in heterogeneous tissues. GEpHIs have been used to extensively measure pH_i in a variety of cell types²³. Early work exploited the inherent pH-sensitivity of Green Fluorescent Protein (GFP) to monitor pH_i in cultured epithelial cells²⁴ but the past two decades have seen GEpHIs used in neurons²⁵, glia²⁶, fungi²⁷, and plant cells²⁸. The combination of the potential for cellular targeting of genetic constructs through the GAL4/UAS expression system¹⁵ and the physiologic accessibility of the Drosophila MT make this an ideal preparation for investigations of pH_i regulation and epithelial ion transport.

pH_i regulation has been studied for decades and is vital to life. The MT preparation offers a robust model to teach physiology of pH_i regulation but also perform sophisticated investigations of pH_i regulation ex vivo and in vivo. This protocol describes quantification of H⁺ movement across the basolateral membrane of the epithelial cells of the Drosophila MT using the NH₄Cl pulse acid loading technique²¹, but as the pH-indicator is genetically encoded, these methods and their theoretical framework can be applied to any preparation amenable to transgenesis and live-imaging.

Protocol

All steps in this protocol comply with the Mayo Clinic (Rochester, MN) animal use guidelines.

1. Fly Husbandry

1. Raise flies and set crosses according to standard husbandry²⁹.
   NOTE: Fluorescent reporter expression by the GAL4/UAS system is proportional to temperature and thus rearing temperature can be adjusted to alter expression level. While high expression levels often lead to a better signal-to-noise ratio this condition is also associated with increased cytosolic and organellar aggregates when using GFP to Red Fluorescent Protein (RFP) fusion constructs such as pHerry^10,30,31. If aggregation is unavoidable, quantification is still possible by performing point calibrations in each experiment and normalizing data such that a fluorescence ratio of 1.0 corresponds to pH_i 7.0 (see step 7.4 Note on calibration below).
2. Set crosses of homozygous capaR-GAL4³² males to homozygous UAS-pHerry¹⁰ virgin females and homozygous c724-GAL4² males to homozygous UAS-pHerry virgin females to permit imaging of pH_i in principal cells and stellate cells of the MT, respectively. Place 6 UAS-pHerry females with 3 GAL4 males into fresh vials of food and leave to mate at 28 °C.
   NOTE: Larvae should be evident within 4 d and adults will begin to eclose around day 10.
3. Collect female flies upon eclosion and set aside to age for 10 d at 28 °C.
   NOTE: The timing of experimentation can be adjusted to correspond to any restrictive behavioral assays (such as the Ramsay secretion assay^17,19) that will be correlated to intracellular live pH imaging. Male flies can be used but tubules from females are often larger and more robust.

2. Preparation of Poly-L-Lysine Slides.

1. Draw a 40 x 20 mm border with a hydrophobic PAP pen around the top of standard 75 x 25 mm slides and set aside to dry for 15 min at RT. Use large coverslips if the slides are not compatible with imaging optics.
2. Transfer 2 mL of stock 0.01% Poly-L-Lysine (PLL) solution onto each slide and set aside for 1 h at RT.
3. Remove excess PLL with a pipette. Save the solution in a 50 mL conical vial for future use. Store at 4 °C.
4. Aspirate any remaining solution with a vacuum line. Run the vacuum line over the entire slide surface to ensure that no solution remains on the slides.
5. Set the slides aside for 1 additional h at RT prior to use. Store the slides dry at RT for up to 1 month in a standard slide book.

3. Preparation of Dissecting Dish and Glass Rods

1. Add 0.5 mL of elastomer curing agent to 4.5 mL of elastomer base in a 35 x 10 mm polystyrene Petri dish at RT to produce a depth of 5 mm. Mix with a disposable pipette tip. Allow elastomer to cure O/N at RT.
   NOTE: Elastomer should be clear and free of bubbles. Clearing of bubbles may be facilitated by keeping elastomer plates in a vacuum jar for 10 - 15 min after pouring.
2. Hold a 5 mm diameter glass rod between hands and melt the center of the rod over a lit Bunsen burner while pulling the ends apart. As the glass melts pull more quickly to produce a thin (0.1 mm) and tapered shaft (Figure 1).
   NOTE: A 45 ° angle at the shank is often helpful in handling tubules. This can be achieved by lowering one hand as the shank is pulled (see Figure 1).
3. Break the thin shaft in the middle with the blunt side of a single-edged carbon steel razor blade. Inspect the thin end of the rod under a dissecting scope to ensure the break is clean.

Figure 1: Fabricating Glass Rods for Handling Malpighian Tubules.
A - E. Process of heating and pulling a glass rod to produce a taper and angle suitable for handling MTs. Arrows denote direction and magnitude of force to be applied. F. Photograph of an appropriately fabricated glass tool. Scale bar = 10 mm. Please click here to view a larger version of this figure.

4. Preparation of Solutions and Perfusion System

NOTE: Perfusion systems differ by manufacturer. This protocol is based around a gravity-fed 8-channel open reservoir with an input flow rate regulator and a vacuum-driven outflow, but the method of mounting MTs as described here can be adapted to work with any perfusion system.

1. Prepare the following solutions:
   1. Aliquot Schneider's medium (40 mL into 50 mL conical vials) and store at 4 °C.
   2. Prepare solutions (i.e. insect Phosphate-Buffered Saline (iPBS) and iPBS with NH₄Cl) at RT as needed according to Table 1). Warm solutions to RT before use on day of experiment.
      NOTE: iPBS and iPBS with 40 mM NH₄Cl can be prepared in large volumes (1 L or more) and stored at 4 °C.
   3. Prepare 8 calibration solutions in 500 mL volumes at pH = 5.0, 6.0, 6.5, 7.0, 7.3, 7.6, 8.0, and 9.0 as indicated in Table 1 and store at 4 °C. Adjust pH of each solution by titration with N-methyl-D-glucamine (NMDG) and HCl.
   4. On the day of experiments, warm 5 mL aliquots of calibration solutions to RT and add stock nigericin solution (20 mM in dimethyl sulfoxide (DMSO)) to produce a final concentration of 10 µM.
      CAUTION: Handle nigericin with gloves. Treat all equipment that comes in contact with nigericin as disposable. Nigericin remains on glass and plastic and will compromise biological preparations if equipment is reused.
2. Perfusion system:
   1. Prime the perfusion system by filling all reservoirs with ddH₂O (Figure 2). Open the channels one at a time to allow all lines proximal to the flow rate regulator to fill.
      NOTE: It may be necessary to clear air in the lines by opening the stalled channel and using a plunger to drive flow from the reservoir.
   2. Open 2 channels and allow ddH₂O to drain. Once the reservoirs are nearly empty, fill the first reservoir with iPBS and the second reservoir with NH₄Cl-pulsed iPBS. Set flow rate to maximum with the flow rate regulator and allow each solution to flow for 1 min to fill the distal lines, then stop the flow (Figure 2).
   3. Position 2 sets of soldering "helping hands" clamps on the imaging microscope stage. Place one clamp on each side of the imaging platform.
   4. Carefully heat the distal 0.5 inches of a piece of capillary glass (inner diameter 1.5 mm, outer diameter 0.86 mm, length 100 mm) over a Bunsen burner. Create a 45 ° bend by allowing the distal end to bend by gravity and removing the glass from the flame once the desired angle is achieved. Repeat this process with a second piece of capillary glass.
   5. Insert the bent glass capillaries in the in-flow line and vacuum-connected outflow line, respectively and mount them in the "helping hands" to align them with the imaging stage of the microscope (Figure 3).

Figure 2: Perfusion System and Imaging Configuration.
Components necessary for the physiological assessment of MT basolateral transport function through simultaneous live fluorescence imaging and rapid solution exchange.Gas lines shown are optional and permit expansion of experiments to the assessment of HCO₃^- transport. Please click here to view a larger version of this figure.

Figure 3: Flow Schematic of Perfusion Apparatus for NH₄Cl Pulse Experiments.
Arrows depict flow path and valve switching points. Solution moves from reservoir to specimen by gravity flow and is drawn from the specimen chamber to the waste flask by vacuum suction. Please click here to view a larger version of this figure.

5. Dissection of Adult Drosophila Anterior Malpighian Tubules.

1. Gather the dissection dish and pulled glass rod from section 3, a PLL-coated slide from section 2, an adhesive perfusion-well divider, vacuum grease, a 4 x 2" strip of sealing film, 2 pairs of #5 fine forceps, and 40 mL aliquots of ice-cold Schneider's medium and RT iPBS.
2. Spread vacuum grease on the sealing tape and press the adhesive perfusion-well divider onto the tape to coat the bottom with grease. Peel off the adhesive perfusion-well divider and place it grease-side-down on top of a PLL coated slide. Remove the perfusion-well divider to leave individual specimen wells traced in hydrophobic grease.
3. Place 200 µL of RT iPBS in the grease-encircled well on the PLL coated slide and move the slide under the stereoscope.
4. Place UAS-pHerry/capaR-GAL4 flies in an empty fly vial and anesthetize them on ice for 10 min.
   NOTE: This method of anesthesia, unlike CO₂, ensures that the flies do not dehydrate.
5. Pour ice-cold Schneider's medium into the dissecting dish and use fine forceps to transfer a single anesthetized female fly into the dish under a dissecting stereoscope.
6. Hold the fly by the thorax with a set of forceps and use the other to gently grip the posterior of the abdomen. Pull open the posterior of the fly using the forceps in short, deliberate motions. Once the hindgut is visible, grip the distal end and free the gut and MTs from the underlying tracheoles by pulling the hindgut away from the body through repetitive, brief tugs.
   NOTE: The anterior and posterior MTs will be visible where they meet the junction of the midgut and hindgut through the ureter. The first pair of MTs to be free will likely be the posterior tubules as they encircle the hindgut. These can be ignored (Figure 4A).
7. Pinch off the anterior MTs at the ureter with fine forceps once the second set of MTs is free of the abdomen. This will separate the anterior MTs from the gut and close the ureter.
8. Pick up the free anterior MTs with the pulled glass rod by sliding the rod under the ureter such that the tubules fall to either side. Lift the MTs straight up out of the solution.
9. Turn the glass rod such that the MTs and ureter are adhered to the underside of the rod and lower the ureter straight down onto the slide. Affix the ureter and seal the distal ends of the MTs by pressing the ureter down onto the glass slide (Figure 4B). Do not manipulate the MTs any more than necessary. The MTs should be floating up in the solution with the ureter anchored to the slide.
10. Use the fine end of the glass rod to gently sweep each tubule across the slide surface. Brace the rod against the slide to avoid crushing the tubule and slide the rod over the top of the tubule, moving distal to proximal, to attach the full length of each tubule to the surface of the PLL-coated slide (Figure 4C).
11. Place the adhesive perfusion-well divider back on the slide to form a small fluid-filled well over the mounted tubule.
12. Place the specimen on the microscope stage. Position the inflow and outflow capillaries over the inlet and outlet opening of the perfusion well, respectively.
    NOTE: The well divider can be left off if an open perfusion chamber is desired. In this case the inflow and outflow capillaries can be aligned to opposite sides of an imaging well.

6. Validation of Imaging Protocol and Tubule Health

Note: This protocol is performed on an inverted wide-field epifluorescent microscope with GFP (SEpH) and RFP (mCherry) filter sets (470/40 nm excitation (ex), 515 nm longpass emission (em), 500 nm dichroic and 546/10 nm ex, 590 nm longpass em, 565 nm dichroic), a 10X/0.45 air objective, a monochromatic camera for live-image capture, and imaging software. The protocol can be adapted for any upright or inverted microscope with automated filter switching between GFP and RFP optics and image acquisition software, although optimal exposure times, light intensity, and binning parameters will vary. In all analysis, the fluorescence intensity should be analyzed as mean pixel intensity in the region of interest (ROI), after background subtraction in each channel using an ROI with contains no fluorescence adjacent to the signal ROI.

1. Turn on the microscope, light source, and imaging system.
2. Open associated imaging software.
3. Look through the eyepiece and manually adjust focus until the lumen of the MT is clearly visible under transmitted light.
4. Click the "Acquisition" tab in image analysis software and select "2x2" in the "Binning" pull-down menu in the "Acquisition Mode" section.
5. Insert a 5% neutral density filter into the light path to reduce illumination light and minimize photobleaching.
6. Click the GFP (SEpH) channel in the "Channels" menu, then click "Live" to observe the fluorescent signal via the camera.
7. Adjust the "Time" slider to set the exposure time such that the brightest pixel values in the intensity histogram are approximately 40% of the maximum value, then click "Stop" to stop the illumination.
8. Repeat steps 6.6 - 6.7 in the RFP (mCherry) channel and confirm the presence of the dilated initial segment of the anterior MT and the absence of cytosolic mCherry aggregates (indicative of tissue damage or overexpression) (Figure 4D).
   NOTE: The dilated segment should be clearly apparent as it is the most proximal segment of the tubule and the diameter of the inner lumen of this segment is ~20 µm greater than that of the adjacent transitional segment. 2 x 2 pixel binning is often sufficient but can be increased to further reduce the required illumination intensity. Typical exposure times are between 150 & 800 ms/channel. Use as little light as possible to minimize photobleaching. Minimizing photobleaching is vital to the use of dual-fluorophore indicators such as pHerry as the two fluorophores can bleach independently, thus invalidating any ratio calibration.
9. Enable a time-lapse imaging protocol by clicking the "Time Series" check box.
10. Adjust the "Duration" in the pull-down menu in the "Time Series" section to 10 min and the "Interval" slider to 0 to set the total capture time with a maximum image acquisition rate. A total acquisition rate of 0.2 Hz is often sufficient.
11. Check both the GFP (SEpH) and RFP (mCherry) boxes in the "Channels" section.
12. Open the iPBS line of the perfusion system by activating the appropriate valve controller and start the imaging protocol by clicking "Start Experiment." After 1 min, switch to NH₄Cl pulse solution for 20 s by opening the appropriate valve and closing the iPBS line, then return to iPBS by closing the NH₄Cl line and reopening the iPBS valve. Allow the full imaging protocol to complete before stopping the perfusion system.
    NOTE: Time-lapse analysis should reveal a stable mCherry signal and a SEpH signal which increases in the presence of NH₄Cl, quenches upon washout, and gradually recovers.
13. Perform a 2-point calibration.
    1. Remove the well divider by peeling it away from the underlying slide and remove the perfusion capillaries and clamps from the imaging well.
    2. Apply 200 µL Calibration iPBS (pH 7.4, 10 µM nigericin) to the imaging well with a 200 µL pipette. Remove the solution from the imaging well with the pipette, then replace with another 200 µL of calibration solution. Repeat this process 4 times to ensure complete solution exchange.
    3. Incubate the preparation in calibration solution for 30 min before imaging. Repeat the imaging protocol using the same parameters determined in steps 6.6 - 6.11, with the modification of only 1 min of image capture.
       NOTE: The perfusion system and capillaries are not needed in this step and should not be attached to the imaging well to avoid exposing the capillaries to nigericin.
    4. Add 200 µL Calibration iPBS (pH 9.0, 10 µM nigericin) to the imaging well with a 200 µL pipette. Remove the solution from the imaging well with the pipette, then replace with another 200 µL of calibration solution. Repeat this process 4 times to ensure complete solution exchange.
    5. Incubate the preparation in the second calibration solution for 10 min before imaging. Repeat the imaging protocol as in step 6.13.3.
    6. Review the captured image stack in image analysis software to confirm that no pixels in either channel are saturated by clicking "Mean ROI" and scrolling though the image stack with the "Frame" slider while observing that no values reported in the intensity histogram reach the maximum detectable value. If any frames contain pixels that reach the maximum detectable intensity, reduce the exposure time or illumination intensity and repeat section 6.
       Note: Once established do not change imaging parameters between experiments or calibration unless point calibrations are to be used in each preparation (see step 8.3).
14. Analyze the image stack to plot fluorescent intensity and fluorescence ratio (SEpH/mCherry) as a function of time.
    1. Click "Mean ROI" and select the freeform tool. Hold left-click to trace a ~50 µm length of MT. Right click to finish drawing the ROI, then repeat in an area adjacent to the MT to define a background ROI (Figure 5A).
    2. Click "Mean Intensity" under "Measurements." Create a table of intensity values by clicking "Export > Data Table > Create."
    3. Click the configuration cogwheel icon and de-select all parameters except "Time" and "Mean Intensity." Right-click the tab for the newly created data table, select "Save As", and export the data as a .csv file.
       NOTE: Similar measurements can also be made using free software such as ImageJ.
    4. Open a spreadsheet table and import the data table by selecting the "Data" tab followed by "From Text".
    5. Use functions in the spreadsheet to subtract the SEpH Background Intensity from the SEpH Signal Intensity at each time point. Repeat this process for the mCherry signal.
    6. Plot each channel intensity as a function of time by selecting the columns containing the Time and background-corrected Intensity data and then clicking "Insert > Scatter (Charts) > Scatter with Straight Lines" (Figure 5B).
    7. Use spreadsheet functions to calculate the SEpH/mCherry fluorescence ratio at each time point.
    8. Plot fluorescence ratio as a function of time by selecting the columns containing the Time and ratio data and then clicking "Insert > Scatter (Charts) > Scatter with Straight Lines" (Figure 5C).

7. Full Calibration of pHerry in Malpighian Tubules Ex Vivo.

1. Dissect and mount a fresh set of anterior MTs as described in section 5.
2. Exchange iPBS for Calibration iPBS (pH 7.4, 10 µM nigericin) as described in step 6.13.2. Incubate for 30 min.
3. Locate the MTs and collect pairs of SEpH/mCherry images as described in steps 6.1 - 6.11. Replace the solution with another stock of Calibration iPBS as described in step 6.13.4, wait 10 min, and image again. Repeat this process until the SEpH/mCherry ratio has been imaged in all solutions. Obtain pH 9.0 images last as the specimen rarely recovers from high pH.
4. Plot fluorescence ratio of SEpH to mCherry from calibrations in eight specimens as a function of imposed pH_i as described in step 6.14.9. Fit the calibration data with a Boltzmann curve to obtain full calibration function according to Equation 1 (Figure 5D). If data are inconsistent, plot calibration sets from each specimen normalized such that a fluorescence ratio of 1.0 corresponds to pH_i 7.0 and re-analyze (Figure 5E).
   NOTE: If the latter process is necessary individual experiments will need their own internal point calibrations³³ (see quantification procedure below (step 8.3)).
5. Equation 1
   
   Where R = SEpH/mCherry ratio and A₁, A₂, x_o, and dx are curve fitting parameters representing minimum fluorescence ratio, maximum fluorescence ratio, pK_a, and width of the function respectively. x_o = apparent pK_a of pHerry, which can vary between 7.1 and 7.4 depending on the cell type and exact calibration conditions.

8. Quantification of Basolateral Acid Extrusion from Ex Vivo Malpighian Tubule Epithelia.

1. Image pHerry-expressing stellate cells and pHerry-expressing principal cells simultaneously.
   1. Dissect anterior MTs from a UAS-pHerry/capaR-GAL4 fly as described in section 5, but do not transfer MTs from the dissecting Schneider's medium to the imaging well.
   2. Dissect anterior MTs from a UAS-pHerry/c724-GAL4 fly in the same dissecting dish using the procedure outlined in section 5.
   3. Transfer the 2 sets of MTs into the same imaging well as described in steps 5.8 - 5.11.
      NOTE: When sweeping the arms of the MTs down to the slide, place the MTs of the UAS-pHerry/c724-GAL4 and the UAS-pHerry/capaR-GAL4 tubules near each other so pHerry-expressing principal and stellate cells can be visualized in the same field (Figure 6A).
2. Apply the NH₄Cl prepulse as described in step 6.12.
   NOTE: If consistent calibration (Figure 4B) could not be achieved, perform a point calibration by setting pH_i to 7.0 at the end of each experiment with Calibration iPBS (pH 7.0, 10 µM nigericin, 30 min incubation) after the adhesive perfusion-well divider and the perfusion equipment have been removed.
3. Calibrate traces from stellate and principal cells of different MT segments (using the absolute or normalized ratio as appropriate) with Equation 2 and analyze the recovery phase after NH₄Cl withdrawal by applying exponential decay functions using statistical analysis software and noting the decay constant (τ) (Figure 6B).
   Equation 2
   
   Where R = SEpH/mCherry ratio and A₁, A₂, x_o, and dx are curve fitting parameters determined by calibration in step 7.4 (Equation 1).
   1. Calculate acid extrusion rate (J_H+, see Equation 3) as a function of pH_i to account for variations in resting pH_i and acid loading between preparations³⁴. Use the exponential functions derived in step 8.3 to calculate the derivative of pH_i with regard to time in each time interval.
      Equation 3
      
   2. Calculate the intrinsic buffering capacity (β_i; Equation 4) of the cytosol at the pH_i from the start of each interval in step 8.3.1 based on previous literature (see Equation 4).
      NOTE: In Drosophila, the most thorough characterization of β_i comes from larval motor nerve terminals³⁵ and these data can be assumed to hold for MT cells in the absence of other available data.
      Equation 4
      
   3. Calculate the product of β_T (from step 8.3.2)_and dpH_i/dt (from step 8.3.1) to determine J_H+ (Equation 3).
      NOTE: In nominally bicarbonate-free solutions such as those described in this protocol, bicarbonate-derived buffering capacity (β_b) is assumed to be ~0 mM. Total buffering capacity (β_T) is the sum of β_i and_βb, and thus β_{i =} β_T in the absence of HCO₃^-/CO₂³⁶.
   4. Plot J_H+ as a function of the pH_i at the beginning of each time interval as outlined in step 6.14.9.
   5. Apply exponential decay functions to the portion of all datasets which overlap in pH_i using statistical analysis software. Compare rates of change of the resulting functions to compare acid extrusion rates between cells and MT segments (Figure 6C).
      NOTE: The most appropriate function used for curve fitting may not always be a single exponential. Other functions can be substituted if they improve the goodness of fit.
   6. Calculate acid flux (see Equation 5) as a function of pH_i to account for variations in cell size and shape.
      Equation 5
      
      NOTE: Cell dimensions can either be measured directly in images or approximated. Principal cells can be represented as halves of a hollow tube with the following dimensions: inner diameter 24 µm; outer diameter 48 µm; height 50 µm. Transitional stellate cells are variable but can be roughly represented as cylinders with heights of 50 µm and diameters of 10 µm. See final paragraph of Representative Results below.
   7. Apply exponential decay functions to the portion of all datasets that overlap in pH_i using statistical analysis software. Compare rates of change of the resulting functions to compare acid fluxes between cells and MT segments (Figure 6D).

Results

Healthy tissues and proper identification of anterior MTs are vital to the success of this protocol. During dissection, care should be taken to not directly touch the MTs and to only handle them by the ureter as gripping the MTs directly will lead to breakage (Figure 4A - B). When MTs are swept flat onto the slide, the tubules must be touched as little as possible and excess motion avoided as this will damage the si...

Discussion

The success of quantification of pH_i in Drosophila MTs depends entirely on the health of extracted MTs and the quality of mounting and dissection (Figure A - C). Thus, the careful handling of tissue as described is imperative. Slides freshly coated in PLL substantially aid MT mounting as they tend to be much more adhesive than slides which have previously been exposed to solution. Careful mounting will also aid in identification of distinct ...

Materials

Name	Company	Catalog Number	Comments
Poly-L-Lysine (PLL) Solution	Sigma-Aldrich	P4832	Store at 4 °C, can be reused.
Nigericin Sodium Salt	Sigma-Aldrich	N7143	CAUTION: Handle with gloves. Store as aliquots of 20 mM stock solution in DMSO at 4 °C.
Adhesive Perfusion Chamber Covers, adhesive size 1 mm, chamber diameter × thickness 9 mm × 0.9 mm, ports diameter 1.5 mm	Sigma-Aldrich	GBL622105	Can be substituted as needed to match perfusion system.
Sylgard 184 Silicone Elastomer Kit	Ellsworth Adhesives	184 SIL ELAST KIT 0.5KG	Available from multiple vendors.
Helping Hands Soldering Stands	Harbor Freight Tools	60501	Available from multiple vendors.
Open Gravity-fed Perfusion System with Valve Controller, 8 to 1 Manifold and Reserviors	Bioscience Tools	PS-8S	Any comparable perfusion system can be used.
Flow Regulator	Warner Instruments	64-0221	Can be substituted as needed to match perfusion system.
Schneider's Medium	Fisher Scientific	21720024	Store at 4 °C in sterile aliquots.
#5 Inox Steel Forceps	Fine Science Tools	11252-20	Can be substituted based on experimenter comfort.
35 x 10 mm polystyrene Petri dish	Corning Life Sciences	Fisher Scientific 08-757-100A	Exact brand and size are unimportant.
75 x 25 mm Microscope Slides	Corning Life Sciences	2949-75X25	Exact brand and size can vary as long as perfusion wells are compatible.
Filimented Borosilicate Capillary Glass, ID 1.5 mm, OD 0.86 mm, thickness 0.32 mm	Warner Instruments	64-0796	Filiment not necessary, glass can be substituted to match perfusion tubing and perfusion wells.
Tygon Tubing, ID 1/16", OD 1/8", thickness 1/32"	Fisher Scientific	14-171-129	Available from multiple vendors, can be substituted to match perfusion system.
Vacuum Silicone Grease	Sigma-Aldrich	Z273554	Available from multiple vendors.
Plastic Flow Control Clamp	Fisher Scientific	05-869	Available from multiple vendors, sterility not required
Glass rods, 5 mm diameter	delphiglass.com	9198	Exact size is personal preference, multiple vendors available
PAP Hydrophobic Pen	Sigma-Aldrich	Z377821	Available from multiple vendors.
Sealing Film	Sigma-Aldrich	P7668	Available from multiple vendors.
15 mL Falcon tube	BD Falcon	352096	Available from multiple vendors.
50 mL Falcon tube	BD Falcon	352070	Available from multiple vendors.
HEPES; 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid	Sigma-Aldrich	H3375	Available from multiple vendors.
MES; 4-Morpholineethanesulfonic acid monohydrate	Sigma-Aldrich	69892	Available from multiple vendors.
TAPS; N-[Tris(hydroxymethyl)methyl]-3-aminopropanesulfonic acid	Sigma-Aldrich	T5130	Available from multiple vendors.
10X/0.45 Air Objective	Zeiss	000000-1063-139	Comparable objectives can be substituted. 40X objectives can be used for single cell imaging.
Dissecting Stereoscope	Zeiss	Discovery.V8	Any dissecting stereoscope can be used.
UAS-pHerry transgenic Drosophila melagnogaster	Available from Romero Lab		First published: Citation 10
capaR-GAL4 driver line Drosophila melagnogaster	Available from Romero Lab		First published: Citation 32
c724-GAL4 driver line Drosophila melagnogaster	Available from Romero Lab		First published: Citation 2
Monochromatic High Sensitivity Digital Camera	Zeiss	Axiocam 506 mono	Exact brand and model can vary, can be replaced with any monochromatic high-sensitivity camera suited to live cellular imaging.
GFP/FITC filter set, 470/40 nm ex., 515 nm longpass em., 500 nm dichroic	Chroma	CZ909	Any GFP/FITC filer set can be substituted.
RFP/TRITC filter set, 546/10 nm ex., 590 nm longpass em., 565 nm dichroic	Chroma	CZ915	Any GFP/FITC filer set can be substituted.
Inverted Epifluoescent Microscope	Zeiss	Axio Observer Z.1	Any comparable microscope with motorized filter switching can be used. Upright microscopes can be used with open perfusion baths and water-immersion objectives.
Statistical Analysis Software	Microcal	Origin 6.0	Any software with comparable functionality can be substituted
Image Analysis Software	National Institutes of Health	ImageJ 1.50i	Any software with comparable functionality can be substituted
Image Acquisition Software	Zeiss	Zen 1.1.2.0	Any software with comparable functionality can be substituted
Single-edged Carbon Steel Razor Blade	Electron Microscopy Sciences	71960	Available from multiple vendors.
Microscopy Slide Folder	Fisher Scientific	16-04	Available from multiple vendors.
Bunsen Burner	Fisher Scientific	50-110-1231	Available from multiple vendors.
Polystrene Drosophila Rearing Vials with Flugs	Genesee Scientific	32-109BF	Comparable items can be substituted.
2.5 L Laboratory Ice Bucket	Fisher Scientific	07-210-129	Available from multiple vendors.
NMDG; N-Methyl-D-glucamine	Sigma-Aldrich	M2004	Available from multiple vendors.
200 uL barrier pipette tips	MidSci	AV200	Available from multiple vendors.
200 μL variable volume pipette	Gilson Incorporated	PIPETMAN P200	Available from multiple vendors.