Measuring the pH, Redox Chemistries, and Degradative Capacity of Macropinosomes using Dual-Fluorophore Ratiometric Microscopy

Summary

We describe protocols for measuring pH, oxidative events, and protein digestion in individual macropinosomes in live cells. An emphasis is placed on dual-fluorophore ratiometric microscopy and the advantages it offers over population-based techniques.

Abstract

In recent years, the field of macropinocytosis has grown rapidly. Macropinocytosis has emerged as a central mechanism by which innate immune cells maintain organismal homeostasis and immunity. Simultaneously, and in contrast to its homeostatic role, it can also drive various pathologies, including cancer and viral infections. Unlike other modes of endocytosis, the tools developed for studying the maturation of macropinosomes remain underdeveloped. Here the protocol describes newly developed tools for studying the redox environment within the lumen of early and maturing macropinosomes. Methodologies for using ratiometric fluorescence microscopy in assessing the pH, production of reactive oxygen species, and the degradative capacity within the lumen of individual macropinosomes in live cells are described. Single organelle measurements offer the advantage of revealing spatiotemporal heterogeneity, which is often lost with population-based approaches. Emphasis is placed on the basic principles of dual fluorophore ratiometric microscopy, including probe selection, instrumentation, calibration, and single-cell versus population-based methods.

Introduction

Macropinocytosis refers to the uptake of large quantities of extracellular fluid into membrane-bound cytoplasmic organelles called macropinosomes^1,2. It is a highly conserved process performed by free-living unicellular organisms, such as the amoeba Dictyostelium spp.³, as well as anthozoans⁴ and metazoans². In most cells, macropinocytosis is an induced event. The ligation of cell-surface receptors induces the protrusion of actin-driven plasma membrane extensions referred to as ruffles. A fraction of those ruffles, by some poorly understood mechanism, seal at their distal tips to form macropinosomes (although beyond the scope of this methods paper, for detailed reviews on the mechanics of macropinocytosis, please refer to references^1,2,5,6,7). The extracellular stimulus inducing macropinocytosis is most often a soluble growth factor^5,8. Accordingly, the macropinocytic event allows for the ingestion of a bolus of extracellular material from which the cell can derive useful metabolites to facilitate growth. Unfortunately, this pathway for nutrient delivery can also drive pathology. Certain cancer cells harbor mutations that result in continuous or constitutive macropinocytosis. The continuous delivery of nutrients facilitates the uncontrolled proliferation of cancer cells and has been linked to particularly aggressive tumors^{9,10,11,12,13}. Similarly, viruses can induce macropinocytosis to gain access to host cells, thereby driving viral pathology¹⁴.

Macropinocytosis also functions in the maintenance of immunity to pathogens. Certain innate immune cells such as macrophages and dendritic cells engage in the constitutive and aggressive sampling of extracellular fluid via macropinocytosis^6,15,16. This mode of macropinocytosis is incredibly active, and a single dendritic cell can engorge itself with a volume of extracellular fluid equivalent to its own weight every hour¹⁷. Despite this constitutive sampling, macrophages and dendritic cells do not replicate uncontrollably as do tumor cells, instead, they seem to process the extracellular material in such a way that information can be extracted to inform upon the presence, or indeed absence, of potential threats. Information is extracted as i) pathogen-associated molecular patterns that can be read by intracellular pathogen recognition receptors and ii) short stretches of amino acids that can be loaded onto major histocompatibility molecules for screening by cells of the adaptive immune system^16,18,19. Whether pathogens subvert this pathway for information processing by immune cells is at present unclear.

Despite these well-defined and critical roles for macropinocytosis in both the maintenance of immunity and homeostasis and in contrast to other more commonly studied modes of endocytosis, little of the inner (luminal) workings of macropinosomes is known. Developing standardized protocols and tools to study the luminal biochemistry of macropinosomes will not only help us to understand their unique biology better but will provide insight that can be leveraged for novel therapeutic strategies, including drug delivery²⁰. This method manuscript will focus on recently developed tools to dissect, at the single organelle level, various aspects of the luminal biochemistry of macropinosomes.

Fluorophores can be used to measure specific biochemistries of organelles if i) they partition preferentially into the compartment of interest and/or ii) they undergo spectral changes in response to the parameter of interest. For example, in the case of pH, fluorescent weak bases, such as acridine orange, cresyl violet, and the LysoTracker dyes accumulate preferentially in acidic organelles. Therefore, their relative intensity is a rough indication that the labeled organelle is acidic. Other pH-responsive fluorophores, such as fluorescein, pHrodo, and cypHer5e, undergo spectral changes upon binding to protons (Figure 1A-C). Changes to the fluorescence emission of pH-sensitive fluorophores can therefore provide a useful approximation of pH. The use of single fluorophores, however, presents a number of disadvantages. For example, changes to the focal plane, photobleaching, and changes to the volume of individual organelles, a common occurrence in macropinosomes²¹, can induce changes to the fluorescence intensity of single fluorophores, and this cannot be easily corrected for²². Single-wavelength assessments, although useful for visualizing acidic compartments, are therefore purely qualitative.

A more quantitative approach is to target the parameter-sensitive fluorophore along with a reference fluorophore to the organelle of interest. The reference fluorophore is ideally insensitive to biochemical changes within the organelle (Figure 1D-F) and can therefore be used to correct for changes in the focal plane, organellar volume, and, to some extent, photobleaching²³. Using this approach, referred to as dual-fluorophore ratiometric fluorescence, correction can be achieved by generating a ratio of the fluorescence emission of the parameter-sensitive fluorophore to the reference fluorophore.

Here, the protocol will be utilizing the principle of dual-fluorophore ratiometric imaging to measure pH, oxidative events, and protein degradation within macropinosomes. In each case, a fluorophore will be selected that is sensitive to the parameter of interest and a reference fluorophore. In order to target the fluorophores specifically to macropinosomes, they will be covalently coupled to 70 kDa dextran, which is preferentially incorporated into macropinosomes²⁴. All assays will be performed in Raw264.7 cells but can be adapted to other cell types. Where possible, the fluorescence ratios will be calibrated against a reference curve to gain absolute values. Importantly, all measurements will be performed in live cells for dynamic and quantitative assessment of the luminal environment of macropinosomes.

When selecting pH-sensitive fluorophores, a number of considerations must be weighed. The first is the pK_a of the fluorophore, which indicates the range of pH values at which the probe will be most sensitive. If it is assumed that shortly after formation, the pH of the macropinosome will be close to that of the extracellular medium (~pH 7.2) and that it will progressively acidify through interactions with late endosomes and lysosomes (~pH 5.0), then a probe with a pK_a that is sensitive within that range (Figure 2C) should be selected. The fluorophore fluorescein, which has a pK_a of 6.4, is optimally sensitive within that range. It has been used extensively to measure other similar organelles, such as phagosomes, and will be the fluorophore of choice in this manuscript^22,25. As a reference fluorophore, tetramethylrhodamine will be used, which is insensitive to pH (Figure 1E). Other fluorophores, such as pHrodo and cypHer5e may be substituted for fluorescein where the spectral properties of fluorescein do match other experimental variables. Some suggested reference fluorophores for pHrodo and cypHer5e are shown in Figure 1.

A second consideration is the method by which the two fluorophores will be targeted specifically to macropinosomes. Dextran of the size 70 kDa, which has a hydrodynamic radius of roughly 7 nm, does not stick non-specifically to cells and is incorporated into macropinosomes, but not clathrin-coated pits or caveolae, and therefore marks macropinosomes (Figure 2A and Figure 3A,B)^16,24,26. In this protocol, fluorescein-labeled 70 kDa dextran and tetramethylrhodamine (TMR)-labeled 70 kDa dextran will be used as the pH-sensitive and reference probes, respectively.

In innate immune cells, macropinocytosis and phagocytosis represent the two major routes for the internalization of exogenous material for processing and subsequent presentation to cells of the adaptive immune response²⁷. The careful and coordinated control of the redox chemistry of the lumen of phagosomes and macropinosomes is critical to the context-specific processing of exogenous material. Perhaps the most well-studied regulator of oxidative events in phagosomes is the NADPH oxidase, a large multi-subunit complex that produces large quantities of reactive oxygen species (ROS) within the lumen of phagosomes²⁸. Indeed, its activity is central to appropriate antigen processing within phagosomes^29,30. Yet, the activity of the NADPH oxidase on macropinosomal membranes has not been explored.

In this protocol, the H2DCFDA succinimidyl ester is used for measuring oxidative events within the macropinosome. This is a modified form of fluorescein (2',7'-dichlorodihydrofluorescein diacetate), which is minimally fluorescent in its reduced form. Upon oxidation, its fluorescence emission increases significantly. It is, however, worth noting a significant caveat of H2DCFDA - as it is based on the fluorophore fluorescein, its fluorescence is also quenched in acidic compartments, and care must be taken to control for this variable when designing experiments²⁸. Similar to the approach for measuring pH, the H2DCFDA succinimidyl ester will be covalently attached to 70 kDa dextran and TMR-labeled 70 kDa dextran will be used as the reference fluorophore (Figure 3A).

Fluorescent ovalbumin will be used to measure protein degradation within macropinosomes. The ovalbumin used here is densely labeled with a 4,4-difluoro-4-bora-3a,4a-diaza-s-indacene (BODIPY) FL dye that is self-quenched. Upon digestion, strongly fluorescent dye-labeled peptides are liberated. As ovalbumin cannot be easily conjugated to 70 kDa dextran, cells with TMR-labeled 70 kDa dextran and fluid-phase ovalbumin will be co-incubated. The TMR signal will be used to generate a macropinosome mask during post-imaging analysis, and the signal liberated from the digested ovalbumin will be measured within the mask (Figure 3B).

Protocol

1. Preparation of cells

1. Grow Raw264.7 cells at 37 °C and 5% CO₂ to 70% confluency in RPMI supplemented with 10% heat-inactivated serum.
2. One day before the assay, seed Raw264.7 cells at a density of 5 x 10⁴ cells per well in a 96-well plate. Ensure that each well contains 100 μL of growth medium. Ensure that the 96-well plate has black sides and a glass-bottom for imaging.
   NOTE: Here, 96-well plates were used for imaging. The 96-well plates allow for the use of smaller quantities of cells and reagents relative to larger imaging chambers. However, any chamber designed for imaging live cells can be used, and reagents can be scaled up accordingly.
3. On the day of the assay, check whether the wells are at least 70% confluent.

2. Measuring macropinosome pH

1. Prepare the cells as in section 1.
2. Wash all the wells with 100 μL of HBSS at 37 °C.
3. Fill each well with 100 μL of HBSS containing 0.025 mg/mL of TMR-labeled 70 kDa dextran and 0.025 mg/mL of fluorescein-labeled 70 kDa dextran.
4. Place the cells into an incubator set to 37 °C for 15 min.
5. Remove the cells from the incubator and wash the cells 6x with 100 μL of HBSS.
6. Fill each well with 100 μL of HBSS at 37 °C.
7. Place the plate on the microscope with a heated stage and chamber.
8. Adjust the excitation/emission parameters for each fluorophore as needed.
   NOTE: Ideal conditions for image acquisition will vary with fluorophores used and individual microscope set-ups. Here, images were acquired on a Leica SP5 laser scanning confocal microscope. TMR was excited with a 543-laser line and emission was collected from 610 nm to 650 nm. Fluorescein was excited with a 488-laser line and emission was collected from 500 nm to 550 nm. A 63x oil immersion objective was used and a 10 μm Z-volume was acquired at 0.5 μm intervals.
9. Acquire an image from each well, alternating between fluorophores in between each well.
10. After the first acquisition, check whether all the wells remained in focus across the entire plate.
11. Acquire images of each well at 1-15 min intervals for the desired length of time.

3. In situ calibration of macropinosome pH

1. During the image acquisition, prepare 1 L of potassium-rich (K⁺-rich) solution containing 140 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, and 5 mM glucose. Supplement one 400 mL volume of the K⁺-rich solution with 25 mM HEPES (from a 1 M, pH 7.2 stock solution) and a separate 400 mL volume with 25 mM MES (from a 0.5 M, pH 6.0 stock solution).
2. Adjust a 50 mL aliquot of the K⁺-rich solution buffered with 25mM HEPES to pH 7.5 using 10 M HCl or 10 M KOH as needed.
3. Adjust three separate 50 mL aliquots of the K⁺-rich solution buffered with 25mM MES to pH 6.5, pH 5.5, and pH 5.0 using 10 M HCl or 10 M KOH as needed.
   NOTE: An acetate-acetic acid buffer may be preferable for lower pH solutions.
4. After the completion of image acquisition, remove the HBSS from the 96-well plate containing the cells and replace it with the K⁺-rich solution that is at pH 7.5.
5. Add nigericin to a final concentration of 10 μg/mL.
   NOTE: Nigericin is an ionophore that exchanges K⁺ for H⁺. By setting the K⁺ concentration of the K⁺-rich solution to a value that approximates the K⁺ concentration of the cytosol, it can be ensured that nigericin will clamp the pH of the macropinosome to that of the cytosol, which in turn reflects the pH of the calibration buffer.
6. Place the plate back on the microscope and acquire images of each well using the same acquisition settings as above.
7. Repeat steps 3.5 and 3.6 for each calibration buffer.

4. Data analysis for macropinosome pH

1. Using FIJI software, subtract the background from both the TMR and the fluorescein channel.
2. Generate a mask on the TMR channel. To generate a mask, convert the image to a binary image by selecting Adjust > Threshold > Apply. Next, highlight the binary image and select Edit > Selection > Create Mask.
3. Apply it to both the TMR and fluorescein images. To do this, highlight the mask and select Edit > Selection > Create Selection. Click on the TMR image and press Shift + E to apply the mask to the TMR channel. Similarly, click on the fluorescein image and press Shift + E to apply the mask to the OB channel.
4. Record the TMR and fluorescein intensity for each macropinosome within the masks.
5. Repeat steps 4.1-4.3 for each time point from the time course.
6. Repeat steps 4.1-4.3 for the images captured in the in situ calibration.
7. Divide the fluorescein intensity by the TMR intensity to generate the fluorescein:TMR ratio.
8. Plot the pH against the fluorescein:TMR ratios for the calibration images.
9. Fit a curve to the calibration data.
10. Interpolate the data for each time point from the time course to get absolute pH values.

5. Measuring oxidative events within macropinosomes

1. Prepare the cells as in section 1.
2. One day before the assay, prepare the H2DCFDA succinimidyl ester labeled 70 kDa dextran by resuspending 10 mg of 70 kDa dextran-amino in 1 mL of 0.1 M sodium bicarbonate solution (pH 8.3). Add 1 mg of the H2DCFDA succinimidyl ester to the dextran-amino solution and incubate for 1 h. Dialyze the H2DCFDA-labeled 70 kDa dextran against PBS. Do not store for more than 1 day as H2DCFDA-labeled 70 kDa dextran will oxidize upon storage.
3. On the day of the assay, wash all the wells with 100 μL of HBSS at 37 °C.
4. Fill each well with 100 μL of HBSS containing 0.025 mg/mL of TMR-labeled 70 kDa dextran and 0.025 mg/mL of H2DCFDA-labeled 70 kDa dextran.
5. Place cells into an incubator set to 37 °C for 15 min.
6. Remove cells from the incubator and wash the cells 6x with 100 μL of HBSS.
7. Fill each well with 100 μL of HBSS at 37 °C.
8. Place the plate on the microscope with a heated stage and chamber and adjust the excitation/emission parameters for each fluorophore as needed.
   NOTE: Ideal conditions for image acquisition will vary with fluorophores used and individual microscope set-ups. Here images were acquired on a Leica SP5 laser scanning confocal microscope. TMR was excited with a 543-laser line, and emission was collected from 610 nm to 650 nm. H2DCFDA was excited with a 488-laser line and emission was collected from 500 nm to 550 nm. A 63x oil immersion objective was used, and a 10 μm Z-volume was acquired at 0.5 μm intervals.
9. Acquire an image from each well, alternating between fluorophores in between each well.
10. After the first acquisition, check whether all the wells remain in focus across the entire plate.
11. Acquire images of each well at 1-15 min intervals for the desired length of time.

6. Data analysis for oxidative events within macropinosomes

1. Using FIJI software, subtract the background from both the TMR and H2DCFDA channels.
2. Generate a mask on the TMR channel. To generate a mask, convert the image to a binary image by selecting Adjust > Threshold > Apply. Next, highlight the binary image and select Edit > Selection > Create Mask.
3. Apply the mask to both the TMR and H2DCFDA images. To do this, highlight the mask and select Edit > Selection > Create Selection. Click on the TMR image and press Shift + E to apply the mask to the TMR channel. Similarly, click on the H2DCFDA image and press Shift + E to apply the mask to the H2DCFDA channel.
4. Record the TMR and H2DCFDA intensity for each macropinosome within the masks.
5. Repeat steps 4.1-4.3 for each time point from the time course.
6. Divide the H2DCFDA intensity by the TMR intensity to generate an H2DCFDA: TMR ratio.
7. Plot the H2DCFDA: TMR ratio against time.

7. Measuring protein digestion within macropinosomes

1. Prepare the cells as in section 1.
2. On the day of the assay, wash all the wells with 100 μL HBSS at 37 °C.
3. Dissolve BODIPY-labeled ovalbumin to a concentration of 4 mg/mL in HBSS.
4. Fill each well with 100 μL of HBSS containing 0.025 mg/mL of TMR-labeled 70 kDa dextran and 0.2 mg/mL of BODIPY-labeled ovalbumin.
5. Place the cells into an incubator set to 37 °C for 15 min.
6. Remove cells from the incubator and wash the cells 6x with 100 μL of HBSS.
7. Fill each well with 100 μL of HBSS at 37 °C.
8. Place the plate on the microscope with a heated stage and chamber and adjust the excitation/emission parameters for each fluorophore as needed.
   NOTE: Ideal conditions for image acquisition will vary with fluorophores used and individual microscope set-ups. Here, images were acquired on a Leica SP5 laser scanning confocal microscope. TMR was excited with a 543-laser line, and emission was collected from 610 nm to 650 nm. BODIPY was excited with a 488-laser line, and emission was collected from 500 nm to 550 nm. A 63x oil immersion objective was used and a 10 μm Z-volume was acquired at 0.5 μm intervals.
9. Acquire an image from each well, alternating between fluorophores in between each well.
10. After the first acquisition, check whether all the wells remain in focus across the entire plate.
11. Acquire images of each well at 1-15 min intervals for the desired length of time.

8. Data analysis for protein digestion within macropinosomes

1. Using FIJI software, subtract the background from both the TMR and BODIPY channels.
2. Generate a mask on the TMR channel and apply it to both the TMR and BODIPY images as in steps 6.2 and 6.3 above (Figure 3B).
3. Record the TMR and BODIPY intensity for each macropinosome within the masks.
4. Repeat steps 4.1-4.3 for each time point from the time course.
5. Divide the BODIPY intensity by the TMR intensity to generate the BODIPY:TMR ratio.
6. Plot the BODIPY:TMR ratio against time.

Results

When measuring macropinosome pH, there is a period of time for which the dynamics of acidification cannot be measured. This period corresponds to the dextran loading phase (gray box in Figure 2C and Figure 3C,D) and will vary depending on the cell type used. The length of the loading phase will vary with (i) the macropinocytic activity of the cells and (ii) the sensitivity of the instrument used. It is recommended to adjust this period at the st...

Discussion

Although there are a number of protocols for both low and high-throughput measurements of macropinocytic uptake in macrophages, fibroblasts, and even Dictyostelium spp.^3,7,31,32,33, very few attempts have been made to measure the luminal biochemistry of these dynamic compartments. This is likely due to a paucity of probes that can be efficiently targe...

Materials

Name	Company	Catalog Number	Comments
Black-walled 96 well plate	PerkinElmer	6005430
CypHer5e, NHS ester	Cytiva	PA15401
Dextran-amino 70 kDa	Invitrogen	D1862
DQ-ovalbumin	Invitrogen	D12053
FITC-dextran 70 kDa	Invitrogen	D1823
HBSS	Gibco	14287
Nigericin	Sigma Aldrich	N7143
OxyBurst Green-SE	Invitrogen	D2935
pHrodo Red, SE	Invitrogen	P36600
Raw264.7 cells	ATCC	TIB-71
RPMI medium	Gibco	11875093
SP5 Confocal Microscope	Leica	-
TRITC-dextran 70 kDa	Invitrogen	D1819
u-Dish	Ibidi	81156