We thank the University of Calgary for its support. We would also like to thank Dr. Robin Yates for access to reagents, equipment, and useful discussions.

1. Preparation of cells
<ol>
	<li>Grow Raw264.7 cells at 37 &#176;C and 5% CO2 to 70% confluency in RPMI supplemented with 10% heat-inactivated serum.</li>
	<li>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 &#956;L 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.</li>
	<li>On the day of the assay, check whether the wells are at least 70% confluent.</li>
</ol>
2. Measuring macropinosome pH
<ol>
	<li>Prepare the cells as in section 1.</li>
	<li>Wash all the wells with 100 &#956;L of HBSS at 37 &#176;C.</li>
	<li>Fill each well with 100 &#956;L of HBSS containing 0.025 mg/mL of TMR-labeled 70 kDa dextran and 0.025 mg/mL of fluorescein-labeled 70 kDa dextran.</li>
	<li>Place the cells into an incubator set to 37 &#176;C for 15 min.</li>
	<li>Remove the cells from the incubator and wash the cells 6x with 100 &#956;L of HBSS.</li>
	<li>Fill each well with 100 &#956;L of HBSS at 37 &#176;C.</li>
	<li>Place the plate on the microscope with a heated stage and chamber.</li>
	<li>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 &#956;m Z-volume was acquired at 0.5 &#956;m intervals.</li>
	<li>Acquire an image from each well, alternating between fluorophores in between each well.</li>
	<li>After the first acquisition, check whether all the wells remained in focus across the entire plate.</li>
	<li>Acquire images of each well at 1-15 min intervals for the desired length of time.</li>
</ol>
3. In situ calibration of macropinosome pH
<ol>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Add nigericin to a final concentration of 10 &#956;g/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.</li>
	<li>Place the plate back on the microscope and acquire images of each well using the same acquisition settings as above.</li>
	<li>Repeat steps 3.5 and 3.6 for each calibration buffer.</li>
</ol>
4. Data analysis for macropinosome pH
<ol>
	<li>Using FIJI software, subtract the background from both the TMR and the fluorescein channel.</li>
	<li>Generate a mask on the TMR channel. To generate a mask, convert the image to a binary image by selecting Adjust &#62; Threshold &#62; Apply. Next, highlight the binary image and select Edit &#62; Selection &#62; Create Mask.</li>
	<li>Apply it to both the TMR and fluorescein images. To do this, highlight the mask and select Edit &#62; Selection &#62; 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.</li>
	<li>Record the TMR and fluorescein intensity for each macropinosome within the masks.</li>
	<li>Repeat steps 4.1-4.3 for each time point from the time course.</li>
	<li>Repeat steps 4.1-4.3 for the images captured in the in situ calibration.</li>
	<li>Divide the fluorescein intensity by the TMR intensity to generate the fluorescein:TMR ratio.</li>
	<li>Plot the pH against the fluorescein:TMR ratios for the calibration images.</li>
	<li>Fit a curve to the calibration data.</li>
	<li>Interpolate the data for each time point from the time course to get absolute pH values.</li>
</ol>
5. Measuring oxidative events within macropinosomes
<ol>
	<li>Prepare the cells as in section 1.</li>
	<li>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.</li>
	<li>On the day of the assay, wash all the wells with 100 &#956;L of HBSS at 37 &#176;C.</li>
	<li>Fill each well with 100 &#956;L of HBSS containing 0.025 mg/mL of TMR-labeled 70 kDa dextran and 0.025 mg/mL of H2DCFDA-labeled 70 kDa dextran.</li>
	<li>Place cells into an incubator set to 37 &#176;C for 15 min.</li>
	<li>Remove cells from the incubator and wash the cells 6x with 100 &#956;L of HBSS.</li>
	<li>Fill each well with 100 &#956;L of HBSS at 37 &#176;C.</li>
	<li>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 &#956;m Z-volume was acquired at 0.5 &#956;m intervals.</li>
	<li>Acquire an image from each well, alternating between fluorophores in between each well.</li>
	<li>After the first acquisition, check whether all the wells remain in focus across the entire plate.</li>
	<li>Acquire images of each well at 1-15 min intervals for the desired length of time.</li>
</ol>
6. Data analysis for oxidative events within macropinosomes
<ol>
	<li>Using FIJI software, subtract the background from both the TMR and H2DCFDA channels.</li>
	<li>Generate a mask on the TMR channel. To generate a mask, convert the image to a binary image by selecting Adjust &#62; Threshold &#62; Apply. Next, highlight the binary image and select Edit &#62; Selection &#62; Create Mask.</li>
	<li>Apply the mask to both the TMR and H2DCFDA images. To do this, highlight the mask and select Edit &#62; Selection &#62; 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.</li>
	<li>Record the TMR and H2DCFDA intensity for each macropinosome within the masks.</li>
	<li>Repeat steps 4.1-4.3 for each time point from the time course.</li>
	<li>Divide the H2DCFDA intensity by the TMR intensity to generate an H2DCFDA: TMR ratio.</li>
	<li>Plot the H2DCFDA: TMR ratio against time.</li>
</ol>
7. Measuring protein digestion within macropinosomes
<ol>
	<li>Prepare the cells as in section 1.</li>
	<li>On the day of the assay, wash all the wells with 100 &#956;L HBSS at 37 &#176;C.</li>
	<li>Dissolve BODIPY-labeled ovalbumin to a concentration of 4 mg/mL in HBSS.</li>
	<li>Fill each well with 100 &#956;L of HBSS containing 0.025 mg/mL of TMR-labeled 70 kDa dextran and 0.2 mg/mL of BODIPY-labeled ovalbumin.</li>
	<li>Place the cells into an incubator set to 37 &#176;C for 15 min.</li>
	<li>Remove cells from the incubator and wash the cells 6x with 100 &#956;L of HBSS.</li>
	<li>Fill each well with 100 &#956;L of HBSS at 37 &#176;C.</li>
	<li>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 &#956;m Z-volume was acquired at 0.5 &#956;m intervals.</li>
	<li>Acquire an image from each well, alternating between fluorophores in between each well.</li>
	<li>After the first acquisition, check whether all the wells remain in focus across the entire plate.</li>
	<li>Acquire images of each well at 1-15 min intervals for the desired length of time.</li>
</ol>
8. Data analysis for protein digestion within macropinosomes
<ol>
	<li>Using FIJI software, subtract the background from both the TMR and BODIPY channels.</li>
	<li>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).</li>
	<li>Record the TMR and BODIPY intensity for each macropinosome within the masks.</li>
	<li>Repeat steps 4.1-4.3 for each time point from the time course.</li>
	<li>Divide the BODIPY intensity by the TMR intensity to generate the BODIPY:TMR ratio.</li>
	<li>Plot the BODIPY:TMR ratio against time.</li>
</ol>

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The authors have no conflicts of interest to disclose.

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...

Macropinocytosis refers to the uptake of large quantities of extracellular fluid into membrane-bound cytoplasmic organelles called macropinosomes1,2. 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 references1,2,5,6,7). The extracellular stimulus inducing macropinocytosis is most often a soluble growth factor5,8. 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 tumors9,10,11,12,13. 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 macropinocytosis6,15,16. 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 system16,18,19. 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 pKa 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 pKa that is sensitive within that range (Figure 2C) should be selected. The fluorophore fluorescein, which has a pKa 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 manuscript22,25. 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 phagosomes29,30. 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&#39;,7&#39;-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).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Black-walled 96 well plate</td>
			<td>PerkinElmer</td>
			<td>6005430</td>
			<td></td>
		</tr>
		<tr>
			<td>CypHer5e, NHS ester</td>
			<td>Cytiva</td>
			<td>PA15401</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran-amino 70 kDa</td>
			<td>Invitrogen</td>
			<td>D1862</td>
			<td></td>
		</tr>
		<tr>
			<td>DQ-ovalbumin</td>
			<td>Invitrogen</td>
			<td>D12053</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC-dextran 70 kDa</td>
			<td>Invitrogen</td>
			<td>D1823</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Gibco</td>
			<td>14287</td>
			<td></td>
		</tr>
		<tr>
			<td>Nigericin</td>
			<td>Sigma Aldrich</td>
			<td>N7143</td>
			<td></td>
		</tr>
		<tr>
			<td>OxyBurst Green-SE</td>
			<td>Invitrogen</td>
			<td>D2935</td>
			<td></td>
		</tr>
		<tr>
			<td>pHrodo Red, SE</td>
			<td>Invitrogen</td>
			<td>P36600</td>
			<td></td>
		</tr>
		<tr>
			<td>Raw264.7 cells</td>
			<td>ATCC</td>
			<td>TIB-71</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI medium</td>
			<td>Gibco</td>
			<td>11875093</td>
			<td></td>
		</tr>
		<tr>
			<td>SP5 Confocal Microscope</td>
			<td>Leica</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>TRITC-dextran 70 kDa</td>
			<td>Invitrogen</td>
			<td>D1819</td>
			<td></td>
		</tr>
		<tr>
			<td>u-Dish</td>
			<td>Ibidi</td>
			<td>81156</td>
			<td></td>
		</tr>
	</tbody>
</table>

measuring the ph, redox chemistries, and degradative capacity of macropinosomes using dual-fluorophore ratiometric microscopy

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.

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...

Watch this Scientific Journal Video about Measuring the pH, Redox Chemistries, and Degradative Capacity of Macropinosomes using Dual-Fluorophore Ratiometric Microscopy at JoVE.com

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

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.

In recent years, the field of macropinocytosis has grown rapidly. Macropinocytosis has emerged as a central mechanism by which innate immune cells ...

measuring-ph-redox-chemistries-degradative-capacity-macropinosomes

Research

JoVE Journal

Immunology and Infection

2.3K Views.  University of Victoria. 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.

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

A Cell Free Assay System Estimating the Neutralizing Capacity of GM-CSF Antibody using Recombinant Soluble GM-CSF Receptor

BACKGROUNDS: Previously, we demonstrated that neutralizing capacity but not the concentration of GM-CSF autoantibody was correlated with the disease severity in patients with autoimmune pulmonary alveolar proteinosis (PAP)1-3. As abrogation of GM-CSF bioactivity in the lung is the likely cause for autoimmune PAP4,5, it is promising to measure the neutralizing capacity of GM-CSF autoantibodies for evaluating the disease severity in each patient with PAP.
Until now, neutralizing capacity of GM-CSF autoantibodies has been assessed by evaluating the growth inhibition of human bone marrow cells or TF-1 cells stimulated with GM-CSF6-8. In the bioassay system, however, it is often problematic to obtain reliable data as well as to compare the data from different laboratories, due to the technical difficulties in maintaining the cells in a constant condition.
OBJECTIVE: To mimic GM-CSF binding to GM-CSF receptor on the cell surface using cell-free receptor-binding-assay.
METHODS: Transgenic silkworm technology was applied for obtaining a large amount for recombinant soluble GM-CSF receptor alpha (sGMR&alpha;) with high purity9-13. The recombinant sGMR&alpha; was contained in the hydrophilic sericin layers of silk threads without being fused to the silk proteins, and thus, we can easily extract from the cocoons in good purity with neutral aqueous solutions14,15. Fortunately, the oligosaccharide structures, which are critical for binding with GM-CSF, are more similar to the structures of human sGMR&alpha; than those produced by other insects or yeasts.
RESULTS: The cell-free assay system using sGMR&alpha; yielded the data with high plasticity and reliability. GM-CSF binding to sGMR&alpha; was dose-dependently inhibited by polyclonal GM-CSF autoantibody in a similar manner to the bioassay using TF-1 cells, indicating that our new cell-free assay system using sGMR&alpha; is more useful for the measurement of neutralizing activity of GM-CSF autoantibodies than the bioassay system using TF-1 cell or human bone marrow cells.
CONCLUSIONS: We established a cell-free assay quantifying the neutralizing capacity of GM-CSF autoantibody.

We designed a cell-free receptor binding assay in order to estimate the binding of granulocyte-macrophage colony-stimulating factor (GM-CSF) to the receptors. It enables us to evaluate competitive inhibition of biotinylated GM-CSF binding to soluble GM-CSF receptor alpha by GM-CSF autoantibody with excellent reproducibility.

BACKGROUNDS: Previously, we demonstrated that neutralizing capacity but not the concentration of GM-CSF autoantibody was correlated with the disease ...

Intravital Imaging of the Mouse Thymus using 2-Photon Microscopy

Two-photon Microscopy (TPM) provides image acquisition in deep areas inside tissues and organs. In combination with the development of new stereotactic tools and surgical procedures, TPM becomes a powerful technique to identify "niches" inside organs and to document cellular "behaviors" in live animals. While intravital imaging provides information that best resembles the real cellular behavior inside the organ, it is both more laborious and technically demanding in terms of required equipment/procedures than alternative ex vivo imaging acquisition. Thus, we describe a surgical procedure and novel "stereotactic" organ holder that allows us to follow the movements of Foxp3+ cells within the thymus.
Foxp3 is the master regulator for the generation of regulatory T cells (Tregs). Moreover, these cells can be classified according to their origin: ie. thymus-differentiated Tregs are called "naturally-occurring Tregs" (nTregs), as opposed to peripherally-converted Tregs (pTregs). Although significant amount of research has been reported in the literature concerning the phenotype and physiology of these T cells, very little is known about their in vivo interactions with other cells. This deficiency may be due to the absence of techniques that would permit such observations. The protocol described in this paper provides a remedy for this situation.
Our protocol consists of using nude mice that lack an endogenous thymus since they have a punctual mutation in the DNA sequence that compromises the differentiation of some epithelial cells, including thymic epithelial cells. Nude mice were gamma-irradiated and reconstituted with bone marrows (BM) from Foxp3-KIgfp/gfp mice. After BM recovery (6 weeks), each animal received embryonic thymus transplantation inside the kidney capsule. After thymus acceptance (6 weeks), the animals were anesthetized; the kidney containing the transplanted thymus was exposed, fixed in our organ holder, and kept under physiological conditions for in vivo imaging by TPM. We have been using this approach to study the influence of drugs in the generation of regulatory T cells.

We have developed novel laboratory tools and protocols for intravital imaging acquisition of the thymus. Our technique should help in the identification of &#x201C;niches&#x201D; within the thymus where T cell development occurs.

Two-photon Microscopy (TPM) provides image acquisition in deep areas inside tissues and organs. In combination with the development of new stereotactic ...

Measuring Local Anaphylaxis in Mice

Allergic responses are the result of the activation of mast cells and basophils, and the subsequent release of vasoactive and proinflammatory mediators. Exposure to an allergen in a sensitized individual can result in clinical symptoms that vary from minor erythema to life threatening anaphylaxis. In the laboratory, various animal models have been developed to understand the mechanisms driving allergic responses. Herein, we describe a detailed method for measuring changes in vascular permeability to quantify localized allergic responses. The local anaphylaxis assay was first reported in the 1920s, and has been adapted from the technique published by Kojima et al. in 20071. In this assay, mice sensitized to OVA are challenged in the left ear with vehicle and in the right ear with OVA. This is followed by an intravenous injection of Evans Blue dye. Ten min after injecting Evans Blue, the animal is euthanized and the dye that has extravasated into the ears is extracted overnight in formamide. The absorbance of the extracted dye is then quantified with a spectrophotometer. This method reliably results in a visual and quantifiable manifestation of a local allergic response.

Allergic responses, characterized by the activation of mast cells and basophils, are driven by the cross-linking of IgE and release of proinflammatory mediators. A quantitative assessment of allergic responses can be achieved by using Evans Blue dye to monitor changes in vascular permeability after allergen challenge.

Allergic responses are the result of the activation of mast cells and basophils, and the subsequent release of vasoactive and proinflammatory mediators. ...

Analysis of Yersinia enterocolitica Effector Translocation into Host Cells Using Beta-lactamase Effector Fusions

Many gram-negative bacteria including pathogenic Yersinia spp. employ type III secretion systems to translocate effector proteins into eukaryotic target cells. Inside the host cell the effector proteins manipulate cellular functions to the benefit of the bacteria. To better understand the control of type III secretion during host cell interaction, sensitive and accurate assays to measure translocation are required. We here describe the application of an assay based on the fusion of a Yersinia enterocolitica effector protein fragment (Yersinia outer protein; YopE) with TEM-1 beta-lactamase for quantitative analysis of translocation. The assay relies on cleavage of a cell permeant FRET dye (CCF4/AM) by translocated beta-lactamase fusion. After cleavage of the cephalosporin core of CCF4 by the beta-lactamase, FRET from coumarin to fluorescein is disrupted and excitation of the coumarin moiety leads to blue fluorescence emission. Different applications of this method have been described in the literature highlighting its versatility. The method allows for analysis of translocation in vitro and also in in vivo, e.g., in a mouse model. Detection of the fluorescence signals can be performed using plate readers, FACS analysis or fluorescence microscopy. In the setup described here, in vitro translocation of effector fusions into HeLa cells by different Yersinia mutants is monitored by laser scanning microscopy. Recording intracellular conversion of the FRET reporter by the beta-lactamase effector fusion in real-time provides robust quantitative results. We here show exemplary data, demonstrating increased translocation by a Y. enterocolitica YopE mutant compared to the wild type strain.

Analysis of Yersinia enterocolitica Effector Translocation into Host Cells Using Beta-lactamase Effector Fusions

Effector translocation into host cells via a type III secretion system is a common virulence strategy among gram-negative bacteria. A beta-lactamase effector fusion based assay for quantitative analysis of translocation was applied. In Yersinia infected cells, conversion of a FRET reporter by the beta-lactamase is monitored using laser scanning microscopy.

Many gram-negative bacteria including pathogenic Yersinia spp. employ type III secretion systems to translocate effector proteins into eukaryotic target ...

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila express gustatory receptors1,2, ion channels3-6, and ionotropic receptors7 that are involved in the detection of volatile and non-volatile sensory cues. These neurons directly contact tastants such as food, noxious substances and pheromones and therefore influence many complex behaviors such as feeding, egg-laying and mating. Electrode recordings and calcium imaging have been widely used in insects to quantify the neuronal responses evoked by these tastants. However, electrophysiology requires specialized equipment and obtaining measurements from a single taste sensillum can be technically challenging depending on the cell-type, size, and position. In addition, single neuron resolution in Drosophila can be difficult to achieve since taste sensilla house more than one type of chemosensory neuron. The live calcium imaging method described here allows responses of single gustatory neurons in live flies to be measured. This method is especially suitable for imaging neuronal responses to lipid pheromones and other ligand types that have low solubility in water-based solvents.

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

The forelegs and proboscis of Drosophila contain a rich repertoire of gustatory sensory neurons. Here, we present a method using calcium imaging to measure physiological responses from sensory neurons in the foreleg and proboscis of live flies upon exogenous application of a gustatory pheromone.

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila ...

Measuring Phagosome pH by Ratiometric Fluorescence Microscopy

Phagocytosis is a fundamental process through which innate immune cells engulf bacteria, apoptotic cells or other foreign particles in order to kill or neutralize the ingested material, or to present it as antigens and initiate adaptive immune responses. The pH of phagosomes is a critical parameter regulating fission or fusion with endomembranes and activation of proteolytic enzymes, events that allow the phagocytic vacuole to mature into a degradative organelle. In addition, translocation of H+ is required for the production of high levels of reactive oxygen species (ROS), which are essential for efficient killing and signaling to other host tissues. Many intracellular pathogens subvert phagocytic killing by limiting phagosomal acidification, highlighting the importance of pH in phagosome biology. Here we describe a ratiometric method for measuring phagosomal pH in neutrophils using fluorescein isothiocyanate (FITC)-labeled zymosan as phagocytic targets, and live-cell imaging. The assay is based on the fluorescence properties of FITC, which is quenched by acidic pH when excited at 490 nm but not when excited at 440 nm, allowing quantification of a pH-dependent ratio, rather than absolute fluorescence, of a single dye. A detailed protocol for performing in situ dye calibration and conversion of ratio to real pH values is also provided. Single-dye ratiometric methods are generally considered superior to single wavelength or dual-dye pseudo-ratiometric protocols, as they are less sensitive to perturbations such as bleaching, focus changes, laser variations, and uneven labeling, which distort the measured signal. This method can be easily modified to measure pH in other phagocytic cell types, and zymosan can be replaced by any other amine-containing particle, from inert beads to living microorganisms. Finally, this method can be adapted to make use of other fluorescent probes sensitive to different pH ranges or other phagosomal activities, making it a generalized protocol for the functional imaging of phagosomes.

Phagosomal pH influences phagosome maturation, oxidant production, phagosomal killing as well as antigen presentation. Here we describe a ratiometric method for measuring time-course and endpoint pH changes in individual phagosomes in living phagocytes using fluorescence microscopy.

Phagocytosis is a fundamental process through which innate immune cells engulf bacteria, apoptotic cells or other foreign particles in order to kill or ...

Ratiometric Imaging of Extracellular pH in Dental Biofilms

The pH in bacterial biofilms on teeth is of central importance for dental caries, a disease with a high worldwide prevalence. Nutrients and metabolites are not distributed evenly in dental biofilms. A complex interplay of sorption to and reaction with organic matter in the biofilm reduces the diffusion paths of solutes and creates steep gradients of reactive molecules, including organic acids, across the biofilm. Quantitative fluorescent microscopic methods, such as fluorescence life time imaging or pH ratiometry, can be employed to visualize pH in different microenvironments of dental biofilms. pH ratiometry exploits a pH-dependent shift in the fluorescent emission of pH-sensitive dyes. Calculation of the emission ratio at two different wavelengths allows determining local pH in microscopic images, irrespective of the concentration of the dye. Contrary to microelectrodes the technique allows monitoring both vertical and horizontal pH gradients in real-time without mechanically disturbing the biofilm. However, care must be taken to differentiate accurately between extra- and intracellular compartments of the biofilm. Here, the ratiometric dye, seminaphthorhodafluor-4F 5-(and-6) carboxylic acid (C-SNARF-4) is employed to monitor extracellular pH in in vivo grown dental biofilms of unknown species composition. Upon exposure to glucose the dye is up-concentrated inside all bacterial cells in the biofilms; it is thus used both as a universal bacterial stain and as a marker of extracellular pH. After confocal microscopic image acquisition, the bacterial biomass is removed from all pictures using digital image analysis software, which permits to exclusively calculate extracellular pH. pH ratiometry with the ratiometric dye is well-suited to study extracellular pH in thin biofilms of up to 75 &#181;m thickness, but is limited to the pH range between 4.5 and 7.0.

A pH-sensitive ratiometric dye is used in combination with confocal laser scanning microscopy and digital image analysis to monitor extracellular pH in dental biofilms in real-time.

The pH in bacterial biofilms on teeth is of central importance for dental caries, a disease with a high worldwide prevalence. Nutrients and metabolites ...

Imaging the Neutrophil Phagosome and Cytoplasm Using a Ratiometric pH Indicator

Neutrophils are crucial to host innate defense and, consequently, constitute an important area of medical research. The phagosome, the intracellular compartment where the killing and digestion of engulfed particles take place, is the main arena for neutrophil pathogen killing that requires tight regulation. Phagosomal pH is one aspect that is carefully controlled, in turn regulating antimicrobial protease activity. Many fluorescent pH-sensitive dyes have been used to visualize the phagosomal environment. S-1 has several advantages over other pH-sensitive dyes, including its dual emission spectra, its resistance to photo-bleaching, and its high pKa. Using this method, we have demonstrated that the neutrophil phagosome is unusually alkaline in comparison to other phagocytes. By using different biochemical conjugations of the dye, the phagosome can be delineated from the cytoplasm so that changes in the size and shape of the phagosome can be assessed. This allows for further monitoring of ionic movement.

This manuscript describes a simple method to measure the phagosomal pH and area as well as the cytoplasmic pH of human and mouse neutrophils using the ratiometric indicator seminaphthorhodafluor (SNARF)-1, or S-1. This is achieved using live-cell confocal fluorescence microscopy and image analysis.

Neutrophils are crucial to host innate defense and, consequently, constitute an important area of medical research. The phagosome, the intracellular ...

Isolation, Characterization, and Purification of Macrophages from Tissues Affected by Obesity-related Inflammation

Obesity promotes a chronic inflammatory state that is largely mediated by tissue-resident macrophages as well as monocyte-derived macrophages. Diet-induced obesity (DIO) is a valuable model in studying the role of macrophage heterogeneity; however, adequate macrophage isolations are difficult to acquire from inflamed tissues. In this protocol, we outline the isolation steps and necessary troubleshooting guidelines derived from our studies for obtaining a suitable population of tissue-resident macrophages from mice following 18 weeks of high-fat (HFD) or high-fat/high-cholesterol (HFHCD) diet intervention. This protocol focuses on three hallmark tissues studied in obesity and atherosclerosis including the liver, white adipose tissues (WAT), and the aorta. We highlight how dualistic usage of flow cytometry can achieve a new dimension of isolation and characterization of tissue-resident macrophages. A fundamental section of this protocol addresses the intricacies underlying tissue-specific enzymatic digestions and macrophage isolation, and subsequent cell-surface antibody staining for flow cytometric analysis. This protocol addresses existing complexities underlying fluorescent-activated cell sorting (FACS) and presents clarifications to these complexities so as to obtain broad range characterization from adequately sorted cell populations. Alternate enrichment methods are included for sorting cells, such as the dense liver, allowing for flexibility and time management when working with FACS. In brief, this protocol aids the researcher to evaluate macrophage heterogeneity from a multitude of inflamed tissues in a given study and provides insightful troubleshooting tips that have been successful for favorable cellular isolation and characterization of immune cells in DIO-mediated inflammation.

This protocol allows a researcher to isolate and characterize tissue-resident macrophages in various hallmark inflamed tissues extracted from diet-induced models of metabolic disorders.

Obesity promotes a chronic inflammatory state that is largely mediated by tissue-resident macrophages as well as monocyte-derived macrophages. ...

Monitoring Extracellular pH in Cross-Kingdom Biofilms using Confocal Microscopy

Cross-kingdom biofilms consisting of both fungal and bacterial cells are involved in a variety of oral diseases, such as endodontic infections, periodontitis, mucosal infections and, most notably, early childhood caries. In all of these conditions, the pH in the biofilm matrix impacts microbe-host interactions and thus the disease progression. The present protocol describes a confocal microscopy-based method to monitor pH dynamics inside cross-kingdom biofilms comprising Candida albicans and Streptococcus mutans. The pH-dependent dual-emission spectrum and the staining properties of the ratiometric probe C-SNARF-4 are exploited to determine drops in pH in extracellular areas of the biofilms. Use of pH ratiometry with the probe requires a meticulous choice of imaging parameters, a thorough calibration of the dye, and careful, threshold-based post-processing of the image data. When used correctly, the technique allows for the rapid assessment of extracellular pH in different areas of a biofilm and thus the monitoring of both horizontal and vertical pH gradients over time. While the use of confocal microscopy limits Z-profiling to thin biofilms of 75 &#181;m or less, the use of pH ratiometry is ideally suited for the noninvasive study of an important virulence factor in cross-kingdom biofilms.

The protocol describes the cultivation of cross-kingdom biofilms consisting of Candida albicans and Streptococcus mutans and presents a confocal microscopy-based method for the monitoring of extracellular pH inside these biofilms.