Protocol

The Use of Spin-Labeled Ligands as Biophysical Probes to Report Real-Time Endocytosis of G ProteinCoupled Receptors in Living Cells

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Science's STKE  11 May 2004:
Vol. 2004, Issue 232, pp. pl9
DOI: 10.1126/stke.2322004pl9

Abstract

Recycling and degradation of plasma membrane receptors and transporters are fundamental mechanisms for regulating cell signaling and metabolic processes. For many membrane proteins, endocytosis reduces the number of molecules available for transport or signal transduction, providing an attenuation response. Fluorescent reporters attached to either the receptor or ligand have been used to monitor the trafficking of internalization; however, these approaches provide poor resolution for the early endocytic response. Here, we describe the use of a spin-labeled ligand for a heterotrimeric guanine nucleotide-binding protein (G protein)-coupled receptor for measuring the kinetics of endocytosis in real time. Included are protocols for designing a nitroxide-labeled ligand and measuring receptor endocytosis in live cells using electron paramagnetic resonance (EPR) spectroscopy. Methods for the evaluation of the receptor binding and activation properties of modified ligands and the generation of a cell line stably expressing high receptor levels are also provided.

Introduction

Endocytosis of heterotrimeric guanine nucleotide-binding protein (G protein)-coupled receptors (GPCRs) is a major mechanism for regulating the signaling levels achieved by this large protein family. The spatial and temporal aspects of endocytosis have been investigated for multiple GPCRs [reviewed in (14)]. Endocytosis serves multiple functions [reviewed in (5)], including sequestration, which is defined as rapid and reversible removal of the receptor, and down-regulation through targeted degradation of the receptor to the lysosome or proteasome. In a more sophisticated process, endocytosis may also modulate signaling pathways through vesicular signaling scaffolds (5).

Historically, endocytosis of cell surface receptors was detected by the use of radioligands (6, 7). In the early 1990s, use of fluorescently labeled ligands allowed for direct observation of endocytosis (7). The application of green fluorescent protein (GFP)-receptor fusions has been particularly useful because the signal can be followed beyond the endosomal compartments. GFP-receptor fusions provide an advantage over the fluorescently tagged ligand approach, because the ligand is generally detached from the receptor upon encountering the acid environment of the endosome; thus, the receptor movement through the endosomal system cannot be followed under the acidic compartments (8, 9).

Fluorescence is a highly sensitive tool for tracking markers of endocytosis. However, kinetic observations require that cells be fixed for each time point. Furthermore, because of resolution limitations, light microscopy may not distinguish between proteins at the cell surface from proteins very early in the internalization process. As a result, many cells must be analyzed individually to obtain significant quantitative kinetics data.

Here, we provide detailed methods and commentary for the real-time monitoring of endocytosis for a GPCR expressed in Chinese hamster ovary (CHO) cells (10). Because it relies on a nitroxide-tagged ligand, the method is sensitive enough to detect early internalization events and does not require cell fixation or destruction. This assay provides kinetics data based on 105 cells in a single sample. In addition, because the reduced nitroxide is electron paramagnetic resonance (EPR)-silent (11), data analysis is straightforward. A detailed protocol is provided for performing quantitative, real-time kinetic endocytosis experiments using EPR.

The reader will find this protocol useful for considerations relevant to the design of a spin-labeled ligand. Because EPR is not as sensitive as fluorescence microscopy, it is critical to create a cell line that expresses levels of protein sufficient to allow observation of the bound spin-labeled ligand. Thus, we also provide a method for establishing a cell line that expresses high levels of the target protein.

Materials

Synthesis and Purification of Spin-Labeled Peptides

1,8-diazabicyclo(5,4,0)undec-7-ene, 2 mmol

Acetonitrile

Amine-reactive spin label: 1-oxyl-2,2,5,5-tetramethylpyrroline-3-carboxylate N-hydroxysuccinimide ester (Toronto Research Chemicals)

Dimethylsulfoxide (DMSO)

Ethylenediaminetetraacetic acid (EDTA)

HCl(1N aqueous)

NaHCO3

Spin-labeled peptide [Peptidogenic Research (http://www.peptidogenic.com/index.html)]

Trifluoroacetic acid

Cell Culture

6-well culture plates

24-well culture plates

Cloning discs (VWR #13196-102)

Cryotube 1.8 ml vials (NUNC brand; VWR, #66021-986)

DMSO, sterile (Sigma-Aldrich, #D-2650)

Dulbecco's phosphate-buffered saline, 1× solution without calcium and magnesium (DPBS) (Cellgro, #MT 21-031-CM)

Dulbecco's phosphate-buffered saline, 1× solution with calcium and magnesium (DPBS+) (Cellgro, #21-030-CV)

Ethanol (EtOH)

Falcon tissue culture dish, 10 cm(100 mm), tissue culture-treated by vacuum gas plasma

Falcon tube with cell strainer cap (Fisher Scientific, #08-771-23)

Fetal bovine serum (FBS) (Invitrogen, #16000-044)

Note: Thaw the FBS in a 37°C water bath and divide it into 25-ml aliquots in sterile, disposable 50-ml conical polypropylene tubes. Store aliquots at −20°C.

FLIP recombinant CHO cell line (Invitrogen, #R758-07)

Flp-In pcDNA 5/FRT vector (Invitrogen, #V6010-20)

Ham’s F-12 medium with glutamine (Mediatech, #MT 10-080-CM)

Hygromycin B, 50 mg/ml (Invitrogen, #10687-010)

L-Glutamine, 200 mM (100×) (Invitrogen, #25030-081)

Note: Divide the L-glutamine into 5-ml aliquots in sterile, disposable 15-ml conical polypropylene tubes. Store aliquots at −20°C.

Lipofectamine 2000 reagent (Invitrogen, #11668-027)

Opti-MEM medium (Invitrogen, #31985-070)

Penicillin, 10,000 U/ml with streptomycin, 10,000 μg/ml (100× P/S) (Mediatech, #MT 30-002-CI)

Note: Thaw and divide the 100× P/S into 5-ml aliquots in sterile, disposable 15-ml conical polypropylene tubes. Store aliquots at −20°C.

pOG44 vector (Invitrogen, #V6005-20)

Trypsin, 0.25% with EDTA, 1 mM (tryp-E) (Invitrogen, #25200-072)

Note: Thaw and divide the tryp-E into 5-ml aliquots in sterile, disposable 15-ml conical polypropylene tubes. Store aliquots at −20°C. Freeze-thaw for use a maximum of two times.

Zeocin, 100 mg/ml (Invitrogen, #R25001)

Radioligand Binding and Membrane Preparation

Bacitracin

Benzamidine (Sigma-Aldrich, #B-6506)

Bio-Rad Dc Protein Assay Kit (Bio-Rad, #500-0111)

Bovine serum albumin (BSA)

Cryotube 1.8 ml vials (NUNC brand; VWR, #66021-986)

Cytoscint scintillation fluid (ICN, #882453)

DMSO, sterile (Sigma-Aldrich, #D-2650)

EDTA

Phosphate-buffered saline (PBS; Cellgro, #MT 21040CM)

Leupeptin (Roche, #1529048)

MgCl2

MultiScreen HTS-GV plates, 96-well (Millipore, #MAGVN2210)

Protease Inhibitor Cocktail Set III, 5 vials (Calbiochem, #539134)

Scintillation vials, 7-ml, polyethylene screw-cap (Fisher Scientific, #0333720)

Staurosporine (Sigma, S4400)

Substance P peptide (Calbiochem, #05-23-0600)

Substance P, 3H-labeled, 50 μCi (Amersham, #TRK86)

Tris base

Trypan Blue stain

Ultrathin gel-loading pipette tip (Eppendorf)

Equipment

Cell culture incubator, humidified, set at 37°C, 5% CO2

Cell scraper/lifter (VWR, #29442-200)

Clay Adams Nutator orbital mixer (Becton-Dickinson)

Cryo 1°C "Mr. Frosty" freezing container (VWR, #55710-200)

Dounce tissue grinder, 15-ml capacity (VWR, #62400-642)

Graph Pad Prism (http://www.graphpad.com)

HPLC C4 column (for example, Phenomenex LUNA 300 A column)

JEOL X-band spectrometer fitted with a loop gap resonator (12, 13) (Molecular Specialties, Milwaukee, WI)

Laminar flow hood dedicated for tissue culture procedures

Lyophilizer

MultiScreen carrier rack for 7-ml vials (Millipore, #MACR08127)

MultiScreen multiple punch (Millipore, #MAMP09608)

MultiScreen vacuum manifold basic kit (Millipore, #MAVM0960R)

Scintillation counter (such as Beckman LS6000TA)

Vacuum aspirator for tissue culture hood

Vacuum centrifuge, tabletop (for example, Speed-Vac)

Recipes

Note: Recipes 1 through 4 apply to synthesis and purification of spin-labeled peptides.
Recipe 1: 5% NaHCO3, pH 9.0
Prepare a 5% solution of NaHCO3 in a solution of 1:1 acetonitrile and water. Adjust pH with 1 M NaOH.
Recipe 2: Aqueous HPLC Solution
Prepare a 0.1% solution of trifluoroacetic acid in water.
Recipe 3: Organic HLPC Solution
Prepare a 0.1% solution of trifluoroacetic acid in acetonitrile.
Recipe 4: 5% NaHCO3, pH 7.5
Prepare a 5% solution of NaHCO3 in a solution of 1:1 acetonitrile and water. Adjust pH with 1 M HCL.
Note: Recipes 5 through 9 refer to cell culture as specifically applied to this Protocol. Use proper sterile technique for making all cell culture solutions (see the "Cell Culture" section below).
Recipe 5: Complete Ham’s F-12 Medium (Ham’s F-12 C)
Ham’s F-12 medium with glutamine500 mlFBS50 ml
100× P/S 5 ml
Thaw the FBS and 100× P/S in a 37°C water bath for about 10 min, then add them to the bottle of Ham’s F-12 medium.
Note: If the medium is older than 2 weeks, add 5 ml of 200 mM L-glutamine.
Recipe 6: Complete Ham’s F-12 Medium with Zeocin (Ham’s F-12 C+Zeo)
Ham’s F-12 C (Recipe 5)555 ml
Zeocin0.555 ml
Pipette the zeocin into the Ham’s F-12 C (Recipe 5). The final concentration of zeocin will be 100 μg/ml.
Note: If the medium is older than 2 weeks, add 5 ml of 200 mM L-glutamine.
Recipe 7: Freezing Medium
Ham’s F-12 medium with glutamine7 ml
FBS2 ml
DMSO, sterile1 ml
Mix solutions in a sterile, disposable 15-ml conical polypropylene tube. Store at −20°C. This solution can be frozen and thawed as often as needed.
Recipe 8: Ham’s F-12 Without Antibiotics
Ham’s F-12 medium with glutamine500 ml
FBS50 ml
Thaw the FBS in a 37°C water bath for about 10 min, then add it to the bottle of Ham's F-12 medium.
Recipe 9: Complete Ham's F-12 Medium with Hygromycin B (Ham's F-12 C+Hyg)
Ham's F-12 C (Recipe 5)555 ml
Hygromycin B666 ml
Pipette the hygromycin into the Ham’s F-12 C (Recipe 5). The final concentration of hygromycin will be 600 μg/ml.
Note: If the medium is older than 2 weeks, add 5 ml of 200 mM L-glutamine.
Note: Recipes 10 through 15 apply to preparation of membranes from CHO cells.
Recipe 10: Antibody Buffer
DPBS+500 ml
100× P/ S 5 ml
Add P/S to DPBS+ and mix.
Recipe 11: 1000× Protease Stock
Benzamidine100 mg
Leupeptin25 mg
Ethanol10 ml
Mix solutions in a sterile, disposable 15-ml conical polypropylene tube. Store at −20°C.
Recipe 12: Lysis Buffer
10 mM Tris-HCl, pH 7.4
1 mM EDTA
The volume of solution to be made will depend on amount of membrane protein desired. Add 1000× Protease Stock (Recipe 11) to a concentration of 1× just before use.
Recipe 13: Storage Buffer
50 mM Tris-HCl, pH 7.4
3 mM MgCl2
The volume of solution to be made will depend on amount of membrane protein desired. Add 1000× Protease Stock (Recipe 11) to a concentration of 1× just before use.
Recipe 14: Binding Buffer
50 mM Tris-HCl, pH 7.4
3 mM MgCl2
0.04 mg/ml BSA
100 mM NaCl
1× Protease Inhibitor Cocktail Set III
The volume of solution to be made will depend on amount of membrane protein desired. Add Protease Inhibitor Cocktail Set III (100× stock) to a final concentration of 1× just before use.
Recipe 15: Rinse Buffer
75 mM Tris-HCl, pH 7.4
12.5 mM MgCl2
1 mM EDTA
The volume of solution to be made will depend on amount of membrane protein desired. This solution can be made in a larger volume than is needed for a single experiment, and the excess stored at 4°C.
Recipe 16: Cell Binding Buffer (CB)
PBS
0.2 mg/ml BSA
1 mM MgCl2
0.1 mg/ml bacitracin
Add the BSA and MgCl2 directly to the desired volume of PBS. Add the powdered bacitracin and mix thoroughly.

Instructions

Designing the Spin-Labeled Peptide

The most straightforward approach for obtaining the peptide is commercial synthesis. Some companies provide custom modification to the synthesized peptides, such as attachment of fluorescent groups. Our collaborator, Peptidogenic Research, synthesized the substance P peptide and attached the amine-reactive spin label to the lysine at position 3 within the peptide. Before undertaking the synthesis, certain requirements must be considered.

Determining the labeling position

Does the peptide contain an amine-reactive side chain, such as lysine? If not, then either the N terminus or a lysine-substituted position (provided that this is functionally tolerated; see below) can serve as adduct sites. In principle, a cysteine could serve as the target labeling site; however, cysteine-containing peptides are difficult to work with because of disulfide bridge formation in oxidative environments.

Biological effect of labeling

A literature search for the chemical sequence modifications to the peptide can reveal sites in the ligand that are critical for binding and activation. For example, the lysine we modified in the substance P peptide had been previously labeled with a danysl-fluorescent group (14). If little information is available about the effects of modification to the ligand of interest, a reasonable first choice may be to modify the N terminus, because its lower pKa allows for selective modification under controlled conditions even in the presence of endogenous lysines.

Determining the amount of labeled peptide required

The amount of peptide needed is largely dependent on where the label is attached. If the label is attached to the N terminus, then synthesis of as little as 1 to 3 mg of peptide should be sufficient. If an internal lysine residue is the target site, then selective labeling is carried out at a lower pH (~7.5) to maintain the N-terminal Fmoc protecting group (15, 16), which can be left in place after peptide synthesis. Unfortunately, labeling at the lower pH is very inefficient and much of the peptide remains unmodified. Therefore, 75 to 100 mg of peptide would be required for ε-amine labeling at a lower pH. Labeling is carried out at 5:1 (label:peptide) ratio.

The presence of a C-terminal amide

It is easy to overlook the fact that many natural peptides contain an amide at the C terminus. This modification can be readily included in the synthesis. The amide group is not reactive toward amine-reactive labels and therefore does not need protection by a blocking group.

Labeling the Peptide

Labeling and purification of peptides labeled on the N terminus

No deprotection step is required for peptides labeled on the N terminus. Do not attempt to spin-label peptide while it is attached to the synthesis resin. Although labeling peptides attached to the resin is convenient and efficient for groups that do not decompose (for example, most fluorescent probes), the use of a strong acid (such as trifluoroacetic acid) to cleave the peptide from the resin readily decomposes nitroxyl groups.

1. Dissolve 1 to 3 mg of the peptide (0.7 to 2.2 μmol) in 2 ml of 5% NaHCO3, pH 9.0 (Recipe 1).

Note: Save a small aliquot containing 10 to 30 μg of the unlabeled peptide, and fractionate by HPLC to determine where unlabeled peptide elutes. Acidify this aliquot as described in step 4.

2. Chill the solution of thoroughly dissolved peptide to 4°C and add 0.2 mmol of amine-reactive spin label.

3. Stir at room temperature for 2 hours.

4. Acidify the reaction mixture to pH 2 by dropwise addition of aqueous 1 N HCl. Check the pH with litmus paper.

5. Purify by HPLC on a Phenomenex LUNA 300 A column with the following conditions. Pump A: Aqueous HPLC Solution (Recipe 2). Pump B: Organic HPLC Solution (Recipe 3). Establish a 5 to 45% gradient from pump B over 60 min. Monitor elution peaks at 220 nm. Unreacted peptide will elute first. Collect all peaks that elute.

6. Send a small sample of each peak(10 to 50 μl) to be analyzed by mass spectrometry.

7. Lyophilize the samples from each peak using a tabletop vacuum centrifuge.

Labeling and purification of peptides labeled on internal residues

This second labeling procedure is based on labeling of Lys3 found within the substance P peptide. In this case, the N terminus must remain blocked with the Fmoc protecting group to prevent the labeling of the N-terminal amine and to achieve the desired labeling of the internal lysine amine.

1. Dissolve 67 mg of crudely synthesized substance P peptide (0.05 mmol) in 2 ml of 5% NaHCO3, pH 7.5 (Recipe 4).

Note: Save a small aliquot containing 10 to 30 μg of the peptide and examine by HPLC to determine where unlabeled peptide elutes. Acidify this aliquot as described in step 4.

2. Once the peptide has dissolved, chill to 4°C and add 0.2 mmol of amine-reactive spin label.

3. Stir at room temperature for 2 hours.

4. Acidify the reaction mixture to pH 2 by dropwise addition of aqueous 1 N HCl. Check the pH with litmus paper.

5. Purify by HPLC on a Phenomenex LUNA 300 A column with the following conditions. Pump A: Aqueous HPLC Solution (Recipe 2). Pump B: Organic HPLC Solution (Recipe 3). Establish a 5 to 45% gradient from pump B over 60 min. Monitor elution peaks at 220 nm. Unreacted peptide will elute first. Collect all peaks that elute.

6. Send a small sample of each peak(10 to 50 μl) to be analyzed by mass spectrometry.

7. Lyophilize the samples from each peak using a tabletop vacuum centrifuge.

8. Remove the Fmoc protecting group from the sample that contains the modified peptide.

Deprotection: Removal of the Fmoc group from the N terminus

1. Dissolve the modified purified peptide in 0.5 ml of DMSO that contains 0.2 mmol of 1,8-diazabicyclo(5,4,0)undec-7-ene.

2. Stir for 5 min at room temperature.

3. Purify using the same HPLC gradient described above in step 5.

4. Analyze peaks by mass spectrometry.

Creating High-Expression Cell Lines

It is essential to determine whether attachment of the spin label, lysine substitution into the peptide, or both, has perturbed the structural and functional properties of the peptide. In the case of the substance P peptide, we performed competition curves for calculating the concentrations at which unlabeled peptide inhibits 50% of the binding of the labeled peptide (IC50) to assess the structural integrity of the modified peptide. To confirm that the functional properties of the modified peptide were not affected, we tested the peptide's ability to activate the substance P receptor, which is called the neurokinin 1 receptor (Nk1R), by monitoring calcium mobilization. This allowed us to calculate concentrations that produce 50% of the maximal response (EC50) to demonstrate that the modified peptide could activate the receptor with the same efficacy as the native unmodified peptide. The nature of the functional assay will depend on the biological system of interest, as well as the cell line used to characterize the peptide. In the Chinese hamster ovary (CHO) cells used in this Protocol, receptors capable of activating phospholipase C (PLC) can be assayed by measuring calcium mobilization (17). For GPCRs that signal through adenylyl cyclase, activation can be observed by adenosine 3′,5′-monophosphate (cAMP) accumulation. From these functional assays, the EC50 values can be determined. Numerous protocols have been developed for determining both IC50 [refer to chapter 1 of (18)] and EC50 (17, 18) of peptide receptors. Graph Pad Prism is a useful software package for generating, analyzing, and fitting competition curves, saturation binding curves, and EC50 curves.

Below, we describe the preparation and analysis of a CHO cell line stably expressing high levels of Nk1R. This cell line can be used for the EPR analysis and for verifying the biological activity of the labeled peptide.

Design of the receptor cDNA

A number of factors should be considered when designing the cDNA for cloning the receptor.

The noncoding sequence should have the appropriate restriction sites for inserting the receptor gene into the vector, in this case the pcDNA 5/FRT vector.

The coding sequence should have the start codon (ATG) as part of a Kozak consensus sequence (ACCATGG) to allow for maximal expression of the receptor (19).

If desired for visualizing expression of the receptor, one of the GFP variants’ coding sequences can be cloned in frame with the receptor's coding sequence. In our experience, GFP has no effect on functional properties of the substance P receptor when fused to the C terminus of the receptor. Binding, activation, and internalization of the receptor are unaffected. In contrast, GFP fused to the N terminus severely decreases cell surface expression. The effects of GFP should be thoroughly investigated before considering using a GFP fusion protein as an alternative to the wild-type receptor. We currently use an eGFP variant, due to its enhanced fluorescence properties. Most of the GFP variants can be purchased in vectors sold by Clontech.

An alternative to a GFP fusion protein for visualizing receptor expression at the cell surface is to add an N-terminal epitope tag. The presence of an epitope tag will allow cell surface binding of fluorescently labeled antibodies to detect cells expressing the receptor. This is a required step when flow cytometry is used to enrich for cells with high receptor expression. Useful epitope tags include the FLAG, myc, and hemagglutinin (HA) peptide sequences. The DNA sequences that encode these small peptides can be found in Genbank. There are numerous commercial suppliers of antibodies directed against these epitope tags. It should be noted that, if one uses the M1 anti-FLAG monoclonal antibody, the first amino acid of the FLAG sequence must have its N terminus exposed. Therefore, we recommend placing a cleavable signal peptide in front of the FLAG sequence, which will expose the N terminus of the first amino acid in the FLAG sequence. Alternatively, the M2 anti-FLAG antibody does not utilize the N terminus of the first amino acid of the FLAG sequence as part of its epitope, so no cleavable signal peptide is required. As with the GFP, confirm that the presence of the N-terminal epitope tag has no effects on binding, activation, and internalization of the receptor.

General cell culture procedures

Creating a cell line that stably expresses high levels of the peptide receptor is the most critical parameter in successfully obtaining an EPR signal corresponding to spin-labeled peptide bound to the receptor on the surface of living cells. We chose a CHO cell line that had been engineered to yield high, stable expression of transfected proteins. Invitrogen developed this CHO line by inserting an integration site next to a transcriptionally active genomic locus. The receptor gene is transfected and becomes site-specifically integrated into the site. Integration is achieved through the presence of a transiently cotransfected recombinase that has recognition sequences in the genomic DNA and the plasmid DNA harboring the receptor gene. Site-specific integration promotes isogenic, high protein expression under the control of a cytomegalovirus (CMV) promoter. Conveniently, upon integration, the engineered cell line becomes sensitive to zeocin and gains resistance to hygromycin. This leads to selective survival of cells harboring the integrated gene, as well as selective killing of cells where site-specific integration has not occurred.

Proper sterile technique

The following are general guidelines for minimizing contamination of cultured cells and for maintaining a sterile work environment. Specific details may vary according to your particular cell line or research system.

Spray hands with 70% EtOH before working in the tissue culture hood.

Before transferring to the tissue culture hood, wipe off all medium bottles that have been prewarmed in the water bath and spray the bottles thoroughly with 70% EtOH, especially the cap area.

Spray the tissue culture hood surfaces with 70% EtOH before and after use.

Spray all tools and containers entering the hood with 70% EtOH and allow to air dry; do not wipe off excess.

Open and close all containers, tubes, and culture dishes in the tissue culture hood.

Minimize the time that the lids or caps are removed from culture dishes or flasks.

Never take the cells out of the tissue culture hood unless the lid or cap is on the dish or flask.

Cell, media, and buffer storage conditions

The following are general guidelines for storing cells and solutions used in most cell culture situations. The cells come in a small plastic "cryotube" on dry ice; store them in a liquid nitrogen freezer or proceed to thawing the cells. Cells are grown in culture medium in a humidified 37°C, 5% CO2 incubator. Cell culture media should be stored at 4°C and discarded after 30 days. Unless indicated in the Recipe, buffers can be stored at room temperature.

Thawing the cells

Note: Before thawing the cells, prewarm Ham’s F-12 C (Recipe 5) in a 37°C water bath for 20 min or until the bottle of medium is warm to touch.

1. Remove the frozen aliquot of cells from the dry ice or liquid nitrogen and swirl the tube in a 37°C water bath so that cells are thawed quickly.

2. Spray the vial with 70% EtOH and place in the tissue culture hood (do not wipe off excess EtOH).

3. Place the cells in a 10-cm culture dish with a sterile 1-ml pipette.

Note: Dishes must be tissue-culture treated or the cells will not properly adhere.

4. Pipette 8 ml of prewarmed Ham’s F-12 C (Recipe 5) into the dish with a sterile 10-ml pipette and gently swirl the dish to distribute the cells uniformly in the dish.

5. Place in the cells in a humidified 37°C, 5% CO2 incubator and allow the cells to recover and become confluent, in about 2 days.

Note: The cells are considered confluent when there is no more room for them to grow. A confluent 10-cm dish will contain about 7 × 106 cells. When cells begin to crowd a dish, or if a culture is unhealthy, many of the cells begin to detach from the substrate. Nonadherent cells are spherical; adherent cells spread out in oblong irregular shapes.

Passaging the cells

When the cells become 100% confluent, they should be divided so that there is sufficient room for new cell growth. Typically, CHO cells are split at a ratio of 1:4, which means 1/4 of the cells on the culture dish are placed into a new 10-cm culture dish, effectively increasing the available surface growing area by a factor of 4. After a 1:4 splitting, the cells will be 25% confluent. Because CHO cell divide every 24 hours, the cells will need to be split every 2 days (25% × 2 × n = 100%, where n = number of days). CHO cells can be split 1:8, but they do not grow as well at this cell density.

1. Prewarm the Ham’s F-12 C+Zeo (Recipe 6), DPBS, and one 5-ml aliquot of tryp-E in the 37°C water bath for 10 to 20 min.

2. Remove the cells from the incubator and place in the tissue culture hood.

3. Tilt the dish of cells to a 30° angle, and, with a vacuum aspirator, remove the medium from the cells by aspirating from the side of the dish.

Note: This technique will prevent the cells from being aspirated with the medium.

4. Add to the dish 5 ml of sterile DPBS, gently swirl the dish a few times, then remove the buffer by aspirating, as above.

Note: This removes the calcium and magnesium from the cells. Either calcium or magnesium will inhibit the trypsin in the tryp-E solution and prevent the cells from detaching from the culture dish.

5. Immediately pipette 1 ml of tryp-E onto the cells and tilt the dish back and forth until the solution covers the entire surface of the dish.

Note: This step will detach the cells and is commonly referred to as trypsinization.

6. Place the cells back into the humidified 37°C, 5% CO2 incubator for 2 to 5 min.

Note: Try to minimize the time spent on this step, because the trypsin is not only digesting cell attachment proteins but also other cell surface proteins. Excessive trypsinization of the cells will cause them to grow very slowly.

7. Take the cells out of the incubator and gently agitate the dish to check whether the cells have detached. Holding the dish above eye level or use of a microscope assists in this observation. If the cells have not detached sufficiently, return to incubator and check for detachment again in 2 min.

Note: Cells may adhere to each other in clumps or small sheets when they detach from the cell culture dish. This is normal. Do not wait for cells to become completely separated and suspended.

8. Add 7 ml of prewarmed Ham’s F-12 C+Zeo (Recipe 6). Mix the medium with the cells by gently pipetting the cells up and down in a 10-ml sterile pipette 5 to 10 times. This will prevent clumping of the cells and ensure even seeding of the cells.

9. Remove 1/4 of the cells(2 ml) and place them into a new 10-cm culture dish that contains 6 ml of prewarmed Ham’s F-12 C+Zeo (Recipe 6). Each 10-cm dish of confluent cells can be split into four new 10-cm dishes.

10. Gently swirl the dishes to evenly distribute the cells onto the surface of the dish.

Note: We refer to splitting cells with respect to the cell surface area on the dish. A 10-cm dish has a surface area of 60 cm2, so a 1:4 split results in increasing the surface area to 4 × 60 cm2 = 240 cm2, or the equivalent of four 10-cm dishes. To pass the cells from a 10-cm dish into a T75 flask (which has a surface area 31% of four 10-cm dishes), then 2.5 ml--or 31% of the total 8 ml cell volume--is transferred into a single T75 flask.

11. Place the four culture dishes back into the incubator and check the cells for confluency after 2 days.

12. Replace the medium with fresh, warmed Ham’s F-12 C+Zeo (Recipe 6) if cells are not confluent, or split the cells 1:4 into sixteen 10-cm dishes if they are confluent.

Note: The medium should be changed every 2 days if the cells are not split. The cells from this step will used to prepare frozen stocks(15 of 16 dishes), and one dish of cells will be passaged for stable transfection.

Making cell stocks

1. Prewarm Ham’s F-12 C+Zeo (Recipe 6), Freezing Medium (Recipe 7) and a 5-ml aliquot of tryp-E.

2. Follow steps 1 through 7 in "Passaging the cells" (above) to detach the cells for all 16 dishes.

3. Continue the remainder of the steps of "Passaging the cells" (above) with one of the 16 dishes, and split the cells 1:4 into four dishes. Return these dishes to the incubator.

4. Add 5 ml of Ham’s F-12 C+Zeo (Recipe 6) to each of the remaining 15 dishes after cell detachment is achieved in step 7 in "Passaging the cells" (above).

5. Combine the cells from the 15 dishes and pipette the cells equally (about 45 ml per tube) into two sterile 50-ml disposable, conical centrifuge tubes.

6. Centrifuge for 5 min at 1000 rpm or 600g.

7. Remove and discard the supernatant by aspirating or pipetting.

8. Resuspend the cells in 15 ml of Freezing Medium (Recipe 7).

Note: Typically we use 1 ml of freezing media for each 10-cm dish of confluent cells. Cells are frozen at a density of 1 to 7 × 106 cells/ml.

9. Pipette 1 ml of cells into each cryotube and tightly screw on the lid.

10. Place the vials into a freezing container and cool the tubes overnight at −80°C.

Note: The freezing container is designed to cool the cells at a rate of −1°C/min. This cooling rate is necessary for successful cryopreservation of cells.

11. Transfer the cells to the liquid nitrogen storage vessel for long-term storage.

Stable transfection of receptor into FlpIn(+) CHO cells

This step will take anywhere from 6 to 9 weeks, depending on how fast the cells grow. Transfections are performed in four 10-cm dishes of confluent cells. One dish serves as a positive control and another as a negative control; the remaining dishes are transfected with the receptor gene of interest (Table 1). The transfection protocol is a modification of the Lipofectamine 2000 protocol for the transfection of four 10-cm dishes.

Table 1.

DNAs for transfection. Dish 3 is the empty vector control (EVC), a positive control for transfection efficiency, and dish 4 is a negative control.

1. Split the cells 1:2 in Ham’s F-12 C+Zeo (Recipe 6) on the morning of the day before transfection.

Note: The goal is to achieve 50% confluency 24 hours before transfection.

2. Grow the four dishes of cells overnight in a humidified 37°C, 5% CO2 incubator.

3. The next morning, aspirate and discard culture medium.

4. Pipette 5 ml of DBPS into each dish and swirl gently.

5. Aspirate DBPS and replace with 15 ml of Ham’s F-12 Without Antibiotics (Recipe 8).

6. In three sterile 15-ml centrifuge tubes, mix 20 μg (~20 μl) of the appropriate vector DNA mixtures (Table 1) with 1.5 ml of Opti-MEM.

Note: The fourth dish is the nontransfected negative control.

7. In three other 15-ml sterile centrifuge tubes, mix 75 μl of the Lipofectamine 2000 reagent with 1.5 ml of Opti-MEM.

8. Combine the DNA solutions with the Lipofectamine-Opti-MEM solutions.

9. Incubate at room temperature for 30 min to allow the DNA to form complexes with the Lipofectamine reagent.

10. Add the proper DNA-Lipofectamine solution to the corresponding dish.

11. Place all four dishes in a humidified 37°C, 5% CO2 incubator and incubate overnight.

12. The next day, passage the cells as described above, and at the detachment step (step 7 of "Passaging the cells"), resuspend the cell suspension in 8 ml of prewarmed Ham’s F-12 C+Hyg (Recipe 9).

Note: At this point, the medium does not contain zeocin anymore, but instead contains hygromycin for selection.

13. Place 8 ml of prewarmed Ham’s F-12 C+Hyg (Recipe 9) into each of 20 fresh 10-cm dishes.

14. Seed the cells into these 20 new dishes at varying cell densities (four dishes at each density): 1:500(16 μl), 1:200(40 μl), 1:100(80 μl), 1:50(160 μl), and 1/20(400 μl) of the cell density of each original dish.

Note: Volumes are for the resuspended cells from step 12. Freshly detached cells will quickly reattach, because the culture medium contains calcium and magnesium; therefore, have the new dishes ready with the Ham’s F-12 C+Hyg.

15. Replace the medium every 2 days with prewarmed Ham’s F-12 C+Hyg (Recipe 9) for the next 2 to 3 weeks.

Note: Individual colonies from single cells should start to be visible by eye around the third week.

16. When the colonies reach 1 to 2 mm in size (about the diameter of the tip of a large Sharpie permanent pen marker), they may be picked and transferred to 24-well plates.

Note: The negative control dishes should have no cells growing on them after 3 weeks, because the cells are hygromycin-sensitive. If any of the negative control dishes that were plated at higher cell densities (for example, 1:20 or 1:50) contain surviving cells, throw out all dishes containing cells that were plated at these densities. This will prevent picking falsely positive colonies.

Picking colonies

After the cell colonies have reached 1 to 2 mm in diameter, or about 200 to 600 cells per colony, they may be transferred to individual wells for further growth in a 24-well plate. After transferring, it can take 1 to 2 weeks before the cells become confluent. Pick 20 colonies for the cells transfected with the receptor cDNA and five colonies for the cells transfected with the empty vector (positive) control (EVC).

1. Remove 25 cloning discs with sterilized forceps and soak them in a 60-mm dish of prewarmed (37°C) tryp-E. Soak the discs for 2 to 3 min just before use in step 5.

2. Aspirate the medium from a 10-cm dish and replace with 5 ml of DPBS; remove the DPBS by aspiration and place the lid back on the plate.

3. Take the plate out of hood and hold it above eye level. At this position, the cell colonies (white spots) should be visible. Using a permanent marker, quickly mark each colony directly under the colony.

Note: Work fast, because the cells can dry out without buffer.

4. Re-wet the dish with another 5 ml of DPBS, then remove the DPBS by aspiration.

5. Place the prewetted cloning discs (one per colony) directly on top of the colonies. Place the lid back on the dish and incubate for 2 to 5 min at 37°C, 5% CO2.

6. Add 0.5 ml of Ham’s F-12 C+Hyg (Recipe 9) to each well of a 24-well plate.

7. Remove each disc with sterilized metal forceps and place it in its own well of the 24-well plate.

Note: Sterilize the metal forceps with ethanol and flame between each disc to avoid mixing cells from the different colonies.

8. Change the medium every 2 days until the cells become confluent (in 1 to 2 weeks).

Quick confirmation of expression (receptor-GFP fusion constructs)

Just before the cells in the 24-well plate are passaged into a larger dish, it is useful to evaluate how many clones are actually expressing the receptor of interest. If the receptor is linked to GFP, then cells can be evaluated under a fluorescence microscope to determine which clones are expressing the receptor.

1. Aspirate medium from cells.

2. Add 0.5 ml DPBS to the cells.

Note: This wash removes phenol red from the medium that may make it difficult to see the GFP fluorescence.

3. Check for fluorescence with 470 nm excitation and 525 emission filters at 200× magnification.

4.Mark the wells containing cells that fluoresce and transfer 5 to 10 of these positive clones.

5. Discard the remainder.

Transfer of cells

When cells in the 24-well plate become confluent, they will need to be transferred to a larger dish. If the receptor is fused to GFP, transfer 5 to 10 of the positive clones identified above. If the receptor is not fused to GFP, keep all clones and transfer them to the 6-well plate, as described below. For the EVC dish, keep three to five clones.

1. Add 2 ml of Ham’s F-12 C+Hyg (Recipe 9) to each well of a 6-well plate (enough for one well per clone).

2. Remove medium from the clones in the 24-well plates by aspiration.

3. Rinse with 0.5 ml of DPBS and aspirate.

4. Add 100 μl of prewarmed tryp-E to each well.

5. Incubate at 37°C, 5% CO2 incubator for 3 to 5 min.

6. Add 0.5 ml of Ham’s F-12 C+Hyg (Recipe 9) to quench the trypsin.

7. Immediately resuspend cells by pipetting 5 to 10 times with a 1-ml pipette and transfer each clone a new well of the 6-well plate.

8. Incubate at 37°C, 5% CO2 for 3 to 5 days until the cells become confluent.

9. Add 8 ml of Ham’s F-12 C+Hyg (Recipe 9) to each 10-cm dish (one dish per clone).

10. Aspirate the medium from the clones in the 6-well plates.

11. Rinse with 5 ml of DPBS and aspirate.

12. Add 300 μl of prewarmed tryp-E to each well.

13. Incubate at 37°C, 5% CO2 for 3 to 5 min.

14. Add 2.0 ml of Ham’s F-12 C+Hyg (Recipe 9) to quench the trypsin.

15. Pipette 5 to 10 times to resuspend the cells, then add to the 10-cm dishes.

16. Incubate cells for about 1 week, replacing the medium every 2 days, until the cells are confluent.

17. Split each 10-cm dish 1:4, preparing four dishes per clone. Use two dishes to prepare a frozen stock of cells, and continue growing one dish as a backup. The last dish is used for cell sorting by flow cytometry.

Sorting the cells by flow cytometry

This step is performed to ensure that all cells are expressing the receptor (or other target protein). If the target protein is not a GFP fusion, a fluorescently tagged antibody or ligand can be used to identify cells expressing the protein. The antibody should be capable of binding to the receptor in live cells. Receptors with N-terminal fusion epitopes (for example, FLAG or Myc or HA) generally work well for this purpose.

Each 10-cm dish of confluent cells will yield ~7 × 106. The optimal density for cell sorting is 1 × 106 cells/ml. Cells from the EVC dish, as well as those from the receptor-transfected clones, should be sorted. The EVC cells will help the flow cytometer operator determine the autofluorescence of the cells that are not expressing the receptor-GFP fusion, or the background fluorescence arising from nonspecific binding of the fluorescently labeled antibody. Because most sorters are able to process ~1000 cells, easily 2 million cells can be processed in a half hour. If 20% of cells per clone are expressing the target protein at high levels, then ~400,000 cells will be collected. The cells expressing the receptor should give a population of cells with a much higher intensity of fluorescence than the EVC cells. This serves as a fivefold enrichment or increase in effective receptor concentration. There should be no need to isolate individual cells, because integration of the receptor gene is uniform, leading to isogenic expression of the receptor.

1. Detach cells, as described above ("Passaging the cells"), from each 10-cm plate to be sorted.

2. Quench the trypsin with 7 ml of DPBS+ to give a concentration of 1 × 106 cells/ml.

3. Resuspend the cells well to prevent clumping.

Note: Flow cytometry may require cell filtration to prevent clumping. Pass the cells through a Falcon tube with a cell strainer cap by pipetting the cells directly onto the strainer cap and allowing the cells to pass through the cap by gravity filtration.

4. If the cells are expressing a receptor-GFP fusion, sort the cells by flow cytometry and proceed to step 9.

5. If the cells must be labeled with antibody, add the fluorescently labeled antibody (using the appropriate dilution) to both cells transfected with the receptor cDNA and cells transfected with the empty plasmid (EVC).

6. Incubate the cells at room temperature for the time suggested by the antibody manufacturer (typically 1 hour).

7. Rinse the cells with 5 ml of DPBS+ and pellet the cells at 500g for 5 min.

8. Resuspend the cells in 7 ml of DPBS+ and pipette well to form a single-cell suspension just before sorting.

9. Collect the highly fluorescing cells.

Note: The volume of collected cells will range from 0.5 to 2 ml. If the volume is greater than 3 ml, pellet the cells at 500g for 5 min and resuspend in 3 ml of Ham’s F-12 C+Hyg (Recipe 9).

10. Seed the cells into a well of a 6-well plate.

Note: Cells should be plated in a dish of a size that will allow cells to become confluent in 1 to 2 weeks. Thus, if 200,000 cells are collected, a well of a 6-well plate allows the cells to be seeded at ~16% confluency (optimal), whereas a 10-cm dish would only be 2.8% confluent (too low).

11. Add 3 ml of Ham’s F-12 C+Hyg (Recipe 9).

12. Gently swirl the cells to evenly distribute them.

13. Incubate overnight.

14. Replace the medium the next day, and then every 2 days until the cells are confluent.

Note: The cells will become confluent in 1 to 2 weeks.

15. Transfer the cells to a 10-cm dish as described above ("Transfer of cells") and grow until confluent.

16. Split the confluent cells 1:4 into four 10-cm dishes.

17. For cells expressing the receptor-GFP fusion, check by fluorescence microscope to confirm that all cells are fluorescing, then proceed to step 21.

Note: Periodically repeat this step to confirm that no loss in receptor expression has occurred.

18. If using fluorescent antibodies, add the appropriate amount of antibody to 8 ml of Antibody Buffer (Recipe 10), and add this directly to one of the 10-cm dishes.

19. Incubate at room temperature for the appropriate amount of time and the remove the buffer.

20. Compare fluorescence of cells from the EVC to the staining of the sorted cells. If all of the sorted cells are fluorescing, then the integration of the receptor is stable.

Note: Periodically repeat this step to ensure that no loss in receptor expression has occurred.

21. Passage the cells enough times to freeze down 20 aliquots of the desired clones and aliquots of the EVC cells.

Characterizing the Cell Line

The expression levels of the stable cell line, as well as the dissociation constant (Kd), should be determined for the cells by performing saturation binding assays using the appropriate radioligand. An excellent, exhaustive source of binding protocols for GPCRs can be found in the online version of Current Protocols in Pharmacology (18). We give our protocol below for the preparation of membranes from the CHO cell line that has been stably transfected with the neurokinin-1-GFP chimera, as well as the saturation binding protocol used. This binding protocol is a good starting point for other receptors; however, binding buffers, ligands, and filters may vary. Start with the same Millipore GV filters described below, and if nonspecific binding is too high (>20 to 30%), try pretreating the filters or try another filter type. The membrane preparation should not need to be modified.

Membrane preparation

As a general rule, we prepare 4 mg of membrane proteins per experiment. A 10-cm dish of confluent cells will give ~1 mg of membrane protein, so four dishes of cells are combined to achieve the desired concentration of membrane proteins(4 mg/ml).

Note: Four confluent 10-cm dishes will be needed. All steps should be done on ice or at 4°C.

1. Rinse dish twice with 15 ml of DPBS+ and remove the final rinse.

2. Add 5 ml of Lysis Buffer (Recipe 12).

3. Using a cell scraper, scrape the cells and transfer the 5 ml of scraped cells to the next dish; continue scraping and transferring the cells until all four plates have been scraped with the same 5 ml of lysis buffer.

4. Transfer the scraped cells to a 15-ml conical tube.

5. Rinse the first dish with an additional 10 ml of Lysis Buffer (Recipe 12) and transfer this rinse sequentially into the other dishes, so that each dish is rinsed with the same 10 ml of lysis buffer.

6. Add the 10-ml rinse to the scraped cells in the 15-ml tube.

7. Pour the 15 ml of scraped cells into a 15-ml Dounce tissue grinder and homogenize the cells(50 to 100 strokes of the piston).

8. Check 20 μl by light microscope to verify that the cells are no longer intact. If they are, give the suspension in the Dounce tissue grinder another 50 piston strokes and check again. Repeat as needed until 80% of the cells are no longer intact.

9. Centrifuge in a 15-ml or 50-ml conical plastic disposable tube in a tabletop centrifuge at 1500 rpm for 10 min.

10. Transfer supernatant into a clean high-speed centrifuge tube, being careful not to disturb the pellet.

11. Centrifuge at 18,000 rpm in a Sorvall SA-800 rotor for 15 min.

12. Discard the supernatant and resuspend membranes in 10 ml of Storage Buffer (Recipe 13).

13. Centrifuge again at 18,000 rpm for 20 min and discard supernatant.

14. Resuspend the membranes in 1 ml of Storage Buffer (Recipe 13) using a 200-μl pipetter until the pellet is completely suspended.

15. Perform a protein assay according to the manufacturer's instructions.

Note: Be sure to use Storage Buffer (Recipe 13) for diluting standards as well as unknowns, so that any additional absorbance from the protease inhibitors is subtracted out.

16. Freeze the membrane proteins in 100-μl aliquots and store at −80°C.

Saturation binding

This Protocol is designed to give triplicate data that can be fit to a saturation binding curve by GraphPad Prism software. Their Web site (http://www.graphpad.com) also provides tools for calculating radioisotope conversions and radioligand binding. This manual should be read thoroughly to avoid or troubleshoot common pitfalls associated with radioligand binding. It will also help in tailoring the binding protocol below to other receptor-ligand systems. The Kd will influence how long the binding assay is performed, as well as the concentration range used for the experiment. In most cases, both parameters can be found in the literature, in addition to specific binding protocols for each receptor (18).

This procedure uses radioactive materials. Proper handling and disposal procedures must be followed.

Note: All solutions should be ice cold.

1. Make stocks of radioligand indicated in Table 2, using Binding Buffer (Recipe 14) to dilute the radioligand.

Table 2.

Radioligand stocks. The designated stock volumes will be sufficient to obtain two sets of triplicate data. One set is used to obtain total binding. This set uses membranes lacking the cold ligand. The second set is used to obtain nonspecific binding values. This second set uses the membranes that have had the cold ligand added to them.

2. Pipette the designated volumes (Table 2) of radioligand stock into ten 7-ml plastic scintillation vials.

Note: Preparation of these vials (steps 2 through 4) for the radioligand can be done while the membranes and radioligand are incubating in step 15 of this procedure, if desired.

3. Add 5 ml of Cytoscint scintillation fluid and vortex briefly to mix.

Note: These samples will provide the counts per minute (CPMs) for the free ligand. If any of the CPMs obtained from binding samples are more than 10% of the CPMs obtained from the corresponding free ligand concentration, then ligand depletion has occurred and true equilibrium binding conditions are not met. If this is the case, repeat the experiment with less membrane.

4. Pipette 5 μl of the commercial stock solution of radioligand into a scintillation vial, add 5 ml of scintillation fluid, and vortex briefly. This will give a working specific activity of the ligand.

5. Prepare a 96-well Millipore GV-plate by filling the wells with 200 μl of water and then removing the water by filtration using the Millipore vacuum filtration unit.

6. Add the appropriate amount of Binding Buffer (Recipe 14) to the designated well (Table 3).

Table 3.

Saturation binding assay. 1Wells A1 through C10 receive the membranes from the total binding tube and wells D1 through F10 receive membranes from the nonspecific binding tube (shown in parentheses). 2Volume of stock ligand used in both the Millipore wells and in the Cytoscint vials for counting.

7. Add the designated amount of stock 3H-radioligand to the corresponding wells.

8. Thaw two 100-μl aliquots of membranes and place on ice. One aliquot will be used for total binding; the other aliquot will be used for nonspecific binding.

9. Dilute each aliquot of membranes to 0.1 mg/ml with Binding Buffer (Recipe 14).

10. To ensure that the membranes are thoroughly resuspended, transfer the suspension to a 15-ml Dounce tissue grinder and homogenize the membranes(30 strokes).

11. Transfer the membranes back to the same tubes.

Note: Do this right before adding the cold ligand to the nonspecific tube and adding the membranes to the wells of the GV-plate.

12. For the "nonspecific" tube, add cold ligand to a final concentration that is at least 1000× higher than the Kd of the radioligand.

Note: For the substance P receptor, we add 8 μl of 1 mM substance P to the 4 ml of diluted membranes for a concentration of 2 μM (this will give a final concentration of 1 μM after the membranes have been added to the binding reaction).

13. Add 100 μl of the diluted membranes to each well of the GV-plate.

Note: Invert the membrane suspension after adding to every 3 to 5 wells to prevent the membranes from settling. Failing to keep the membranes well suspended is a major source of error.

14. Mix each well thoroughly with a 12-tip multipipetter.

Note: The same tips can be used for wells A1 through C10. The tips should be changed and then this second set of tips can be used for D1 through E10. Inconsistent pipetting and mixing are the biggest sources of error, so be sure to pipette and mix each sample in a consistent manner. For mixing, mix the same number of times for each sample. For pipetting, always dispense the entire volume.

15. Incubate the plate for 1 hour at room temperature.

Note: This time period is sufficient to reach equilibrium for all concentrations used in the case of substance P binding to the neurokinin 1 receptor. Times may need to be adjusted for other ligand-receptor combinations.

16. Chill the plates at 4°C for 10 min.

17. Quickly filter the membranes by placing the 96-well plate on the Millipore vacuum manifold that is connected to a standard laboratory vacuum line.

18. Keeping the plate under vacuum, immediately rinse each well (use a 12-tip multipipetter) three times with 200 μl of ice-cold Rinse Buffer (Recipe 15). Allow each rinse to go completely through the well before adding the next rinse. This will minimize background counts.

19. Remove the plastic backing from the bottom of the plate and gently blot off the underside of the filters and allow the filter to dry.

20. Punch the filters out of each well with the MultiScreen Multiple Punch, and place them into 7-ml scintillation vials containing 5 ml of Cytoscint that have been placed in the MultiScreen Carrier Rack.

21. Count the vials (from steps 3, 4, and 20 of this procedure) the next day.

Binding assay analysis

1. Average the CPMs for the triplicate data.

2. Subtract nonspecific CPMs from total CPMs to obtain specific binding.

Note: Our specific counts ranged from 15 to 1200 CPMs.

3. Convert from CPMs to pmol or fmol using the working specific activity (divide by the value of the radioligand’s working specific activity in CPMs/pmol).

4. Divide the number of moles by the amount of membrane protein used in the assay(10 μg) and convert to fmol/mg to convert the values to the amount of ligand bound/mg of membrane protein.

Note: Alternatively, calculate the concentration of bound ligand in the assay by dividing the number of pmol of bound ligand by the assay volume(200 μl) and convert to nM.

5. Using nonlinear regression, fit data to either a one-site or two-site hyperbola (saturation curve).

6. Repeat the binding experiment two more times, and fit the average of the three independent experiments.

EPR Analysis with Spin-Labeled Peptide

Binding of spin-labeled peptide ligand to live cells

This step will confirm specific binding of the spin label to the receptor on live cells. EPR spectra should be acquired for cells expressing and cells not expressing the receptor. There should be no signal from cells lacking the receptor. In most cases, the signal from the bound spin-labeled ligand should display more hyperfine anisotropy, and be distinguishable from the signal of the free ligand (that is, the bound spectrum is expected to be broader than the free spectrum). A chase sample is also used to calculate the concentration of spin-labeled ligand bound to the cell surface. In the chase sample, the EPR signal should change from the bound spectrum to a sharper spectrum resembling the free ligand (Fig. 1). In addition, the total number of surface receptors must be determined to design control experiments for the mechanism of nitroxide reduction (described in detail below in "Calculation of Bound Spin-Labeled Ligand Concentration").

Fig. 1.

EPR scans of the different binding conditions of spin-labeled substance P (SP). (A) The free ligand displays a sharp spectrum with little anisotropy. (B) In contrast, the spectrum from the ligand bound to live cells expressing Nk1R-GFP is broadened because of its immobilization (black trace). Spin-labeled peptide is completely chased-off the Nk1R-GFP when unlabeled SP is added to the cells in excess (gray trace). Spectra of samples at room temperature(20 to 22°C) were obtained by a single 120-s scan over 100G at a microwave power of 2 mW, a receiver gain of 500 and a modulation amplitude optimized to the natural line width of the individual spectrum.

We use a conventional X-band EPR spectrometer fitted with a loop gap resonator. This configuration provides high sensitivity with minimal sample volume (~4 μl). Dielectric resonators from Bruker provide similar advantages. For instrumental parameters, samples were scanned over 100 G at a microwave power of 2 mW, using a scan time of 60 s, a time constant of 0.1 s, a modulation width optimized to the natural line width of the signal (typically on the order of 1 G).

Use two 10-cm dishes confluent with cells stably expressing the target protein, and two confluent dishes of CHO cells lacking the protein.

1. Rinse the cells in the dishes two times with 15 ml of DPBS.

2. Add 5 ml of DPBS containing 1 mM EDTA and place the cells in a 37°C incubator for 5 min.

Note: All subsequent steps should be performed on ice or in a 4°C cold room.

3. Scrape the plates and combine the cells into a 15-ml tube.

4. Centrifuge the cells at 600g for 5 min and discard the supernatant.

5. Rinse the cells by resuspending in 5 ml of CB (Recipe 16).

6. Centrifuge the cells at 600g for 5 min and discard the supernatant.

7. Resuspend the cells at a density of 4 × 106 cells/ml in CB (Recipe 16).

8. Place 0.5 ml of cells into a 1.5-ml microcentrifuge tube, centrifuge at 600g for 5 min, and discard the supernatant.

9. Resuspend the cells in 1 ml of CB (Recipe 16) containing 1 μM of spin-labeled ligand.

10. Gently rock the cells on a Nutator orbital mixer at 4°C for 2 hours.

11. Centrifuge the cells at 600g for 5 min and discard the supernatant.

12. Rinse the cells by resuspending in 1 ml of CB (Recipe 16), centrifuging at 600g for 5 min, and discarding the supernatant.

Note: More than one rinse may be required to remove any signal due to nonspecific binding, which is measured by an EPR signal from the cells not expressing the protein. We do not observe an EPR signal from nontransfected cells after one rinse.

13. Resuspend the cells in 20 μl of CB (Recipe 16).

Note: This will be a very thick suspension.

14. Using an ultrathin gel-loading pipette tip, remove 10 μl of cell suspension and load it immediately into a flame-sealed capillary of appropriate dimensions for the EPR resonator.

15. Spin the capillary at 500g in a chilled centrifuge for 3 min. The cells should be loosely packed at the bottom of the capillary and should be separated from the buffer.

16. Acquire the EPR spectra for the samples while maintaining the samples at 4°C.

17. Add unlabeled ligand to the remaining 10 μl of cell suspension for a final concentration of 1 to 10 μM and incubate 30 to 60 min on ice.

Note: This is the chased sample.

18. Using an ultrathin gel-loading pipette tip, remove 10 μl and load the cell suspension immediately into a flame-sealed capillary of appropriate dimensions for the EPR resonator.

19. Spin the capillary at 500g in a chilled centrifuge for 3 min. The cells should be loosely packed at the bottom of the capillary and should be separated from the buffer.

20. Acquire the EPR spectrum at the same setting used for step 16 while maintaining the samples at 4°C.

Calculation of bound spin-labeled ligand concentration

1. Acquire an EPR spectrum of 5 μM spin-labeled free ligand in CB (Recipe 16). This is the standard for calculating signal intensity per μM spins.

2. Double-integrate the standard spectrum twice to obtain the intensity (in arbitrary units) of the spectra and double integrate the spectrum obtained for the chased sample described above in step 19.

3. For the standard, calculate the intensity units per μM spin-labeled ligand by dividing the chased sample by double integrated chased value to obtain a μM concentration of free ligand.

4. Calculate the number of surface receptors per cell based on the total free ligand in the chased sample.

Note: This calculation is based on the assumption that the spin-labeled ligand had saturated all surface receptors. We recommend that an additional cell count be obtained directly from the chased EPR sample.

Endocytosis assay by EPR

Cell interiors possess a reducing environment, and the importation of nitroxides results in a conversion of the paramagnetic nitroxyl into the EPR-silent hydroxylamine form (11, 2022). This reduction provides a simple and direct method for measuring the rate of surface protein internalization. As described in (10), our EPR measurements of endocytosis were compared to confocal images of receptor-GFP distribution in CHO cells. Although confocal microscopy does not have the spatial resolution to detect early internalization, it is a useful method for verifying the expected behavior of control samples.

Use three types of controls. Control 1 will be receptor-expressing cells exposed to a general inhibitor of endocytosis. Control 2 will be receptor-expressing cells maintained at 4°C, a condition where no endocytosis is expected. Control 3 will be nontransfected cells and will serve to ensure that nonspecific reduction is not occurring on the surface of the cells. Four 10-cm dishes of confluent cells expressing the receptor and four 10-cm dishes of nontransfected cells will be required.

1Rinse the cells in the dishes two times with 15 ml of DPBS.

2. Add 5 ml of DPBS containing 1mM EDTA and place the cells in a 37°C incubator for 5 min.

3. Scrape the plates on ice and combine the receptor-expressing cells into one 15-ml tube and the nontransfected cells into a second tube and place both tubes on ice.

4. Centrifuge the cells at 600g for 5 min at 4°C and discard the supernatant.

5. Rinse the cells by resuspending in 10 ml of ice-cold CB (Recipe 16).

6. Centrifuge the cells at 600g for 5 min at 4°C and discard the supernatant.

7. Resuspend the cells at a density of 4 × 106 cells/ml in ice-cold CB (Recipe 16) containing 1 μM of spin-labeled ligand. There should be about 17 ml of cells.

8. Gently rock the cells on a Nutator orbital mixer at 4°C for 2 hours.

Note: Step 9 should be performed during this incubation time .

9. Half an hour (30 min) before the binding complete, prepare Control 1 by transferring 5 ml of receptor-expressing cells to a 15-ml tube and adding staurosporine to a final concentration of 1 μM. Place these cells back on the Nutator orbital mixer to complete the binding step.

10. Divide the remaining 12 ml of receptor expressing cells into two 6-ml aliquots in two 15-ml tubes (Control 2 and Endocytosis Cells) on ice.

11. Transfer two 6-ml aliquots of nontransfected cells into two 15-ml tubes and place on ice.

Note: These are the Control 3 samples.

12. Pellet Control 1, Control 2, Control 3, and Endocytosis Cells by centrifugation at 600g for 5 min at 4°C.

13. Rinse all of the samples with equal volumes of ice-cold CB (Recipe 16), in the following volumes. 5 ml for Control 1; 6 ml for Control 2 and Endocytosis Cells; and 6 ml for the two Control 3 samples. Pellet by centrifugation at 600g for 5 min.

14. Prepare the EPR instrument to take readings.

15. Resuspend the Control 2 in 6 ml of ice-cold CB (Recipe 16) and keep on ice.

Note: This sample will remain on ice and EPR spectra taken at the same time points as the Endocytosis Cells.

16. Resuspend the Control 3 samples each in 120 μl of either 4°C or 37°C CB (Recipe 16) containing an appropriate concentration of spin-labeled ligand as determined from the chase experiment (~1 μM), and maintain each at the appropriate temperature.

17. Resuspend the Endocytosis Cells and the Control 1 cells each in 6 ml of 37°C prewarmed CB (Recipe 16) and place the cells in a 37°C incubator.

18. Remove 0.5-ml samples from the Endocytosis Cells and each of the control samples at time = 0, 3 min, 5 min, 10 min, 30 min, and 60 min and place in 1.5-ml microcentrifuge tubes on ice as each time point is reached.

Note: Be sure the cells are well suspended before removing the 0.5-ml samples by inverting the tube 5 to 10 times before pipetting.

19. Acquire EPR spectra of the samples as soon as possible after the time point has been reached.

20. Quantify the viability of the cells after taking the EPR spectra by Trypan Blue staining.

Analysis of endocytosis data

1. For each scanned time point, measure the peak-to-peak height of the central (MI = 0) line ("I" in Fig. 2).

Fig. 2.

EPR signal strength as a function of time following agonist binding. A rapid signal (I) loss was obtained when cells prebound with spin-labeled substance P were incubated at 37°C (diamonds), compared with cells that were maintained at 4°C (squares). Free SPSL(1 μM) incubated with nontransfected CHO does not exhibit any loss in signal intensity over 30 min at 37°C (triangles). Nitroxide reduction is substantially lowered in the presence of 1 μM staurosporine (circles). The spectrum for the free label is shown to the right, with the intensity (I) of the EPR signal measured as the peak-to-peak height for the central line of first derivative of the spectra.

2. Calculate the fractional amplitude (I/I0) by dividing each time point intensity by the intensity at t = 0 min (I0).

3. Plot I/I0 versus time to obtain a plot that shows the kinetics of endocytosis (Fig. 2).

Notes and Remarks

Briefly, we outline the expected results of a successful EPR endocytosis assay, a sample of which is plotted in Fig. 2. Cells expressing the receptor that were incubated at 37°C should exhibit a disappearance of the signal from bound ligand, indicating internalization and subsequent reduction of the free radical. Cells in the Control 1 sample, which were expressing receptor and were incubated at 37°C in the presence of 1 μM staurosporine, should show a diminishing signal over time. However, the rate of signal loss should not be as steep as that observed for the cells not treated with staurosporine. After treatment, most of the staurosporine-treated cells (>80%) will stain positive with Trypan Blue, indicating that the cells are dead or very unhealthy. Thus, although staurosporine may not completely block internalization, at least some of the reduction seen in this control can be attributed to leakage of the cytoplasmic contents from dying cells. Cells in the Control 2 sample, which were expressing the receptor and were incubated at 4°C, should show almost no loss of the signal.

Cells in the Control 3 sample, which were not expressing the receptor and were incubated at either 37°C or 4°C, should exhibit no loss of signal. We have not observed any nonspecific cell surface reduction in either of these controls.

Viability is maintained during the EPR analysis in our experimental conditions because the cells are removed from aerobic conditions at the specified time points and briefly exposed (< 5 minutes) to a sealed capillary to obtain the EPR measurement. The viability of the cells after the EPR reading does not change. However, if continuous EPR measurements (at a controlled temperature) from the same sample are desired, conditions can be adjusted that improve cell viability. For example, samples can be examined in the presence of cell culture media. In addition, a gas-permeable TPX or Teflon (Zeus Industrial Products, Orangeburg, SC) capillary tube can be used to maintain optimal atmospheric conditions.

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