Enrichment of Phosphopeptides with Captivate™ Microscale Phosphopeptide Isolation Kit

• Captivate ferrofluid phosphopeptide-binding reagent (Component A), 1 mL of a 0.5 mg/mL solution
• Binding/wash buffer (Component B), 20 mL
• Elution buffer (Component C), 10 mL
• Barium hydroxide (Component D), 1.3 g
• Methylamine (Component E), 100 µL of a 12 M solution
• Phosphopeptide control mixture (Component F), 100 µL, 20 pmol/µL of each peptide (see Table 1)

• Captivate™ magnetic separator for six microfuge tubes (Molecular Probes’ product C24703) or for 96-well microplates (Molecular Probes’ product C24702), or a similar device
• 25–50% acetonitrile/0.1% trifluoroacetic acid (TFA)
• 1 M ammonium sulfate

Sample Preparation

1. Adjust the sample pH. Dilute the sample in the binding/wash buffer (Component B) to a final amount of approximately 10 pmol of phosphopeptide in a volume of 1–5 µL, or adjust the sample to pH 4 by adding glacial acetic acid.

 
 
Ferrofluid Preparation

1. Resuspend the Captivate ferrofluid phosphopeptide-binding reagent. Vortex mix the vial of binding reagent (Component A) or pipet the reagent up and down to thoroughly mix the binding reagent. The presence of irregularly sized clumps indicates the material has aggregated. If the mixture has aggregated, sonicate the reagent in a water bath for 5–10 minutes until the clumps have dispersed into a homogeneous mixture of fine particles.


2. Wash the Captivate ferrofluid particles. Remove the total amount of ferrofluid phosphopeptide-binding reagent needed for all samples (~20 µL/sample), and transfer the binding reagent to a microfuge tube. Place the microfuge tube in the magnetic separator to separate the ferrofluid particles from the supernatant. Wait 1–2 minutes or until the supernatant has cleared. Remove and discard the supernatant. Remove the microfuge tube from the separator and add the binding/wash buffer (Component B) to attain the original volume. Pipet the solution up and down to disperse the pellet. Repeat this wash step once.


3. Aliquot the binding reagent. Dispense 20 µL aliquots of the binding reagent into microfuge tubes. Small-volume microtubes, e.g., 0.6 mL tubes, are recommended.

 
Phosphopeptide Binding                                                                                                                                                 

1. Prepare the phosphopeptide control mixture. Thaw the phosphopeptide mixture (Component F) on ice, and dilute the desired amount fourfold in the binding/wash buffer (Component B) to have a final concentration of 5 pmol of each peptide per µL. Return the unused portion of the control mixture to <–20°C storage as soon as possible. Note that this control mixture contains four phosphopeptides and three non-phosphopeptides (Table 1).


2. Add the samples to the ferrofluid reagent. Add 1–5 µL of the sample containing approximately 10 pmol of phosphopeptides (prepared in step 1) to a 20 µL aliquot of the ferrofluid phosphopeptide-binding reagent (prepared in step 3). Add 1 µL of the diluted phosphopeptide control mixture (prepared in step 1) to another aliquot of the binding reagent. Vortex mix the sample and the control for 10 minutes, keeping the particles in suspension.


3. Separate the phosphopeptides. Place the tubes on the magnetic separator for 5 minutes. Transfer the supernatants to new tubes, and keep the tubes on ice or at <–20°C prior to analysis. The supernatants contain the non-phosphorylated peptides.


4. Wash the ferrofluid particles. Remove the microfuge tubes from the magnetic separator, and add 100 µL of the binding/wash buffer to each. Pipet the solution up and down to disperse the particles, and place the microfuge tubes back in the magnetic separator. After the suspensions have cleared, remove the supernatants. Repeat this washing step twice. Additional washes with 500 mM NaCl or with 20% acetonitrile may reduce nonspecific binding due to acidic or hydrophobic interactions, respectively. A more acidic wash (e.g., 50 mM glacial acetic acid, pH 3) may reduce the nonspecific binding due to acidic residues, but may lower the phosphopeptide recovery.


5. Elute the phosphopeptides. Add 5–10 µL of the elution buffer (Component C) to each tube. Rinse the ferrofluid particles by pipetting the solution up and down. Perform this step quickly (~30 seconds). Place the microfuge tubes in the magnetic separator, and after the supernatants have cleared, transfer the eluates to new tubes. The eluates contain the phosphorylated peptides. Neutralize the eluates with 1 µL of glacial acetic acid. Keep the samples on ice or at <–20°C prior to analysis.


6. Dilute the samples, if desired. The eluates must be diluted or desalted prior to MALDI analysis to remove interfering components from the elution buffer. If the phosphopeptide concentration is >5 pmol/µL, the samples may be diluted tenfold with 25–50% acetonitrile/0.1% TFA. Desalting and simultaneous concentration of the samples is recommended for lower analyte concentrations. See MALDI-MS Analysis, below, for more information.

 
 
ß-Elimination and Methylamine Addition

A ß-elimination reaction, catalyzed by barium hydroxide, followed by alkylation with methylamine will distinguish between the three different types of peptide phosphorylation. The ß-elimination step will remove phosphates bound to threonine or serine residues, but not those bound to tyrosine residues. The alkylation with methylamine will add a methyl group only to dephosphorylated serine residues. The three groups can be distinguished by predictable changes in mass following these reactions. During this reaction, O-linked carbohydrates can also undergo b-elimination, but under the suggested reaction conditions, the rate is 20-fold slower than for peptides, thus carbohydrate reactions are not likely to confound the results.
Before performing these reactions, disulfides should be reduced and the cysteine residues should be alkylated in protein-digest samples to protect these sites from b-elimination. Standard protocols for disulfide reduction and cysteine alkylation can be followed.
 

1. Prepare the barium hydroxide solution. Dissolve 32 mg of barium hydroxide (Component D) in 1.0 mL of deionized water (dH₂O). To prevent the formation of barium carbonate, degas the solution by bubbling nitrogen or an inert gas through the solution for about 1 minute.


2. Bind the phosphopeptides. Follow the above protocol through steps 3.1–3.4 (up to the addition of the elution buffer). Do not add elution buffer.


3. Elute the phosphopeptides with barium hydroxide. Add 5–10 µL of the barium hydroxide solution (prepared in step 4.1) to each sample, still bound to the ferrofluid particles. Pipet the sample solutions up and down, making sure that all of the particles are in the buffer. This step should be performed quickly (~30 seconds). Place the microfuge tubes in the magnetic separator, and after the suspensions have cleared, transfer the eluates to new tubes. The eluates contain the phosphopeptides.


4. Add methylamine. Add 1 µL of 12 M methylamine (Component E) and 10–20 µL of dH₂O to each microfuge tube containing the eluates from step 4.3. The methylamine concentration will be ~0.5 M. Vortex mix, then incubate the samples at 30°C for 1 hour. Under these conditions, ≥90% of the phosphoserine residues unb-elimination (–98 Da) and methylamine addition (+31 Da), which results in a net mass shift of –67 Da. Phosphothreonine residues undergo b-elimination (–98 Da) with little or no methylamine addition. Phosphotyrosine residues are not modified.


5. Stop the reaction and remove Ba2+ from the samples. Add 3–5 µL of 1 M ammonium sulfate to each sample to form a barium salt. Precipitate the salt by centrifuging the samples for 30 seconds. Transfer the supernatants containing the samples to new tubes. 


6. Acidify the samples. Add 1 µL of glacial acetic acid for each 10 µL of sample from step 4.5. Eluates must be acidified to < pH 4 prior to desalting.


7. Desalt the samples. It is necessary to desalt the samples prior to MALDI analysis. See MALDI-MS Analysis, below.

MALDI-MS Analysis                                                                                                                                                                 

Salt Removal:   Generally, salt concentrations above 10 mM will interfere with MALDI analysis. Desalting is labor intensive but allows for maximum recovery of most peptides in very small volumes free from interfering salts. If the analyte concentrations are sufficiently high, samples may be diluted to reduce the salt concentration to acceptable levels prior to spotting on the MALDI plate. Otherwise, desalt the samples using ZipTips™ (Millipore), or the equivalent reversed-phase cleanup devices, following the manufacturer’s instructions and eluting with 50% acetonitrile/0.1% TFA. Samples must be acidified to pH ≤4 with glacial acetic acid (step 4.6) prior to reversed-phase desalting/concentration. One drawback with reversed-phase desalting is that very hydrophilic peptides may not be recovered.
 
Dried-Droplet MALDI Spotting Method:    Spot 0.5–1 µL of a sample onto a target plate. (Desalted samples may be eluted with 50% acetonitrile/0.1% TFA directly onto the plate.) Spot an equal volume of 5–10 mg/mL a-cyano-4-hydroxycinnamic acid (HCCA) dissolved in 50% acetonitrile/0.1% TFA on top of the sample, and allow the spot to air-dry. This simple and fast method is suggested for samples containing at least 0.5 pmol of phosphopeptide.
 
Nitrocellulose MALDI Spotting Method:    Prepare a 40 mg/mL solution of HCCA in acetone and a 20 mg/mL solution of nitrocellulose in acetone. Vortex mix each solution for 5–10 minutes. Prepare a matrix solution by mixing 200 µL of the HCCA solution with 100 µL of the nitrocellulose solution and 100 µL of isopropanol. Next, mix 1.5 µL of a sample with an equal volume of the matrix solution, and deposit 1.5 µL onto the MALDI target. Allow the spot to dry at ambient temperature. Next, place 5 µL of 5% formic acid on top of the spot for 10 seconds, and then pipet off the excess formic acid. Allow the spot to dry. Finally, apply 5 µL of dH2O to wash the spot for 10 seconds, remove the excess dH2O, then allow the spot to dry again. This method markedly enhances peptide detection for samples with <1 pmol of material. It also allows for eliminating salts from the sample directly on the MALDI plate.
 
Note:   Enhanced detection of phosphopeptides may be achieved by the addition of 25–50 mM ammonium salts (e.g., ammonium sulfate, ammonium citrate or ammonium acetate), but this effect is peptide-specific. Techniques include mixing the ammonium salt 1:1 with the sample or with the matrix solution, or spotting an equal volume on top of the sample on the target plate. Also, some phosphopeptide signals may be enhanced relative to other peaks by analysis in the negative ion mode.