High-Throughput Capillary Electrophoresis–Mass Spectrometry: From Analysis of Amino Acids to Analysis of Protein Complexes

Abstract

Recent advances in capillary electrophoresis–mass spectrometry (CE-MS) interfacing using porous tip is leading to commercialization of CE-MS with a sheathless interface for the first time. The new sheathless interface in conjunction with CE capillary coatings using self-coating background electrolytes (BGE) has significantly simplified CE-MS analysis of complex mixtures. CE-MS, with its high separation efficiency, compound identification capability, and ability to rapidly separate compounds with a wide range of mass and charge while consuming only nanoliters of samples, has become a valuable analytical technique for the analysis of complex biological mixtures. These advances have allowed a single capillary to analyze a range of compounds including amino acids, their D/L enantiomers, protein digests, intact proteins, and protein complexes. With these capabilities, CE-MS is poised to become the multipurpose tool of separation scientists. More recently, an eight-capillary CE in conjunction with an 8-inlet mass spectrometry has allowed 8 CE-MS analyses to be performed concurrently, significantly increasing throughput.

Appendices

Appendix 1. Standard Operating Procedure for Making Porous Capillary Using HF

Substance: hydrogen fluoride, HF

Identification numbers: CAS number 7664-39-3

Main risks and target organs: Hydrogen fluoride is highly corrosive to all tissues. Skin: burns, necrosis; underlying bone may be decalcified. Eyes: burns

The course of the skin burns from hydrogen fluoride depends on concentration; more dilute solutions cause potentially serious but delayed (24 h) systemic effects. Absorption of the fluoride ion is a significant hazard mainly due to hypocalcaemia. Hydrogen fluoride burns to the eye are corrosive and require immediate flushing and ophthalmologic consultation.

1.1 Storage

Hydrogen fluoride should be stored in a cool, dry, well-ventilated area in tightly sealed containers. Containers of hydrogen fluoride should be protected from physical damage and should be stored separately from metals, concrete, glass, strong bases, sodium hydroxide, potassium hydroxide, and ceramics. Glass containers should not be used for storage as hydrogen fluoride will dissolve glass. Explicit warnings in appropriate languages and graphics should be maintained on all containers and work areas.

1.2 Personal Hygiene Procedures

If hydrogen fluoride contacts the skin, workers should flush the affected areas immediately with plenty of water for 15 min, followed by washing with soap and water.

Clothing contaminated with hydrogen fluoride should be removed immediately, and provisions should be made for the safe removal of the chemical from the clothing. Persons laundering the clothes should be informed of the hazardous properties of hydrogen fluoride, particularly its potential for causing irritation.

A worker who handles hydrogen fluoride should thoroughly wash hands, forearms, and face with soap and water before eating, using tobacco products, using toilet facilities, applying cosmetics, or taking medication.

Workers should not eat, drink, use tobacco products, apply cosmetics, or take medication in areas where hydrogen fluoride or a solution containing hydrogen fluoride is handled, processed, or stored.

1.2.1 Initial Medical Treatment

Thoroughly irrigate with water. Massage calcium gluconate gel into the burned skin for a minimum of 30 min and for as long as the pain persists; at times pain persists up to 4 h. If no gel is available, use a calcium (20 g Ca²⁺ in about 2 L of water) or magnesium solution for soaking.

1.2.2 Specific Preventive Measures

Appropriate latex or rubber gloves should be used when working with hydrogen fluoride. Frequent checks of glove integrity or changes of the gloves should be performed. Any suspected exposure should be immediately decontaminated. Eye goggles, aprons, sleeves, and other protective devices should be used in accord with the conditions of use. Concentrated and anhydrous hydrogen fluoride should only be used in the hood.

Use nitrile gloves, apron, face mask, and rubber gloves when working with HF solution and nitrile gloves and safety glasses when dealing with etched capillaries. Work under the hood when dealing with HF. Replace gloves each time your hands need to be outside the hood. Change gloves without touching skin.

1.2.3 Procedure for Making the Porous Tip Using the Teflon Container

• 1. Prepare a 500-ml saturated solution of calcium carbonate in a beaker and 500 mL of DI water in another beaker. These solutions will be used for neutralization of HF-contaminated glassware and fused-silica capillaries.

• 2. Fill Teflon container with fresh 48% HF, which is in a plastic container under a well-ventilated hood. To do this you may have to discard the old HF in the container by removing ∼100 μL of HF solution from the HF container and slowly pouring it into the carbonate solution. Wash the container with DI water by filling the container with water and discard the water using disposable pipettes. Pour methanol into the empty Teflon container and discard it into the carbonate container.

• 3. Wait a few minutes for the leftover methanol in the HF container to evaporate.

• 4. Gradually (300–400 μL at a time) fill the dry Teflon container with fresh HF using a plastic disposable pipette (or a glass pipette if no plastic pipette is available). Fill the HF container to the top.

• 5. Replace gloves and etch a capillary by first burning about 4–5 cm of the capillary using a lighter. Second, remove the burnt debris at the capillary tip by using Chemwipes and methanol.

• 6. Attach the inlet of the capillary to the air hose in the hood using an Upchurch union.

• 7. Insert the bare fused-silica end of the capillary (the outlet) into the HF container all the way without touching the bottom of the container, and adjust it to prevent it from touching the wall of the HF container.

• 8. Wait appropriate amount of time for the outlet tip to become porous.

• 9. Using fresh gloves, remove the capillary outlet from the HF container and neutralize it by immersing it into the carbonate solution for a few seconds and then in water for a few seconds. Disconnect the fused-silica capillary from the air pressure and wash it under the faucet.

• 10. Replace the HF container top.

• 11. Decontaminate the area using the carbonated solution.

Appendix 2. Methods and Techniques for Analyzing Red Blood Cells

The chemical contents of RBCs were analyzed using three different sampling methods:

• (1) A solution of lysed RBCs was injected into the capillary using the pressure injection mode. The solution was prepared by diluting 5 μL of fresh blood from a healthy adult male to 50 μL using saline solution, followed by centrifugation and removal of the supernatant, leaving only intact cells without the plasma. After adding 50 μL of water to lyse the cells, the lysed cells were then diluted either 5× or 200× with water to final dilutions of 50× and 2,000×, respectively.

• (2) Intact RBCs (one or three cells) were drawn into the capillary inlet using suction and a 200× microscope for observation.

• (3) Intact RBCs suspended in a saline solution were injected into the CE capillary using the pressure injection mode of the CE instrument. The solution of suspended intact RBCs was prepared in a manner almost identical to method 1, except that saline solution was used instead of water to maintain intact cells. To homogenize the solution, the sample vial was hand-shaken immediately before sampling. It should be noted that it was necessary to use fresh blood to separate the Hb chains; when a 1-week-old blood sample was analyzed using the 0.1% acetic acid solution, only one peak with an average molecular weight of ∼65,000 was observed. For intact cell analysis, best overall performance (separation and sensitivity) was achieved using the 0.1% acetic acid solution without any organic additives. However, when lysed blood was analyzed, the best overall performance was achieved using a 0.1% acetic acid solution containing 50% acetonitrile. Under these conditions, the presence of acetonitrile in the CE BGE reduced the EOF, and consequently, the four major proteins of RBCs were all separated (Fig. 18). Moreover, addition of acetonitrile also reduced the background chemical noise and adduct formation due to the incomplete desolvation of liquid droplets and, therefore, improved the S/N of CAI (Fig. 18, panel b). The improvement of S/N due to the latter BGE was further confirmed by analyzing a standard mixture of CAI and CAII (0.1 mg/mL) using a 0.1% acetic acid solution and a 0.1% acetic acid solution containing 50% acetonitrile. It was observed that while the mass spectra of these compounds had similar intensity on the absolute scale, both noise level and adduct formation were significantly lower for the BGE containing 50% acetonitrile. Other possible factors for the higher sensitivity of these proteins include better solubility and less adsorption to the capillary wall when the acetonitrile-containing BGE was used.

Appendix 3. Successful CE-MS analysis of protein complexes depends on three major factors

• 1. The use of ammonium acetate at pH 7.4. Ammonium acetate at pH ∼7 was ideal for the analysis of protein complexes that we have studied. This is because it keeps the complexes intact and it is a mass-friendly background electrolyte. It was made by titrating a 0.01% acetic acid solution with a 0.01% ­ammonium hydroxide solution until desired pHs were reached. At pH of 7.4 the concentration of the ammonium acetate (NH4OAc) buffer was ∼1.7 mM. Polybrene (PB) was added to these buffers as the self-coating reagent 2 with a final concentration of PB at ∼0.1%.

• 2. Soaking the protein complex in ammonium acetate for a few hours before CE-MS analysis. More accurate MWs (∼0.01% error) were obtained for all major protein–protein and protein–metal complexes when they were diluted in ammonium acetate buffer and left in that solution for a few hours. The longer the protein complexes stayed in ammonium acetate solution before CE-MS analysis, the more intense and narrow the higher charge states of the protein complexes became. This was because longer times improved the ion exchange between the nonspecific, nonvolatile salts (mostly sodium) that were associated with the interior of the protein complexes under native conditions and the volatile ammonium ions of the BGE. Ion exchange between ammonium ions in the solvent and nonspecific sodium adducts of the protein–protein and protein–metal complexes slightly unraveled the complexes. CE/ESI-MS further enhanced this charge exchange and separated the nonspecific salt adducts from protein complexes, resulting in formation of higher charge-state protein complexes free from nonspecific salt adducts. Deconvolution of these higher charge-state ions provided more accurate (∼0.01% error) average MW of intact protein–protein and protein–metal complexes for better identification of the constituents of the complexes.

• 3. Operation of the CE under forward polarity. Operation of the CE under forward polarity mode was essential for detection of intact protein–protein and protein–metal complexes. Intact protein complexes with minimal fragmentation can only be observed in forward polarity. This was because of the electrochemical nature of CE and ESI processes. As we have explained before, CE and ESI-MS represent two electrical circuits, each with two sets of electrodes, CE inlet and outlet electrodes, ESI emitter, and MS inlet electrodes. The CE/ESI-MS overlays these two separate circuits so that the CE outlet electrode and the ES emitter electrode are shared between the two circuits (CE outlet/ESI shared electrode). Therefore, under CE/ESI-MS, at the shared electrode two electrochemical reactions occur simultaneously. Depending on the polarity and magnitude of the voltage of the shared electrode compared with that of the CE inlet and MS inlet electrodes, electrochemical reactions at the shared electrode can be both reductive, both oxidative, or one reductive and the other oxidative. For example, under positive electrospray ionization mode, where +1.5 kV is applied to the shared electrode and 0–100 V is applied to the MS inlet electrode, two possibilities exist: (1) Under reverse polarity CE, where −30 kV is applied to the CE inlet electrode, the shared electrode is anodic with respect to both CE inlet electrodes and the MS inlet electrode. (2) Under forward polarity CE, where +30 kV is applied to the CE electrode, the shared electrode is cathodic with respect to CE inlet electrode and anodic with respect to MS inlet electrode. At the anode, the major oxidation reaction for aqueous buffer solutions is typically the electrochemical oxidation of water (2H₂O ↔ 4 H⁺  +  O₂  +  4e⁻). The extent of the electrochemical reactions and therefore the pH change depends on the magnitude of the current that flows through each circuit. Under experimental conditions used here, CE current was ∼1 μA, while ESI current was in the nanoampere range. Under reverse polarity mode in which the electrochemical reactions at the shared electrode are anodic for both circuits, the pH of the solution decreases enough to dissociate all the tetramer. The pH at the ESI tip under reverse polarity is estimated to decrease from pH 7.4 to 5, the pH at which some protein complexes dissociate to dimer. However, under forward polarity mode, the shared electrode is only anodic in the ESI circuit, which has a lower current, and its effect is mostly canceled by the cathodic reaction at this electrode in the CE circuit causing minimal dissociation of the tetramer. Since this dissociation (in both cases) is happening close to the outlet of the CE capillary, all dissociation products comigrate and show up as a single electrophoretic peak.