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Department of Cell and Molecular Pharmacology & Experimental Therapeutics

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Protein Mass Spectrometry

The history of mass spectrometry in the Department of Pharmacology at MUSC dates back to 1972 with the establishment of a GC-MS facility as part of an NIH funded Pharmacology Center Grant Program. The mass spectrometry activity has steadily grown and since the late 1980s has been increasingly focused upon protein studies. The Mass Spectrometry Facility is housed within the Department of Pharmacology and directed by Pharmacology faculty. It serves as a university research resource facility and as a component of the MUSC Proteomics Center.

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An Overview of Protein Mass Spectrometry

New mass spectrometric methods developed over the last several years have led to the establishment of mass spectrometry as a powerful technique for protein structure determination. Key to this development was the introduction of fast atom bombardment (FAB) ionization which permitted production of intact molecular ions from peptides and small proteins as well as from other labile biological molecules. Other ionization methods (electrospray ionization [ESI], matrix assisted laser desorption ionization [MALDI]) have subsequently been developed which have been demonstrated to permit measurement of protein molecular weights in excess of several hundred thousand daltons.

In combination with these new ionization methods, tandem mass spectrometry permits amino acid sequencing of peptide fragments of proteins as well as characterization of posttranslational modifications. Mass spectrometry therefore provides a means of determining the complete primary structure of a protein. Use of chemical modifications in conjunction with mass spectrometric analysis provides a means of gaining information on the three dimensional structure as well. The use of mass spectrometry in the study of the various aspects of protein structure is discussed in more detail on subsequent pages.

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Molecular Weight Determination

All three of the major approaches to mass spectrometric determination of the molecular weights of intact proteins are available in the Medical University of South Carolina Mass Spectrometry Laboratory. These include FAB (the term as used here also includes cesium ion bombardment), electrospray ionization (ESI), and matrix assisted laser desorption ionization (MALDI). The JEOL HX110/HX110 tandem mass spectrometer is equipped with a high voltage (35 kV.) cesium gun and has a mass range of 14,500. Molecular weights of proteins in excess of 10,000 daltons have been successfully determined on this magnetic sector type instrument using FAB ionization. The resolution and mass accuracy of the JEOL instrument is such that the molecular weights can be determined to better than a single mass unit. In practice the protein is applied as a solution (on the order of 1 microliter) to the tip of a sample probe and mixed with a matrix liquid (e.g. glycerol, thioglycerol or nitrobenzyl alcohol). The probe is then inserted into the mass spectrometer and the sample bombarded with high energy cesium ions to sputter off intact (usually protonated) molecular ions of the protein in the sample. The ions are then accelerated into the mass spectrometer and the mass to charge ratio determined. Since one is usually observing singly charged ions, the result is a direct measurement of molecular weight. A pure sample is not necessary, and it is possible to measure molecular weights of multiple components of a mixture although competition effects can cause some components to be observed more easily than others. Buffer salts and detergents can impede the observation of sample components; optimal samples are those collected from a reversed phase HPLC separation.

Electrospray ionization permits measurement of much larger molecular weights by virtue of the fact that it produces multiply charged ions. Since mass spectrometers normally measure mass to charge ratio, increasing the number of charges on the ion effectively multiplies the mass range of the instrument. Mass measurement accuracy on the order of 0.01% is possible with this technique. In practice the sample is introduced into the ion source in a liquid stream containing an organic modifier and an acid delivered from a pump driven microliter syringe or as an HPLC effluent. A variety of acids and concentrations are used in ESI; a typical solvent system for discrete samples is 50% methanol in water containing 5% acetic acid. Buffer salts and ionic detergents must be avoided since they can dominate the ion current and prevent observation of the protein. Since electrospray ionization gives multiple peaks for each protein, mixtures can give very complex spectra. Algorithms have been developed to resolve the complex mixtures of multiply charged ion peaks into a spectrum with a true mass scale and one peak per component, but the technique works best with relatively pure samples.

Matrix assisted laser desorption (MALDI) mass spectrometry permits measurement of still larger molecular weights. In this technique, the sample is mixed with a UV absorbing matrix compound (commonly sinapinic acid) typically in a 30% aqueous acetonitrile solution, and 1-3 microliters of the solution is dried on the end of a sample probe or as a spot on a sample plate. The dried sample is then inserted into the mass spectrometer and the sample mixture exposed to the focussed beam from a pulsed UV laser. Absorption of the laser energy by the matrix results in ejection of intact molecular ions of the protein. The ions produced are typically singly charged, therefore the mass analyzer must be of a different type to measure very large masses. A time-of-flight analyzer is used to measure mass by measuring the time it takes for an ion, after electrostatic acceleration, to travel from the ion source to the detector. Heavier ions travel slower, hence, by measuring long times, the mass range of this type of analyzer is theoretically unlimited. Molecular weights as high as 160,000 have been measured on our home built instrument with mass measurement errors less than 1% (less than 0.1% for masses under 20,000). The newly installed (11/96) commercial instrument will provide higher performance. The MALDI technique is applicable to mixture analysis and is less affected than ESI by the presence of salts or ionic detergents. (However, some detergents are more troublesome than others; see the section below on MALDI and detergents.)

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Determination of Primary Structure

The procedure for determination of the primary structure of a protein is outlined in the figure below. The first step (1. in the figure) is to determine the molecular weight of the intact protein. This can be done using either FAB, electrospray, or matrix assisted laser desorption mass spectrometry. Knowledge of the molecular weight of the intact protein is necessary to be sure that the entire protein is accounted for in analyzing fragments of the protein. The next step is to cleave the protein into smaller fragments and determine the amino acid sequence of each fragment (2. in the figure). If the protein contains cysteine residues, it is normally subjected to reduction and carboxymethylation of the cysteine sulfhydryl groups prior to cleavage (alternatively carboxamidomethyl or pyridylethyl derivatives may be prepared). Cleavage is then performed either chemically or enzymatically.

Large proteins are usually cleaved in two stages: a first cleavage is performed at relatively low abundance sites (step 2a, e.g. cyanogen bromide cleavage at methionine residues) to yield large fragments which are separated and then cleaved at more abundant sites (step 2b, e.g. trypsin cleavage at lysines and arginines). Smaller proteins can be initially cleaved at higher abundance sites. The cleavage mixtures are then separated into simpler mixtures by HPLC (step 2c). Unlike with Edman degradation where it is necessary to individually purify each degradation product, the mass spectrometric approach can be carried out with mixtures of cleavage fragments. The fractionation into simpler mixtures is done to overcome potential competitive effects in the ionization process to increase the probability of observing all of the fragments in the mass spectrometer. The fragment mixtures are first subjected to single stage mass spectrometry to determine the molecular weights of the fragments (step 2d) and then analyzed by tandem mass spectrometry (step 2e) to determine the amino acid sequence of each fragment.

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Posttranslational Modifications

We examine the posttranslational modifications to the most abundant lens membrane protein, Aquaporin 0 (also known as the lens major intrinsic protein, MIP). Numerous modifications have been identified (see figure) by mass spectrometry. A goal of this research is to determine those modifications which lead to lens opacification through structural studies of human lenses and through functional assays of recombinant modified protein. Additional areas of interest involve protein-protein interactions of lens proteins with Aquaporin 0, changes in the lens membrane proteome with age and cataractogenesis, and lens protein oxidation.

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Membrane Proteins

As stated above, integral membrane proteins present greater challenges not only for the mass spectrometry, but also for the preceding peptide chemistry. The MUSC laboratory has a particular interest in membrane receptor proteins. Work is underway to develop improved procedures for protein cleavage and sample preparation for mass spectrometry of integral membrane proteins and their peptide cleavage products. Recent developments in HPLC separations of cyanogen bromide cleavage products promise to greatly expedite our work on membrane proteins

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Sample Requirements

The molecular weight of a protein can usually be obtained with a few picomoles by electrospray or laser desorption techniques if the protein has favorable solubility properties. Sample quantity requirements in these cases are usually dictated more by sample manipulation requirements than by the low femptomole actual sample consumption in the measurement. Hydrophobic proteins, particularly integral membrane proteins, present greater difficulty; however, we have developed methodology which permits analysis of intact integral membrane proteins by MALDI. Molecular weight measurement of smaller proteins and peptides (under ca. 2000 mass units) by FAB ionization can often be obtained with low picomole amounts of material. Larger species usually require larger amounts due to the greater difficulty of producing ions from larger molecules. Extremely hydrophobic or extremely hydrophilic molecules tend to be difficult to ionize by FAB, but the problem can often be overcome by forming a more favorable chemical derivative. ESI and MALDI generally require less sample. Tandem mass spectrometry requires significantly more sample than that required for molecular weight measurement. Present collision dissociation technology limits the ability to obtain complete sequences of peptides by tandem mass spectrometry to peptides under molecular weight of about 2500-3000 (approximately 25-30 amino acids). Partial sequence information can often be obtained, however, for larger peptides. With a peptide of molecular weight 2000 for which the molecular ion could be measured with picomole amounts, as much as two orders of magnitude more sample could be needed to obtain a good tandem mass spectrum.

Use of the array detector on the four sector instrument and use of the new ion trap instrument should significantly reduce these sample requirements. Since the current technology for tandem mass spectrometric sequencing requires peptide fragments of 25-30 amino acids or less, sequencing of a protein requires that it be cleaved into smaller fragments. The sample size requirement for sequencing a protein is therefore dictated not only by the mass spectrometry but also by the preceding peptide chemistry. Due to the practical aspects of sample manipulations in cleaving and isolating peptide fragments one should expect to require as much as nanomole to tens of nanomole amounts for small proteins in the 10-15,000 molecular weight range (tens to hundreds of micrograms). In theory a larger protein should not require any more sample on a molar basis, but the increasing complexity of the problem with larger proteins can lead to need for more sample.

It should be emphasized that while state of the art Edman sequencing can claim smaller sample requirements, the mass spectrometric methods do not require individual purification of each peptide fragment (which leads to a practical requirement for more sample). Furthermore, most Edman sequencing facilities do not operate at state of the art sensitivity for real samples. The above discussion of sample quantity requirements for tandem mass spectrometry relates primarily to sector type tandem instruments employing a point detector. The new scanning array detector on the MUSC four sector instrument is expected to be fully operational soon (11/96). Use of an array detector can increase sensitivity by as much as two orders of magnitude. The new (11/96) LCQ instrument is also expected to require significanly less sample. Two orders of magnitude improvement in instrument sensitivity does not necessarily directly translate to two orders of magnitude lower sample requirement, however, since sample manipulation limitations become more significant with extremely small samples. The answer to the question of how much sample is required is therefore not a firm figure. Every sample is a different molecular species with different properties, and the technology is constantly evolving. It can safely be said, however, that even with the current state of the art in protein and peptide mass spectrometry, the limitations lie more in the practical aspects of sample handling and peptide chemistry than with the mass spectrometry itself.

Another issue with respect to sample size requirement is that of the question being approached. For example, if the objective is to obtain some partial sequences for DNA probe construction, the sample requirement can be far less than would be required for full sequencing.

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Department of Cell and Molecular Pharmacology & Experimental Therapeutics at MUSC

173 Ashley Avenue
BSB 358 MSC 509
Charleston, SC 29425

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