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Overview of Mass Spectrometry

Mass spectrometry (MS) measures the mass-to-charge ratio of ions to identify and quantify molecules in simple and complex mixtures. MS has become invaluable across a broad range of fields and applications, including proteomics. The development of high-throughput and quantitative MS proteomics workflows within the last two decades has expanded the scope of what we know about protein structure, function, modification and global protein dynamics.

This overview outlines the role of mass spectrometry in the field of proteomics and reviews MS methodology and instrumentation and touches on sample preparation and liquid chromatography-based separation prior to MS analysis.

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Mass Spectrometry Sample Preparation Handbook

Introduction to Mass Spectrometry

Proteomics is the study of all proteins in a biological system (e.g., cells, tissue, organism) during specific biological events (1, 2). Although complementing genomics, proteomics is considerably more difficult to study than genomics or even transcriptomics because of the dynamic nature of proteins expression. Additionally, the majority of proteins undergo some form of post-translational modification (PTM), which further increases the proteomic complexity. The broad scope of proteomics has only begun to be realized within the last 15 years due in large part to technological developments in mass spectrometry.

Mass spectrometry is a sensitive technique used to detect, identify and quantitate molecules based on their mass and charge (m/z). Originally developed almost 100 years ago to measure elemental atomic weights and the natural abundance of specific isotopes (3), MS was first used in the biological sciences to trace heavy isotopes through biological systems and later to sequence oligonucleotides and peptides and analyze nucleotide structure (4).

The development of methods of macromolecule ionization, including electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI), enabled the study of protein structure by MS (5, 6, 7), which allowed scientists to obtain protein mass "fingerprints" that could be matched to proteins and peptides in databases to predict the identity of unknown proteins. Methods of isotopic tagging have led to the ability to quantitate target proteins both in relative terms and absolute quantities. Technological advances have provided methods to analyze samples in solid, liquid or gas states, and the sensitivity of current mass spectrometers allows one to detect analytes at concentrations in the attomolar range (10-15) (10).

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Overview of Post-Translational Modifications (PTMs)

Mass Spectrometry Applications

Mass spectrometry is now used across a broad range of disciplines and settings, including academic research, biotechnological development, pharmaceutical discovery, clinical testing, environmental analysis and geological testing. Some of the common applications for MS are listed below:

Common applications and fields that use mass spectrometry.
Field of Study Applications
Proteomics
  • Determine protein structure, function, folding and interactions
  • Identify a protein from the mass of its peptide fragments
  • Detect specific post-translational modifications throughout complex biological mixtures
  • Quantitate (relative or absolute) proteins in a given sample
  • Monitor enzyme reactions, chemical modifications and protein digestion
Drug Discovery
  • Determine structures of drugs and metabolites
  • Screen for metabolites in biological systems
Clinical Testing
  • Perform forensic analyses such as confirmation of drug abuse
  • Detect disease biomarkers (e.g., newborns screened for metabolic diseases)
Genomics
  • Sequence oligonucleotides
Environment
  • Test water quality or food contamination
Geology
  • Measure petroleum composition
  • Perform carbon dating
 

Methodology

All mass spectrometers have an ion source, a mass analyzer and an ion detector, although the nature of these components varies based on the type of mass spectrometer, the type of data required and the physical properties of the sample. Samples are loaded into the mass spectrometer in liquid or dried form and then vaporized and ionized by the ion source (e.g., ESI, MALDI) .

Schematic of the basic components of a mass spectrometer
Schematic of the basic components of a mass spectrometer.

The charge that these molecules receive allows the mass spectrometer to accelerate the ions throughout the remainder of the system. The ions encounter electrical and/or magnetic fields from mass analyzers, which deflect the paths of individual ions based on their mass and charge (m/z). Commonly used mass analyzers include time-of-flight [TOF], quadrupoles and ion traps, and each type has specific characteristics. Mass analyzers can be used to separate all analytes in a sample for global analyses, or they can be used essentially like a filter to properly deflect only specific ions towards the detector.

Ions that have successfully been deflected by the mass analyzers then hit the ion detector. Most often, these detectors are electron multipliers or microchannel plates, which emit a cascade of electrons when each ion hits the detector plate (4). This cascade results in amplification of each ion hit for improved sensitivity. This entire process is performed under an extreme vacuum (10-6 to 10-8 torr) (8) to remove contaminating non-sample ions, which can collide with sample ions and alter their paths or produce non-specific reaction products (9).

Mass spectrometers are connected to computers with software that analyzes the ion detector data and produces graphs that organize the detected ions by their individual m/z and relative abundance. These ions can then be processed through databases to predict the identity of the molecule based on the m/z.

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Simple diagram of a mass spectrometer
Diagram of a sector mass spectrometer. A sample is injected into the mass spectrometer, and the molecules are ionized and accelerated. The ions are then separated by mass and charge by the mass analyzer via electromagnetic deflection, and the ions that are properly aligned are then detected and amplified. The entire system is under intense vacuum during the entire procedure. After signal amplification, the ion hits are analyzed, and data is generated that yields the relative abundance of each ion based on the mass-to-charge (m/z) ratio.
Although sector instruments have decreased in use in recent years due to improvements in mass analyzers (e.g., quadrupole, ion trap), the simplified diagram conveys a key principle of mass spectrometry—the ability to select and analyze specific ions in a complex sample.

Tandem mass spectrometry (MS/MS) offers further information about specific ions. In this approach, distinct ions of interest are selected based on their m/z from the first round of MS and are fragmented by one of a number of methods of dissociation, such as colliding the ions with a stream of inert gas, as in collision-induced dissociation (CID) or higher energy collision dissociation (HCD). Other methods of ion fragmentation include electron-transfer dissociation (ETD) and electron-capture dissociation (ECD).

These fragments are then separated based on their individual m/z ratios in another round of MS. MS/MS is commonly used to sequence proteins and oligonucleotides, as the fragments can be used to match predicted peptide or nucleic acid sequences, respectively, that are found in databases such as IPI, RefSeq and Swis-Prot. These sequence fragments can then be organized in silico into full-length sequence predictions.

 
Diagram of tandem mass spectrometry (MS/MS)
Diagram of tandem mass spectrometry (MS/MS). A sample is injected into the mass spectrometer, ionized and accelerated and then analyzed by mass spectrometry (MS1). Ions from the MS1 spectra are then selectively fragmented and analyzed by mass spectrometry (MS2) to give the spectra for the ion fragments.While the diagram indicates separate mass analyzers (MS1 and MS2), some instruments can utilize a single mass analyzer for both rounds of MS.

Biological samples are often quite complex and contain molecules that can mask the detection of the target molecule, such as when the sample exhibits a large dynamic concentration range between the target analyte(s) and other molecules in the sample. Therefore, methods of separation are often employed to partition the target analyte(s) from the other molecules in a sample.

Gas chromatography (GC) or liquid chromatography (LC) are common methods of pre-MS separation used when analyzing complex gas or liquid samples by MS, respectively.

High performance liquid chromatography (HPLC) is the most common separation method to study biological samples by MS or MS/MS (termed LC-MS or LC-MS/MS, respectively), because the majority of biological samples are liquid and nonvolatile. LC columns have small diameters (e.g., 75μm; nanoHPLC) and low flow rates (e.g., 200nL/min), which are ideal for minute samples (2). Additionally, "in-line" liquid chromatography (LC linked directly to MS) provides a high-throughput approach to sample analysis, as multiple analytes that elute through the column at different rates are immediately analyzed by MS. For example, 1-5 peptides in a complex biological mixture can be sequenced per second by in-line LC-MS/MS (2).

Example of in-line LC-MS-MS
Example of in-line LC-MS/MS system. Thermo Scientific LTQ Orbitrap Velos Mass Spectrometer with in-line HPLC.
 

Quantitative Proteomics

While mass spectrometry can detect very low analyte concentrations in complex mixtures, MS is not inherently quantitative because of the considerable loss of peptides and ions during analysis. Therefore, peptide labels or standards are concomitantly analyzed with the sample and act as a reference point for both relative or absolute analyte quantitation, respectively. Commercial products are now available that allow the detection and quantitation of multiple proteins in a single reaction, demonstrating the high-throughput and global analytical platform that MS is becoming in the field of proteomics.

Relative quantitation strategies include stable isotope labeling using amino acids in cell culture (SILAC) and tandem mass tagging (TMT). In these approaches, proteins or peptides are labeled with stable isotopes that give them distinct mass shifts over unlabeled analytes. This mass difference can be detected by MS and provides a ratio of nonlabeled analyte levels to those of labeled analyte. These approaches are often used in discovery proteomics, where many proteins are identified across a broad dynamic range using different-sized labels.

Absolute quantitation is performed in targeted proteomic experiments and increases the sensitivity of detection for a limited number of target analytes. These approaches require spiking a sample with known amounts of synthetic peptides containing heavy stable isotopes, which act as internal quantitative standards for absolute quantitation of the corresponding natural peptides in the sample.

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Quantitative Proteomics

Sample Preparation

All samples require some form of preparation prior to study by MS to remove detergents, reduce the complexity of the sample when focusing on specific proteins and/or tag proteins for identification/quantitation. Proper sample preparation is critical for MS analysis, because the quality and reproducibility of sample extraction and preparation significantly impact results from MS instruments. Sample preparation encompasses a wide range of techniques that includes lysate preparation, protein or peptide enrichment, sample clean-up and protein digestion, which are described in detail on a separate page dedicated to sample preparation.

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Sample Preparation for Mass Spectrometry

References

  1. Kuster B. et al. (2005) Scoring proteomes with proteotypic peptide probes. Nat Rev Mol Cell Biol. 6, 577-83.
  2. Mallick P. and Kuster B. (2010) Proteomics: A pragmatic perspective. Nat Biotechnol. 28, 695-709.
  3. Willard H. H. (1988) Instrumental methods of analysis. Belmont, Calif.: Wadsworth Pub. Co. xxi, 895pp.
  4. Finehout E. J. and Lee K. H. (2004) An introduction to mass spectrometry applications in biological research. Biochem Mol Biol Educ. 32, 93-100.
  5. Chowdhury S. K. et al. (1990) Electrospray ionization mass spectrometric peptide mapping: A rapid, sensitive technique for protein structure analysis. Biochem Biophys Res Commun. 167, 686-92.
  6. Fenn J. B. et al. (1989) Electrospray ionization for mass spectrometry of large biomolecules. Science. 246, 64-71.
  7. Barber M. et al. (1981) Fast atom bombardment of solids as an ion source in mass spectroscopy. Nature. 293, 270-5.
  8. Bakhtiar R. and Tse F. L. (2000) Biological mass spectrometry: A primer. Mutagenesis. 15, 415-30.
  9. Hoffmann E. d. and Stroobant V. (2001) Mass spectrometry : Principles and applications. Chichester ; New York: Wiley. xii, 407pp.
  10. Forsgard N. et al. (2010) Accelerator mass spectrometry in the attomolar concentration range for 14c-labeled biologically active compounds in complex matrixes. Journal of Analytical Atomic Spectrometry. 25, 74-8.
 
Written and/or reviewed by Jared Snider, Ph.D.

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