Analytical Development and Application of Mass Spectrometry to skeletal muscle proteomics and Identification of Structure
Modifications
Dissertation
zur Erlangung des akademischen Grades eines Doktors der Naturwissenschaften
vorgelegt von Bogdan Ioan Bernevic
an der
Naturwissentschaftliche Sektion Fachbereich Biologie
Konstanz, 2011
Tag der mündlichen Prüfung: Donnerstag, den 8. September 2011 1. Referent: Prof. Dr. Iwona Adamska
2. Referent: Prof. Dr. Dr. h.c. Michael Przybylski
Vorsitzender der Prüfungskomission: Prof. Dr. Wolfram Welte
“I don't know driving in another way which isn't risky. The danger sensation is exciting.
Each driver has its own limit. My limit is a little bit further than other's. You will never know the feeling of a driver when winning a race. Being second is to be the first of the
one’s to lose….”
Ayrton Senna (1960-1994)
For my wonderful parents Mariana and Gheorghe Bernevic and for my dear brothers Dragos and Georgian
The current work has been performed in the time frame from October 2006 to October 2010 in the Department of Biology and Laboratory of Analytical Chemistry and Biopolymer Structure Analysis of the University of Konstanz, under the supervision of Prof. Dr. Iwona Adamska and Prof. Dr. Dr. h. c. Michael Przybylski.
I would like to address very special thanks to:
Prof. Dr. Iwona Adamska and Prof. Dr. Dr. h. c. Michael Przybylski for the entire support, scientific advice and suggestions which have had an enormous contribution to my scientific development;
I am very grateful to all our collaborators especially Prof. Dr. Karl Schellander and Prof.
Dr Michael Wicke who kindly provided me with the muscle samples for proteomics and mass spectrometric analysis. Also, special thanks to Prof. Dr. Gerd Döring and Dr.
Martina Ulrich for providing me the cystic fibrosis sputum sample.
Special thanks to Dr. Andreas Marquardt for friendly collaboration and help with the mass spectrometric analysis.
All members of the group are acknowledged for scientific discussions and intresting advices during my work, and to Alexandra Hajnic for all the kind help she gave me.
Last but not least I wish to thank and to express my deep gratitude to my family, my parents and my brothers Dragos and Georgian for supporting and encouraging me during this time.
The following publications have been resulting from this dissertation:
1. Bernevic B., Petre B.A., Galetskiy D., Werner C., Wicke M., Schellander K., Przybylski M., (2010) Degradation and oxidation post-mortem of myofibrillar proteins in porcine skeleton muscle revealed by high resolution mass spectrometry proteome analysis Int. J. Mass Spectrom. 305, (2-3), 217-227
2. Iuliana Susnea, Bogdan Bernevic, Eliska Svobodova, Diliana Dancheva, Simeonova, Michael Wicke, Carsten Werner, Bernhard Schink, and Michael Przybylski (2010) Mass spectrometric protein identification from two-dimensional gel separation with stain-free detection and visualization using native fluorescence Int. J. Mass Spectrom. 301, (1-3), 22-28 3. Marquardt A, Bernevic B. and Przybylski M. (2007) Identification, and affinity characterization and biological interactions of lectin-like peptide-carbohydrate complexes derived from human TNF-α using high-resolution mass spectrometry J. Peptide Sci.13, (12), 803-810
4. Iuliana Susnea, Bogdan Bernevic, Michael Wicke, Li Ma, Shuying Liu, Karl Schellander and Michael Przybylski (2011) Application of MALDI-TOF-Mass Spectrometry to Proteome Analysis using Stain-free Gel Electrophoresis Topics in Current Chemistry - submitted
Conference presentations:
1. Bogdan Bernevic Third International Symposium Oxidative Post-Translational Modifications of Proteins in Cardiovascular Disease (OPTM conference), Boston, September (2008) “Pig muscle protein degradation and oxidation post-mortem revealed by high resolution mass spectrometric proteome analysis” - Oral presentation
2. Brindusa-Alina Petre, Bogdan Bernevic, Martina Ulrich, Gerd Doering and Michael Przybylski, Molecular identification of nitro-tyrosine modification in human eosinophil
proteins by proteolytic affinity extraction-mass spectrometry (PROFINEX), Proteome binder workshop/DGMS Fachgruppe Affinity-MS, Konstanz, December (2009), - Poster presentation 3. Bogdan Bernevic Deutsche Gesellshaft für Massenspectrometrie FTMS Fachgruppen Symposium Mülheim an Ruhr September (2010) The identification of post-mortem pig muscle L. dorsi protein degradation by high resolution FT-ICR mass spectrometry – Oral presentation 4. Bogdan Bernevic, Brînduşa-Alina Petre, Dmitry Galetskiy, Andreas Marquardt, Carsten Werner, Michael Wicke , Karl Schellander, and Michael Przybylski Degradation and oxidation post-mortem of myofibrillar proteins in porcine skeleton muscle revealed by high resolution mass spectrometric proteome analysis American Society for Mass Spectrometry (ASMS) Denver, June (2011) – Poster presentation
TABLE OF CONTENTS
1 INTRODUCTION... 1
1.1 From Genomics to Proteomics... 1
1.2 Analytical approaches for protein identification ... 2
1.3 Mass spectrometric methods for proteome analysis ... 5
1.3.1 Ionization methods ... 5
1.3.2 Mass analysers ... 7
1.4 Proteome analysis of skeletal muscle proteins... 10
1.5 Post-translational modifications in proteins ... 13
1.6 Scientific goals of the thesis ... 18
2 RESULTS AND DISCUSSION... 19
2.1 Mass spectrometric methods for proteome analysis of post-mortem changes of skeletal muscle proteins ... 19
2.1.1 Methods of high resolution mass spectrometry for proteome analysis... 19
2.1.2 Protein visualization using native fluorescence and mass spectrometric identification after two-dimensional gel separation... 21
2.2 Application of mass spectrometry for identification of post-mortem protein changes of porcine skeletal muscle proteins... 29
2.2.1 Muscle protein changes during post-mortem storage ... 29
2.2.1.1 2-D PAGE protein separation and identification of post-mortem protein degradation by high resolution MALDI FT-ICR mass spectrometry ... 32
2.2.1.2 High resolution muscle proteome analysis for identification of pH-dependent post-mortem changes ... 42
2.3 Immunological and mass spectrometric characterization of post-mortem protein carbonylation... 49
2.3.1 Carbonylation structure analysis by mass spectrometry ... 60
2.3.2 Identification of in vivo carbonylation in α-actin ... 62
2.3.3 Identification of in vivo carbonylation sites in muscle proteins... 69
2.4 Mass spectrometric identification of oxidative modification structures in proteins ... 82
2.5 Mass spectrometric characterization of phosphorylated muscle proteins ... 88
2.6 Proteome analysis of nitration and related oxidatively modifications in cystic fibrosis... 92
2.6.1 Protein nitration in cystic fibrosis ... 92
2.6.2 Affinity binding studies of anti-3-nitro tyrosine antibody to nitrated peptides by Dot-blot ... 94
2.6.3 Detection and identification of oxidatively modified proteins using anti-3-nitro tyrosine antibody after 2-D gel separation... 99
2.6.4 Affinity-mass spectrometric characterization of anti-3-nitro tyrosine antibody binding to nitrated peptides ... 106
2.6.5 Mass spectrometric identification of oxidatively modified proteins after affinity- SDS PAGE purification ... 109
3 EXPERIMENTAL PART... 116
3.1 Materials and reagents... 116
3.2 Sample preparation for proteome analysis... 117
3.2.1 Muscle sample preparation using denaturing conditions... 117
3.2.2 Muscle sample preparation using non-denaturing conditions ... 117
3.2.3 Sample preparation of sputum from Cystic Fibrosis patients ... 118
3.3 Labelling methods of protein carbonyl groups for immunological detection 118 3.3.1 Pre-isoelectric focusing labelling of protein carbonyl using 2,4-dinitrophenyl hydrazine ... 118
3.3.2 Post-isoelectric focusing labelling of protein carbonyl using 2,4- dinitrophenyl hydrazine ... 119
3.4 Chromatographic and electrophoretic methods for protein separation and staining procedure... 120
3.4.1 Reverse Phase High Performance Liquid Chromatography... 120
3.4.2 Sodium dodecyl sulphate – polyacrylamide gel... 121
3.4.3 Two- dimensional gel electrophoresis ... 123
3.4.4 Colloidal Coomassie staining ... 124
3.4.5 Silver staining... 126
3.4.6 BioAnalyzer Gel reader for native fluorescence detection... 127
3.5 Chemical modification reaction and enzymatic fragmentation of proteins... 128
3.5.1 Reduction and alkylation of disulfide bonds in gel matrix ... 128
3.5.2 Proteolytic digestion of protein in-gel following silver staining ... 128
3.5.3 Proteolytic digestion of proteins in-gel following Coomassie-blue staining.. 129
3.6 Desalting and concentration of peptides and proteins prior to mass
spectrometric analysis... 129
3.6.1 ZipTip cleanup procedure... 129
3.6.2 Desalting and concentration of peptide samples using Microcon centrifugal filter devices ... 130
3.7 Immuno-analytical methods ... 131
3.7.1 Dot blot assay ... 131
3.7.2 Western blot assay... 131
3.7.3 Preparation of monoclonal anti-3-nitrotyrosine antibody affinity column ... 133
3.8 Mass spectrometry... 134
3.8.1 MALDI – TOF MS... 134
3.8.2 MALDI – FTICR MS ... 135
3.8.3 ESI - Ion Trap MS/MS analysis ... 136
3.8.4 Linear ion trap (Orbitrap) mass spectrometry... 137
3.9 Bioinformatic tools for proteome analysis... 139
3.9.1 PD Quest 2-D gel analysis software... 139
3.9.2 Protein identification based on MS and MS/MS data by using different database search engines ... 140
3.9.3 GPMAW ... 141
4 SUMMARY... 142
5 ZUSAMMENFASSUNG ... 146
6 REFERENCE LIST ... 150
7 APPENDIX ... 167
1 INTRODUCTION
1.1 From Genomics to Proteomics
The term proteomics describes the study and characterization of a complete set of proteins present in a cell, organ, or organism at a given time [1]. In the last years the emphasis has been shifted to the proteome complement of the human organism since the completion of the human genome sequence in 2003. The central dogma of molecular biology states that a gene is transcribed into RNA and then translated into a protein. A subsequent paradigm has been that the genome (all genes in an organism) gives rise to the transcriptome (the complete set of mRNA in cell), which is then translated to produce proteome (the complete collection of genes in any given cell) [2]. In general the genome remains unchanged over time, whereas the proteome is changing by turning on and off certain genes as a response to environmental stimuli. In contrast, both the transcriptome and proteome are dynamic entities whose content can vary dramatically under different conditions due to the regulation of the transcription, RNA processing, protein synthesis and protein modifications. The dynamic nature of the proteome has been termed functional proteomics to describe all the proteins produced by a cell in a single time frame. The transcriptome and proteome are much more complex then the genome, because a single gene can produce many different RNAs and proteins. Different transcripts can be generated by alternative splicing, alternative promoter or poly adenylation site usage, and by special processing strategies such as RNA editing. Many proteins can be generated by alternative use of start and stop codons and the proteins synthesized from these RNAs can be modified in various ways during or after translation [3].
Usually proteins are primarily involved in biochemical processes of both normal and disease organisms and direct analysis of these proteins could lead to a better understanding of diseases. For example, the porcine genome harbours about 20000 protein encoding genes, while the human genome harbours approximately 29000 protein encoding genes [4]. The total number of human protein products, including splice variants and essential post-translational modifications, has been estimated to be close to one million [5, 6]. Thus, muscle genome can generate a large number of protein species encoded by a small number of muscle-specific
genes. Although as an example, the sarcoendoplasmic reticulum Ca2+ ATPase (SERCA)-type Ca2+ proteins from slow and fast muscle are encoded by three genes, the isoform diversity of this ion pump protein is drastically increased by alternative splicing of the transcripts and a number of post-translational modifications [7], resulting more than 10 different SERCA isoforms [8].
Genomics can not predict post-translational modifications that proteins undergo although it provides a vast amount of information linking gene activity with a disease. There are a number of reasons why gene sequence information and the pattern of gene activity in a cell do not provide a complete profile of a protein’s abundance and its final structure and state of activity. After synthesis on ribosomes, proteins are cleaved to eliminate initiation, transit, and signal sequences, as well as simple chemical groups or complex molecules that are attached. The gene transcript can be spliced in different ways prior to translation into protein. Following translation most of proteins are chemically changed by post-translational modifications, notably through structural addition of carbohydrate and phosphate groups. Untill now more than 300 types of post-translational modifications have been documented, among the static, and dynamic types such as phosphorylation, sulfation, glycosylation, acetylation, deamidation and palmitoylation. Moreover, a large class of post- translational modifications are originated by reactive oxygen species giving rise to the oxidative post-translational modifications. Such modifications play an important role in modulating the functions of many proteins, but they are not coded by genes.
1.2 Analytical approaches for protein identification
Protein changes in expression levels and the post-translational modifications can be identified and characterized by the development of proteomics approaches.
Proteome analysis offers the unique possibility of delineation of global changes in expression patterns resulting from transcriptional and post-transcriptional control, post-translational modifications, and shifts between cellular compartments [9].
Analysis of post-translational modifications such as phosphorylation, glycosylation, carbonylation or nitration is very important for understanding of protein activities, stabilities, and turnover. In spite of new technologies, the analysis of complex biological mixtures, techniques for quantifying separated protein species, sufficient
sensitivity for analysis of proteins of low abundance, quantification over a wide dynamic range, the analysis of protein complexes, and high throughput applications are only partially developed. A wide range of proteomic approaches are available such as gel-based applications including one- and two-dimensional polyacrylamide gel electrophoresis [10, 11], and large-scale Western blot assays [12].
Two-dimensional electrophoresis (2-DE) was developed two decades before the term proteomics was coined [13]. The 2-DE entails the separation of complex protein mixtures by molecular charge in the first dimension and by mass in the second dimension. 2-DE analysis provides several types of information about hundreds of proteins, including molecular weight, pI and quantity, as well as some information on possible post-translational modifications. 2-DE is extensively used, but mostly for qualitative experiments, and this method falls short in reproducibility and capability to detect low abundant and hydrophobic proteins. There are two ways to study post-translational modifications by means of 2-DE; first, post-translational modifications that alter the molecular weight and/or pI of a protein are reflected in a shift in location of the corresponding protein spot on the proteomic pattern. Second, in combination with Western blotting, antibodies specific for post-translational modifications can reveal spots on 2-DE patterns containing proteins with these modifications [14]. Protein extraction and solubilization are key steps for proteomic analysis using 2-DE. In order to enhance protein extraction and solubilization, different treatments and conditions are necessary to efficiently solubilise different types of protein extracts [15]. The major challenge for protein visualization in 2-DE is the compatibility of sensitive protein staining methods with mass spectrometric analysis. Therefore, several staining methods have been developed for the visualization of 2-DE patterns including Coomassie Blue [16], silver staining [17] or stain-free [18].
Several MS approaches including so called “buttom-up” and “top down” [19]
are now applied for protein structure determination. In the “buttom up” approach the intact protein is digested using different proteases. Trypsin is the most often used enzyme, which cleaves the protein at the C-terminal side of lysine and arginine residues. In some particular cases proteases with other specificities can be used.
“Top-down” MS refers to an approach, where the intact molecule is introduced into the mass spectrometer without enzymatic fragmentations and structural information
is obtained from the fragmentation pattern of the intact molecule inside the mass spectrometer. Proteins separated by 2-DE are identified based on their chemical structure measured by mass spectrometry (Figure 1.1).
Figure 1.1. Schematic representation of the experimental approach to identify a protein spot.
Generally, the spot of interest is digested with trypsin and analysed by MS. The protein is identified based on the peptide mass fingerprint using MS-specific database search
In combination with 2-DE, the analysis of peptide mixtures by high resolution mass spectrometry such as FTICR-MS and LTQ Orbitrap technologies provided a series of masses with high accuracies suitable for identification of intact proteins [20, 21] and their degradation products [22] with those of genomic databases [23].
Several protein sequence databases are available; for example, Swiss-Prot which is maintained by The Swiss Institute of Bioinformatics and The European Bioinformatics Institute. A number of search engines can also be accessed over the Internet such as: ProFound, Fit MS or MASCOT at Matrix Science Server.
1.3 Mass spectrometric methods for proteome analysis 1.3.1 Ionization methods
The characteristic features of mass spectrometry have raised it to an outstanding position amongst analytical methods: unequalled sensitivity, low detection limits, speed and diversity of applications. In analytical chemistry, the recent applications are mostly oriented towards biochemical problems, such as proteomic, metabolomic, high throughput analysis in drug discovery and metabolism, and characterization of biomacromolecular complexes [24-27]. Other analytical applications are applied in pollution control, food control, forensic science, natural products or process monitoring. The ability to ionize large molecules was further improved with the development of electrospray ionization (ESI) by Fenn and coworkers since 1988 [28]. The electrospray ionization source was easily connected on-line to liquid chromatography (LC), which enable the analysis of complex mixtures. The development of matrix-assisted laser desorption/ionization (MALDI), was the result of pionners work by several groups such as Hillenkamp, Karas, and coworkers [29] and by Tanaka and coworkers [30]. Like ESI, the MALDI ion source is capable of ionizing and vaporizing large molecules such as proteins. ESI and MALDI mass spectrometry have progressed very rapidly during the last decade. This progress has led to the advent of new instruments. New atmospheric pressure sources were developed [31], existing analysers were perfected and new hybrid instruments were realized by new combinations of analysers. The analysers based on new concepts are Fourier transform ion cyclotron resonance mass spectrometry (FTICR MS) [32] and LTQ Orbitrap [33].
Electro-spray Ionization (ESI) is today the widely used atmospheric pressure ionization application. It has undergone remarkable growth in recent years and it is frequently used for LC/MS of thermally labile and high molecular weight compounds.
The electrospray is created by applying an electric potential between a metal inlet needle and the first skimmer in an atmospheric pressure ionization source (Figure 1.2). The mechanism for the ionization process is not well understood and, there are several different theories that explain this ionization process. One theory is that as the liquid leaves the nozzle, the electric field induces a net charge on the small droplets. As the solvent evaporates, the droplet shrinks and the charge density at the
surface of the droplet increases. The droplet finally reaches a point, where the coulombic repulsion from this electric charge is greater than the surface tension holding it together. This causes the droplet to explode spontaneously and to produce multiply charged analyte ions. A typical ESI spectrum shows a distribution of molecular ions with different charge numbers. Because electrospray produces multiply charged ions, high molecular weight compounds are observed at lower m/z value. This increases the available mass range of the analyzer, so that higher molecular weight compounds may be analyzed with a less expensive mass spectrometer. An ion with a mass of 5000 amu and a charge of +10 can be observed at m/z 500 and is easily analyzed with an inexpensive quadrupole analyzer.
Atmospheric Pressure Vacuum
Tailor cone
Spray Solution with
Analyte
+ Electric field Capillary
Solvated
macromolecular ion
10 µm 10 nm 1 nm
Aerosol droplets
+ + +
+ +
+ + + + +
+ + + +
+ + + +
++
+ + + +
Desolvated
macromolecular ion (3+) Atmospheric Pressure Vacuum
Tailor cone
Spray Solution with
Analyte
+ Electric field + Electric field Capillary
Solvated
macromolecular ion
10 µm 10 nm 1 nm
Aerosol droplets
+ + + +
+ +
+ +
+ + + + +
+ + + + +
+ + + +
+ + + + +
+ +
+ + +
+ + + +
++ + + ++ +
+ + + + +
+ + +
Desolvated
macromolecular ion (3+)
Figure 1.2. Schematic representation of electrospray ionization. Positive ions are formed by applying high positive potential to the capillary (anode). The excess ions accumulate at the liquid surface then adopt the shape of a cone called “Taylor cone” from the tip of which a fine jet of liquid is ejected.This jet disintegrates into charged droplets whose diameter ranges between 10 nm -1 µm
Matrix Assisted Laser Desorption/Ionization (MALDI) is a technique, which directly ionizes and vaporizes the analyte from the condensed phase. MALDI is often used for the analysis of synthetic and natural polymers, proteins, and peptides.
Analysis of compounds with molecular weights up to 200,000 daltons is possible and this high mass limit is continually increasing. In MALDI, both desorption and ionization is induced by a single laser pulse (Figure 1.3). The sample is prepared by mixing the analyte and a matrix compound chosen to absorb the laser wavelength.
The mixture is placed on a probe tip and dried. A vacuum lock is used to insert the probe into the source region of the mass spectrometer. A laser beam is then focused on this dried mixture and the energy from a laser pulse is absorbed by the matrix.
Figure 1.3. The MALDI source. The laser is pulsed at a mixture of matrix and sample molecules that have been co-crystallized. When the laser strikes the matrix crystals, the energy deposition cause a rapid heating of the crystals, which may lead to the sublimation of the matrix crystals and the expansion of the matrix and analyte in the gas phase. Ions are formed through gase-phase proton transfer reactions in the expanding gas phase plume
This energy ejects analyte ions from the surface so that a mass spectrum is acquired for each laser pulse. The detailed mechanism for this process is not well understood and it is the subject of much controversy in the literature. This technique is more universal (works with more compounds) than other laser ionization techniques, because the matrix absorbs the laser pulse. With other laser ionization techniques, the analyte must absorb at the laser wavelength. Typical MALDI spectra include the molecular ion, some multiply charged ions, and only few fragments.
1.3.2 Mass analysers
In the present work two high performance instruments were used, namely FT-ICR MS and LTQ-Orbitrap. Of all mass spectrometric methods Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS) combines high resolution, high mass-accuracy, non-destructive multichannel detection, long ion-observation times, the possibility of performing gas-phase reactions on trapped ions, and, most importantly, tools for structural analysis of large biomolecules and complexes [27, 34]. The heart of FTICR-MS is the analyzer cell, which can be cubic or cylindric in shape placed in a strong magnetic field. The ICR cell is made of three opposing pairs of plates forming the cube, which are positioned such that one pair (trapping plates)
lies perpendicular and the other two pairs (excitation and detection plates) parallel to the magnetic field lines. In one of the trapping plates is an orifice through which ions can enter the cell (Figure 1.4).
Figure 1.4. The cyclotron motion and the excitation process shown in the cubic analyzer cell
After injection into the cell, ions undergo harmonic oscillations in the electric field between the trapping plates (trapping motion). Simultaneously, ions in the analyzer cell are exposed to the strong magnetic field and undergo stable cyclic motion in a plane perpendicularly to the magnetic field, the so-called cyclotron motion as a result of the Lorentz force and the centrifugal force acting in opposite directions.
The angular frequency of this motion is given by ωc = qB0 /m
where ωc is the unperturbed cyclotron frequency dependent on the magnetic field B0 and the mass-to-charge ratio m/z. Movement of ions parallel to the magnetic field is not influenced by this field. Each ion rotates with its typical frequency in respect to its mass-to-charge-ratio (m/z), the so-called cyclotron frequency. With increasing magnetic field strength however, the performance of the FTICR-MS instrument improves [20, 35-38]. Thus, the mass resolution is directly proportional to the magnetic field strength, while the highest mass-to-charge ratio that can be determined in the analyzer cell increases with the square of the magnetic field.
Routinely, using a FT-ICR MS instrument a resolution of 180,000 can be achieved
with the isotopic peaks well-resolved providing direct information about the macro- molecular structure.
The analytical capabilities for the identification of protein modifications rely strongly on the sensitivity and dynamic range of the mass spectrometric methods employed. In the last few years, mass spectrometry based on the Orbitrap technology was proven to be ideal for the identification of protein post-translational modifications (e.g. phosphorylations), since this technology provides highest sensitivity as well as high mass accuracy. An orbitrap mass analyzer is the most recent addition to the set of tools that can be applied to identification, characterization and quantitation of components in biological systems [33, 39]. With its ability to deliver low-ppm mass accuracy and high resolution, all within a time scale compatible with nano-LC separations, the orbitrap has become an instrument of choice for many proteomics applications [40]. Unlike FT-ICR MS ions are not trapped inside the ICR cell, but the moving ions are trapped in an electrostatic field [41]. The electrostatic attraction towards the central electrode is compensated by a centrifugal force that arises from the initial tangential velocity of ions like a satellite on the orbit.
The analysis of oxidative post-translational modifications is often quite difficult, since these modifications occur at low stoichiometric levels. A key technology for the analysis of proteins and their modifications is liquid chromatography coupled to high-resolution mass spectrometry (LC-MS) using collision induced dissociation (CID) [42] (described in detail in the “Experimental part”) emerged as a powerful tool for identification and molecular characterization of oxidative post-translational modifications, providing information about their site- occupancy and site-specific microheterogeneity. The fragmentation spectra of peptides for database searches might not be necessarily required with such a high precision. Recording MS/MS spectra with the linear ion trap detector delivers comfortably 3–5 spectra per second. Usually both mass analyzers work in parallel;
while a high resolution/ mass accuracy spectrum of the precursor is acquired in the orbitrap, the fast linear ion trap carries out fragmentation and detection of MS/MS (or higher order MSn) spectra of selected peptides. True parallel operation is achieved, because the initial small part of the high resolution spectrum could be used for data- dependant selection of precursors, which are then fragmented while high resolution spectrum is still being acquired. In real-life applications, an orbitrap spectrum
acquired at resolving power of 60,000, and 3–5 linear ion trap fragmentation spectra are obtained within approximately 1 second [43].
1.4 Proteome analysis of skeletal muscle proteins
Skeletal muscle tissue represents the cellular units responsible for coordinated excitation-contraction-relaxation cycles involved in voluntary movements and postural control [44]. Skeletal muscle plays an important role in heat homeostasis and has the capacity for insulin-mediated uptake of glucose in the body, making the muscle tissues important organ in carbohydrate metabolism and that integrates various biochemical pathways. Proteomics represents a key technology in biochemistry and a thorough approach for a detailed analysis of heterogenous types of tissue, such as muscle. The main goal of skeletal muscle proteomics is the identification, structural and biochemical characterization of the entire protein complement of voluntary contractile tissues in both normal and disease [45].
In the last few years, mass spectrometry-based proteomics has been successfully applied for identification of several hundred of most abundant and soluble muscle-associated proteins [46, 47] leading to better understanding of muscle development, maturation and aging [48, 49], as well as the analysis of contractile tissues undergoing physiological adaptations during physical exercise and chronic muscle transformation [50, 51]. Biomedical research into the proteome-wide alterations of skeletal muscle tissue was also used to development of new biomarker signatures of neuromuscular desease. Such markers were determined for muscle- associated disease such as diabetes-related contractile weakness [52], hypokalemic myopathy [53], sepsis [54], dystrophinopathy [55] and obesity [56]. Muscle proteomics has been applied successfully for the identification and characterization of protein changes during post-mortem storage, particularly protein degradation related to meat tenderness [57, 58] or proteomic evaluation of muscle hypertrophy in chicken, sheep, pig and cow muscles [59-62].
Despite of the advanced technology now available, it is still extremely difficult to achieve an accurate detection and identification of skeletal muscle proteome, because of the limited availability of all-encompassing protein analytical capabilities.
At the moment, no biochemical techniques can efficiently separate and consistently
detect the total protein content from a given cell type. Two-dimensional gel electrophoresis represents the most powerful tool for separation of protein mixtures extracted from different cells, although, there are some limitations, which are important to be taken into consideration: (i), 2-D PAGE separation of cell extracts usually underestimates the presence of low abundant proteins, (ii), proteins with extreme pI values are not well resolved at the edge of the gels, when the IPG strips with broad pI range are used. This problem can be overcome by using narrow-range pI IPG strips or by employing overlapping gel systems covering several pI ranges [63], (iii), distortion of the protein spots due the the presence of highly abundant proteins or protein isoforms with extensive post-translational modifications [64].
Particularly, the presence in muscle sample of highly abundant muscle proteins such as actin, myosin heavy chains and light chans, troponins and tropomyosins can distort those specific zones within 2-D gel separation and, thus, contaminating neighbouring protein spots. In this case, the identification and densitometric analysis of that spot located in this gel region might be difficult. However, in spite of these limitations 2-D gel electrophoresis yields excellent coverage of soluble and abundant muscle proteins involved in regulation and execution of the contraction-relaxation cycle, energy metabolism and the cellular stress response [65].
A classic proteomics experiment involves several steps: (i), muscle sample preparation for extraction of soluble proteins, distinct subcellular fractions including membrane-associated elements or isolated complexes, (ii), separation of proteins by one-dimensional gel-electrophoresis or high resolution 2-D gel electrophoresis and/or protein transfer on PVDF membranes, (iii), determination of altered expression levels in protein maps by densitometric gel analysis (iv) the identification of different proteins by mass spectrometric analysis of tryptic peptides based on the mass fingerprint [66]. The experimental steps of a proteomics analysis are summarized in Figure 1.5.
Skeletal muscle
Extraction of muscle protein content
Protein separation
1-D PAGE 2-D IEF/PAGE Western blot
Determination of abundance protein changes of muscle proteins
Mass spectrometric identification of specific muscle proteins
Interpretation of mass spectrometric data Skeletal muscle
Extraction of muscle protein content
Protein separation
1-D PAGE 2-D IEF/PAGE Western blot
Determination of abundance protein changes of muscle proteins
Mass spectrometric identification of specific muscle proteins
Interpretation of mass spectrometric data
Figure 1.5. Schematic representation of analytical and immunological methods used for the identification and characterization of muscle proteins. In the first step muscle proteins are extracted using different lysis buffers and after 2-D gel separation proteins are visualized by staining methods such as Coomassie and silver, or by "stain-free" native fluorescence. Changes in protein expression can be determined by using PDQuest software. Following 2-D gel electrophoresis proteins can also be transferred onto membranes for Western blotting experiments using specific antibodies. Protein spots are excised from gels, digested usually with trypsin, and subsequently analysed by mass spectrometry, then relevant spectra are manually verified
1.5 Post-translational modifications in proteins
Post-translational modifications of proteins regulate protein functions by causing changes in protein activity, their cellular locations and interactions with other proteins. These kinds of protein modifications are transient and reversible involved in signalling pathways from membrane to nucleus in response to external stimuli.
Besides performing catalytic functions, signaling proteins modified by phosphorylation, myristoylation, farnesylation, cysteine oxidation, ubiquitination, acetylation, methylation, nitrosylation, etc, serve as scaffolds for the assembly of multiprotein signaling complexes, as adaptors, as transcription factors and as signal pathway regulators [67]. Phosphorylation and dephosphorylation on S, T, Y and H residues are the best known modifications involved in reversible, activation and inactivation of enzyme activity and modulation of molecular interactions in signaling pathway [68]. Acetylation regulates many diverse functions, including DNA recognition, protein-protein interaction and protein stability. Acetylation and deacetylation in N-terminal and K-residue are suggested as rival to phosphorylation [69]. As well as phosphorylation, glycosylation is another regulatory post-translational modification which occurs on serine and threonine residues of cytosolic and nuclear proteins [70]. The glycans are distinguished by the type of linkage atom involved as N-, O-, C- or S-glycans. Determination of glycoprotein structure modification is difficult because of the number of monosaccharides and amino acid residues involved in the O-glycan linkages [71]. Previous reports show that there is a sequence consensus for N-glycosylations, Asn-Xaa-Ser/Thr/Cys where the Xaa may be any amino acid except Pro [72].
Furthermore, biological functions of many proteins may be altered by ubiquitination and deubiquitination, sumoylation and desumoylation through covalent attachment to the polypeptide modifiers. Ubiquitin plays a key role in targeting proteins for degradation by the proteasome [73]. In recent reports it is suggested that ubiquitination, sumoylation, acetylation and methylation of lysine residues link specific covalent modification of the transcriptional apparatus to their regulatory function [74]. The most common post-translational modifications are summarized in Table 1.1.
Table 1.1. Summary of the post-translational modification found in proteins PTM types Mass
increament Modified amino
acid residuesa Positionb Remarks
Phosphorylation 79.9 Y, S, T, H, D Anywhere Reversible, regulate protein function Glycosylation Anywhere Reversible, cell-cell interaction O-linked > 800 S, T
(O-Glc-NAc) 203.2
N-linked > 800 N
Acetylation 42.0 S N-term Reversible, protein activity, stability
K Anywhere
Deamidation 0.9 N, Q Anywhere N to D, Q to E
Methylation Anywhere Regulation of gene expression
Monomethylation 14.0 K
Dimethylation 28.0 K
Trimethylation 42.0 K
Acylation Cellular localization to membrane
farnesylations 204.3 C C-term
myristoylation 210.3 G N-term
K Anywhere
palmitoylation 238.4 C (S, T, K) Anywhere Carbonylation
(4-HNE) 156.1 C, H, K Anywhere Oxidative damage
Nitration 45.0 Y
S-Nitrosylation 29.0 C
Cys oxidation Anywhere Oxidative regulation of proteins Disulfide bond -2.0 C
Glutathionylation 305.3 C Sulfenic acid 16.0 C Sulfinic acid 32.0 C
Ubiquitination K Anywhere Reversible/ireversible
Sumoylation K [ILFV]KD
Hydroxyproline 16.0 P Protein stability
Pyroglutamic
acid -17 Q N-term
a- amino acid residues where the modification has occurred b- the location of the modified amino acid in the protein sequence
A large class of post-tranlational modifications are caused by reactive oxygen species which induce oxidative post-tranlational modifications. ‘‘Oxidative stress’’
occurs when the balance of formation of oxidants exceeds the ability of antioxidant systems to remove reactive oxygen species, when inflammatory phagocytes (e.g., neutrophils and macrophages) are activated to undergo an oxidative burst by exposure to a foreign agent. Under these conditions, biomolecules become subjected to attack by excess reactive oxygen species and significant molecular and physiological damage can occur [75, 76]. Reactive oxygen species encompass a variety of diverse chemical species including superoxide anions, hydroxyl radicals and hydrogen peroxide. Some of these species, such as superoxide or hydroxyl
radicals, are extremely unstable, whereas others, like hydrogen peroxide, are freely diffusible and relatively long-lived. Because there are so many mechanisms for induction of protein oxidation and because all of the amino acyl side chains can become oxidatively modified, there are numerous different types of protein oxidative modification. The thiol (-SH) moiety on the side chain of the amino acid cysteine is particularly sensitive to redox reactions and is an established redox sensor. As for cysteine residue-specific oxidative post-translational modification of the protein (S- thiolation), it is found to act as a switch regulating biological function like phosphorylation-dephosphorylation [77]. With Met residues, the major product under biological conditions is methionine sulfoxide [78]
Direct protein carbonylation can occur through a variety of reactions.
Oxidation of amino acid side chains with metals and hydrogen peroxide is known to cause the formation of semialdehyde amino acids, with the majority of these reactions occurring with lysine, arginine, and proline residues [79]. According to their proposal, lysine, proline and/or arginine from myofibrillar protein are oxidized in the presence of Fe3+ and H2O2 to yield aminoadipic semialedehyde. The reaction is initiated by OOH● radicals derived from the reaction between Fe3+ and H2O2. The oxidative deamination from the intermediate radical molecule occurs in the presence of Fe3+ and yields the semialdehyde. The resulting Fe2+ could propagate the oxidative degradation to new amino acid residues by reacting with H2O2 to form further hydroxyl radicals. A recent study [80] has confirmed the H2O2-activated myoglobin and metal ions such as Cu2+ are also able to promote the formation of aminoadipic semialedehyde from myofibrillar proteins. Aminoadipic semialedehyde is thought to account for approximately 70% of the total amount of protein carbonyls formed in oxidized animal proteins [81]. It is worth noticing that the formation of these semialdehydes does not require a previous cleavage of the peptide bond as protein- bound amino acids can be degraded into their corresponding semialdehydes.
Aminoadipic semialedehyde has already been detected and employed as indicators of protein oxidation in raw meat and a large variety of processed muscle foods such as cooked patties, frankfurters and dry-cured meats [82, 83].
Alternatively, protein carbonylation can result from an indirect mechanism involving the hydroxyl radical-mediated oxidation of lipids. Polyunsaturated acyl chains of phospholipids or polyunsaturated fatty acids such as arachidonic acid and
linoleic acid are highly susceptible to peroxidation and breakdown through non- enzymatic Hock cleavage, forming a variety of lipid-derived aldehydes and ketones [84]. Lipid peroxidation products can diffuse across membranes, allowing the reactive aldehyde-containing lipids to covalently modify proteins localized throughout the cell and relatively far away from the initial site of reactive oxygen species formation. The most reactive aldehydes generated from polyunsaturated fatty acid oxidation are 4- hydroxy-2-nonenal (HNE) [84], 4-oxo-2-nonenal [85], and acrolein [86]. Because of the presence of electron-withdrawing functional groups, the double bond of 4-HNE or 4-ONE serves as a site for Michael addition with the sulfur atom of cysteine, the imidizole nitrogen of histidine, and, to a lesser extent, the amine nitrogen of lysine.
The modification can take place by the 1,4-addition (Michael addition) of the nucleophilic groups in cysteine, histidine or lysine residues of the protein, respectively, onto the electrophilic double bond of HNE, giving an increase in the protein’s molecular mass by 156 amu with each molecule of HNE being added. After forming Michael adducts, the aldehyde moiety may in some cases undergo Schiff base formation with amines of adjacent lysines, producing intra- and/or intermolecular cross-linked amino acids [87, 88]. 4-HNE exerts a potentially detrimental effect to proteins by forming covalent adducts, resulting in diminished protein function, altered physicochemical properties [89] and induction of antigenicity [90]. In addition to these reversible oxidative post-translational modifications, it becomes clear that irreversible oxidative modification of histidine, lysine, and cysteine residues involves the generation of abnormal protein associated with the etiology of lifestyle related diseases.
Tyrosine nitration is a covalent protein modification resulting from the addition of a nitro- (NO2) group onto one of the two equivalent carbons CE1 and CE2 in the ortho position relative to the hydroxyl group of tyrosine residue and it is believed to depend on the simultaneous availability of tyrosyl (Tyr•) and nitrogen dioxide (•NO2) radicals [91, 92]. Since nitration has the effect of decreasing the pKa of the phenoxyl groups (from ~ 10 to 7.5 in free tyrosine), this modification may not only change the protein conformation, but may also affect the redox and signalling properties of tyrosines, thus contributing to peroxynitrate-mediated cell signalling [93]. There has been increasing interest in the effects of tyrosine nitration on changes in protein structure in diverse pathogenesis [94]. Protein tyrosine nitration usually occurs near
basic residues in loop regions [95] in areas free of steric hindrances. Also, the presence of amino acids that compete for nitrating agents proximal to tyrosine residues, including tryptophan (Trp), cysteine (Cys), and methionine (Met), may prevent tyrosine nitration by removing the nitrating agents [96]. The location of tyrosine residues in favorable environments for nitration within the secondary and tertiary protein structure may also influence site-specific nitration.
1.6 Scientific goals of the thesis
Structural and metabolic changes in the muscle during different physiological/pathophysiological conditions as well as structural changes during post- mortem storage have been extensively studied with the focus on the genes and proteins. Proteomic profiling has been applied to define the molecular signature of denervation, immobilization, exercise-training, age-related muscle disease, insulin- resistance, muscular dystrophy and meat quality. Sometimes, determination of structural changes of metabolic enzymes and contractile proteins and their corresponding post-translational modification and/or oxidative post-translational modifications is very difficult, because such structural modifications are present at very low stoichiometrical level. Such problems can be overcome by development of separation techniques coupled with advanced mass spectrometric methods. The major goals of the present thesis were a comprehensive study of muscle proteins structural changes by application of high resolution 2-D gel electrophoresis in combination with high resolution mass spectrometric methods.
The scientific goals of the present dissertation are summarized as follows:
Analytical development and bioanalytical application of high resolution 2-D gel electrophoresis and mass spectrometric methods for identification and structural characterization of meat quality biomarkers: (i), Proteomics analysis of proteome changes during post-mortem storage of muscle samples with different pH values and (ii) structure determination of post-mortem protein degradation products by high resolution MALDI FT-ICR mass spectrometry.
Identification and characterization of oxidation structures occurring in muscle protein by high performance liquid chromatography in combination with collision induced dissociation mass spectrometry: Determination of muscle protein carbonylation sites occurred via (i), lipid peroxidation (ii), metal catalysed oxidation (MCO) and (iii) non-carbonylation protein oxidation
Isolation and separation of sputum proteins extracted from cystic fibrosis patients for identification of physiological protein nitration and oxidation. In this part, a combination of immunologic and affinity-mass spectrometric methods was employed for the identification of specific protein oxidation using specific antibodies.
2 2.1
RESULTS AND DISCUSSION
Mass spectrometric methods for proteome analysis of post-mortem changes of skeletal muscle proteins
2.1.1 Methods of high resolution mass spectrometry for proteome analysis In proteome applications, the various cell proteins are separated by 2-D gels and in-gel digested. MS and MS/MS analyses are used for the protein identification via database searches [23]. In functional proteomics protein complexes containing 2 up to 50 components do not need 2-D gels, but for a safety analysis via SDS-PAGE of small and very large components the analysis of digests of intact isolated complexes is important. Mass spectrometers with high mass resolution and accuracy are taken into consideration for complex protein mixture analysis because of lower requirements for the preceding separation. Proteins can now be analysed by MS to reveal elemental composition, complete or partial amino acid sequence, post- translational modifications, protein-protein interaction sites, and even provide insight into conformational aspects. With these pieces of information new data into the molecular behaviour of individual molecules or molecular ensembles can be obtained directly from mass spectrometric data. As a result, mass spectrometry has evolved into an enabling discipline that plays an increasingly important role in many areas of science, particularly, in characterization of proteins function in many different biological systems.
Of all mass spectrometric methods Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS) offers a unique combination of analytical qualities.
An FTICR mass spectrometer combines high resolution, high mass accuracy, non- distructive, multichannel detection, long ion-obsevation times and, most importantly, tools for structural analysis of large molecules. The most important advantages of FT-ICR as a mass analyser is that the ion mass-to-charge ratio is experimentally manifested as a frequency. Because the frequency can be measured more accurately than any other experimental parameter, ICR-MS, offers inherently higher resolution (and thus higher mass accuracy) than any other type of mass measurement. The introduction of FT techniques to ICR MS, by Comisarow and Marshall [35] has brought the advantages of increased speed (factor of 10,000), or increased sensitivity (factor of 100), but also the advantage of fixed magnetic field;
namely, increased mass resolution (factor of 10,000) and increased mass range (factor of 500). Applications that derive from these advantages include determination of chemical formulas, particular in complex mixtures and detection limit in attomole range [20].
However, the high complexity and cost of the FT-ICR, as well as the relatively low space-charge capacities of both analyzers, suggests why new approaches to ion trapping are welcome in tackling the increasingly complex problems in biological mass spectrometry. Therefore, a more compact, less costly, easier to maintain analyzer with comparable performance (for relatively short acquisition time ≤1.8 sec) was desired to supplement the FT-ICR. This technological gap was filled by the LTQ-Orbitrap hybrid mass spectrometer. The coupling liquid chromatography and mass spectrometry improved significantly, the efficiency of proteome analysis by providing sensitivity, resolution and mass accuracy. Hence, the possibility to perform tandem MS experiments (MS/MS) enhanced the capability of structure determination. Despite its relatively recent commercial introduction, the LTQ-Orbitrap has already proven to be an important analytical tool with a wide range of applications. The high resolving power (>150,000) and excellent mass accuracy (specified as ~2–5 ppm, but demonstrated to be as low as 0.2 ppm under favourable conditions) [39] significantly reduce false positive peptide identifications in bottom-up protein analyses. The most important benefit of using LC-MS/MS is the extended dynamic range [40]; complex peptide mixture can be simultaneously analysed containing a wide range of concentrations. These high performance features of the orbitrap can facilitate unambiguous determination of the charge states of fragment ions, as well as identification of oxidative post-translational modifications. The interpretation of LC-MS data is performed routinely in a widely automated fashion as a result of recent development of bioinformatics tools for data acquisition and analysis, although all relevant spectra need to be verified manually.
2.1.2 Protein visualization using native fluorescence and mass spectrometric identification after two-dimensional gel separation
A variety of protein detection and visualisation techniques of protein bands or spots from one and two-dimensional gel electrophoretic separations have been developed and employed in mass spectrometric proteomics. Well established staining procedures for visualization of proteins in gels have used dyes such as Coomassie Brilliant Blue, silver salts and fluorescent dyes (Flamingo, Sypro®Ruby) [16, 97]. While several of these approaches provide high detection sensitivities of proteins, major problems are frequently encountered with the compatibility of staining procedures with the mass spectrometric analysis, background arising from polar staining materials, and the need for applying destaining procedures of isolated proteins. Several procedures have been recently explored in order to overcome these problems, by using unstained gels in gel electrophoretic separations [98, 99].
Fluorescence detection of proteins has been evaluated with pre- or post- electrophoretic incorporation of halogenated compounds such as trichloromethane, trichloroethanol and trichloroacetic acid, which react with tryptophane residues upon treatment with UV light yielding products that show emission in the visible light range suitable for visualization of protein bands [99]. A direct UV fluorescence detection method for unstained proteins in gels was first developed by Roegener et al. who used laser excitation with 280 nm UV light (35 mJ/cm2) and showed the visualisation of proteins in both 1-D and 2-D gel separations with low detection limits (1-5 ng) [98].
A commercial gel bioanalyzer based on native fluorescence has been recently developed (LaVision-BioTec; Bielefeld, Germany) [100]. Native fluorescence detection of proteins was developed in stain-free one and two-dimensional gel electrophoretic separations as a sensitive and efficient approach for mass spectrometric identifications in proteome analysis [18]. Following 1-D or 2-D gel separations, proteins were visualized using sensitive colloidal Coomassie staining and silver staining as described using a GS-800 calibrated imaging densitometer (Bio-Rad, München, Germany), or scanned with the Gel-BioAnalyzer (BAG). The components of the Gel-Bioanalyzer (LaVision-Biotec GmbH, Bielefeld, Germany) are schematically shown in Figure 2.6.
Figure 2.6. Scheme of the Gel-BioAnalyzer (LaVision- Biotec; Bielefeled, Germany), adapted after http://www.lavisionbiotec.com/en/microscopy-products/gelreader/ used for stained free mass spectrometric protein identification
The experimental setup is based on a UV excitation source and a detection system within the UV range. The UV excitation light was generated by a 300 W xenon lamp (265 - 680 nm). The irradiation area was set to 1 cm2 at 35 mW/cm2 and imaged by three lenses onto a photomultiplier detector. A UV bandpass filter (280- 400 nm) is incorporated to block the excitation light from the detection system. From four filter positions (one for UV excitation, three for visible fluorescence), the UV filter transmitting light at λ = 343 ± 65/2 nm was employed. The large reading area (30 x 35 cm2) provided scanning of both 1-D and 2-D gels. The instrument has a removable gel tray and is equipped to read unstained as well as stained protein gels.
In the present study only scanning of unstained gels was applied. High precision polycarbonate tools for localisation and isolation of protein spots were prepared by the Department’s mechanical workshop; after fixation in position on the gel tray, localisation and isolation of gel spots was carried out by moving the gel tray, with positioning and scanning of the gel controlled by the LaVision- Biotec scanning
software. SDS-PAGE separations of several model proteins were initially tested by comparison of Colloidal Coomassie, silver staining, and stain-free detection using native fluorescence and fluorescence upon fixation with halogenated derivatives.
Previously, it has been shown the incorporation of halogenated compounds in polyacrylamide gels either prior to polymerization [99] or subsequent to the electrophoretic separation [101], followed by UV illumination provide fluorescent protein derivatives. However, our model studies showed that protein fixation using halogenated derivatives after SDS-PAGE can be omitted, as illustrated by SDS- PAGE separations of lysozyme and myoglobin without fixation (Figure 2.7a), and with fixation (30 min) in 12 % trichloroacetic acid (TCA) (Figure 2.7b).
Figure 2.7. Comparison of native fluorescence detection for 12 % SDS-PAGE separation of 5 µg hen eggwhile lysozyme (lane 1) and 5 µg myoglobin (lane 2) with and without fixation with trichloroacetic acid. For each gel a 5 µl aliquot of molecular weight marker (10– 200 kDa) was used (lane m). (a), no protein fixation was performed; (b), proteins were fixed for 30 min with a 12 % aqueous trichloroacetic acid solution
Upon scanning with the gel bioanalyzer, protein bands were detected with and without fixation, however bands with approximately 30-50 % higher abundance were observed without protein fixation for the model proteins studied. A further increased abundance was obtained by washing the gel band with water which leads to reduced background fluorescence; however, a washing step was found to cause decreased stability of the protein fluorescence. We observed an increased stability (slower decrease) of the fluorescence intensity in proteins within 48 hours after
fixation in 12 % TCA (data not shown), which may be explained by a more stable fluorescence emission of UV-reaction products of tryptophane with TCA. The MALDI- mass spectrometric identification of the gel band of myoglobin isolated from the gel presented in Figure 2.7a (lane 2, without fixation) is shown in Figure 2.8 (see details of protein localisation and isolation below). The band was excised, in-gel digested with trypsin, and the digest mixture analysed by MALDI-TOF-MS, followed by database search employing the MASCOT peptide mass fingerprinting (PMF) search engine. The database search provided unequivocal identification of myoglobin.
Figure 2.8. MALDI-TOF mass spectrum of horse heart myoglobin (lane 2 in Figure 2.7) identified after stain-free gel detection and in-gel digestion with trypsin
For the gel scanned with the Bioanalyzer, no destaining step was required, which provided high sensitivity and considerably lower sample preparation time compared to staining procedures. For the 320 ng protein band, identification was obtained with a score of 83 (64 % sequence coverage). Thus, these model studies clearly showed that gel separation of proteins detected by native fluorescence without a fixation step represents an efficient and sensitive approach for mass spectrometric identification, providing sufficiently fast mass spectrometric analysis of the gel separated proteins.
Comparative gels scanned with the gel Bioanalyzer and stained with Coomassie blue and silver were performed in order to test the sensitivity of the stain- free native fluorescence detection. Corresponding gel separations of mixtures of the two model proteins, immunoglobulin-G and bovine serum albumin at concentrations of 320 - 5 ng/band are compared in Figure 2.9.
Figure 2.9. Sensitivity of stain-free fluorescence detection and visualization in comparison with silver visualization. Protein samples, IgG (150 kDa heavy and light chain dimer) and BSA (67 kDa) were separated in 7 lanes at 320 - 5 ng. Gel areas presented are zoomed regions from 12 % SDS-PAGE separations. (a), silver stained gels; (b), native fluorescence gels
These results indicated comparable sensitivities for the UV fluorescence detection and silver staining with detection limits of approximately 1-5 ng for IgG, while the corresponding Coomassie-stained gel band was not detectable at this concentration (data not shown). The detection limit in the low nanogram range observed is in good agreement with sensitivity data reported by Roegener et al.[98].
In order to evaluate the isolation and localization of protein bands using the stain-free detection, protein extracts from pig muscle were separated and analysed by 2-D gel electrophoresis. Corresponding 12 % separation gels of muscle protein extract were prepared and visualized by native fluorescence and Coomassie staining (Figure 2.10a, b). Using the Bio-Rad PDQuest software approximately 600 µg proteins were detected in the 2-D gel with native fluorescence (Figure 2.10a), while the detection of approximately 350 protein spots was estimated in the Coomassie- stained gel (Figure 2.10b).
Figure 2.10. Protein identification after 2-D gel separation (12 % SDS-PAGE) of a post-mortem porcine muscle sample (1.5 mg total protein per gel). (A) Gel visualized by native fluorescence (B) Gel stained with Coomassie. MALDI-TOF identification of (C) α-actin (spot 1), (D) Creatine kinase (spot 2), (E) Triosephospate isomerase (spot 3), (F) adenylate kinase (spot 4), (G) Myosin regulatory light chain 2 (spot 5)
This suggests comparable detection sensitivity of the protein spots with native fluorescence and Coomassie staining. Since the fluorescence signal depends on the abundance of aromatic amino 12 acids in the protein, proteins with low abundance in aromatic amino acid residues may not be visualized using this method.
Figure 2.10 shows five examples of stained-free gel spots detection using gel Bioanalyser. Following tryptic digestion of isolated gel spots, the MALDI-MS analysis provided the identification of five proteins summarized in Table 2.2.
Table 2.2. Protein identifications in proteome applications using native fluorescence for visualization of gel separated proteins.
Spot Protein Score Peptides
matched Sequence
coverage % Accession number
1 Skeletal α-actin 92 16 70 P68137
2 Creatine kinase M chain 78 22 90 Q5XLD3
3 Triosephosphate isomerase 78 12 28 Q29371
4 Adenylate kinase isoenzyme 1 98 9 49 P00571
5 Myosin regulatory light chain 2 76 9 44 P02608
Using the stain-free gel bioanalyzer enabled the detection and mass spectrometric identification of proteins from gel spots at detection limits in the low nanogram range, similar to silver staining. Moreover, this approach does not require any post-electrophoretic manipulation by destaining and fixation, thus providing advantages for mass spectrometric analysis by reduced background and time needed for sample preparation. The use of fluorescence detection with 2-D gel electrophoresis suggests that this technique can be developed for automated, high- throughput technologies of proteome analysis. Thus, the stain-free fluorescence visualization should be a useful complement to staining techniques of gel electrophoresis for mass spectrometric protein analysis.
2.2 Application of mass spectrometry for identification of post-mortem protein changes of porcine skeletal muscle proteins
2.2.1 Muscle protein changes during post-mortem storage
Multiple factors, including palatability, water-holding capacity, color, nutritional value and safety, determine meat quality. There is a large variation in both the rate and extent of post-mortem tenderization of meat, and this result in the inconsistency of meat tenderness found at the consumer level. It has been known for a long time that meat tenderness improves during cooler storage, and it was suggested almost a century ago that this is due to enzymatic activity [102].
The three factors that determine meat tenderness are background toughness, the toughening phase and the tenderization phase. While the toughening and tenderization phases take place during the post-mortem storage period, background toughness exists at the time of slaughter and does not change during the storage period. The background toughness of meat is defined as ‘‘the resistance to shearing of the unshortened muscle’’ [103], and variation in the background toughness is due to the connective tissue component of muscle. Muscle is made up of many myofibrils. This aspect of the myofibrils is due to the presence of two types of thick and thin filaments which have an order, in that they overlap, forming a repetitive configuration of bands with identical characteristics.
Each myofibril therefore contains a repetitive series of dark and clear bands.
The wide bands of proteins of the muscular fiber, designated A (anisotropic) bands, contain a clear central area, an H zone, which in turn presents a dense M line, while clear bands, called I (isotropic) bands, are each divided in half by a Z line. The distance between two Z lines is known as a sarcomere. Immediately after slaughtering the very well-organized muscle structure begins to desintegrate. A variety of studies have shown that weakening of the myofibers is the key event in tenderization. The most consistently reported ultrastructural change associated with tenderization is breaking at the junction of the I band and Z-disk [104] (Figure 2.11).
