Anal. Chem. 1997, 69, 29R-57R
Separation and Analysis of Peptides and Proteins Andreas F. R. Hu 1 hmer, Gabi I. Aced, Melissa D. Perkins, R. Neslihan Gu 1 rsoy, D. S. Seetharama Jois, 1 neich* Cynthia Larive,† Teruna J. Siahaan, and Christian Scho
Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, Kansas 66047 Review Contents Liquid Chromatography Separation of Proteins and Aggregates by Size Exclusion Chromatography Structural Characteristics of Peptides Probed by RP-HPLC Separation and Analysis of Synthetic Peptides Analysis of Proteins and Peptides in Biological Matrices Peptide Mapping Miniaturization of Equipment and Micromanipulation of Peptides Protein Analysis Using Hyphenated Techniques Separation of Isoenzymes Separation and Analysis of Membrane-Bound Proteins Preparative Chromatography Isolation of Proteins from Tissue and Cultured Cells Purification of Genetically Modified Proteins Large-Scale Purifications Purification of Viruses Mass Spectrometry Sequencing Location of Cysteine and Cystine Residues in Proteins Locating Sites of Protein Phosphorylation Glycoproteins, Glycopeptides, and Carbohydrates Epitope Mapping Interactions between Proteins and Proteins, Oligonucleotides, and Metals Analysis of Cells and Bacteria Fourier Transform Infrared (FT-IR) Spectroscopy Nuclear Magnetic Resonance Structure Determination Pulsed-Field Gradient Methods Combinatorial Chemistry HPLC/NMR Protein-Ligand Binding and Dynamics Protein Hydration Analysis Solid-State NMR Circular Dichroism Nonenzymatic Posttranslational Modifications of Proteins Chemical Modification of Asn and Asp Protein-Associated Carbonyls, Low-Density Lipoprotein (LDL), and Advanced Glycation End Products (AGEs) Protein Modification by Nitric Oxide and Nitric Oxide-Derived Species Literature Cited
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This review covers applications of separation and characterization of peptides and proteins by liquid chromatography (LC), mass spectrometry (MS), Fourier transform infrared (FT-IR) spectros†
Department of Chemistry, University of Kansas, Lawrence, KS 66045.
S0003-2700(97)00003-6 CCC: $14.00
© 1997 American Chemical Society
copy, circular dichroism (CD), and NMR, mainly published between October 1994 and October 1996. This is by no means a comprehensive coverage of all publications but rather a selection of articles that we feel show important new strategies and applications which may be useful to improve existing methodologies. New developments in instrumentation and theory are not covered in this article, and the reader is referred to separate reviews that are published biannually in this journal. Many articles reviewed by us reflect the trend of converging separation techniques and detection methodologies that give some kind of structural information. Especially after the introduction of new ionization techniques in the field of MS, many developed protocols contain some form of chromatographic or electrophoretic separation combined with the detection by MS. To assist the reader in finding a useful solution for a separation problem in a particular area, we structured the review to consider a specific analytical problem and its solution rather than dividing it into applications of individual separation modes. In the part covering preparative chromatography, we included purification procedures employing fast protein liquid chromatography (FPLC) since the increased stability of the soft chromatographic support materials used in FPLC now allows separation speeds similar to high-perfomance liquid chromatography (HPLC). As we approach the post-genome area, the progress in sequencing of the human genome will create an overwhelming amount of information. As the entire human genome sequence becomes available, the characterization of the encoded proteins and their physiological functions in the organism demand the development of rapid methods to separate and analyze newly expressed proteins and peptides. New approaches based on hyphenated methods with reliable performance at the laboratory bench, and progress in the development of new chromatographic support materials, should also be major contributing factors. Advances in other areas, such as computing and database accessibility, will support this progress. This is particularly obvious in the mass spectrometric identification of protein digests, and the characterization of proteins based on only a few identified peptide fragments. Indicators of such an effort are parallel search algorithms against multiple protein sequences and the partial identification of fragments unique for a particular protease that can be unambiously identified by computer on-line searches of respective databases (A1). An important task is not only the identification of proteins and confirmation of their sequences but also the characterization of posttranslational modifications. Enzymatic modifications such as phosphorylation or glycosylation are mainly dealt with in the chapter on mass spectrometry. Nonenzymatic alterations such as hydrolytic and oxidative modifications are dealt with in a separate chapter at the end of this article. Analytical Chemistry, Vol. 69, No. 12, June 15, 1997 29R
LIQUID CHROMATOGRAPHY The separation and analysis of proteins and peptides by HPLC continues to be the method of choice. The well-established principles and methods of polypeptide separation by HPLC will evolve as an important tool for every molecular biology laboratory. A short introduction into the various modes of HPLC that may be useful for a molecular biology laboratory is given by Sheehan, discussing in particular the practical use of anion-exchange chromatography (A2). For a general introduction into reversedphase (RP)-HPLC, the reader is referred to an article by Neville (A3). Standard procedures of eluent preparation and standard RP-chromatographic conditions for the chromatography of peptides are discussed in a short overview by Nirenberg (A4). Other topics on the analysis of proteins by chromatography, such as hydrophobic interaction chromatography (HIC) (A5) or RP-HPLC (A6) are available in the same compendium. Separation of Proteins and Aggregates by Size Exclusion Chromatography. Due to the importance of size exclusion chromatography (SEC) for the determination and estimation of protein size and conformation, an extented chapter in the Handbook of Size Exclusion Chromatography is dedicated to this issue (A7). Size exclusion HPLC on silica-based columns was applied to probe the interaction of molecular chaperones with their substrate molecules and to characterize the complexes formed between chaperones and other molecules (A8). Basic fibroblast growth factor (bFGF) and its multimers formed by exposure to air or to dithiobisnitrobenzoic acid (DTNB) were characterized by SEC in combination with multiangle laser light scattering (MALLS) (A9). This form of analysis provided the absolute molecular mass and the mean square radius for each eluted aggregate. The results indicated the formation of rodlike rigid structures that are formed by the sequential binding of bFGF monomers through disulfide bond formation. A series of short amphiphilic helical peptides was designed to investigate the components necessary for the formation of helical bundles, and the aggregation state of the different multimer complexes was characterized by SEC (A10). A single fraction, isolated by SEC, of a synthetic Alzheimer β-amyloid peptide analog, β22-35, eluted as a series of oligomers when further analyzed by RP-HPLC (A11). The existence of this peptide as an octamer in solution was proposed. It is commonplace that at high concentations polypeptides tend to aggregate in their unfolded state. Productive folding of proteins must, therefore, take place at low concentrations of polypeptides. A new SEC-based method that improves refolding yields and simultaneously concentrates the refolded proteins in a single step was described by Batas and Chaudhuri (A12). Reduced diffusion of proteins in gel filtration media suppressed the nonspecific interaction of partially folded proteins, thereby decreasing aggregation. Applying the method to hen egg white lysozyme and bovine carbonic anhydrase, buffer exchange and elimination of aggregation due to an enhanced productive folding reaction was performed in a single step. Rissler and Engelmann described the purification of 125I-labeled insulin by SEC. Reaction side products were removed by means of a gel filtration step. The homogeneity of the fraction containing only pure labled insulin was confirmed by RP-HPLC analysis (A13). The rapid characterization of recombinant proteins derived from the genome project is often not paralleled by an analysis of the biological activity of these proteins in their natural enviroment. During the isolation of proteins from tissue, cofactors and natural inhibitors 30R
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do not always copurify and may get lost. SEC can be used to probe the size of cofactors or inhibitors required for the specific activity or inhibition of a protein. Rapid fractionation of a crude mixture can be obtained often without loss of protein activity while the specific activity of the desired protein in the separated fractions can conveniently be tested. By SEC on Superose 12, Krick et al. determined that two factors are responbible for the inhibition of the chloride ion channel from the cytosol of human placenta. A low molecular weight inhibitor was separated whereas a high molecular weight inhibitor coeluted with the protein (A14). The careful monitoring of the specific activity of calcineurin during its purification by gel filtration from crude brain supernatant revealed that a small heat-stable factor can modulate the in situ activity of calcineurin. Wang et al. identified the factor as the protein superoxide dismutase that couples the Ca2+-dependent regulation of cellular processes to the redox state of the Fe-Zn center in calcineurin (A15). Similarly, a new form of the substance P (SP)-hydrolyzing enzyme from the human cerebrospinal fluid was discovered by Eriksson et al. (A16). Using ion-exchange chromatography and SEC, the specific hydrolytic activity was monitored for each fraction in order to locate the endoproteinase activity that yields SP-7 fragments. Various aspects of the application of SEC to the analysis of denaturation and unfolding of globular proteins were reviewed by Uversky (A17). SEC can be used not only to obtain denaturation curves but also to quantitatively estimate molecular dimensions of different confirmational states. The term gel permeation chromatography is a synonym for SEC describing the quantitative aspect of that technique in the analysis of proteins (A18). The fractionation of higher molecular weight complexes of the human serine/threonine protein kinase Mst1 by gel filtration served to analyze the importance of the dimerization domain of Mst1 (A19). Sarkar and DasGupta used SEC to measure the Stoke’s radius of a guanidinium hydrochloride (GdnHCl)-unfolded intermediate of the RTEM-β lactamase of Escherichia coli. The kinetics and the energetics of unfolding were studied by CD and fluorescence spectroscopy (A20). Chang et al. described the use of size exclusion HPLC for the analysis of recombinant bovine somatotropin (rbST) monomer and dimer from bulk drug substance (A21). The heat-induced formation of a dimer from recombinant human erythropoietin (rHuEPO) through an intermolecular disulfide linkage was shown by isolation of the high molecular fraction by gel filtration and subsequent peptide mapping (A22). SEC combined with three different detection modes, on-line light scattering, UV absorbance, and refractive index, is a reliable method to probe the stoichiometry of protein complexes and to identify protein-protein interactions (A23). This approach allows the molecular weight measurement independent of the elution volume and can exclude the contributions from existing carbohydrate moieties. Using this approach, it was shown that brainderived neurotrophic factor (BDNF) and neurotrophin 3 (NT-3) dimerize their respective soluble receptors, STrkB and sTrklC, respectively, and that an anti-sHer2 monoclonal antibody dimerizes sHer2 (A24). SEC methods for the characterization of the stoichiometries and energetics of protein-protein association equilibria were extensively reviewed by Beckett and Nenortas (A25). Structural Characteristics of Peptides Probed by RP-HPLC. The separation of protein molecules on RP columns can be accompanied by a variety of changes of the molecules, depending
on the separation conditions such as column temperature, chain length of the stationary phase, and residence time of the solute on the column. Such conformational effects can lead to inhomogeneities during the chromatography. When gramicidin S analogs containing 6-14 residues were analyzed by RP-HPLC (A26), the formation of a cyclic conformer resulted in the appearance of double peaks in the chromatogram. An additional D-Leu residue preceding an ornithine (Orn) residue caused a moderate stabilization of the observed conformer that was separated by RP-HPLC. Chromatographic conditions can affect the secondary structure of proteins and promote the formation of dynamically interconverting conformers. Changes in secondary structure during the elution process can be monitored by the combination of HPLC with FT-IR detection where quantitative and semiquantitative measurements of the major amide bands can facilitate the assessment of the secondary structure content of an eluting protein. In particular for lysozyme, partial structural transitions from R-helix to β-sheet occurred when the mobile phase contained a high content of an organic modifier (A27). Often, but not always, minor alterations caused by the chromatographic conditions are reversible upon removal of the organic modifier. Changes in secondary structure induced by binding to the RP support can be utilized to monitor conformational properties of synthetic peptides and their analogs. For example, the differences in retention times of two 18-mer model peptides and neuropeptide Y analogs can be plotted as a function of methionine or methionine sulfoxide residues at various positions in the peptides. The locations of amphipathic helical structures were determined in the respective peptides (A28). Replacements in nonamphipathic R-helical domains caused local preferential binding areas and led to sequence-dependent retention time profiles. Specific interactions between the leucine zipper domains of Max and c-Myc were investigated by RP-HPLC. Upon heterodimerization of Max and c-Myc to form the basic helix-loop-helix leucine zipper motif (b-HLH-zip), DNA can bind to the complex and perform its oncogenic activity. The dimerization behavior of the synthesized leucine zipper domains was studied using RP-HPLC and CD spectroscopy. Putative buried electrostatic interactions between a His residue in Max and two glutamic acid side chains in the c-Myc at the heterodimer interface may be the main features of interaction (A29). Purcell et al. studied the chromatographic properties of four different insulin isoforms as a function of temperature (A30). The results demonstrated that, in addition to the conformational changes induced by the chromatographic RP support, the temperature can further enhance such changes and assist in the separation through an increase in selectivity. Despite a very high degree of sequence homology of the four different insulin isoforms, a significant selectivity change could be induced when the temperature was varied. A differential contribution of chain A and chain B of insulin to the retention behavior under several chromatographic conditions could be explained as a result of the progressive unfolding of the protein in the presence of the hydrocarbon ligands of the chromatographic support (A31). Conformational changes of the dimeric transforming growth factor b3 (TGF-b3) on a RP column, induced by a change of chromatographic conditions (especially temperature), were described by Lambert and Stamper (A32). Sometimes the fast interconversion of protein conformers at higher temperatures can mask the presence of additional conformers in solution. However, low temperatures can be adjusted to increase the
selectivity during the separation process. Rotationally hindered peptide conformers like proline containing β-casomorphins could be separated at low temperatures by isocratic RP-HPLC elution (A33). The order of retention with the trans conformer eluting before the cis conformer was unambiguously established when the fractions were isolated and subsequently identified by NMR at -15 °C. Separation and Analysis of Synthetic Peptides. In recent years the demand for synthetic peptides for research and pharmaceutical applications has grown extensively. Solid-phase peptide synthesis (SPPS) combined with HPLC for the purification and analysis of the products has been the method of choice. RPHPLC separation and purification of the peptides with aqueous mobile phases containing acetonitrile (ACN) and trifluoroacetic acid (TFA) are common standard procedures. A high selectivity of the RP column is critical for the isolation of the product from the frequently structurally closely related impurities. An extensive review of the chromatographic properties of 21 synthetic low molecular weight peptides and protein fragments is available from Millet and Rivier (A34). The development of analytical and preparative HPLC over the last 20 years and its application in the isolation of natural products and their synthetic constructs is reviewed in a historical context. The effect of different chromatographic conditions on the purification and analysis of triple-helical peptides (THPs or “minicollagens”) was investigated by Fields et al. (A35). The best resolution of THPs from impurities was achieved with diphenyl or nonporous C18 reversed phases using water-acetonitrile gradients, reflecting the conditions under which the native conformations of the collagen-like THPs are most likely conserved. The absolute purity of synthetically produced peptides is of great concern, especially when they are used in biological or pharmaceutical studies. Complementary selectivities of separations conducted with RP-HPLC and capillary electrophoresis can assist in monitoring the intermediate and final products. Janaky et al. achieved a more complete control of their synthetic work when separations by conventional RP-HPLC were complemented with electrophoretic separation methods (A36). Chemiluminescent nitrogen detection as a new method for the direct quantification of a crude synthetic peptide without the need for an analytical standard was reported by Bizanek et al. (A37). The method described was applied to the purification and analysis of the proinsulin chain C peptide. The purification of a poorly soluble polypeptide or the application of a crude preparation from SPPS can be facilitated when the material is preadsorbed on an appropriate preparation-grade chromatography adsorbent. The support material with the adsorbed peptide can be dry-packed into a guard column cartridge which is connected to a semipreparative HPLC column. The product can be selectively eluted with a shallow gradient avoiding the precipitation of insoluble materials (A38). Short cartridge-type columns were also used in a reversedphase liquid chromatography purification procedure described by Cooley and Meyer (A39). Purification methods were developed that exactly matched the type of cartridge that was selected for the individual peptide purification. This alternative procedure can substitute standard peptide purification by LC and accelerate peptide sythesis. Conditions for a single-step purification of synthetic gramicidin A and other tryptophan-substituted analogs was achieved using RP-HPLC with water-methanol-based mobile phases. The position of a given Trp f Phe substitution had no influence on the peptide retention, but more than one Trp f Phe Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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substitution required the addition of 2-propanol to the mobile phase for complete elution of the peptides (A40). The separation of isomers of synthetic peptides may require the modification of the standard separation conditions based on aqueous ACN and TFA. For example, an aqueous methanol-based mobile phase was necessary to achieve the baseline resolution of 23 epimeric pairs of protected tripeptides of the type LLL/LDL-Z-Ala-Xaa-Val -OMe (A41). In another method, the separation of enantiomeric and diastereoisomeric dipeptides was achieved by complexation of the peptides on an Amex Prepsil column or by separation on a conventional RP column when coordinated as ternary Co(III) complexes (A42). A rapid and accurate protocol for the detection of chiral purity of each amino acid in a purified pentapeptide, thymopentin, was described by Goodlett et al. (A43). The racemic peptide mixture was hydrolyzed in a mixture of deuterated hydrochloric acid and acetic acid and subsequently derivatized with Marfey’s reagent (1-fluoro-2,4-dinitrophenyl-5-L-alanine amide) to yield the corresponding diastereomers. During hydrolysis in deuterated solvent, racemizing amino acids were labeled with deuterium at the R-carbon.The racemic mixture of D-/L-amino acids could be separated by conventional RP-HPLC, and the subtile changes in mass were detected by on-line electrospray ionization (ESI)-MS. The use of strong cation-exchange (SCX) chromatography for the general analysis of peptides and for the separation of N-terminal-blocked peptides was discussed by Crimmins (A44). Especially for the analysis of disulfide-linked homo- and heterodimers, this technique was far superior as compared to standard RP-HPLC. Analysis of Proteins and Peptides in Biological Matrices. The development of multidimensional separations has been mostly driven by separation of complex mixtures of structurally related proteins and by the need for the characterization of small amounts of samples in complex matrices. Hemorphin-related peptides isolated from the filter membranes of uremic patients that were subjected to hemofiltration were analyzed using various chromatographic modes and conditions subsequent to acidic extraction of the peptides from the dialysis membranes (A45). Characterization of an opioid-active sequence was achieved by N-terminal sequencing and ESI-MS. A far less time consuming method for the quantitative determination of hemorphins and other naturally occurring bioactive peptides was reported by Zhao et al. (A46). Based on diode array detection, second-order-derivatized spectra were obtained for every peak, permitting the identification of aromatic amino acid residues in peptides even in complex biological matrices. It was demonstrated that the nondestructive quantitative detection of bioactive hemorphins in a total bovine hemoglobin hydrolysate was feasible without further sample preparation. The human serum R1-acid glycoprotein (AGP) was separated and analyzed for its carbohydrate content with the goal to use AGP as a tumor marker. The procedure employed an improved sample preparation method based on solvent extraction. Ion-exchange chromatography and RP-HPLC were applied to detect differences in the AGP levels in the serum of cancer patients (A47). This method allowed the detection of the differences in the composition of the fucose content of the carbohydrate moiety in AGP, which was established as a possible marker for malignant disease. A complication in the analysis of peptides arises when the desired component and its metabolic fragments are present in a complex biological matrix such as blood or cerebrospinal fluid. A strategy for the successful isolation and characterization of 32R
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biologically derived MHC class I peptides was outlined by Tomlinson et al. (A48). The presence of several thousand peptides at subfemtomolar concentrations in a mixture represented a substantial analytical challenge. After immunoaffinity concentration of the MHC class I peptides, RP separation of the mixture yielded two fractions of either hydrophilic or hydrophobic character. These coarse fractions were loaded onto a specially designed membrane preconcentration capillary and were subsequently separated according to their charge/mass ratio by capillary electrophoresis (CE). A 60 fmol sample of a peptide was ultimately characterized by sequencing by CE-MS/MS. Frequently, only a limited amount of biological sample is available. On-line trace enrichment on a cation-exchange column with subsequent gradient elution with ACN and voltammetric electrochemical (EC) detection enabled Kilts et al. to resolve the tridecapeptide neurotensin and six of its fragments with a detection limit of 1-5 ng of peptide per injection (A49). The development of general strategies to identify and characterize peptides in tissues and body fluids may involve the combination of HPLC and a radioimmunoassay (RIA). The combination of these techniques takes advantage of the sensitivity of the RIA and the resolution of the HPLC separation. Smith and Lew demonstrated the application of this technique to the characterization of peptide-processing pathways, such as the process of expression and secretion of peptides influenced by physiological regulators (A50). A method for the resolution of 17 opiate or “anti-opiate” and other synthetic neuropeptides was described by Partilla et al. (A51). A single RP-HPLC run resolved most of the peptides such as enkephalins, dynorphins, and β-endorphins, and a second HPLC separation was only needed to distinguish among the different forms of dynorphins. Using the described sample preparation, this method can be extended for the recovery and analysis of a wide variety of neuropeptides from body liquids. The determination of SP from human nasal lavage fluid was described by Schultz et al. (A52). Several femtomoles of SP per milliliter of fluid could be reliably measured after RP-HPLC separation. The analysis of cecropin B from biological fluids was described by Moore et al. (A53). Cecropin B existed in a protein-bound form and could only be analyzed after the dissociation from the proteins by citric acid. The lytic peptide cecropin B was recovered by ultrafiltration and subsequently analyzed by RP-HPLC. Peptide Mapping. Peptide mapping has become the predominant form of protein characterization in research and the pharmaceutical industry. A review by Hoff and Chloupek focuses on tryptic peptide mapping illustrating the principles and strengths of this technique (A54). Subtleties of peptide mapping were extensively review by Ling et al. (A55), considering sample preparation as well as detection limits. Complex mixtures may result in poor resolution during initial separation steps. The simultaneous alteration of temperature and gradient conditions is a convenient and effective means to achieve higher selectivity and band spacing. A series of protein digests, including those of tissue plasminogen activator, cereal storage protein, calmodulin (A56), and recombinant human growth hormone (A57), were investigated in this regard. Furthermore, computer simulations of separation patterns of complex mixtures often facilitate the optimization of separation strategies and are helpful in interpreting achieved separations (A56). HPLC at elevated temperatures is often overlooked but undoubtly an important variable that increases the speed and selectivity of protein separations. The
instrumental aspects of exact temperature control as well as applications for the ultrafast separation of proteins were reviewed by Ooms (A58). Peptide mapping can also assist in the characterization of the phosphorylation state and sites of protein isozymes. First the peptides are separated by RP-HPLC according to their hydrophobicity and the appropriate fractions are collected. Then the phosphorylated peptide fragments in the fraction are easily distinguished from the unphosphorylated peptide by capillary zone electrophoresis (CZE) through their charge differences, caused by the additional phosphate group(s). The complimentary resolving power of both methods was applied to the R-chymotrypsin digest of swine pepsin. It was further demonstrated that this approach was applicable to obtain the ratio of phosphorylated and dephosphorylated forms of pepsin isozymes in human gastric cancer patients (A59). A fully automated method for proteolytic mapping of proteins was described by Nadler et al. (A60). Reduction of cysteine thiols with reducing agents and subsequent alkylation was achieved by a computer-controlled autosampler routine prior to the digestion. Immobilized enzyme columns coupled in tandem with the analytical RPLC column generated the protein digests and reproducibly separated the obtained fragments. The system was successfully applied to proteins, such as bovine insulin, horse heart cytochrome c, human serum albumin, and ribonuclease A, containing up to seven disulfide bridges. The unequivocal assignment of disulfide bonds in proteins is the most difficult challenge faced after the primary sequence of a novel peptide is obtained. Often, peptide mapping under nonreducing conditions gives the desired information. The differences in reactivity of several disulfide bonds toward a flourescent alkylating reagent were exploited by a gradual alkylation of Cys residues in the thyroid-stimulating hormone β-subunit (A61). The development of capillary HPLC allows the characterization of small quantities of proteins. In contrast to the conventional wide-bore columns, capillary HPLC offers increased mass sensitivity at lower flow rates. A detailed description of a procedure that improves conventional HPLC systems including the fabrication and operation of slurry-packed capillary columns was described by Mortiz et al. (A62). The chromatographic performance of these columns was applied to a series of standard proteins and a V8 protease digest of the recombinant murine interleukin-6. An extension of the capillary technique was investigated as part of a multidimensional HPLC system. The sequential coupling of columns was applied to the isolation of four proteins from fetal calf serum (A63). The rapid separation within less than 10 min on conventional silica-based RP packings supported the characterization of peptide maps of phosphorylase b, obtained from colecteral cancer cell lines subsequent to twodimensional electrophoretic separation (A64). The limit of conventional RP separation is determined by the slow masstransfer kinetics of the solute during the separation process. Nonporous particles can reduce the band-broadening effect introduced by the slow diffusion into the pores and facilitate faster equilibration. Optimization strategies for the ultrafast separation of proteins on RP supports have been outlined by Hanson et al. (A65). Perfusion media (flow-through particles) allow the mobile phase to penetrate the particles and overcome the problems of stagnant mobile phase transfer without loss of separation speed and column capacity (A66). Porous microspherical reversedphase silica materials (2 µm) were tested for the fast HPLC of peptides and proteins. The support material facilitated the fast
separation of polypeptide mixtures with very high resolutions in a relatively short time (A67). In particular, the excellent recovery of late-eluting hydrophobic and large proteins such as ovalbumin or apoferritin demonstrates superior characteristics as compared to conventional materials due to the reduced residence time of the solutes in the stationary phase (A68). Rapid separation of the major bovine whey proteins by perfusion HPLC was investigated by Torre et al. (A69). The application of this technique to R-lactalbumin, serum albumin, and the genetic variants of β-LG A and β-LG B allowed the quantification of these proteins within 1.5 min. Miniaturization of Equipment and Micromanipulation of Peptides. Small sample amounts usually permit only few sample handling steps without substantial loss of the analyte. For a general introduction into the principles and the use of micropreparative HPLC for the purification of microgram or nanogram quantities of proteins and peptides, the reader is referred to a comprehensive overview by Nice (A70). Considerations for the successful micromanipulation of proteins and peptides and their recovery in small eluent volumes at high concentrations for the subsequent use with highly specific and analytical techniques such as amino acid analysis, MS, or analysis by biosensors are discussed. The treatment of a number of proteins, specifically growth factors and their receptors, are outlined in detail. A review of many modern microchemical approaches to the purification, identification, and primary structural analyses of peptides and proteins is given by Inglis et al. (A71). The discussion covers not only two-dimensional polyacrylamide gel electrophoresis (PAGE), microbore, and capillary HPLC but also combinations of these with MS and automated amino acid sequencers. The characterization and quantification of labile aromatic amino acids in proteins and peptides during amino acid analysis constitutes a problem commonly overcome by various extended procedures. Recent developments of high-resolution photodiode array detectors allows the simultanous detection of the ratios of three aromatic residues by derivative UV spectrophotometry during RPHPLC analysis of peptides in real time with an error of less than 5% (A72). Due to the power and the ease of use of gel electrophoresis, many combinations of gel electrophoresis with subsequent protein digestion and RP-HPLC analysis of the peptide maps have been described. The critical step involves the transfer of separated proteins and peptides from the gel to the HPLC column. Several strategies have been suggested. FernandezPatron et al. described the electrophoretic transfer of reverse stained gels onto a RP support in a self-made minicartridge. The minicartridge was then connected to HPLC, and the proteins eluted free of gel contaminations for further characterization (A73). Similarly, bands of electrophoretically separated proteins were excised and the gel pieces were minced individually into micrometer-sized particles. After in situ digestion, the slurry was transferred into a funnel and the peptides were transferred onto a minicolumn, filled with RP-material, by application of nitrogen pressure. Separation and analysis of the peptides was then achieved under standard conditions with the minicolumn connected to the analytical column (A74). Mincing of the gel before the in situ digestion usually allows a better access of the protease(s) to the proteins and ensures complete digestion. The reduced amounts of sample handling in this case allowed automation of large parts of the protocol, leading to an increased peptide recovery. Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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Protein Analysis Using Hyphenated Techniques. Off-line peak collection from enzymatically digested proteins for subsequent MS analysis often suffers from low sample recovery. Peptide mapping can be significantly improved by on-line coupling to MS detectors. Poor performance of the LC-ESI-MS caused by signal suppression effects of strong acid modifiers in the LC mobile phase such as TFA has been addressed and improvements for the reliable performance of LC-MS have been sugggested (A75). An extensive review of the coupling of LC to ESI-MS, in particular focusing on the analysis of recombinant human glycoproteins, was written by Guzzetta and Hancock (A76). Posttranslational glycosylation of Chinese hamster ovarian cell (CHO)derived interleukin-4 (IL-4) was analyzed by Tsarbopoulos et al. using a combination of enzymatic deglycosylation of IL-4 and proteolytic digestion of IL-4 followed by on-line HPLC-MS characterization (A77). Sugar ion diagnostics subsequent to chromatographic separation were amended by a high orificeinduced fragmentation which allowed the characterization of glycopeptides coeluting with other peptide fragments. Chromatographic modes employing nonvolatile mobile phases are in general not compatible with MS detection. For more details on the analysis of glycoproteins see the section on Mass Spectrometry (vide infra). A volatile buffer system allowed the use of a cationexchange step for the separation and elution of reaction products of substance P(1-7) (A78). This new mode of LC should allow the addition of a second dimension to conventional RPLC-MS. Employing this new separation strategy, the investigators studied the susceptibility of synthetic peptides to peptidases in body fluids (A79). SEC was connected to ESI-MS that enabled selective detection of dynorphin A, dynorphin B, and R-neoendorphin and their respective proteolytic fragments. By monitoring the rate of peptide conversion, the resistance of these prodynorphin-derived brain peptides to proteolytic cleavage was monitored. Using similar chromatographic conditions with volatile buffers, this strategy should also be useful in the identification of other peptidyl components from body fluids. In this context, the reader may enjoy a personal recollection by Desiderio (A80) of the historical events of the development of mass spectrometry, HPLC-MS and MS/MS for the analysis of brain peptides. An LC-MS assay for the analysis of enzymatically degraded human β-endorphin from the biological matrix of endothelial cells was developed by Brudel et al. (A81). The incubation of β-endorphin with cultured endothelial cells yielded 10 fragments that were prepurified and enriched using a small RP perfusion column followed by RPLCMS/MS analysis. Separation of Isoenzymes. The characterization of posttranslational modifications, e.g., such as formed under conditions of oxidative stress, requires the analysis of proteins that may be structurally very similar. A clear separation of structural isomers of proteins and peptides, as well as their purification to homogeneity, is an absolute necessity before any analysis of their particular biological characteristics can begin. Separation strategies for such isoforms should consider the heterogeneities that differentiate these polypeptides such as charge differences, differences in hydrophobicity, and preferential affinities. Recently, the characterization of histones has received much attention. The biological function of the histones is closely related to the extent of posttranslational acetylation. Separation of the histone variants is therefore critical in the elucidation of their biological function. Hydrophilic interaction chromatography (HILIC) was introduced 34R
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by Lindner et al. to separate various variants of the histone class H2A and H4 proteins from butyrate-treated Friend erythroleukemic cells (A82). The different classes of the proteins were first fractionated using RP-HPLC. Selective elution of hyperacetylated histone H4 and H2A variants was achieved from a weak cationexchange column using an increasing sodium perchlorate gradient in the presence of 70% acetonitrile at pH 3.0. The purity of the collected fractions of the monoacetylated and multiply acetylated forms of the histone proteins was determined by established methods using CE and acid-urea-Triton gel electrophoresis (AUT). Three variants of the histone H1 from porcine thymus are different in amino acid composition. Their separation was easily achieved by RP-HPLC. As part of an assay system, this separation allowed the measurement of the annealing feature of the individual variants (A83). Variants of the β-amyloid peptide Aβ1-40 present in brains of Alzheimer’s patients have been difficult to purify due to their close structural similarity. A combination of SEC and a novel high resolution separation by RPLC resulted in homogenous fractions of pure Aβ1-40 variants (A84). A variety of chromatographic modes in combination with other separation techniques must often be applied to achieve the desired separation. Human myelin basic protein (HMBP) occurs in multiple forms. A procedure that separates three isoforms was described by Deibler et al. (A85). During purification by ion-exchange chromatography (IEX), component 1 (HMBP-18.5 kDa) and a fraction containing monophosphorylated and deamidated component 3 were obtained. The latter could be further separated into two fractions by cation-exchange FPLC into a fraction containing HMBP-17.2 kDa and the posttranslationally modified forms of component 3. Limited thrombic digestion of the second fraction yielded a homogenous monophosphorylated, uncleaved, HMBP component 3 and the digestively cleaved, unphosphorylated component 3. The pure monophosphorylated form of component 3 was further separated from the mixture by SEC. Recombinant proteins retrieved from mammalian cell culture systems sometimes display unexpected charge heterogeneities This heterogeneities can be a result of C-terminal truncations of Arg or Lys residues by basic carboxypeptidase and can easily be detected by IEX. Case studies for the identification of such variants have been reviewed by Harris (A86). Microheterogeneities in glycoproteins can be caused by charge differences in the attached oligosaccaride moiety. Human transferrin isoforms, containing sialic acid in addition to oligosaccharides, can be separated by pellicular anion-exchange chromatography at high pH. Rohrer and Avdalovic used pulsed amperometric detection to reveal that protein retention is directly related to the sialylated oligosaccharide, with longer retained fractions having a greater sialylated oligosaccharide content (A87). An alternative method for the separation by SEC of isoenzymes and similarly sized proteins from a potato homogenate was discussed by Ovalle (A88). To sort the proteins on the basis of pI rather than molecular weight, a zwitterionic buffer was used. Different hydrodynamic properties of multiple forms of circulating chromogranin A (CgA) allowed the analysis of three different forms of CgA polypeptides from pheochromocytoma patients by SEC. The results suggested that CgA consists of at least three forms of polypeptides with different conformations and shapes but with similar molecular weight (A89). Selectivity in the separation of closely related peptides may also be enhanced by the right choice of organic modifier. The separation of the closely related peptides human angiotensins
I and III was investigated as a function of mobile-phase additive in an isocratic elution from porous RP columns (A90). The differences in selectivity and resolution observed with HCl and phosphoric acid can be attributed to the differences in the ion pair association between the ionized peptides and the counterions in the mobile phase. In contrast, the selectivity in the peptide separation with TFA as the modifier was a function of pH at increasing concentrations of the additive. Separation and Analysis of Membrane-Bound Proteins. In general, the purification of membrane-bound proteins is more problematic than the purification of water-soluble proteins. The isolation of biologically active membrane-bound proteins can be successful if short purification times and few delipidating steps are involved. As a general introduction to this field the Guide to Membrane Protein Purification by von Jagow and Schaegger is highly recommended (A91). The choice of a fast protein liquid chromatography step in the purification protocol can be an asset for the viability of the final purified product. Ion-exchange perfusion chromatography allows such a rapid isolation. Previously introduced for soluble proteins, Roobol-Boza and Andersson now described this technique for the isolation of the oxygenevolving photosystem II core complexes and the photosystem II reaction center particles from the thylakoid membranes of spinach chloroplasts (A92). The use of perfusion chromatography in the isolation of other hydrophobic membrane proteins is also discussed. The solubilization problems and the formation of membrane aggregates require high detergent concentrations during the sample preparation and purification. Conventional purification strategies, such as chromatofocusing, ion-exchange, and hydrophobic interaction chromatography (HIC) methods usually employ detergent concentrations beyond their critical micelle concentration (cmc) to isolate membrane-bound proteins. Conventional methods were unsuccessful for the purification of the membranebound proteinase from Bacillus cereus compromised by an irreversible binding of the lipoprotein to the chromatographic support. However, Frick et al. described the utilization of a tentacle ion exchanger for the isolation of the membrane-bound proteinase that effectively reduced the hydrophobic interacions between the proteinase and the matrix (A93). Casein-cleaving membrane proteinase (CCMP) and insulin-cleaving membrane proteinase (ICMP) could be eluted with a 2-propanol gradient, leaving the latter irreversibly denatured. Membrane-bound proteins are often reconstituted into their respective environment in vesicles by dialysis. Reconstitution yields an inhomogeneous vesicle preparation with different biological activities. Na+,K+ATPase was labeled quantitatively with a fluoresence dye, and protein-containing vesicles were separated by ion-exchange chromatography. The respective lipid and protein content of each fraction was characterized by fluorescence detection during the elution process from the column (A94). This protocol allowed the investigation of the process of vesicle reconstitution by dialysis under different conditions. A combination of HPLC and CE was used to characterize the human red cell glucose transporter (Glut1). This transmembrane protein was purified by ionexchange chromatography in the presence of nonionic detergent and subsequently dialyzed to remove excess ionic strength. The detergent concentration was increased and carrier ampholytes were added for the focusing in a cellulose-coated glass capillary. Phosphorus analysis of the electrophoretically separated zones revealed that at pH 8 Glut1 existed as a monomer whereas the
more acidic zones represented oligomers of the transporter protein (A95). The results exemplify that capillary isoelectric focusing of hydrophobic membrane proteins is possible. A general discription of SEC for the analysis of lipoproteins differing in size was given by Barter (A96). Many membrane proteins display their biological activity in molecular aggregates of interacting counterparts. The analysis of these aggregates remains difficult since intermolecular complexes dissociate and artificially aggregate or recombine during the isolation procedure. A new approach to characterize such protein aggregates was described by Loester et al. (A97). SEC under nondenaturing conditions combined with the chemical cross-linking of the proteins and SEC under denaturing conditions represents a methodolgy that determines the composition of molecular aggregates. The method was applied to R1β1-integrin dipeptidyl aminopeptidase IV as well as the cell adhesion molecule 105. Preparative chromatographic techniques that are useful in the isolation and purification of lipoproteins are reviewed by Tadey and Purdy (A98). The SEC separation of lipoproteins according to their size was discussed but, in general, found less useful because of lower specificities as compared to the separation by affinity chromatography. Far superior to these methods, however, is the use of RP-HPLC, which offers the best performance in terms of speed and resolution of structural variants. RP-HPLC purified macrophage-stimulating material (MDHM) from Mycoplasma fermentans, analyzed by Muehlradt et al. (A99), revealed that the macrophage-stimulating activity might reside in the lipopeptide moiety of a lipoprotein after proteinase K treatment and RP-HPLC analysis. A 110000-fold purification of the membrane-associated phosphatidylinositol phosphate 5-kinase from sheep brain was achieved by a procedure that uses several chromatographic steps. After extraction of the membrane with 1 M sodium chloride and ammonium sulfate fractionation, the specific binding characteristics of the proteins to phosphocellulose were exploited to separate the active fraction. The sequential separation of the fractions by hydroxyapatite chromatography, heparin Sepharose, polymer cation exchange and gel filtration finally gave the pure product (A100). Minuth et al. described a successful coupling of selective extraction techniques with conventional anionexchange chromatography (A101). Cholesterol oxidase from Nocardia rhodochrous was first isolated by solubilizing the integral membrane protein with mixed nonionic surfactant. After a temperature-induced phase transition of the unclarified broth of N. rhodochrous, a liquid phase suitable for injection onto an ionexchange column was generated. 2-Methylpropan-1-ol was employed to recycle the surfactant from a coazervate (surfactantenriched) phase used in the initial extraction step. This method gave a 160-fold-purified enyzme suitable for analytical applications with 80% yield. The isolation protocol should be useful on a bigger scale due to the resource-friendly recycling of the detergent. Conditions used for the isolation of exchangable apolipoproteins, particularly apoA-I, and quantitation of the protein using analytical RP-HPLC columns were described by Anantharamaiah and Garber (A102). A general approach for the isolation and characterization of large quantities of hydrophobic outer membrane proteins was described by Lee et al. (A103). The procedure was applied to bacterial membrane proteins and utilized precipitation from the detergent solubilizing phase with 90% ethanol before the protein, resolubilized in formic acid, was separated by RP-HPLC. An alternative to the standard precipitation assay for the analysis of Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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total cholesterol (TC) was described by Haginaka et al. based on DEAD-glucomannan gel chromatography (A104). The method described the complete separation of high-, low-, and very lowdensity lipoproteins from human plasma coupled with a derivatization procedure for fluorometric detection. PREPARATIVE CHROMATOGRAPHY Isolation of Proteins from Tissue and Cultured Cells. Isolation of novel proteins for subsequent characterization usually requires a several thousandfold purification. The rapid isolation of proteins with high yields from natural sources or cultured cells but without interfering contaminating activities can be easily achieved by preparative chromatography. Multiple chromatographic steps are often necessary, beginning with a mode of separation that separates the bulk impurities from the desired active fraction. A final, highly specific and selective chromatography step such as affinity chromatography or RPLC under mild conditions will yield the pure product. A rapid two-step procedure for the isolation of milligram quantities of CD26/dipeptidyl peptidase IV was reported by De Meester et al. (A105). This procedure yields CD26/dipeptidyl peptidase IV from human tissue without contaminating amino peptidase activity and can also be used for the purification of the CD26 from other mammalian sources. A combination of HP-SEC and subsequent HP-immobilized cholesterol affinity chromatography was the method of choice to recover highly active cholesterol-esterifying enzymes from rat liver cytosol (A106). Mortensen et al. reported a twostep FPLC purification procedure for the isolation of the E. coli translation termination factor RF-3. Utilization of an anionexchange step at the beginning of the isolation protocol allowed the high enrichment of crude mixtures which could be further fractionated by strong cation-exchange chromatography to yield 20 mg of protein/L medium of high specific activity (A107). The purification of the antigen for the mouse monoclonal antibody 703D4, the heterogeneous nuclear ribonucleoprotein-A2 (hnRNPA2), was described (A108). Purification steps included ionexchange chromatography and preparative isoelectric focusing. A minor co-purifying immunoreactive protein (hnRNP-B1), a splice variant of hnRNP-A2, was successfully removed in a final RP-HPLC step on a C4 column. A form of a prion protein-induced brain disorder, spongiform encephalopathies, is hard to study unless bigger quantities of the presumed pathogene can be isolated and characterized. Pergami et al. published a procedure that should enable researchers to elucidate the molecular basis of the pathogenesis of the prion disease. The semipreparative chromatographic procedure allows the purification of a normal, noninfectious, cellular isoform of prion protein (PrPC) from hamster brain. PrPC was solubilized by the nonionic detergent n-octyl β-glucopyranoside from synaptosomal and microsomal membranes and separated on a cation-exchange HPLC column (A109). Sequential elution from an immobilized Co(II) ion-affinity column followed by wheat germ agglutinin-affinity chromatography or SEC yielded a fraction containing a 95% pure monomeric form of PrPC. PrPC was in its native conformation, as determined by unfolding in guanidinium hydrochloride and SEC characterization. The isolation of the activity-dependent neurotrophic factor protein by sequential chromatography on ion-exchange, SEC, and HIC columns led to the characterization of a novel 14 kDa molecule which is a close homolog to the intracellular heat shock protein 60 (hsp60). A particular 14-amino acid peptide sequence 36R
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from this novel protein was synthesized and shown to exhibit neuroprotection at femtomolar concentrations in neuronal cells (A110). A recombinant form of the human neu differentiation factor was isolated from E. coli cultures (A111). Characterization of the molecule by peptide mapping and sequencing revealed that the bacterially produced recombinant product was properly folded and contained the correct disulfide structure. Very hydrophobic proteins and peptides can be easily retained on reversed-phase material and the hydrophilic portion of a crude preparation can be eluted. Subsequent purification steps involve more specific interactions such as ion-exchange or affinity columns. In this way an oviposition-inducing peptide was isolated for functional studies (A112). Similarly, recombinant soluble human E-selectin (rshEselectin), expressed in CHO cells, became available through a single RP-HPLC purification step at a purity of >95% (A113). In contrast to the conventionally purified material that had high mannose components with variable content, rshE-selectin separated by RP-HPLC reproducibly had a uniform carbohydrate composition. The above described procedure should also be amenable to other mammalian selectins such as P-selectin and L-selectin. Proteins may show specific structural properties such as, for example, hydrophobic binding pockets, that can be utilized to assist in the purification procedures. Proteases usually contain hydrophobic sites that allow the application of HIC and affinity chromatography on immobilized dye ligands. A review of several protocols for the purification of proteases with these specific support media is given by Ibrahim-Granet and Bertrand (A114). Some emphasis is placed on the separation of proteases with cyclic peptide antibiotics as general affinity ligands. Similarly, due to their mild hydrophic nature, the purification of hydroxysteroid dehyrogenases is very efficient with HIC. A review by Lin et al. shows that conventional HIC steps can be improved when adenylyl-containing affinity supports are employed in the isolation procedure (A115). High yields of pure and biologically active calcineurin, (calmodulin-dependent phosphatase 2B) can be obtained by a three-step purification scheme suggested by Nairn et al. (A116). The separation of calmodulin (CaM) from all other CaM-binding proteins is achieved by ion-exchange chromatography in a first purification step. Quantitative retention of the CaMdependent phosphodiesterase by Affi-gel blue chromatography allows complete separation from the otherwise chromatographically similarly behaving calcineurin. A final CaM-affinity chromatography step yields the pure product calcineurin at nearly 100% homogeneity. This simple scheme can be further extended to obtain other CaM-dependent enzymes. Slight net charge differences in exogenously expressed (foreign) and endogenous tobacco calmodulin from a prepurified HIC fraction were recognized by Oh et al. (A117) on the basis of anion-exchange chromatography. The principles and the application of HIC in the preparation of proteins in general and the purification of aspartate aminotransferase from Thermus aquaticus in particular were dicussed by O’Farell (A118). A protocol for the purification of three basic proteins, M1, M2A, and M2B from yellow mustard seeds that are substrates for the plant Ca2+-dependent protein kinase was devised by Neumann et al. (A119). The purification involved batchwise chromatography on CM-cellulose, cationexchange HPLC, and RP-HPLC. Identifcation of the novel proteins through Edmann sequencing identified these proteins as γ-thionins, which are potent antifungal factors. An alternative purification procedure for the isolation of the myelin basic protein (MBP)
was outlined by Sedzik et al. (A120). The mild conditions of the procedure using an ion-exchange continuous polymer bed in the presence of ethylene glycol and salt allowed the purification of MBP in its native state. Poly(ethylene glycol)-modified Sepharose was successfully employed to recover large amounts of lipase from Chromobacterium viscosum with 80% of lipolytic activity (A121). An alternative HIC support was even more successful in the isolation of the lipase. Using an epoxy-activated spacer arm, the influence of mobile-phase composition on the chromatographic behavior was investigated, and conditions were established that provided higher recovery of the lipase lipolytic activities, i.e., >90% from the bacterium (A122). Usually a salt gradient is employed for the elution of the desired components from an ion-exchange column, separating proteins according to their pI. In a report by Mhatre, a pH gradient for the elution of antibodies from cationexchange resins was used instead (A123). This purification method allowed the separation of Fab fragments and other proteins that showed differences in the isoelectric points as small as 0.1. Purification of Genetically Modified Proteins. By means of molecular biology techniques it is possible to build specific recognition sites into a desired recombinant polypeptide. This approach is especially useful if the resulting product would otherwise show no or little specific retention characteristics in subsequent purification steps. A strategy frequently employed is the His-tag technique. This approach exploits the specific affinity of a poly(His)-containing “tail” at the polypeptide to immobilized metal affinity supports, a technique called immobilized metal affinity chromatography (IMAC). Several proteins were sucessfully isolated by this technique, such as the polypeptide of the trigger factor of E. coli carrying the cis/trans isomerase catalytic activity (A124). Sydow et al. employed the histidine-tag technique with subsequent IMAC to purifiy the N-methyl-D-aspartate (NMDA) receptor subunit 1b, overexpressed in insect cells (A125). The potential use of genetically fused poly(histidine) tags on lactate dehydrogenase, β-glucuronidase, galactose dehydrogenase, and protein A for the simultaneous use with affinity precipitation, IMAC purification, and site-specific immobilization was extensively investigated by Carlsson et al. (A126). The β-amyloid peptide Aβ1-42, responsible for the plaques observed in Alzheimer’s disease, has an extreme tendency to aggregate and form insoluble fibers. This peptide is, therefore, not amenable for purification by HPLC in its monomeric form. A genetically altered β-amyloid peptide containing a His-tag tail facilitated the solubilization and purification of the monomeric peptide. The peptide was subsequently immobilized on a RP column, and the His-tag tail was specifically cleaved with CNBr without altering the peptide itself. The Aβ1-42 peptide monomer purified with this method was aggregation competent (A127). Sometimes the isolation of proteins from the cell culture medium is assisted by the generation of a fusion protein which can be subsequently cleaved. This is especially useful if the host itself contains similar proteins that could otherwise not be distinguised. The human insulin-like growth factor I (IGF-I) was modified to contain a truncated β-galactosidase “tail” linked with a sequence that could be specifically cleaved by NH2OH. After collection of the inclusion bodies and cleavage of the fusion protein by NH2OH, the IGF-I was purified by cation-exchange chromatography, refolded, and purified to homogeneity by RP-HPLC. The IGF-I was indistinguishable from the native IGF-I. These results suggest that the
expression and simple purification of hIGF-I may be possible in large-scale production (A128). An HPLC assay for the quantitative determination of recombinant human interleukin-11 fusion protein (rhIL-11 FP) from E. coli lysate utilized two-dimensional chromatography. SEC allowed the initial removal of high and low molecular mass impurities, and the final quantitative determination of rhIL-11 Fp in the lysate was done by RP-HPLC. The rapid recovery of the product from the lysate assisted in the optimization of the sample preparation conditions. The assay was linear over the range between 0.0294 and 0.235 mg of protein/mL (A129). Large-Scale Purifications. Large-scale purification is the final production step in the manufacturing of therapeutic proteins. The high standards for product quality provide a considerable challenge to any large-scale purification scheme. Selective separation of a therapeutic protein from relatively impure starting materials can reduce the number of separation steps and promote the fast concentration at a large scale. The isolation of thymosin β4 and thymosin β9 on a large scale uses a chromatofocusing-based method that allows processing of several kilograms of bovine tissue. A solid-phase extraction step through a LiChroprep RP18 material was included before chromatofocusing on PBE 94modified Sepharose gave the final pure product (A130). Protein purification for therapeutic use in humans puts extrastringent rules on the isolation procedure. A method for a highly purified, functional, and stable preparation of activated protein C (APC) from cryoprecipitation-poor human plasma was established by Orthner et al. (A131). The report described the process and the performance of a pilot plant. The process consists of three chromatographic steps and an enzymic conversion of an intermediate to yield the product APC. Metal chelation affinity chromatography on copper-charged chelating Sepharose was evaluated as a mild procedure for the purification of factor IX from human plasma on a large scale. No degradation of protein during the procedure was detected while unwanted proteins and detergent reagent, included to inactivate lipid-enveloped viruses, were removed (A132). The ability to carry out the simultaneous concentration and purification of the protein in a single step has a significant advantage for the downstream processing of pharmaceuticals. The economical savings obtained with such an efficient approach are obvious. However, the major obstacle for the implementation of the displacement method has been the lack of suitable displacer compounds (A133). Jayaraman et al. discovered that low molecular weight dendritic polymers can be successfully employed as displacers for protein purification in ionexchange chromatography and replace high molecular weight polyelectrolytes (A134). Even very strongly retained proteins such as lysozyme could be readily displaced when another form of displacer was employed (A135). Low molecular weight antibiotics such as neomycin B and streptomycin A showed high efficacy in the displacement of proteins in cation-exchange chromatography. To overcome the restriction of relatively low flow rates imposed in the preparative use of displacement chromatography, a perfusive support was selected for the rapid ion-exchange diplacement of β-lactoglobulin. Displacement of β-lactoglobulin and separation into β-lactoglobulins A and B was achieved in 90 s on an analytical scale under heavily overloaded conditions yielding 18 mg of pure β-lactoglobulin A and B (A136). The recovery of polypeptides on a large scale has limitations not encountered in a laboratory preparative scale. Every step during the purification procedure entails expensive equipment and is cost Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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intensive. The use of a Streamline DEAE material enabled the direct application of osmotically lysed E. coli on an expanded Streamline column without prior clarification of the cell extract. Purification of modified Pseudomonas aeruginosa exotoxin A from several kilograms of E. coli was feasable with a higher and more concentrated recovery than with conventional methods (A137). The recovery of the purified product from the collected chromatography fraction can be as time consuming as the separation itself. The recovery of insoluble proteins and aggregates from brain tissue homogenates (e.g., prion proteins) by aqueous twophase partition (ATPS) circumvents the labor-intensive conventional ultracentrifugation. Walker et al. described the use of poly(ethylene glycol) (PEG) and its manipulation to couple the ATPS with conventional HIC used in the final purification step (A138). A simple method to reduce the amount of volume after the purification that may also be useful in a conventional laboratory was described by Gu et al. (A139). Removal of the majority of ACN from the effluent of RP-HPLC separated proteins such as human growth hormone (hGH) was achieved without any further chromatographic step by lowering of the temperature of the ACN/ water effluent to -17 °C, yielding an ACN-rich top phase and a nonfrozen hGH-containing low, highly aqueous bottom phase. This appears to be an easy and energy efficient method to remove the majority of organic solvents after a RP-HPLC separation. Purification of Viruses. Human gene therapy trials offer new challenges such as the analysis of viruses. The use of column chromatography for the purification of type 5 recombinant adenovirus (encoding p53) was investigated by Huyghe et al. (A140). Anion-exchange, HIC, and metal-chelating resins were tested, and the results suggested that use of column chromatography is an appropriate alternative to ultracentrifugation, saving considerable amount of time. MASS SPECTROMETRY MS plays an increasingly important role in the characterization of peptides and proteins. This is mainly due to soft ionization methods such as electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) which enable the ionization of large intact or fragmented biopolymers with high efficiencies. A comprehensive review of all publications dealing with the MS of peptides and/or proteins would be impossible. We have attempted to focus on some specific areas and, largely, on the characterization of posttranslational modifications. Many reviews have been published on various applications of MS as well as the coupling of MS to separation systems. Interested readers are referred to current reviews dealing with the following topics: recent advances in mass spectrometry (B1B4), liquid chromatography/mass spectrometry (B5-B8), capillary electrophoresis (CE)/mass spectrometry (B9-B11), ESI-MS (B12-B14), and MALDI-time of flight (TOF) MS (B15-B17). Sequencing. Amino acid sequence information is crucial for the characterization of novel peptides and proteins, synthetic products, and peptides isolated from protein digests. MS has become a powerful tool for the sequencing of small amounts of peptides and proteins (B18-B20). Chait et al. (B21) developed a new method, referred to as “protein ladder sequencing”, for the N-terminal sequencing of peptides that utilizes multiple steps of partial Edman degradation chemistry prior to the analysis of the reaction mixture by MALDI-TOF MS. This method permitted the location of a phosphoserine residue in a phosphopeptide. Practical 38R
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aspects of combined Edman chemical/MALDI-TOF MS peptide microsequencing methodologies, including the requirements for and strategies of sample preparation, were described by Tempst et al. (B22). C-Terminal sequence information is of interest for the investigation of N-terminal-blocked peptides and proteins. The sequencing of substance P, glucagon, angiotensinogen, the insulin B chain, and myoglobin was possible by enzymatic digestion with the exopeptidases carboxypeptidase Y and P in combination with monitoring of the C-terminal peptide ladder by MALDI-TOF MS (B23). Because of identical molecular weights it was not possible to differentiate between the pairs of leucine/isoleucine and lysine/ glutamine. Ramsey et al. developed a method for the differentiation of Leu and Ile in peptides by means of the negative-ion mass spectra following the [M - H]- ions of phenylthiohydantoin derivatives of Leu and Ile. The spectrum of the Leu derivative showed a major loss of propane, while that of the Ile derivative showed elimination of both methane and ethane (B24). A C-terminal digestion procedure with carboxypeptidase Y involving the use of microliter wells machined into a MALDI plate for the concentration-dependent digestion of peptides was developed by Patterson et al. (B25). Using an on-plate concentration-dependent digestion strategy, 22 peptides of various amino acid composition, size, charge, and polarity were digested to explore the generality of the technique. Cytotoxic T cells (CTL) recognize small peptide fragments of cytoplasmic proteins bound to the major histocompatibility complex (MHC) class I on cell surfaces. Tumor antigens are processed and presented in a manner similar to viral antigens. The identification of peptides recognized by tumor-specific cytotoxic T cells would provide valuable information about their parent proteins. Woods et al. combined MALDI-TOF MS with on-slide exopeptidase digestion to identify and directly sequence a model tumor-specific peptide antigen derived from an integrated viral gene (B26). The enhanced sensitivity (femtomolar range) of this technique permitted the sequencing of specific MHC-bound peptides derived from as few as 1 × 109 cells. Further results on the sequencing of peptides presented by MHC class I and MHC class II have been reported by Gulden et al. (B27) and McCormack et al. (B28). Sequence-dependent peptide fingerprints can be rapidly obtained subsequent to partial hydrolysis of peptides with HCl followed by analysis by MALDI-TOF MS (B29). When synthetic peptides were treated with 3 M HCl for 5 min at 110 °C, amino acids can be released from the C-terminus and/or the N-terminus, sufficient for the confirmation of the identity of peptides during peptide mapping. The exposure of a polypeptide to perfluoroacyl anhydride vapor at -20 °C for 30-60 min caused sequential chemical degradation of the molecule from the C-terminus. Fast atom bombardement (FAB) MS of the resultant mixture of C-terminally truncated molecules permitted the detection of the C-terminal sequence by simple calculation of the mass differences of the molecular ions (B30). New Edman-type protein sequencing reagents such as 4-[3-(pyridinylmethylaminocarboxy)propyl]phenyl isothiocyanate have been shown to lower the detection levels to the low femtomole range using LC/electrospray mass spectrometry (B31, B32). Vladimirov et al. (B33) fractionated 40S ribosomal proteins from human placenta with RP-HPLC. Each chromatographic fraction was characterized by two-dimensional polyacrylamide gel electrophoresis (2D PAGE) and N-terminal sequencing. N-Terminal-blocked proteins were cleaved with
endoproteinase Lys-C, and suitable peptides were sequenced. They were able to identify all proteins from the 40S human ribosomal subunit in the HPLC elution profile. ESI-MS has been applied to the high-sensitivity sequencing of short peptides, but technical difficulties have prevented similar success with proteins isolated from gels. A simple and robust technique for the sequencing of proteins isolated by PAGE, using nanoelectrospray tandem mass spectrometry, was described by Wilm et al. (B34). Amounts as low as 5 ng of protein applied to the gel can be sequenced. The authors have demonstrated this method by sequencing and cloning of a protein that inhibits the proliferation of capillary endothelial cells in vitro which may have potential antiangiogenic effects on solid tumors. A silver-stained one-dimensional gel of a fraction from yeast proteins was also analyzed by nanoelectrospray tandem mass spectrometry. Silver staining allows a substantial shortening of sample preparation time and may be preferable over Coomassie Blue staining (B35). Small peptides were used to compare postsource decay (PSD) fragmentation of positive and negative peptide ions in a reflectron MALDITOF mass spectrometer (B36). Peptide cations produced predominantly fragment ions from the N-terminus, whereas anion dissociation gave primarily C-terminal fragment ions. The results suggest that cation and anion PSD yields complementary information on peptide primary sequences. Collisionally activated dissociation (CAD) studies of linear peptides were performed by MALDI-TOF MS. Helium, nitrogen, argon, or xenon was introduced into a gas cell to obtain high-energy CAD information. The fragments characteristically obtained in conventional high-energy CAD tandem mass spectrometry were observed also in these spectra. A difference between CAD spectra and postsource decay product-ion spectra has been demonstrated by Kosaka et al. (B37). Location of Cysteine and Cystine Residues in Proteins. The redox equilibrium between cysteine (Cys) and cystine (disulfide) plays an important role in the redox activation of certain proteins as well as in the stabilization of the three-dimensional structure of a protein. The chemical modification of a protein prior and subsequent to the reduction of protein disulfide bonds, followed by MS analysis, permits the quantitative determination of the number of cystine and cysteine moieties within a protein. Zaluzec et al. (B38) reported the use of a microscale derivatization procedure for protein thiols with p-hydroxymercuribenzoate (pHMB), combined with a sample adsorption onto Zetabind transfer membranes to facilitate excess reagent removal. Mass shifts observed in the MALDI-TOF mass spectra obtained before and after cysteine derivatization with pHMB permitted the determination of the number of free sulfhydryl groups. The total sulfhydryl content can be determined by disulfide reduction and subsequent derivatization. A competing reaction sometimes observed was the modification of cysteine with mercuric ion that may be present in the derivatization reagent. With time, even low concentrations of mercuric ions can displace the organomercurial reagent. Thus, the purification of pHMB to remove Hg2+ salts prior to its use was essential for a quantitative derivatization of the thiols of peptides or proteins. Wu et al. (B39) developed a simple method to characterize the number and the locations of free cysteines and cystine groups by a chemical modification with 2-nitro-5-thiocyanobenzoic acid (NTCB). This reagent was used to selectively cyanylate cysteinethiols in a protein under nonreducing conditions. The N-terminal peptide bond of the modified cysteinyl residue can be cleaved under alkaline conditions with
tris(2-carboxyethyl)phosphine hydrochloride to form an aminoterminal peptide and a series of 2-iminothiazolidine-4-carboxyl peptides. They can then be mapped by MALDI-TOF MS. With the above experiments repeated under reducing conditions, the total number of cysteine residues can be determined. The experiments were performed both in solution and on Nylon-based Zetabind membranes. A similar method for the determination of cysteine and cystine was shown by Watson et al. (B40). Picomole quantities of starting materials were reacted with NTCB and tris(carboxyethyl)phosphine, followed by MALDI-TOF MS analysis. Denslow et al. (B41) demonstrated procedures for the cleavage of proteins bound to PVDF membranes. The cleavage was performed at cysteine residues after cyanylation with NTCB, and the masses of the obtained fragments were determined by MALDITOF MS. Ming et al. (B42) employed 4-vinylpyridine and N-ethylmaleimide for the chemical modification of protein sulfhydryl groups, followed by determination of the molecular weights of the modified products by MALDI-TOF MS or ESI-MS. MS has been employed to demonstrate that protein folding may be associated with disulfide bond formation. Ruoppolo et al. (B43) studied the refolding pathway of reduced and denatured RNase A using MS strategies that allowed the identification of the formation and rearrangement of disulfide bonds during the folding process. It was shown that the formation of the disulfide bonds does not occur randomly even when reoxidation takes place under denaturing conditions. Crimmins et al. (B44) reported on the in situ MALDI-TOF MS analysis and assignment of disulfide pairs in heteropeptide dimers. Disulfide-linked synthetic heterodipeptides analyzed by MALDI-TOF MS displayed a mass of the dipeptide as well as of the individual constituent monomer peptides while similar analysis by ESI-MS resolved only the dipeptide. They concluded that the fragmentation during MALDI-TOF MS bears some resemblance to light-induced homolytic cleavage of aqueous solutions of the amino acid cystine. This approach provides a simple method for mapping disulfide bonds in proteins. Sun et al. (B45) reported several methods for the location of disulfide bonds in proteins using FAB MS, on-line microbore/capillary HPLC/ion spray MS, and MALDI-TOF MS. Locating Sites of Protein Phosphorylation. Among the proteins expressed in mammalian cells, as many as one-third are thought to be phosphorylated (B46). Mass spectrometry is becoming an important tool for the characterization of phosphorylation sites in proteins. Annan and Carr (B47) identified and sequenced phosphopeptides at the subpicomole level by MALDITOF MS in the reflectron mode. It was possible to distinguish tyrosine phosphorylation from serine and threonine phosphorylation for peptides containing a single phosphate group. Phosphopeptides were identified in the positive-ion MALDI-TOF reflector spectrum by the presence of [MH - H3PO4]+ and [MH - HPO3]+ fragment ions. PSD was shown to be a viable technique for sequencing phosphopeptides. Carr et al. (B48) decribed a new ESI-MS procedure that enabled the selective detection and sequencing of peptides phosphorylated at Ser, Thr, and Tyr, present in protein digests at the low femtomole level. Huddleston et al. (B49) used HPLC/MS techniques which take advantages of ion-source collision-induced dissociation (CID) to produce phosphate-specific marker ions. The primary structure of rat protein kinase C βII was probed by HPLC on-line coupled to ESIMS and by high-energy CID analysis to identify in vivo phosphorylation sites (B50). Four phosphopeptides were identified: the Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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major phosphorylation site (>95%) in the sequences Thr500-Lys520 and Glu490-Lys520 was Thr500; His636-Arg649 was phosphorylated at Thr641, and Asn650-Lys672 at Thr660. Watts et al. (B51) applied microbore HPLC-ESI MS to the identification of in vivo sites of tyrosine phosphorylation on the lymphocyte-specific protein tyrosine kinase ZAP-70 and the T-cell receptor ζ (TCR) subunit, following stimulation of T cell via their TCR. The approach is applicable to the investigation of signal transduction pathways of peptides recovered from two-dimensional phosphopeptide maps. A technique that employs a protein tyrosine phosphatase enzyme microreactor coupled on-line to either CE or HPLC and ESI-MS was used for the determination of tyrosine-phosphorylated peptides (B52). Phosphocysteine was identified by ESI-MS combined with Edman degradation (B53). Sulfated peptides could be distinguished from phosphorylated peptides on the basis of their negative ion MALDI-TOF spectra, as described by Talbo and Mann (B54). Glycoproteins, Glycopeptides, and Carbohydrates. Mass spectrometry plays an important role in the structural characterization and analysis of glycoproteins. The information is especially useful in the pharmaceutical industry, where knowledge of the structure is necessary to make cost-effective, safe, and effective drugs. Recently MS of intact plasma proteins has been used as a diagnostic tool in the diabetes field by determining the extent of glycan oxidation (B55, B56). ESI and MALDI-TOF MS are both widely used tools for the characterization of glycoproteins. With MALDI-TOF MS there is some metastable fragmentation and adduct formation when reflectron instruments are used. This may be largely due to the matrixes used. Matrixes have been shown to influence the degree of fragmentation in large glycoproteins (B57, B58). Karas et al. (B58) demonstrated that 3-hydroxypicolinic acid was highly superior to other matrixes for the MALDI-TOF MS analysis of proteins. To characterize glycoproteins, the glycoprotein is first chemically or enzymatically cleaved into glycopeptides and characterized using a myriad of analytical tools such as HPLC, CE, and MS to determine the site, extent, or heterogeneity of glycosylation. If needed for identification, the amino acid sequence of each glycopeptide/peptide can be determined, for example, by MS analysis and Edman degradation. In addition, the carbohydrates (oligosaccharides or glycans) can be proteolytically released from the protein or peptide and mapped using MS and/or NMR. Recent reviews (B59-B61) discuss the analysis of RP-HPLC peptide maps of recombinant human glycoproteins using MS. Apffel et al. (B62) demonstrated the identification of N-linked glycosylation patterns in Desmodul salvary plasminogen activator using HPLC, ESI-MS, and MALDITOF MS in combination with selective enzymatic modifications. They also described the characterization of the Desmodul salvary plasminogen activator by CE, ESI-MS, and MALDI-MS (B63). The development of CE-ESI conditions for the characterization of protein glycoforms was reported by Kelly et al. (B64). The glycoforms were separated by CE using acidic buffers and Polybrene-coated capillaries. Greis et al. (B65) and Hunter and Games (B66) used novel MS methods to rapidly locate sites in heterogenic glycoprotein preparations. CID was used to generate and detect ions from the carbohydrate moieties. Multiple sequential fraction collection of peptides and glycopeptides by CE under applied high voltage was reported by Boss et al. (B67). Characterization of carbohydrates (oligosaccharides or glycans) after release from glycoproteins and purification to homogeneity 40R
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requires the determination of linkage positions, anomeric configuration, and identification of monosaccharide building blocks. Direct MS analysis without further chemical pretreatment of the glycoconjugates can give monosaccharide sequence, molecular weight, and, sometimes, glycosidic linkage positions. The sequence and linkage of individual carbohydrates usually requires the species to be treated with a series of exoglycosidases to perform digests, followed by MS analysis. Several reviews (B68B70) outline the various techniques of glycoprotein analysis by MS and the chemical pretreatments necessary for complete characterization. Several other publications have reported the characterization of the carbohydrate moieties of glycopeptides and proteins (B71-B75). When oligosaccharides are analyzed with MALDI-TOF MS, the matrix is an important parameter for achieving low detection limits and limited fragmentation. Harvey et al. (B76) examined complex oligosaccharides with MALDI-TOF MS and various matrixes and found that 2,5-dihydrooxynaphthalene-2-carboxylic acid (2,5-DHB) proved to be the most appropriate matrix. Papac et al. (B77) examined alternate matrixes and referred to 2′,4′,6′-trihydroxyacetophenone (THAP) as the preferred matrix. If individual samples are not sufficiently clean for effective MALDI-TOF MS analysis, on-the probe sample cleanup strategies are available (B78). Epitope Mapping. Antibodies can recognize linear (continuous) epitopes or nonlinear (conformational) epitopes. Zhao and Chait (B79) developed a mass spectrometric method for the rapid mapping of linear epitopes in proteins that are bound to monoclonal antibodies. An antigenic protein can be digested by a proteolytic enzyme to produce an appropriate set of peptide fragments. The peptide fragments containing the linear epitope were selected from the pool of peptide fragments by immunoprecipitation with the monoclonal antibody. The immunoprecipitated peptides were then identified with MALDI-TOF MS. The method was established for an antibody with a relatively low binding affinity (10-6 M) and should, therefore, be applicable to most antibodies that bind linear epitopes. The characterization of antibody-antigen interactions by mass spectrometry was reviewed by Zhao and Chait (B80), describing affinity-directed mass spectrometry for epitope mapping to several monoclonal antibodies with binding affinities in the range of 10-6-10-9 M. The digestion of an antigen before affinity binding can miss discontinuous epitopes and epitopes that contain enzymatic cleavage sites. Therefore, MALDI-TOF MS in combination with proteolytic protection assays was validated by application to a system with the known epitope of gastrin-releasing peptide, recognized by the anti-bombesin monoclonal antibody (B81). Parker et al. identified the functional epitope on HIV-1IIB p26 recognized by a monoclonal antibody (B82). To identify the epitope, the intact protein was affinity bound under physiological conditions to an immobilized monoclonal antibody. A combination of cleavages by different proteolytic enzymes was then performed to remove unprotected residues. The part of the antigen in contact with the antibody and protected from proteolysis was identified by MALDI-TOF MS. An important feature of this method is that the antigen remains in its native conformation so that conformational (discontinuous) epitopes can be determined. Macht et al. (B83) applied both limited proteolysis of antibody-bound antigen followed by removal of nonbound peptides, and enzymatic digest of an antigen followed by extraction of the antigenic peptides with the antibody. For the characterization of epitope structures of markers for myocar-
dial infarction, myoglobin, and troponin T, they presented a method that did not require prior immobilization of the monoclonal antibody. The separation of nonepitope peptides from antibodybound peptides was carried out by ultrafiltration. Both fractions were analyzed by MALDI-TOF MS without further purification. Interactions between Proteins and Proteins, Oligonucleotides, and Metals. MALDI-TOF MS combined with proteolytic digestion has been used to analyze digests of the kinase inhibitory domain of the cell cycle regulatory protein (p21-B) in free solution and a 1:1 complex of p21-B with cyclin-dependent kinase 2 (Cdk2) (B84). The analysis of proteolytic digests of the p21-B/Cdk2 complex, simplified by the use of both natural isotope abundance and 15N-labeled p21-B, revealed a segment of 22-36 amino acids of p21-B that was protected from trypsin cleavage, suggesting that this fragment constitutes the Cdk2 binding site of p21-B. Earlier, this method was applied to the analysis of a protein-DNA complex (B85). The results of Cheng et al. (B86) demonstrate the stoichiometric characterization by ESI-MS of protein-oligonucleotide complexes, i.e., of gene V protein from bacteriophage f1 with several oligonucleotides. Jensen et al. (B87) developed a protocol based on MALDI-TOF MS and ESI-MS that combines mass spectrometrical methods with UV light-induced photochemical cross-linking of proteins to nucleic acids to locate and identify amino acid and nucleotide residues that are covalently attached. However, MALDI-TOF MS was also used to explore nonspecific interactions between proteins and oligonucleotides (B88). The formation of noncovalent complexes was dependent on the nature of the oligonucleotide bases and the amino acid composition of the proteins. Przybylski et al. (B89) analyzed enzyme-substrate and -inhibitor complexes by negative-ion ESI MS. Ions specific of noncovalent protein and oligonucleotide complexes can be selectively dissociated by changing the solution conditions and by increasing the desolvation potential. ESI-MS opens new analytical perspectives for the direct characterization of noncovalent supramolecular complexes. MALDI-TOF mass spectra of peptide-metal ion complexes formed by a zinc finger peptide of the transcription factor IIIA (Cys2-His2) type and zinc and cobalt ions were obtained by Woods et al. (B90). Analysis of Cells and Bacteria. In recent years, the direct chemical analysis of single cells has received considerable attention. Hofstadler et al. (B91) presented results that demonstrate the feasibility of using capillary electrophoresis/electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry (CE/ESI-FTICR) for the analysis of proteins acquired directly from small populations (i.e., 5-10) of intact living cells. The human erythrocyte was chosen as a model system. The authors demonstrated the on-line acquisition of high-resolution mass spectra (average resolution >45.000 fwhm) of both R- and β-chains of hemoglobin (Hb) acquired from the injection of 10 human erythrocytes (4.5 fmol of Hb). The presented technique should be adaptable to the study of larger mammalian cell systems. Valaskovic et al. (B92) investigated 8-27 kDa proteins by CE/MS at a resolution of ∼60 000 for attomolar injections with errors of 30 kDa) with many proton resonances, even with complete solvent suppression, standard analog-to-digital converters do not have sufficient dynamic range to properly digitize the protein signals in the FID. One solution to this problem is to sample at rates faster than that required by the sampling theorem. In some cases, this method, called oversampling, can decrease the digitization noise and, therefore, increase the signal from the protein. The benefits that can be derived from oversampling in NMR have been analyzed, and guidelines to help the analyst evaluate the utility of this method for individual applications have been reported (D12). Spin diffusion can result in inaccurate interproton distance measurements, leading to distortions in protein structures calculated from NOE-derived data. Normally, a short mixing time NOESY experiment is performed to minimize the effects of spin diffusion. Unfortunately, short mixing times also restrict the Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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observation of long-range NOEs. Therefore, several techniques have been developed to overcome distance inaccuracies resulting from multispin effects, including magnetization exchange network editing (MENE) (D13), continuum approximation of relaxation pathways between dilute spins (CROWD) (D14), selective pulses (D15), and a statistical approach to eliminate the contribution of spin diffusion (D16). MENE spectra acquired with the blockdecoupled or the complementary block-decoupled NOESY (CBDNOESY) sequence (D17) give more precise input data than does the regular NOESY sequence because of the reduction of spin diffusion effects. The MENE approach was used to produce a three-dimensional structure of turkey ovomucoid in good agreement with the X-ray structure with a level of precision that could not be achieved with the standard NOESY experiment (D13). Accurate distance measurements using selective NOE measurements to avoid spin diffusion may open new avenues to evaluate the dynamic behavior of two proton pairs in proteins; however, in these studies, the effects of relaxation during the selective pulse must be considered (D15). A statistical approach has been developed to eliminate the spin diffusion contribution in calculated interproton distances to achieve a high-quality NMR structure (D16). It remains to be seen whether or not the experimental effort expended in improving the accuracy or precision of NOE constraints will lead to significant improvement in the quality of the resulting structure. Pulsed-Field Gradient Methods. Magnetic field gradients have become an indispensable tool for high-resolution NMR experiments, including applications in water suppression, measuring proton exchange, and high-quality shimming (D18). A rapid method has been developed to evaluate gradient nonlinearity; it should find widespread application since gradient linearity is a requirement for many experiments (D19). New schemes using high-resolution pulsed-field gradient methods have been developed to improve NOESY experiments (D20, D21). An interesting pulsed-field gradient method that places all of the gradient pulses before the read pulse has been reported (D21). This method, called spatial population sculpting, offers acceptable levels of artifact suppression without the signal losses normally associated with gradient-enhanced experiments. An area of NMR spectroscopy that has grown substantially in the number of new techniques and applications in recent years is the measurement of diffusion coefficients using pulsed-field gradient methods. Fourier transform pulsed gradient spin echo (FT-PGSE) is routinely used in the determination of aggregation and diffusion coefficients. The data sets measured with this experiment are often complicated with overlapped band shapes and poor signal-to-noise ratios. Typically data sets from such experiments consist of 16 or 32 different magnetic field gradient settings and 10-1000 frequencies. A new method, componentresolved (CORE) NMR spectroscopy, has been developed for processing data from FT-PGSE experiments (D22). It uses a global minimization approach to produce spectra with improved signal-to-noise ratios from the raw data. CORE-NMR is particularly useful for the study of aggregation, binding of polymers, and solutions of surfactants (D22). An improved pulse sequence for the measurement of diffusion coefficients that employs bipolar gradient pulses to reduce the effects of eddy currents has been reported (D23). Several authors have introduced diffusion as the third dimension in NMR experiments by coupling a pulsed-field gradient pulse 44R
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sequence with a standard two-dimensional pulse sequence. This approach has produced diffusion-COSY (D24), diffusion-TOCSY (D25), and diffusion-NOESY (D20) pulse sequences, all of which promise to be useful for the analysis of mixtures of peptides and proteins. The diffusion-COSY experiment was illustrated by the spectral separation of alanine, glutamine, and arginine. The diffusion-TOCSY experiment may be more useful than the diffusion-COSY experiment in future applications for the measurement of diffusion coefficients in complex samples because of the larger number of cross peaks and the absorptive in-phase line shape of TOCSY spectra (D25). The use of the diffusion-TOCSY pulse sequence for the spectral resolution and measurement of diffusion coefficients in a mixture of five esters illustrates the potential of this experiment. The diffusion-NOESY pulse sequence is less useful for mixture analysis but offers an advantage over the transferred NOE experiment for the measurement of NOEs in binding experiments since a large molecular weight difference in the molecules involved in binding is not required (D20). The principles of diffusion-ordered spectroscopy (DOSY) have been described in a recent review (D26). This is an automated method of data analysis and display which enables “at a glance” recognition of components in complex mixtures. NMR experiments incorporating diffusion provide the analyst with an extra analytical handle that can be effectively applied to the analysis of complex mixtures. For example, diffusion coefficients were used to evaluate the formation of β-cyclodextrin inclusion complexes by the cis and trans isomers of phenylalanylproline (D27). Spectral editing based on diffusion and relaxation has been applied for the simplification and assignment of the 1H NMR resonances in spectra of human plasma (D28). Although relaxation-based spectral editing methods have been employed for the detection of small-molecule analytes in biological fluids for some time, this method is unique since it permits the detection of plasma proteins by suppressing the resonances of small molecules based on their larger diffusion coefficients (D28). An important application of pulsed-field gradient experiments is the analysis of protein aggregation based on diffusion coefficients (D29, D30). The measurement of the diffusion coefficients of the monomeric globular proteins lysozyme and ubiquitin and the dimeric proteins interleukin-10 and monocyte chemotactic protein-1 demonstrates the utility of this method for the unambiguous determination of oligomerization states in the structural analysis of proteins (D29). Diffusion coefficients were used to study insulin aggregation in solutions containing a mixture of dimeric and tetrameric metal-free insulin (D30). Alternative approaches for studying protein aggregation based on the measurement of rotational correlation times have also been reported. (D31, D32) Off-resonance rotating-frame 13C NMR experiments were used to study aggregation of the bovine lens protein γ-crystallin (D31). Tumbling rates for a dimeric leucine zipper peptide have been determined using new pulse sequences employing polarization transfer for the sensitive measurement of 13C relaxation rates in AX spin systems (D32). 2 Combinatorial Chemistry. Some of the most impressive advances in NMR spectroscopy in the period since the previous review have been in the development of NMR methods that address the considerable analytical challenges of combinatorial chemistry. Detection of NMR spectra of compounds bound to solid-phase resins has been demonstrated using magic-angle spinning (MAS) to narrow the analyte resonances. The influence
of resin composition, tether length, and solvent on the quality of 1H NMR spectra that can be obtained for compounds bound to solid-phase synthesis resins has been reported (D33). The unambiguous identification of compounds bound to the beads is enhanced by heteronuclear experiments that provide information about proton-carbon connectivities (D34, D35). An impressive demonstration of the potential for these methods in the analysis of products of combinatorial chemistry is the identification of a compound bound to a single solid-phase-synthesis bead (D34). HMQC and TOCSY spectra were used to completely assign the NMR spectrum of protected lysine bound to solvent-swollen resin (D36). Similar MAS HMQC NMR experiments were employed along with MS/MS and capillary electrophoresis analysis to completely characterize a combinatorial peptide library (D37). HPLC/NMR. The high information content of the NMR spectrum and the power of this technique for structure elucidation makes NMR a potentially attractive detector for chromatographic separations. Historically, the wide-spread adoption of HPLC/NMR as an analytical tool has been hampered by difficulties that include the relatively low sensitivity of NMR, problems associated with solvent suppression, particularly in separations using gradient elution, and lack of commercially available HPLC/NMR accessories. Recent developments have made considerable progress in addressing these problems, and HPLC/NMR has been employed for a variety of applications in pharmaceutical and biomedical research. A summary of experimental challenges encountered in HPLC/NMR, recent technical developments in this field and a comprehensive range of examples of applications have been described in a recent review of this area (D38). The geometry and dimensions of the rf coil and detection cell have important ramifications for the sensitivity of the HPLC/NMR experiment and the quality of the resulting spectrum. The factors influencing the signal-to-noise ratio for a cell with a volume of 50 nL using a solenoidal rf coil design have been analyzed in detail (D39). These authors report the separation and detection of microgram quantities of several amino acids and peptides. Online detection in a 75 µm i.d. reversed-phase capillary column was reported using a 1.2 cm long saddle-type rf coil (D40). This configuration also produced a 50 nL volume detection cell. The authors report separation and on-line detection of nanomole quantities of dansyl amino acids. A simple modification to adapt 500 MHz 1H NMR probes for use in HPLC/NMR has been reported (D41). This method will be of great interest to laboratories that have an NMR instrument for which the manufacturer does not offer HPLC/NMR hardware or when the cost of a commercial probe is not justifiable. One of the greatest challenges in HPLC/NMR is the detection of the 1H resonances of small quantities of analyte in the presence of protonated solvents. Although most investigators choose to use deuterated solvents, this does not resolve the solvent interference for experiments employing gradient elution where even the 13C satellites of the organic component of the mobile phase may be more intense than the analyte resonances. An improved method of solvent suppression for one and two-dimensional HPLC/NMR experiments (WET) is exceptionally effective at suppressing the HOD and acetonitrile resonances, with a level of suppression for acetonitrile reported as >50 000 (D42). Protein-Ligand Binding and Dynamics. In addition to secondary structure determination, NMR is a useful method for the analysis of protein binding. One comprehensive review has
focused on the role of combinatorial synthesis and NMR studies in developing an improved understanding of protein-ligand interactions (D43). The importance of extended conformations for molecular recognition in peptide/protein complexes has also been reviewed (D44). In a similar vein, a recent report highlights the importance of considering protein flexibility and internal motions in structure-based design of target ligands (D45). A new method for identifying high-affinity ligands called SAR (structureactivity relationship) by NMR has been reported (D46). This method uses changes in the chemical shift measured with 2D NMR in response to addition of a ligand to an 15N-labeled protein (D46). The spectra can be obtained rapidly, making it possible to screen a large number of compounds. Chemical shifts and chemical shift nonequivalence due to folding have been shown to be useful for structure characterization rather than simply a necessary prerequisite for multidimensional NMR (D47). The dynamics of the protein backbone and side chains are important for many biological functions including receptormediated signal transduction and enzyme catalytic activity. Recently, many attempts have been made to understand the mobility of protein backbone and side chains in the presence and absence of ligands (D48-D50). For example, the backbone dynamics of chymotrypsin with inhibitor fragments (D51) and dihydrofolate reductase with folate (D48) have been studied. Because biochemical events such as turnover rates, ligand binding and dissociation rates, and the rates of conformational change often occur near the time scale of low-frequency structural motion, it is possible that these slower protein motions affect enzyme function. Multinuclear NMR has played a major role in studying internal motion of proteins in the presence and absence of ligands. 15N T1 and T2 relaxation times and {1H}-15N heteronuclear NOEs of 15N-labeled proteins were used to evaluate the dynamics of the backbone and side chains of proteins (D48, D49, D51-D53). The dynamic motion of each of the backbone amide nitrogens was determined by the order parameter (S2) and the effective correlation time for internal motion (te) (D49, D51). Motional time scales were evaluated using an effective correlation time for internal motion (te) or 15N-exchange broadening (Rex) contributions for each NH resonance. Values of S2 and te provide information on motion on the picosecond-to-nanosecond time scale and Rex provides information on motions in the microsecond-to-millisecond region (D49, D52). “Model-free” analysis was used to determine the amplitude of dynamic motion of the amine nitrogen via an order parameter (S2) reflecting the degree of motional restriction of the NH bond by back calculation of the relaxation data (T1, T2) and NOE data (D50). Because protein side chains are often involved in ligand binding, the structural dynamics of side chains in the presence and absence of ligand may be useful for evaluating the proteinligand binding mechanisms, kinetics, and selectivity. For example, interactions between the phosphotyrosine peptide (PY1021) and four arginine residues of the binding pocket of the C-terminal SH-2 domain of phospholipase C-γ (PLCC SH2) were characterized by measuring the relaxation of 15N and 13Cξ spins and the exchange rates of H nuclei of the arginine residues (D53). Quantitative information on the flip rates about the partial double bonds of arginine was obtained from the natural-abundance 1H[15N] HMQC spectra (D54). The characterization of arginine guanidinium groups is important because a motionally restricted arginine residue may be involved in a salt bridge, in binding with substrate, Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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or in maintaining a hydrogen-bonding network in the threedimensional structure. Another method to evaluate the side-chain dynamics of amino acid residues that contain methyl groups (i.e., Leu, Thr, Met) in proteins involves the use of fractionally deuterium-labeled methyl groups (CH2D) and the measurement of 2H T1 and T1F relaxation times indirectly from a series of 2H(13C-1H) constant time spectra (D50). This method has been used to analyze the dynamic motion of the side chain of the free state of PLCC SH2 domain and its complex with pY1021. The motion of the methyl group containing amino acids in the phosphotyrosine-binding region of this protein is restricted; in contrast, the hydrophobic binding site that is responsible for recognition of sequences C-terminal to the p-Tyr displays significant motional disorder (D50). Hydrogen bond interactions between side chains are frequently found in protein structures and even fairly weak interactions produce significant effects on the stability of helical regions. A method to measure these side-chain interactions in peptide helices by NMR is described and applied to glutamine-aspartate (i, i + 4) hydrogen bond interactions, important in stabilizing peptide helices in water (D55). Protein Hydration Analysis. Pulsed-gradient NMR experiments were conducted for the measurement of diffusion coefficients as an indirect indicator of protein hydration in solution, although this method does not distinguish between the bound and the free water present (D56). The ratio of water bound to macromolecules to the total amount of water has been determined for aqueous solutions of bovine serum albumin and for red blood cells in osmotically altered sizes utilizing triple-quantum-filtered 17O NMR spectroscopy and simultaneous analysis of transverse and longitudinal triple-quantum NMR experiments (D57). Time domain reflectometry (TDR) and NMR experiments were used to determine the amount of water bound to the albumin surface (D58). The results of these experiments indicated that the TDR method gave the amount of water bound to the albumin surface whereas the NMR method included the amount of water that can exchange with the tightly bound water (D58). 2D NMR spectroscopy has been utilized to determine the binding of water molecules to HIV-1 protease-DMP323 complex in solution by detecting NOE and ROE interactions between water and protein protons (D59). 1H and 13C NMR techniques as well as other rheological procedures were employed to determine the cause for the unusual rheological behavior of aqueous solutions of lysozyme in contact with some organic solvents to be the change in protein conformation due to the contact between aqueous and organic solvents (D60). Solid-State NMR. Solid-state NMR (D61) and the application of MAS (D62) have been recently reviewed. The use of MAS solid-state NMR techniques to measure distances between pairs of spin 1/2 nuclei has been described from an experimental standpoint (D62). A review of solid-state NMR studies to determine the structures of peptides and proteins in membranes has been reported (D63). The amide N-H couplings of peptide bonds in proteins are essential in studies of protein structures. In a recent study, the amide 1H chemical shift, 1H-15N dipolar coupling, and 15N chemical shift tensors in the powder pattern spectrum of a model dipeptide, Ala-[15N]Leu, were presented in a three-dimensional fashion, which provides simultaneous information about the three aforementioned parameters (D64). The 2D 1H-13C HETCOR 46R
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experiment has been shown to provide valuable information about the structure of amino acids and peptides (D65). Rotational echo double-resonance (REDOR) NMR spectroscopy has been used to determine the internal reorganization of the conformations of large proteins and to investigate the effect of serine binding at the β-subunit of tryptophan synthase (Trp-S) on the protein (D66). Compared to other solid-state NMR methods, double-quantum spectroscopy has the advantage of requiring incorporation of only one doubly 13C-labeled amino acid residue into the peptide (D67). The separated-local-field double-quantum (SELFIDOQ) NMR technique has been reported as a new method for determination of solid peptide and polypeptide structure (D67). The backbone torsion angle, ψ, is the only relevant parameter in the simulations that can be determined by correlating the CR-HR bond direction with the carbonyl tensor orientation in a 2D NMR spectrum (D67). CIRCULAR DICHROISM In recent years, CD spectroscopy has been used to (a) study the effect of an electric field on conformational changes in proteins (E1), (b) evaluate the effect of aspartate bond isomerization on the conformation of short- and medium-sized synthetic peptides (E2), (c) demonstrate the formation of R-helix polypeptide assemblies in lipid monolayers (E3), (d) compare the R-helical break by proline residues in water and in membrane environments (E4), (e) study the conformation of native and modified disulfide bonds in proteins (E5, E6), and (f) analyze CD-active drugs in urine for pharmacokinetic studies (E7). Magnetic circular dichroism spectroscopy has been used to probe the spin state, oxidation state, and axial ligand identity in histidine-ligated iron-containing proteins (E8). In this review, we will focus on recent topics related to the CD spectroscopy of peptides and proteins, including new algorithms to evaluate secondary structure, correction of solvent effects, evaluation of helices screw sense in solution and the solid state, vibrational CD (VCD) spectroscopy, time-resolved CD (TRCD), and stopped-flow CD. Many developments have focused on new algorithms to (a) obtain an accurate percentage of secondary structures, (b) differentiate transmembrane and peripheral helices, and (c) analyze turn structures in peptides and proteins. An excellent source for these methods and their applications has been presented by Fasman (E9). Secondary structural analysis using the primary sequence has been improved by incorporating CD data constraints (E10). Application of this method to an immunosuppressant binding protein indicates that this type of analysis gives a more accurate result than prediction based on the sequence alone (E10). Several new methods have been developed to analyze the secondary structures of peptides and proteins from CD data. Greenfield (E11) compares various methods including linear combination (LINCOMB), multilinear regression (MLR), convex constraint analysis (CCA), neural network program (E12), single-value decomposition (SVD), CONTIN program for rigid regression analysis, and the self-consistent method (SELCON) to obtain secondary structural information from CD data. Each method has been shown to have its own advantages and pitfalls; for example, the wavelength range of data acquisition and the precision of protein concentration determination have been shown to affect the reliability of each method. Most methods were developed by comparing the CD experimental curve with the reference protein using a least-squares fit or best fit. Unfortunately, extraction of a secondary structure using the best fit does
not necessarily give appropriate secondary structural elements for β-sheet or β-turn structures. Dalmas et al. (E13) attempted to solve this problem by constructing a basis set using a large number of reference proteins with known secondary structures. The basis set was constructed using neural network and Monte Carlo simulations. This study found that using the basis set and subtracting the contribution of the nonchromophore group prior to spectral deconvolution improves the secondary structure prediction results (E13). The protein microenvironment in solution, including solvent effects, influences secondary structural determination of peptides and proteins. The effect of the solvent dipole moment on CD spectra has been investigated by Cascio and Wallace (E14). This information is useful for the interpretation of the CD spectra of peptides and proteins in nonaqueous environments such as membranes (E14). The solvent effect is usually not considered in the secondary structural determination of peptides and proteins since the X-ray crystal structure is used as a reference. However, the change in solvent environment can cause a change in the transition energy which leads to blue or red shifts of the CD maxima (E15). These shifts contribute to a discrepancy in the secondary structure result. Therefore, Cascio and Wallace (E15) developed a Gaussian curve fit method to correct the effect of the blue or red shift caused by solvent effect. To test the reliability of this method, crambin was studied in different solvent microenvironments. This method resulted in a secondary structure of crambin close to the X-ray crystal structure, whereas other methods (i.e., CCA, SVD) underestimated the secondary structure of crambin. This method is also useful for studying membrane proteins because their native conformation is associated with a hydrophobic environment (E15). Like other methods, this method has disadvantages; it only accurately predicts R-helices in proteins and it assumes that all peptide bonds are in the same microenvironment, which in reality may not be true (E15). CD has been used to differentiate between the contribution of the 310-helix and the R-helix using the ratio of ellipticity at 222 and 208 nm (E16). Three derivatives of (L-(R-Me)Val)8 with different forms of the NR-blocking group have been shown to acquire 310-helical conformations in solution and solid states (E17). The right-handed 310-helical peptides display a negative band at 207 nm accompanied by a shoulder centered near 222 nm. The helical screw sense of these peptides can also be determined in the solid state by CD using a KBr disk (E18). A p-bromobenzamido chromophore linked to the N-terminus was used to assign the screw sense of the 310-helical peptides in the solid state (E18). The CD spectra of these peptides were dominated by the p-bromobenzamido chromophore at 238-240 nm (E18). The results from solid-state CD were in agreement with those already reported for these peptides in solution using electronic and vibrational CD (E18). Solid-state CD has advantages over X-ray in determining the screw sense of an R-helix because it does not require a single crystal and is faster than X-ray crystallography. VCD spectroscopy is an established technique for monitoring the local sensitive region of a protein, while electronic CD spectroscopy gives information about the overall structure. Athough VCD is sensitive to backbone conformation, it is not sensitive to change in the side-chain conformation. The applications of VCD spectroscopy for analysis of the secondary structure of peptides and proteins as well as nucleic acids has been reviewed (E9, E19). This method monitors the well-resolved vibrational
transition of amide bands; these bands are spectrally well resolved and very sensitive to structural changes. Therefore, protein structural changes can be monitored using these amide bands, which is more sensitive than electronic CD. Structural changes in R-lactalbumin as a function of pH and solvents that were not observed by electronic CD were monitored by VCD (E20). Structural studies by VCD spectroscopy have focused on the amide I′ band, which is due to the CdO stretch on the amide group (1670-2000 cm-1) for N-deuterated peptides. The amide II band, which results from the deformation of NH and the CN stretch (1550-1500 cm-1), is very sensitive to structural changes. Several algorithms have been developed to deconvolute and analyze VCD spectra in order to quantitatively estimate protein secondary structures; however, some of these algorithms are not sensitive enough to follow the changes in VCD spectra (E20). One algorithm was developed based on neural network analysis using amide I′ or II′ bands independently as well a combination of both bands (E21). Estimation of the secondary structure was improved using combined amide band data. Later, a more advanced method of processing VCD and FT-IR data for the prediction of protein secondary structure was developed by Baumruk et al. (E22). Because VCD is a weak phenomenon, a high concentration of protein is required for analysis which may lead to protein aggregation. Little variation in VCD spectra has been detected over a small range of protein concentrations. Conformational changes and relaxation of proteins can be studied by time resolved circular dichroism (TRCD). Photolysis of the hemoglobin-CO complex (HbCO) has been monitored by TRCD at room temperature in the near-UV spectral region (E23). The photolyzed HbCO shows a striking negative ellipticity at 258 nm due to the aromatic residues at the interdimer interfaces of the T quartenary form (E23). The appearance of this band in TRCD gives strong evidence of a rapid shift at the R1β2 interface to a more T-like conformation, with a time constant of ∼500 ns (E23). Relaxations of protein structures are more evident in TRCD than in corresponding absorption spectra. The pathway of protein folding can be determined by the relative stability of the intermediates or the relative height of the activation energy barrier leading to these intermediates (E24). Stopped-flow CD spectroscopy has been utilized to determine the folding processes of several polypeptides (E25) and to investigate the kinetics and transient intermediates of refolding proteins (E26). Induced folding and unfolding of Staphylococcal nuclease (SNase) was studied using stopped-flow CD and differential scanning microcalorimetry (DSC) (E24). The data show that the kinetics in the time range between 2 ms and 500 s are triphasic processes (E24). This supports the sequential mechanism for SNase folding: U3 T U2 T U1 T N0, where U1, U2, and U3 are states of the unfolded protein and N0 is the native state (E24). Although the U1, U2, and U3 states are nearly isoenergetic, no random walk occurs among these states during the folding; therefore, the pathway of folding is unique and sequential (E24). The folding pathway is not dictated by the relative stability of the folding intermediates. In contrast, the folding process follows an energy descent through the lowest activation energy path toward the global free-energy minimum of the native state (E24). Therefore, barrier avoidance leads to the folding path, and the height of the energy barrier limits the rate of folding (E24). In vitro experiments have shown that there are a few proteins in this size range that can refold within milliseconds or seconds after Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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unfolding, while other proteins of similar size may never be able to refold (E24). The leucine zipper peptide GCN4-p1 provides a simple model system for studying both the intramolecular and intermolecular interactions that govern protein folding by stopped-flow CD (E27). The unfolding and refolding kinetics of GNC4-p1 were monitored as a function of peptide concentrations and final denaturant concentrations (E27). The equilibrium of an unfolding curve is accurately predicted from these rate constants, providing further support for the validity of the two-state kinetic model (E27). Thermodynamic and kinetic analyses of the induced unfolding and refolding of the GCN4-p1 dimer show that this simple system is well described by a two-state model (E27). Only the native dimer and unfolded monomer are required to explain the equilibrium kinetic behavior of this peptide (E27). The absence of stable transient folding intermediates, which could provide clues to the mechanism, differentiates this simple dimeric systems from many larger proteins (E27). NONENZYMATIC POSTTRANSLATIONAL MODIFICATIONS OF PROTEINS There is a large variety of nonenzymatic pathways that lead to the covalent modification of proteins in vivo (e.g., during biological aging or conditions of oxidative stress) and in vitro (e.g., during processing and strorage). The characterization of these processes, their reaction stoichiometries and mechanisms, relies on many different experimental techniques some of which have been described in more detail in the preceding chapters. This part of the review summarizes recent work on the hydrolytic and oxidative modifications of proteins in vivo and in vitro. Chemical Modification of Asn and Asp. Asn and Asp are involved in a variety of hydrolytic/nucleophilic degradation mechanisms involving deamidation, isomerization, and peptide backbone cleavage. These reactions affect peptides and proteins in solution as well as in lyophilized forms. Chang et al. developed a stable freeze-dried formulation of recombinant human interleukin-1 receptor agonist (rhIL-1ra) in their need to avoid competitive aggregation and deamidation (F1), and the physical parameters such as glass transition temperature (Tg) and the presence of amorphous stabilizers on protein conformation and stability were evaluated (F2). A chemically stable formulation of human epidermal growth factor (hEGF) was developed at neutral pH (Tris buffer) in the presence of nonionic surfactants or polymers at low salt concentrations (F3). An important intermediate during the deamidation of Asn or the isomerization of Asp is a cyclic imide (succinimide). In general, the formation kinetics of succinimide are strongly influenced by the sequence and structure of peptides and proteins (for review, see ref F4). In recent examples, Brennan and Clarke found a significant acceleration of succinimide formation for Asp when preceded by a His residue (F5), and “hot spots” for the formation of succinimide, isoAsp, and Asp were characterized for several protein sequences such as of a recombinant antibody to human IgE, Asp32-Gly33 (D6), recombinant erythropoietin receptor (EPOr), Asn163-Gly164 (D7), and polyanionstabilized acidic fibroblast growth factor (aFGF; FGF-1), Asn8Tyr9 (D8). Recently, progress has been made toward the understanding of an apparent heterogeneity of preparations of creatine kinase. Though apparently homogenous according to sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDSPAGE), isoelectric focusing has resolved creatine kinase into three 48R
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bands. By means of high-resolution tandem mass spectrometry, these three bands were characterized as corresponding to native and potentially deamidated protein isoforms (F9). Darrington and Anderson established a catalytic effect of the deprotonated C-terminus of the A-chain of human insulin on the deamidation of Asn at position A-21 (F10). However, the intermediate succinimide reacts not only with water (deamidation) but also with the N-terminus of the B-chain, leading to covalent dimers (F11, F12). It is the ring opening of the succinimide that leads to Asp or isoAsp, respectively. Capasso et al. established a change in the rate-determining step of ring opening of the succinimide in the pH region 4-5 (F13). This has now been confirmed by isotopic labeling experiments in H218O measuring the incorporation of 18O into succinimide using an oxygen isotope effect on the chemical shift of the carbonyl 13C-resonance during 13C-NMR spectroscopy (F14). Besides ring opening, the succinimide serves as a precursor for yet another process, racemization of L-Asp to D-Asp. Recent theoretical studies have revealed that the C-H bonds of R-carbons of succinimide residues are much more acidic than those of amides, e.g., of 2-pyrrolidone, providing a basis for accelerated racemization at CR (F15). Deamidation and isoAsp formation are relevant to protein aging not only in vitro but also in vivo. In mammalian brain, a highmass methyl-accepting protein (HMAP) was characterized which displayed a 50-fold higher level of isoAsp as compared to the average protein in brain cytosol (F16). When rat PC12 cells were incubated in the presence of a methyltransferase inhibitor, there was a significant increase of the isoAsp content of tubulin as well as the formation of two types of covalent aggregates (F17). Whereas one of the aggregates was identified as disulfide the other was suggested to form via the reaction of a succinimide intermediate with Lys. In some cases, deamidation can mark proteins for accelerated turnover, investigated in detail for mammalian triosephospate isomerases (F18, F19). Interestingly, in the triosephosphate isomerase of Bacillus stearothermophilus, one Asn residue usually labile toward deamidation has been substituted by His (F20). The crystal structure of the recombinant protein shows that the Nδ2 nitrogen of the His residue is placed in the same position as the Nδ of Asn in other isoforms of the protein, maintaining hydrogen-bonding capacity without presenting a deamidation “hot spot” (F20). Thus, the enzyme is active and stable at significantly higher temperatures. The conversion of Asn to Asp or isoAsp can have significant structural consequences for proteins. For example, the crystal structure of bovine pancreatic ribonuclease in which Asn67 had been replaced by isoAsp displayed a perturbation of the H-bond network responsible for the rigidity of the protein in that region (F21). Miesbauer et al. reported an analysis of water-soluble crystallins isolated from the lens of young human adults (F22). Among other modifications, they characterized a deamidation process for Asn101 of R-Acrystallin. However, whenever in vivo deamidation sites are characterized by means of tryptic mapping, one has to be cautious not to promote deamidation by the conditions of the tryptic digestion per se (here: pH 8.2, 4 h, 37 °C). It should be noted that the formation of succinimide and deamidation is not always a biologically deleterious process. For example, protein splicing appears to rely on amino succinimide formation by an Asn residue which is located at the C-terminus of the intervening sequence (intein) (F23, F24).
Protein-Associated Carbonyls, Low-Density Lipoprotein (LDL), and Advanced Glycation End Products (AGEs). The quantitative assessment of oxidative protein modification in vivo frequently relies on the determination of protein-associated carbonyls. Several procedures have been developed among which the derivatization with 2,4-dinitrophenylhydrazine (2,4-DNPH) is probably most commonly used (F25, F26). Derivatization with 2,4-DNPH yields the corresponding hydrazones which can be monitored by UV spectroscopy (λ ) 360-370 nm), size exclusion chromatography coupled to a UV detector (or diode array detector), or Western blotting (F25, F26). It should be noted, however, that in particular for in vivo samples the spectrophotometric and the chromatographic method need to be carefully controlled for possible errors due to contamination by excess 2,4DNPH or nucleotides (F27). The loss of protein during washing steps appears to be another problem. In fact, recently Lyras et al. (F28) attempted to reproduce earlier results on elevated carbonyls as a result of motor neuron disease (F29, F30) and demonstrated great variability depending on the method used. The most specific assay for protein-associated carbonyls appears to be Western blotting using a mono- or polyclonal antibody against the 2,4-dinitrophenyl moiety of the derivatizing agent (F25). With this methodology, Shacter et al. have identified oxidized plasma proteins and established that levels of 1 pmol of protein-associated carbonyls could be detected (F31). An interesting result of these studies was the finding that fibrinogen showed a significantly higher level of protein-associated carbonyls compared to the other modified plasma proteins (F31). Western blotting of rat erythrocyte proteins revealed that R-spectrin and β-spectrin are major targets for the formation of protein-associated carbonyls (F32). Among the proteins of the soluble fraction of rat liver homogenate, one protein was particular rich in proteinassociated carbonyls, identified as carbonic anhydrase III (F33). However, high levels of carbonyls were present regardless of whether the protein was isolated from 2, 10, or 18 month old rats, indicating that in vivo aging did not correlate with an increasing accumulation of carbonyls on the protein for the investigated age groups. It was proposed that the protein functions in an oxidative environment responsible for the formation of high levels of carbonyls even on protein isolated from young animals. As the expression levels of protein decreased with aging, the authors had no possibility to examine liver homogenate of old rats (e.g., g24 months) for the levels of protein-associated carbonyls. However, many other reports have appeared showing that proteins accumulate carbonyl groups with increasing age and during acute oxidative stress (F34-F53). Protein-associated carbonyls can form via the direct chemical conversion of an amino acid side chain or a peptide (amino acid) N-terminus or via oxidative peptide cleavage (F54). The oxidation of Lys, Arg, and Pro has been shown to yield carbonyls (F54). However, in general, carbonyl formation is possible for alkyl side chain(s) of any amino acid. Suitable precursors for carbonyl formation are amino acid-bound hydroperoxides which can be reduced by electron transfer (e.g., from reduced transition metals) to yield alkoxyl radicals which subsequently suffer R-β fragmentation. A recent example has been given by Davies et al. for the reductive fragmentation of γ-hydroperoxyglutamic acid into aldehyde and •CO2- (F55). The •CO - radical anion rapidly reduces molecular oxygen to super2 oxide (F56), which can initiate further oxidation mechanisms of proteins (most likely after dismutation to hydrogen peroxide). The
exposure of amino acids and proteins to free radicals has yielded peroxides (hydroperoxides) on the amino acid side chains of Glu, Ile, Leu, Lys, Pro, and Val (F57), and especially the structures of valine hydroperoxides have been characterized (F58, F59). It was proposed that, after chemical reduction to Val hydroxides, these oxidation products of Val may serve as biological markers for protein oxidation. Alternatively to a direct chemical conversion of amino acids, lipid peroxidation products such as malondialdehyde (MDA) or 4-hydroxy-2-nonenal (HNE) may covalently attach to nucleophilic amino acid side chains of Lys, Cys, or His (for review, see ref F60), thereby attaching carbonyl groups to a protein. Whether or not the carbonyl-containing forms of such adducts can be isolated in vivo depends on their stability in particular with regard to the formation of cross-links with other available Lys residues. Antibodies to HNE-modified protein sequences have been prepared for the detection of HNE-protein adducts in biological samples (F61-F63). The interactions of lipid peroxidation products with proteins are particularly important for membrane proteins and LDL. Much evidence has been provided that oxidized/modified LDL is an atherogenic agent though the in vivo oxidation mechanisms have not been clearly established, and in vitro mechanisms may only partly apply to in vivo conditions (F64). An antibody against oxidized LDL has been prepared which specifically recognizes oxidized phosphatidylcholine (PC) (F65). The immunoreactivity was enhanced in the presence of proteins such as bovine serum albumin (BSA), suggesting that the epitopes be complexes between polypeptides and oxidized PC. More recently, the authors established that peroxide and aldehyde products of PC were particularly antigenic, probably through covalent bond formation with polypeptides such as apolipoprotein B (apoB) (F66). A sensitive assay for the quantitative analysis of oxidized LDL in humans was developed using this antibody in a sandwich ELISA method (F67). Using an HNE-specific antibody, Uchida et al. had established that oxidized LDL contains HNE adducts at Lys and His (F68). The specific His residues of apoB modified by conjugation with HNE have now been identified as His21 (or His2940), His763, His916, His2074, His3281, His3948, and/or His3960 (F69). All of these His residues reside on the surface of the LDL particle, and an effect of His modification on the interaction between modified LDL and macrophage-bound receptors has been suggested. Identification of the modified His residues was possible by tryptic digestion of oxidized apoB, followed by electrospray tandem mass spectrometry including a screening procedure for precursors of a fragment ion of m/z 266 which corresponds to an immonium ion derived from HNE-modified His. This method had recently been developed and tested for the characterization of HNE-modifed His residues of apomyoglobin (F70). Interestingly, the incubation of LDL with glucose resulted in a rapid formation of HNE and 4-hydroxyhexenal as analyzed by gas chromatography/mass spectrometry (GC/MS), indicative of lipid peroxidation (F71). The reactions of amines with aldehydes such as HNE appear to be responsible for the age-related formation of lipofuscine pigments, which show distinct fluorescence characteristics. Derivatives of 2-hydroxy-1,2-dihydropyrrol-3-one represent possible structures that may be responsible for the observed lowwavelength spectroscopic properties (e.g., excitation at 360 nm, emission at 430 nm) (F72). Fu et al. reported that lipid peroxidation causes the formation of N-(carboxymethyl)lysine (CML) on LDL (F73), quantitatively monitored, after hydrolysis of the Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
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protein, by GC/MS as trifluoroacetyl methyl ester derivative. CML is one of the few well-characterized AGEs, which are a hallmark for protein modifcation during normal biological aging as well as complications of diabetes and renal insufficiency. The interesting observation was that CML formed much faster as a result of lipid peroxidation compared to glucose oxidation, suggesting that lipid peroxidation may be a primary cause of CML formation in vivo (F73). On the other hand, other prominent AGEs such as pentosidine were not formed during lipid peroxidation, suggesting that both lipid and glucose oxidation contribute to different extents to the formation of various AGEs in vivo (F73). On the basis of immunochemical methods and microsequencing, potential sites of AGE modifications of LDL have recently been characterized as Lys residue(s) located within a 67-amino acid region between residues 1388 and 1454 of the primary sequence of apo B (F74). By means of boronic acid affinity chromatography, glycated proteins can be conveniently detected, e.g., in human atherosclerotic tissue (F75). Several antibodies to AGEs have been prepared (F76). By means of a specific monoclonal antibody, 6D12, the presence of AGEs in atherosclerotic lesions of human aorta was demonstrated (F77). The major epitope recognized by polyclonal anti-AGE antibodies (F78) as well as 6D12 (F79) is a CMLprotein adduct. We note that, besides conjugate formation, LDL can suffer various other covalent modification on apo B. For example, for Cu2+-oxidized LDL, cysteic acid and methionine sulfoxide were located within a sequence corresponding to Glu4187-Arg4195 of apo B using HPLC and FAB mass spectrometry (F80). The presence of hypochlorous acid (HOCl)-modified LDL in human lesions was demonstrated by means of a monoclonal antibody specifically raised against HOCl-modifed LDL (F81). Leeuwenburgh et al. quantified by GC/MS that levels of 3-nitrotyrosine are significantly (90-fold) higher in LDL isolated from aortic atherosclerotic intima as compared to LDL from plasma of healthy subjects (F82) (a more detailed discussion of 3-nitrotyrosine follows below). Fluorescence spectroscopic measurements revealed that the incubation of LDL with Cu2+ results in the modification of Trp which is not affected by the presence of vitamin E (F83). The gross structural changes of LDL during oxidation were continuously monitored by neutron solution scattering (F84) and synchrotron X-ray solution scattering (F85), demonstrating aggregation of the protein and a preferential oxidation of the outer shell of the surface phospholipids at early time points followed by the modification of apo B and the internal neutral lipid core. A considerable effort has been placed on the chemical characterization of AGEs as well as the specific mechanisms according to which they form in vitro and in vivo (for a review on mass spectrometrical approaches to the characterization of AGEs see ref F86). The major products of glucose oxidation, available for protein modification, were determined to be glyoxal and arabinose (F87). A new series of AGE products was identified as originating from a reaction of glyoxal (or methylglyoxal) with Lys peptides, forming imidazolysine derivatives responsible for Lys-Lys cross-links (F88-F90). The stoichiometry of this reaction was suggested as involving 2 mol of Lys peptide and 2 mol of glyoxal producing 1 mol each of imidazolysine, formic acid, and water (F88). Other identified structures include 2-acetamido6-[3-(1,2-dihydroxyethyl)-2-formyl-4-hydroxymethyl-1-pyrrolyl]hexanoic acid (formyl threosyl pyrrole), originating from the reactions of degradation products of ascorbate (F91), levuglandin E2-derived 50R
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protein-bound pyrrole (F92), and pyrraline [-2-[formyl-5-(hydroxymethyl)pyrrol-1-yl]-L-norleucine] (F93). In a mechanistic study on the formation of AGEs, preformed protein-bound Amadori products on collagen were exposed to glucose in phosphate buffer. It was shown that the major source of AGEs at high concentrations of phosphate buffer (200 mM) was the oxidizing glucose but not further conversion of the preformed Amadori products. At low, physiologically more relevant phosphate concentrations (10 mM), both glucose oxidation and further conversion of the preformed Amadori products contributed to the formation of AGEs. Experimentally, these results were achieved by incubating collagen-bound Amadori products with [13C6]glucose and monitoring the formation of [13C2]CML (F94). Similar findings on the origin of CML were reported by Glomb and Monnier (F95). However, using the rat tail tendon breaking time assay (TBT), Elgawish et al. could demonstrate that preglycation of protein or the presence of small amounts of Amadori products can catalyze glycoxidation and protein cross-linking in an oxygendependent but glucose concentration-independent manner (F96). This pathway involves hydrogen peroxide, which may form as a result of oxidation of the 2,3-enol tautomer of an Amadori product. Thus, the mechanisms of AGE formation are complex and may be further complicated by the findings of Khalifah et al., showing that high concentrations of ribose can inhibit the breakdown of Amadori products to AGEs (F97). The decomposition of a model Amadori compound, NR-formyl-N-fructoselysine (fFl), was significantly influenced by the nature of the buffer, e.g., phosphate vs N-(2-hydroxyethyl)piperazine-N′-2-ethanesulfonic acid (HEPES), and the presence of oxygen (F98). An interesting approach toward a more detailed understanding of the role(s) of Amadori products in the formation of AGEs would be to selectively remove Amadori products from incubation media. This can be achieved by Amadori product-degrading enzymes of which several have now been described though their mechanisms of action are not fully understood (F99-F101). Free-radical reactions of glyoxal (methylglyoxal) may play an important role in the formation of AGEs. For example, the formation of methylglyoxal radical anions and methylglyoxal dialkylimine radical cations were observed by means of ESR spectroscopy during the reaction of methylglyoxal with the amino acid Ala (F102). The methylglyoxal radical anions reacted further with molecular oxygen to produce superoxide, whereas the radical cations were suggested to be potential precursors for yellow fluorescent AGE products. Metal ions and molecular oxygen did not play a role in the initial electron-transfer reaction between methylglyoxal and methylglyoxal dialkylimine as the reaction proceeded anaerobically in the presence of metal chelators. Whereas glucose and its degradation products are precursors for the extracellular glycation and formation of AGEs, intracellular pathways of AGE formation are much less understood. One important precursor has been shown to be ADPribose, as it effectively forms AGEs in its reaction with histones (F103). As evident from this chapter, most of the experimental work on the structural characterization of AGEs has been performed with model compounds in vitro. However, recently Vasan et al. presented a potential concept for the elucidation of structures of glucose-derived protein cross-links which may be useful both in vitro and in vivo (F104). A synthetic reagent, N-phenacylthiazolium bromide (PTB), cleaves R-diketone groups present in cross-links, which may allow the further identification
of the obtained fragments. After treatment with PTB, collagen isolated from diabetic rats was susceptible to digestion by cyanogen bromide, comparable to untreated collagen from nondiabetic rats. Even more interestingly, the treatment of fibrillar aggregates of AGE-modified β-amyloid peptide (βAP), a hallmark of Alzheimer’s disease, resulted in some disaggregation of βAP, raising the hope for some clinical application of AGE-cleaving agents. Much research effort is currently devoted to the elucidation of the mechanisms according to which βAP is involved in the pathology of Alzheimer’s disease (F105). It appears that βAP is neurotoxic based on its ability to promote lipid peroxidation and modifications of proteins (F106-F112). Recently, it has been suggested that one pathway for the induction of cellular oxidative stress is mediated by binding of βAP to RAGE, a cellular receptor for advanced glycation end products (F113). Protein Modification by Nitric Oxide and Nitric OxideDerived Species. Nitric oxide is an important biological messenger with a great variety of biochemical functions (F114). One potential problem associated with increased levels of NO under certain physiological conditions is its diffusion-controlled reaction with superoxide which yields peroxynitrite (ONOO-), a highly reactive species which can modify proteins and lipids (F115). A specific product of the reaction of peroxynitrite with proteins is the formation of 3-nitrotyrosine, which can be monitored by various standard analytical techniques such as UV spectrophotometry, amino acid analysis (HPLC and GC/MS) (F116, F117), Western blotting (F116, F118), and peptide mapping in conjunction with mass spectrometry (F119). Based on the detection of 3-nitrotyrosine in vivo, an increasing number of papers support the hypothesis that ONOO- is involved in oxidative cell injury under conditions of oxidative stress (F82, F120-F127) as well as biological aging (F128). At this point it should be noted, however, that nitration of tyrosine may also be caused by other nitric oxide-derived nitrating species such as ClNO2 (F129). The nitration of Tyr may involve catalysis by transition metal complexes including superoxide dismutase (F118), catalysis by adduct formation with CO2 (F130-F133), and free-radical mechanisms with intermediary formation of Tyr radicals (F130, F134). In its complex chemistry, peroxynitrite may react as cis and trans isomers of its ground state and several excited states, resulting in complex kinetics and various different products with different substrates such as Tyr (F118, F130-F139), Trp (F140), Cys (F141, F142), Phe (F138), Met (F143), and selenomethionine (F144) in their free or protein-bound forms. The reactions of nitric oxide with proteins are more or less restricted to metalloproteins (F145) and to protein cysteine residues (F114). An important observation by Jia et al. (F146) was that hemoglobin cysteine residues are nitrosylated in vivo, depending on ligand binding and the spin state of the heme group (see also comments in ref F147). Mechanistically, the formation of nitrosothiol from NO and a thiol requires the participation of either transition metals (F114) or, possibly, molecular oxygen (F148). Andreas F. R. Hu 1 hmer is a third-year graduate student in the Department of Pharmaceutical Chemistry at the University of Kansas. He received his Diplom in chemistry at the Free University of Berlin, Germany. During that time he also joined the group of Dr. H. J. Mo¨ckel at the Hahn-Meitner-Institut Berlin where he gained experience in chromatography. His current research involves the purification of proteins and studies on their reactions with reactive oxygen species.
Gabi I. Aced received her Diplom in chemistry in 1985 at the University of Merseburg, Germany, and her Ph.D. in chemistry in 1989 at the Technical University of Berlin, Germany. Between 1989 and 1993 she worked for Wissenschaftliche Gera ¨ tebau Dr. Knauer GmbH before she joined the Hahn-Meitner Institut Berlin, Germany, for the time between 1993 and 1994. From 1994 to 1996 she was Research Assistant Professor at the Higuchi Biosciences Center at the University of Kansas, Lawrence, KS, and in 1997 she accepted a position as Senior Scientist at Oread Laboratories, Inc. Melissa D. Perkins is a second-year graduate student at the University of Kansas, where she is pursuing her Ph.D. in Pharmaceutical Chemistry under the direction of Professor Susan Lunte. She received her bachelor’s degree in biology and chemistry from the University of Kansas in 1992. After graduation, she worked in the Analytical Pharmaceutical Chemistry Department of Oread Laboratories, Inc. for three and a half years. Her research interests include the analysis of estrogens using CEEC and glycoprotein characterization using CE-PAD and off-line analysis by MALDI-TOF mass spectrometry. Reyhan Neslihan Gu 1 rsoy is a doctoral candidate in the department of Pharmaceutical Chemistry at the University of Kansas, Lawrence, KS. She received her B.S. degree in Pharmacy from Hacettepe University, Ankara, Turkey, in 1992. After receiving a five-year sponsorship from the Turkish Government, she joined the Pharmaceutical Chemistry Department at the University of Kansas in 1993 where she earned an M.S. degree in 1995. She is currently involved in the structural, biophysical, and biological studies of LFA-1/ICAM-1 peptides, which participate in T-cell adhesion. Dr. Seetharama Jois received his Ph.D. from the Molecular Biophysics Unit, Indian Institute of Science, Bangalore, India, in 1992 under the supervision of Professor K. R. K. Easwaran. Currently, he is working in Professor Siahaan’s laboratory at the University of Kansas as a postdoctoral scientist. Dr. Jois is involved in conformational studies of ICAM/LFA-1 peptides, which are responsible for modulating T-cell adhesion. Dr. C. L. Larive received her Ph.D. degree from the University of California, Riverside under the supervision of Dallas Rabenstein. She began her appointment as an Assistant Professor of Chemistry at the University of Kansas in 1992. Dr. Larive’s research involves bioanalytical applications of NMR spectroscopy and includes the application of NMR measurements of diffusion to study dynamic processes such as binding and aggregation, the relationship between aggregation and conformation in model peptides, and the development of improved methodology for analysis of metabolism in cell culture using in vivo NMR spectroscopy directly coupled with HPLC. Dr. T. J. Siahaan received his Ph.D. from the Department of Chemistry, University of Arizona, under the supervision of Professor Robert B. Bates and postdoctoral training at the Department of Chemistry, University of California, Santa Barbara, with Professor Bruce Lipshutz. He joined the Department of Pharmaceutical Chemistry, University of Kansas, in 1991 as an Assistant Professor. Dr. Siahaan’s research interest is in the area of modulation of cell-matrix and cell-cell adhesions by peptides and the conformational analysis of cell adhesion peptides by NMR, CD, and FTIR to elucidate their mechanisms of activity, stability, and delivery. Christian Scho1 neich is Assistant Professor at the Department of Pharmaceutical Chemistry at the University of Kansas. He received his Diplom in chemistry in 1987 at the Free University Berlin, Germany, and his Ph.D. in chemistry in 1990 at the Technical University Berlin, Germany. After two years of postdoctoral research at the Hahn-Meitner Institut Berlin, Germany, and the Department of Pharmaceutical Chemistry at the University of Kansas, he joined the Department as an Assistant Professor in 1992. His research interests include reaction mechanisms of reactive oxygen and reactive nitrogen species with peptides and proteins in vitro and in vivo.
LITERATURE CITED LIQUID CHROMATOGRAPHY (A1) Taylor, J. A.; Kennth, A.; Johnson, R. S. Rapid Commun. Mass Spectrom. 1996, 10, 679-87. (A2) Sheehan, D.; FitzGerald, R. Methods Mol. Biol. 1996, 59, 14550. (A3) Neville, B. Methods Mol. Biol. 1996, 59, 277-92. (A4) Nirenberg, U. Methods Mol. Biol. 1994, 36, 23-35. (A5) Wu, S.-L.; Karger, B. L. Methods Enzymol. 1996, 270, 27-47. (A6) Aguilar, M.-I.; Hearn, M. T. W. Methods Enzymol. 1996, 270, 3-26. (A7) Himmel, M. E.; Baker, J. O.; Mitchell, D. J. In Handbook of Size Exclusion Chromatography; Wu, C.-S., Ed.; Marcel Dekker: New York, 1995. (A8) Palleros, D. R.; Shi, L.; Reid, K.; Fink, A. In Techniques in Protein Chemistry VI; Crab, J., Ed.; Academic Press: San Diego, CA, 1995; pp 467-74. (A9) Astafieva, I. V.; Eberlein, G. A.; Wang, Y. J. J. Chromatogr., A 1996, 740, 215-29. (A10) Lutgring, R.; Lipton, M.; Chmielewski, J. A. Amino Acids 1996, 10, 295-304.
Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
51R
(A11) Vyas, S. B.; Duffy, L. K. Protein Pept. Lett. 1996, 3, 89-94. (A12) Batas, B.; Chaudhuri, J. B. Biotechnol. Bioeng. 1996, 50, 1623. (A13) Rissler, K. Engelmann, P. J. Chromatogr., B 1996, 679, 219. (A14) Krick, W.; Disser, J.; Rabe, A.; Froemter, E.; Hansen, C. P.; Roch, B.; Kunzelmann, K.; Greger, R.; Fehlhaber, H.-W.; Burckhardt, G. Cell. Physiol. Biochem. 1995, 5, 259-68. (A15) Wang, X.; Culotta, V. C.; Klee, C. B. Nature 1996, 383, 4347. (A16) Eriksson, U.; Andren, P. E.; Caprioli, R. M.; Nyberg, F. J. Chromatogr., A 1996, 743, 213-220. (A17) Uversky, V. N. Int. J. Bio-Chromatogr. 1994, 1, 103-14. (A18) Ctuler, P. Methods Mol. Biol. 1996, 59, 255-67. (A19) Creasy, C. L.; Ambrose, D. M.; Chernoff, J. J. Biol. Chem. 1996, 271, 21049-53. (A20) Sarkar, D.; DasGupta, C. Biochim. Biophys. Acta 1996, 1296, 85-94. (A21) Chang, J. P.; Ferguson, T. T.; Record, P. A.; Dickson, D. A.; Kiehl, D. E.; Kennington, A. S. J. Chromatogr., A 1996, 736, 97-104. (A22) Derby, P. L.; Strickland, T. W.; Rohde, M. F.; Stoney, K.; Rush, R. S. Int. J. Pept. Protein Res. 1996, 47, 201-8. (A23) Wen, J.; Arkawa, T.; Tsutomu, P.; John, S. Anal Biochem. 1996, 240, 155-66. (A24) Wen, J.; Arkawa, T.; Talvenheimo, J.; Welcher, A. A.; Horan, T.; Kita, Y.; Tseng, J.; Nicolson, M.; Philo, J. S. In Techniques in Protein Chemistry VII; Marshak, D. R., Ed.; Academic Press: San Diego, CA, 1995; pp 23-31. (A25) Beckett, D.; Nenortas, E. Adv. Biophys. Chem. 1995, 5, 23362. (A26) Tamaki, M.; Akabori, S.; Muramatsu, I. J. Chromatogr., A 1994, 685, 237-41. (A27) Turula, V. E.; de Haseth, J. A. Anal. Chem. 1996, 68, 62938. (A28) Rothemund, S.; Krause, E.; Beyerman, M.; Bienert, M.; Sykes, B.; Sonnichsen, F. D. Biopolymers 1996, 39, 207-19. (A29) Lavigne, P.; Kondejewski, L. H.; Houston, M. E., Jr.; Sonnichsen, F. D.; Lix, B.; Sykes, B. D.; Hodges, R. S.; Kay, C. M. J. Mol. Biol. 1995, 254, 505-20. (A30) Purcell, A. W.; Aguilar, M. I.; Hearn, M. T. W. J. Chromatogr., A 1995, 711, 249-56. (A31) Purcell, A. W.; Aguilar, M. I.; Hearn, M. T. W. J. Chromatogr., A 1995, 711, 71-9. (A32) Lambert, W. J.; Stamper, G. F. J. Chromatogr., A 1995, 709, 249-56. (A33) Kalman, A.; Thunecke, F.; Schmidt, R.; Schiller, P. W.; Horvath, C. J. Chromatogr., A 1996, 729, 155-71. (A34) Miller, C.; Rivier, J. Biopolymers 1996, 40, 265-317. (A35) Fields, C. G.; Grab, B.; Lauer, J. L.; Fields, G. B. Anal. Biochem. 1995, 231, 57-64. (A36) Janaky, T.; Szabo, E.; Balaspiri, L.; Adi, B.; Penke, B. J. Chromatogr., B 1996, 676, 7-12. (A37) Bizanek, R.; Damon, J.; Fujinari, M.; Fujinari, E. M. Pept. Res. 1996, 9, 40-4. (A38) Thompson, P. E.; Hearn, M. T. W. J. Biochem. Biophys. Methods 1995, 30, 153-61. (A39) Cooley, R. S.; Meyer, R. Isol. Purif. 1994, 2, 83-91. (A40) Salom, D.; Abad, C. J. Chromatogr., A 1996, 725, 315-22. (A41) Griehl, C.; Merkel, S. Int. J. Pept. Protein Res. 1995, 45, 21724. (A42) Aldrich-Wright, J. R.; Newman, P. D.; Rao, K. R. N.; Vagg, R. S.; Williams, P. A. J. Chromatogr., A 1995, 693, 366-70. (A43) Goodlett, D. R.; Abuaf, P. A.; Savage, P. A.; Kowalski, K. A.; Mukherjee, T. K.; Tolan, J. W.; Corkum, N.; Goldstein, G.; Crowther, J. B. J. Chromatogr., A 1995, 707, 233-44. (A44) Crimmins, D. L. Methods Mol. Biol. 1996, 36, 53-64. (A45) Nyberg, G.; Sanderson, K.; Andren, P.; Thoernwall, M.; Einarsson, M.; Danielson, B.; Nyberg, F. J. Chromatogr., A 1996, 723, 43-9. (A46) Zhao, Q.; Sannier, F.; Garreau, I.; Lecoeur, C.; Piot, J. M. J. Chromatogr., A 1996, 723, 35-41. (A47) Kremmer, T.; Boldizsar, M.; Kovacs, J.; Paulik, E.; Bencsik, K.; Szajani, B. J. Liquid. Chromatogr. 1995, 18, 1207-18. (A48) Tomlinson, A. J.; Jameson, S.; Naylor, S. J. Chromatogr., A 1996, 744, 237-78. (A49) Kilts, C. D.; Knight, D. L.; Nemeroff, C. B. Life Sci. 1996, 59, 911-20. (A50) Smith, A. I.; Lew, R. A. Methods Neurosci. 1995, 23, 125-39. (A51) Partilla J. S.; You, J.; Rothman, R. B. J. Chromatogr., B 1995, 668, 219-31. (A52) Schultz, K.-D.; Furkert, J.; O’Conner, A.; Boettcher, M.; Schmidt, M. Baumgarten, C. R.; Kunkel, G. Neuropeptides 1996, 30, 117-24. (A53) Moore, A. J.; Loadman, P. M.; Devine, D. A.; Bibby, M. C. J. Chromatogr., B 1995, 672, 225-31. (A54) Hoff, E. R.; Chloupek, R. C. Methods Enzymol. 1996, 271, 5168. (A55) Ling, V. T.; Eng, M. L.; Lee, P.-J.; Keck, R. G.; Keyt, B. A.; Canova-Davis, E. In New Methods in Peptide Mapping for the Characterization of Proteins; Hancock, W. S., Ed.; CRC: Boca Raton, FL, 1996; pp 1-30. (A56) Chloupek, R. C.; Hancock, W. S.; Marchylo, B. A.; Kirkland, J. J.; Boyes, B. E.; Snyder, L. R. J. Chromatogr., A 1994, 686, 45-59. 52R
Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
(A57) Hancock, W. S.; Chloupek, R. C.; Kirkland, J. J.; Snyder, L. R. J. Chromatogr., A 1994, 686, 31-43. (A58) Ooms, B. LC-GC 1996, 14, 306. (A59) Hynek, R.; Kasicka, V.; Kucerova, Z.; Kas, J. J. Chromatogr., B 1996, 681, 37-45. (A60) Nadler, T.; Blackburn, C.; Mark, J.; Gordon, N.; Regnier, F. E.; Vella, G. J. Chromatogr., A 1996, 743, 91-8. (A61) Fairlie, W. D.; Stanton, P. G.; Hearn, M. T. W. Biochem. J. 1996, 314, 449-55. (A62) Moritz, R. L.; Reid, G. E.; Ward, L. D.; Simpson, R. J. Methods 1994, 6, 213-26. (A63) Ronk, M.; Davies, M. T.; Lee, T. D.; Shively, J. E.; Hefta, S. A. In Techniques in Protein Chemistry V; Crabb, J. W., Ed.; Academic Press: San Diego, CA, 1994; pp 259-68. (A64) Moritz R. L.; Eddes, J.; Simpson, R. J. In Methods in Protein Structure Analysis; Atassi, M., Zouhair, A. E., Eds.; Plenum Press: New York, 1995. (A65) Hanson, M.; Unger, K. K.; Mant, C. T.; Hodges, R. S. Anal. Chem. 1996, 15, 102-10. (A66) Staby, A.; Mollerup, J. J. Chromatogr., A 1996, 734, 205-12. (A67) Moriyama, H.; Anegayama, M.; Kato, Y. J. Chromatogr., A 1996, 729, 81-5. (A68) Nimura, N. Itoh, H. Mol. Biotechnol. 1996, 5, 11-6. (A69) Torre, M.; Cohen, M. E.; Corzo, N.; Rodriguez, M. A.; DiezMasa, J. C. J. Chromatogr., A 1996, 729, 99-111. (A70) Nice, E. C. Biopolymers 1996, 40, 319-41. (A71) Inglis, A. S.; Reid, G. E.; Simpson, R. J. EXS 1995, 73, 14171. (A72) Perrin, E.; Miclo, L.; Driou, A.; Linden, G. J. Chromatogr., B 1995, 664, 267-76. (A73) Fernandez-Patron, C.; Madrazo, J.; Hardy, E.; Mendez, E.; Frank, R.; Castellanos-Serra, L. Electrophoresis 1995, 16, 91120. (A74) Eckerson, C.; Grimm, R. Electrophoresis 1996, 17, 899-906. (A75) Apffel, A.; Fischer, S.; Goldberg, G.; Goodley, P. C.; Kuhlmann, K. E. J. Chromatogr., A 1995, 712, 177-90. (A76) Guzzetta, A. W.; Hancock, W. S. In New Methods in Peptide Mapping for the Characterization of Proteins; Hancock, W. S., Ed.; CRC: Boca Raton, FL, 1996; pp 181-217. (A77) Tsarbopoulos, A.; Pramanik, B. N.; Nagabushan, T. L.; Covey, T. R. J. Mass Spectrom. 1995, 30, 1752-63. (A78) Li, Y.-M.; Brostedt, P.; Hierten, S.; Nyberg, F.; Silberring, J. J. Chromatogr., B 1995, 664, 426-30. (A79) Nylander, I.; Tan-No, K.; Winter, A.; Silberring, J. Life Sci. 1995, 57, 123-29. (A80) Desiderio, D. M. Biopolymers 1996, 40, 257-64. (A81) Brudel, M.; Kertscher, U.; Schroeder, D.; Melzig, M. F.; Mehlis, B. W. J. Chromatogr., A 1995, 712, 169-75. (A82) Lindner, H.; Sarg, B.; Meraner, C.; Helliger, W. J. Chromatogr., A 1996, 743, 137-44. (A83) Uno, T.; Uchikawa, T.; Iwasaki, T.; Aizono, Y. Biosci. Biotechnol. Biochem. 1995, 59, 420-4. (A84) Naeslund, J. Karlstroem, A. R.; Tjernberg, L. O.; Schierhorn, A.; Terenius, L.; Nordstedt, C. J. Neurochem. 1996, 67, 294301. (A85) Deibler, G. E.; Burlin, T. V.; Stone, A. L. J. Neurosci. Res. 1995, 41, 819-27. (A86) Harris, R. J. J. Chromatogr., A 1995, 705, 129-34. (A87) Rohrer, J. S.; Avdalovic, N. Protein Expression Purif. 1996, 7, 39-44. (A88) Ovalle, R. Anal. Biochem. 1995, 229, 1-7. (A89) Corti, A.; Gasparri, A.; Chen, F.-X.; Pelagi, M.; Brandazza, A.; Sidoli, A.; Siccardi, A. G. Br. J. Cancer 1996, 73, 924-32. (A90) Corradini, D.; Cannarsa, G. J. Liq. Chromatogr. 1995, 18, 3919-31. (A91) Von Jagow, G., Scha¨gger, G., Eds. A Practical Guide to Membrane Protein Purification; Academic Press: San Diego, 1994. (A92) Rooblo-Boza, M.; Andersson, B. Anal. Biochem. 1996, 235, 127-33. (A93) Fricke, B.; Buchmann, T.; Friebe, S. J. Chromatogr., A 1996, 715, 247-58. (A94) Ivashchuk-Kienbaum, Y. A.; Apell, H.-J. Biochim. Biophys. Acta 1994, 1196, 29-37. (A95) Englund, A.-K.; Lundahl, P.; Elenbring, K.; Ericson, C.; Hjerten, S. J. Chromatogr., A 1995, 711, 217-22. (A96) Barter, P. J. In New Trends in Lipid and Lipoprotein Analysis; Sebedio, J.-L., Perkins, E. G., Eds.; AOCS Press: Champaign, IL, 1995; pp 337-46. (A97) Loester, K.; Baum, O.; Hofmann, W.; Reutter, W. J. Chromatogr., A 1995, 711, 187-99. (A98) Tadey, T.; Purdy, W. C. J. Chromatogr., B 1995, 671, 23753. (A99) Muehlradt, P. F.; Meyer, H.; Jansen R. Biochemistry 1996, 35, 7781-6. (A100) Kwok, F.; Humpage, A.; Cheng, C. H. K. Prep. Biochem. Biotechnol. 1996, 26, 1-19. (A101) Minuth, T.; Thoemmers, J.; Kula, M.-R. Biotechnol. Appl. Biochem. 1996, 23, 107-16. (A102) Anantharamaiah, G. M.; Garber D. W. Methods Enzymol. 1996, 263, 267-82. (A103) Lee, R. P.; Doughty, S. W.; Ashman, K.; Walker, J. J. Chromatogr., A 1996, 737, 273-9. (A104) Haginaka, J.; Yamaguchi, Y.; Kunitomo, M. Anal. Biochem. 1995, 232, 163-71.
(A105) DeMeester, I.; Vanhoof, G.; Lambeir, A.-M.; Scharpe, S. J. Immunol. Methods 1996, 189, 99-105. (A106) Hradec, J.; Franek, F.; Dufek, P. J. Chromatogr., B 1996, 681, 55-62. (A107) Mortensen, K. K.; Hansen, F.; Grentzmann, G.; Buckingham, R. H. Eur. J. Biochem. 1995, 234, 732-6. (A108) Zhou, J. Mulshine, J. L.; Unsworth, E. J.; Scott, F. M.; Avis, I. M.; Vos, M. D.; Treston, A. M. J. Biol. Chem. 1996, 271, 10760-6. (A109) Pergami, P.; Jaffe, H.; Safar, J. Anal. Biochem. 1996, 236, 6373. (A110) Brenneman, D. E.; Gozes, I. J. Clin. Invest. 1996, 97, 2299307. (A111) Hara, S.; Liu, N. Meng, S.-Y.; Lu, H. S. Biochim. Biophys. Acta 1996, 1292, 168-76. (A112) Li, D.; Tsutsui, K.; Muneoka, Y.; Minakata, H.; Nomoto, K. Endocrinology 1996, 1618-26. (A113) Burrows, S. D.; Franklin, S. G.; Brighan-Burke, M. R.; Brooks, I. S.; McNulty, D. E.; Feild, J. A.; Anumula, K. R.; O’Shannessy, D. J. J. Chromatogr., B 1995, 668, 219-31. (A114) Ibrahim-Granet, O.; Bertrand, O. J. Chromatogr., B 1996, 684, 239-63. (A115) Lin, S.-X.; Xie, B.; Li, B.; Zhou, M. J. Chromatogr., B 1996, 684, 99-105. (A116) Nairn, A. C.; Sihra, T. S.; Andjus, P.; Craig, A. M.; Miyawaki, A.; Kloppenburg, P.; Lin, Z.; Pouzat, C. NeuroProtocols 1995, 6, 105-7. (A117) Oh, S.-H.; Cha, Y.-S.; Lee, T.-K. J. Biochem. Mol. Biol. 1995, 28, 306-10. (A118) O’Farrell, P. Methods Mol. Biol. 1996, 59, 151-5. (A119) Neuman, G. M.; Condron, R.; Polya, G. M. Int. J. Pept. Protein Res. 1996, 47, 437-46. (A120) Sezdik, J.; Mohammad, J.; Hjerten, S. Neurochem. Res. 1995, 20, 651-8. (A121) Queiroz, J. A.; Garcia, F. A. P.; Cabral, J. M. S. J. Chromatogr., A 1996, 734, 213-9. (A122) Queiroz, J. A.; Garcia, F. A. P; Cabral, J. M. S. J. Chromatogr., A 1996, 707, 137-42. (A123) Mhatre, R.; Nashabeh, W.; Schmalzing, D.; Yao, X.; Fuchs, M.; Whitney, D.; Regnier, F. J. Chromatogr., A 1995, 707, 22531. (A124) Stoller, G.; Tradler, T.; Reucknagel, K. P.; Rahfeld, J.-U.; Fischer, G. FEBS Lett. 1996, 284, 117-22. (A125) Sydow S.; Koepke, A. K. E.; Blank, T. Spiess, J. Mol. Brain Res. 1996, 41, 228-40. (A126) Carlsson, J.; Mosbach, K.; Buelow, L. Biotechnol. Bioeng. 1996, 51, 221-8. (A127) Doebeli, H.; Draeger, N.; Huber, G.; Jakob, P.; Schmidt, D.; Seilheimer, B.; Stueber, D.; Wipf, B.; Zulauf, M. Bio/Technology 1995, 13, 988-93. (A128) Kim, S. O.; Lee, Y. I. J. Biotechnol. 1996, 48, 97-105. (A129) Amari, J. V.; Mazsaroff, I. J. Chromatogr., A 1996, 729, 11324. (A130) Roboti, A.; Livaniou, E.; Evangelatos, G. P.; Tsoupras, G.; Tsolas, O.; Ithakissios, D. S. J. Chromatogr., B 1994, 662, 2734. (A131) Orthner, C. L.; Ralston, A. H.; Gee, D.; Kent, R.; Kolen, B.; McGriff, J. D.; Drohan, W. N. Vox Sang. 1995, 69, 309-18. (A132) Feldman, P. A.; Bradbury, P. I.; Williams, J. D.; Sims, G. E.; McPhee, J. W.; Pinnell, M. A.; Harris, L.; Crombie, G. I.; Evans, D. R. Blood Coagulation Fibrinolysis 1994, 5, 939-48. (A133) Kundu A.; Vunnum, S.; Jayaraman, G.; Cramer, S. M. Biotechnol. Bioeng. 1995, 48, 452-60. (A134) Jayaraman, G.; Li, Y.-F.; Moore, J. A.; Cramer, S. M. J. Chromatogr., A 1995, 702, 143-55. (A135) Kundu A.; Vunnum, S.; Cramer, S. M. J. Chromatogr., A 1995, 707, 57-67. (A136) Gerstner, J. A.; Morris, J.; Hunt, T.; Hamilton, R.; Afeyan, N. B. J. Chromatogr., A 1995, 695, 195-204. (A137) Johansson, H. J.; Jaegersten, C.; Shiloach, J. J. Biotechnol. 1996, 48, 9-14. (A138) Walker, S. G.; Dale, C. J.; Lyddiatt, A. J. Chromatogr., B 1996, 680, 91-6. (A139) Gu, T.; Gu, Y.; Zheng, Y.; Wiehl, P. E.; Kopchick, J. J. Sep. Techol. 1994, 4, 258-60. (A140) Huyghe, B. G.; Liu, X.; Sutjipto, S.; Sugarman, B. J.; Horn, M. T.; Shepard, H. M.; Scandella, C. J.; Shabram, P. Hum. Gene Ther. 1995, 6, 1403-16. MASS SPECTROMETRY (B1) Burlingame, A. L.; Boyd, R. K.; Gaskell, S. J. Anal. Chem. 1996, 68, 599R-651R. (B2) Caprioli, R. M.; Suter, M. J. F. In Introduction to Biophysical Methods for Protein and Nucleic Acid Research; Glasel, J. A., Deutscher, M. P., Eds.; Academic Press: San Diego, CA, 1995; pp 147-204. (B3) Bowers, M. T.; Marshall, A. G.; McLafferty, F. W. J. Chromatogr., A 1996, 100, 12897-910. (B4) Nguyen, D. N.; Becker, G. W.; Riggin, R. M. J. Chromatogr., A 1995, 705, 21-45. (B5) Niessen, W. M. A.; Tinke, A. P. J. Chromatogr., A 1995, 703, 37-57. (B6) Gelpı´, E. J. Chromatogr., A 1995, 703, 59-60. (B7) Tomer, K. B.; Moseley, M. A.; Deterding, L. J.; Parker, C. E. Mass Spectrom. Rev. 1994, 13, 431-57.
(B8) Yates, J. R., III; McCormack, A. L.; Link, A. J.; Schielz, D.; Eng, J.; Hays, L. Analyst 1996, 121, 65R-76R. (B9) Cai, J.; Henion, J. J. Chromatogr. 1995, 703, 667-92. (B10) Tomlinson, A. J.; Guzman, N. A.; Naylor, S. J. Capillary Electrophor. 1995, 2, 247-66. (B11) Wahl, J. H.; Udseth, H. R.; Smith, R. D. In New Methods in Peptide Mapping for the Characterization of Proteins; Hancock, W. S., Ed.; CRC: Boca Raton, FL, 1996; pp 143-79. (B12) Farmer, T. B.; Caprioli, R. M. NATO ASI Ser., Ser. C 1996, 475, 61-88. (B13) Saru, F.; Montanarella, L. Rev. Anal. Chem. 1995, 14, 20526. (B14) McLafferty, F. W.; Senko, M. W.; Little, D. P.; Wood, T. D.; O’Connor, P. B.; Speir, J. P.; Chorush, R. A.; Kelleher, N. L. Adv. Mass Spectrom. 1995, 13, 115-28. (B15) Zaluzec, E. J.; Gage, D. A.; Watson, J. T. Protein Expression and Purification 1995, 6, 109-23. (B16) Zenobi, R. Int. J. Mass Spectrom. Ion Processes 1995, 145, 5177. (B17) Vestling, M. M.; Fenselau, C. Mass Spectrom. Rev. 1995, 14, 169-78. (B18) Patterson, S. D.; Aebersold, R. Electrophoresis 1995, 16, 1791814. (B19) Spengler, B. Methods Mol. Biol. 1996, 61, 43-56. (B20) Wilm, M.; Houthaeve, T.; Talbo, G.; Kellner, R.; Mortensen, P.; Mann, M. In Mass Spectrometry in the Biological Sciences; Burlingame, A. L., Carr, S. A., Eds.; Humana: Totowa, NJ, 1996; pp 245-65. (B21) Chait, B. T.; Wang, R.; Bevais, R. C.; Kent, S. B. H. Science 1993, 262, 89-92. (B22) Tempst, P.; Erdjument-Bromage, H.; Casteels, P.; Geromanos, S.; Lui, M.; Powell, M.; Nelson, R. W. In Mass Spectrometry in the Biological Sciences; Burlingame, A. L., Carr, S. A., Eds.; Humana: Totowa, NJ, 1996; pp 105-33. (B23) Thiede, B.; Wittmann-Liebold, B.; Bienert, M.; Krause, E. FEBS Lett. 1995, 357, 65-9. (B24) Ramsay, S. L.; Steinborner, S. T.; Waugh, R. J.; Dua, S.; Bowie, J. H.; Rapid Commun. Mass Spectrom. 1995, 9, 1241-3. (B25) Patterson, D. H.; Tarr, G. E.; Regnier, F. E.; Martin, S. A. Anal. Chem. 1995, 67, 3971-8. (B26) Woods, A. S.; Huang, A. Y. C.; Cotter, R. J.; Pasternack, G. R.; Pardoll, D. M.; Jaffee, E. M. Anal. Biochem. 1995, 226, 1525. (B27) Gulden, P. H.; Hackett, H.; Addona, T. A.; Guo, L.; Walter, C. B.; Sherman, N. E.; Shabanowitz, J.; Hewlett, E. L.; Hunt, D. F. In Mass Spectrometry in the Biological Sciences; Burlingame, A. L., Carr, S. A., Eds.; Humana: Totowa, NJ, 1996; pp 281305. (B28) McCormack, A. L.; Eng, J. K.; DeRoss, P. C.; Rudensky, A. Y.; Yates, J. R., III ACS Symp. Ser. 1996, No. 619, 207-225 (Biochemical and Biotechnological Applications of Electrospray Ionization Mass Spectrometry). (B29) Knierman, M. D.; Coligan, J. E.; Parker, K. C. Rapid Commun. Mass Spectrom. 1994, 8, 1007-10. (B30) Takamoto, K.; Kamo, M.; Kubota, K.; Satake, K.; Tsugita, A. Eur. J. Biochem. 1995, 228, 362-72. (B31) Bures, E. J.; Nika, H.; Chow, D. T.; Morrison, H. D.; Hess, D.; Aebersold, R. Anal. Biochem. 1995, 224, 364-72. (B32) Hess, D.; Nika, H.; Chow, D. T.; Bures, E. J.; Morrison, H. D.; Aebersold, R.; Anal. Biochem. 1995, 224, 373-81. (B33) Vladimirov, S. N.; Ivanov, A. V.; Karpova, G.; Musolyamov, A. K.; Egorov, T. A.; Thiede, B.; Wittmann-Liebold, B.; Otto, A. Eur. J. Biochem. 1996, 239, 144-9. (B34) Wilm, M.; Shevchenko, A.; Houthaeve, T.; Breit, S.; Schweiger, L.; Fotsis, T.; Mann. M. Nature 1996, 379, 466-9. (B35) Shevchenko, A.; Wilm, M.; Vorm, O.; Mann, M. Anal. Chem. 1996, 68, 850-8. (B36) Jainhuknan, J.; Cassady, C. J. Rapid Commun. Mass Spectrom. 1996, 10, 1678-82. (B37) Kosaka, T.; Jshikawa, T.; Kinoshita, T. Rapid Commun. Mass Spectrom. 1995, 9, 1342-4. (B38) Zaluzec, E. J.; Gage, D. A.; Watson, J. T. J. Am. Soc. Mass Spectrom. 1994, 5, 359-66. (B39) Wu, J.; Gage, D. A.; Watson, J. T. Anal. Biochem. 1996, 235, 161-74. (B40) Watson, J. T.; Wu, J. Polym. Prepr. 1996, 37, 318. (B41) Denslow, N. D.; Nguyen, H. P. In Techniques in Protein Chemistry VII; Marshak, D. R., Ed.; Academic Press: San Diego, 1996; pp 241-248. (B42) Ming, D.; Markley, J. L.; Hellekant, G. BioTechniques 1995, 18, 808-10. (B43) Ruoppolo, M.; Torella, C.; Kanda, F.; Panico, M.; Pucci, P.; Marino, G.; Morris, H. R. Folding Des. 1996, 1, 381-90. (B44) Crimmins, D. L.; Saylor, M.; Rush, J.; Thoma, R. S. Anal. Biochem. 1995, 226, 355-61. (B45) Sun, Y.; Bauer, M. D.; Keough, T. W.; Lacey, M. P. Methods Mol. Biol. 1996, 61, 181-210. (B46) Hubbard, M. J.; Cohen, P. Trends Biochem. Sci. 1993, 18, 172-7. (B47) Annan, R. S.; Carr, S. A. Anal. Chem. 1996, 68, 3413-21. (B48) Carr, S. A.; Huddleston, M. J.; Annan, R. S. Anal. Biochem. 1996, 239, 180-92. (B49) Huddleston, M. J.; Annan, R. S.; Bean, M. F.; Carr, S. A. J. Am. Soc. Mass Spectrom. 1994, 4, 710-7. (B50) Tsutakawa, S. E.; Medzihradszky, K. F.; Flint, A. J.; Burlingame, A. L. J. Biol. Chem. 1995, 270, 26807-12.
Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
53R
(B51) Watts, J. D.; Affolter, M.; Krebs, D. L.; Wange, R. L.; Samelson, L. E.; Aebersold, R. ACS Symp. Ser. 1996, No. 619, 381-407 (Biochemical and Biotechnological Applications of Electrospray Ionization Mass Spectrometry). (B52) Amankwa, L. N.; Harder, K.; Jirik, F.; Aebersold, R. Protein Sci. 1995, 4, 113-25. (B53) Weigt, C.; Korte, H.; von Strandmann, R. P.; Hengstenberg, W.; Meyer, H. E. J. Chromatogr., A 1995, 712, 141-7. (B54) Talbo, G.; Mann, M. In Techniques in Protein Chemistry V; Crabb, J. W., Ed.; Academic Press: San Diego, 1994; pp 10513. (B55) Lapolla, A.; Fedele, D.; Seraglia, R.; Catinella, S.; Baldo, L.; Aronica, R.; Traldi, P. Diabetologia 1995, 38, 1076-81. (B56) Lapolla, A.; Fedele, D.; Aronica, R.; Garbeglio, M.; D’Alpaos, M.; Seraglia, R.; Traldi, P. Rapid Commun. Mass Spectrom. 1996, 10, 1133-5. (B57) Tsarbopoulos, A.; Pramanik, B. N. J. Mass Spectrom. 1995, S207-S209. (B58) Karas, M.; Bahr, U.; Strupat, K.; Hillenkamp, F.; Tsarbopoulos, A.; Pramanik, B. N. Anal. Chem. 1995, 67, 675-9. (B59) Guzzetta, A. W.; Hancock, W. S. In New Methods in Peptide Mapping for the Characterization of Proteins; Hancock, W. S., Ed.; CRC: Boca Raton, FL, 1996; pp 181-217. (B60) Burlingame, A. L. Curr. Opin. Biotechnol. 1996, 7, 4-10. (B61) Chankel, J. A.; Apffel, J. A.; Hancock, W. S. LC-GC 1995, 13, 866, 868, 870, 872, 874, 875-876. (B62) Apffel, A.; Chakel, J. A.; Hancock, W. S.; Souders, C.; M’Timkulu, T.; Pungor, E., Jr. J. Chromatogr., A 1996, 732, 27-42. (B63) Apffel, A.; Chakel, J. A.; Udiavar, S.; Hancock, W. S.; Souders, C.; Pungor, E. ACS Symp. Ser. 1996, No. 619, 432-471. (B64) Kelly, J. F.; Locke, S. J.; Ramaley, L.; Thibault, P. J. Chromatogr., A 1996, 720, 409-27. (B65) Greis, K. D.; Hayes, B. K.; Comer, F. I.; Kirk, M.; Barnes, S.; Lowary, T. L.; Hart, G. W. Anal. Biochem. 1996, 234, 38-49. (B66) Hunter, A. P.; Games, D. E. Rapid Commun. Mass Spectrom. 1995, 9, 42-56. (B67) Boss, H. J.; Rohde, M. F.; Rush, R. S. Anal. Biochem. 1995, 230, 123-9. (B68) Nilsson, B. Mol. Biotechnol. 1994, 2, 243-80. (B69) Harvey, D. J. J. Chromatogr., A 1996, 720, 429-46. (B70) Lapolla, A.; Fedele, D.; Catinella, S.; Traldi, P. NATO ASI Ser., Ser. C 1996, 475, 317-49. (B71) Oliver, R. W. A.; Green, B. N.; Harvey, D. J. Biochem. Soc. Trans. 1996, 24, 917-27. (B72) Kroon, D. J.; Freedy, J.; Burinsky, D. J.; Sharma, B. J. Pharm. Biomed. Anal. 1995, 13, 1049-54. (B73) Reinhold, V. N.; Reinhold, B. B.; Costello, C. E. Anal. Chem. 1995, 67, 1772-84. (B74) Siciliano, R. A.; Amoresano, A.; Alving, K.; Cantilena, A.; Pucci, P. Chim. Oggi 1995, 13, 17-22. (B75) Settineri, C. A.; Burlingame, A. L. Tech. Protein Chem. 1993, 5, 97-104. (B76) Harvey, D. J.; Rud, P. M.; Bateman, R. H.; Bordoli, R. S.; Howes, K.; Hoyes, J. B.; Vickers, R. G. Org. Mass Spectrom. 1994, 29, 753-65. (B77) Papac, Damon I.; Wong, Anissa; Jones, Andrew J. S. Anal. Chem. 1996, 68, 3215-23. (B78) Rouse, Jason C.; Vath, James E. Anal. Biochem. 1996, 238, 82-92. (B79) Zhao, Y.; Chait, B. T. Anal. Chem. 1994, 66, 3723-6. (B80) Zhao, Y.; Chait, B. T. Methods Mol. Biol. 1996, 66, 129-36. (B81) Papac, D. I.; Hoyes, J.; Tomer, K. B. Protein Sci. 1994, 3, 1485-92. (B82) Parker, C. E.; Papac, D. I.; Trojak, S. K.; Tomer, K. B. J. Immunol. 1996, 157, 198-206. (B83) Macht, M.; Fiedler, W.; Kuerzinger, K.; Przybylski, M. Biochemistry 1996, 35, 15633-9. (B84) Kriwacki, R. W.; Wu, J.; Siuzdak, G.; Wright, P. E. J. Am. Chem. Soc. 1996, 118, 5320-1. (B85) Cohen, S. L.; Ferre-D’Amare, A. R.; Burley, S. K.; Chait, B. T. Protein Sci. 1995, 4, 1088. (B86) Cheng, X.; Harms, A. C.; Goudreau, P. N.; Terwilliger, T. C.; Smith, R. D. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 7022-7. (B87) Jensen, O. N.; Barotsky, D. F.; Young, M. C.; von Hippel, P. H.; Swenson, S.; Seifried, S. E. In Techniques in Protein Chemistry V; Crabb, J. W., Ed.; Academic Press: San Diego, 1994; pp 27-37. (B88) Tang, X.; Callahan, J. H.; Zhou, P.; Vertes, A. Anal. Chem. 1995, 67, 4542-8. (B89) Przybylski, M.; Kast, J.; Glocker, M. O.; Duerr, E.; Bosshard, H. R.; Nock, S.; Sprinze, M. Toxicol. Lett. 1995, 82/83, 56775. (B90) Woods, A. S.; Buchsbaum, J. C.; Worrall, T. A.; Berg, J. M.; Cotter, R. J. Anal. Chem. 1995, 67, 4462-5. (B91) Hofstadler, S. A.; Swanek, F. D.; Gale, D. C.; Ewing, A. G.; Smith, R. D. Anal. Chem. 1995, 67, 1477-80. (B92) Valaskovic, G. A.; Kelleher, N. L.; McLafferty, F. W. Science 1996, 273, 1199-202. (B93) Holland, R. D.; Wilkes, J. G.; Rafii, F.; Sutherland, J. B.; Persons, C. C.; Voorhees, K. J.; Lay, J. O., Jr. Rapid Commun. Mass Spectrom. 1996, 10, 1227-32. (B94) Cain, T. C.; Lubman, D. M.; Weber, W. J., Jr. Rapid Commun. Mass Spectrom. 1994, 8, 1026-30. (B95) Van Strien, F. J. C.; Jespersen, S.; van der Greef, J.; Jenks, B. G.; Roubos, E. W. FEBS Lett. 1996, 379, 165-70. 54R
Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
(B96) Jimenez, C. R.; Li, K. W.; Smit, A. B.; Van Minnen, J.; Janse, C.; Van Veelen, P.; Dreisewerd, K.; Zeng, J.; Van Der Greef, J. In Mass Spectrometry in the Biological Sciences; Burlingame, A. L., Carr, S. A., Eds.; Humana: Totowa, NJ, 1996; pp 22743. (B97) Berkenkamp, S.; Karas, M.; Hillenkamp, F. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 7003-7. FOURIER TRANSFORM INFRARED SPECTROSCOPY (C1) Haris, P. I.; Chapman, D.; Benga, G. Eur. J. Biochem. 1995, 233, 659-64. (C2) Rahmelow, K.; Huebner, W. Appl. Spectrosc. 1996, 50, 795804. (C3) Ende, M. T.; Peppas, N. A. Pharm. Res. 1995, 12, 2030-5. (C4) Keiderling, T. A.; Wang, B.; Urbanova, M.; Pancoska, P.; Dukor, R. K. Faraday Discuss. 1995, 99, 263-85. (C5) Ludlam, C. F. C.; Arkin, I. T.; Liu, X. M.; Rothman, M. S.; Rath, P.; Aimoto, S.; Smith, S. O.; Engelman, D. M., Rothschild, K. J. Biophys. J. 1996, 70, 1728-36. (C6) Liu, X. M.; Sonar, S.; Lee, C.-P.; Coleman, M.; RajBhandary, U. L.; Rothschild, K. J. Biophys. Chem. 1995, 56, 63-70. (C7) Ludlam, C. F. C.; Sonar, S.; Lee, C.-P.; Coleman, M.; Herzfeld, J.; RajBhandary, U. L.; Rothschild, K. J. Biochemistry 1995, 34, 2-6. (C8) Martin, I.; Schaal, H.; Scheid, A.; Ruysschaert, J.-M. J. Virol. 1996, 70, 298-304. (C9) Baenziger, J. E.; Methot, N. J. Biol. Chem. 1995, 270, 2912937. (C10) Dong, A.; Hyslop, R. M.; Pringle, D. L. Arch. Biochem. Biophys. 1996, 333, 275-81. (C11) Remmele, R. L., Jr.; Stushnoff, C. Biopolymers 1994, 34, 36570. (C12) Griebenow, K.; Klibanov, A. M. Proc. Natl. Acad. Sci. U.S.A. 1995, 92, 10969-76. (C13) Costantino, H. R.; Langer, R.; Klibanov, A. M. J. Pharm. Sci. 1994, 83, 1662-9. (C14) Dong, A.; Prestrelski, S. J.; Allison, S. D.; Carpenter, J. F. J. Pharm. Sci. 1995, 84, 415-24. (C15) Costantino, H. R.; Griebenow, K.; Mishra, P.; Langer, R.; Klibanov, A. M. Biochim. Biophys. Acta 1995, 1253, 69-74. (C16) Turula, V. E.; de Haseth, J. A. Anal. Chem. 1996, 68, 62938. (C17) Goormaghtigh, E.; Cabiaux, V.; Ruysschaert, J. M. Subcell. Biochem. 1994, 23, 405-50. (C18) Oberg, K. A.; Link, A. L. In Techniques in Protein Chemistry VI; Crabb, J. W., Ed.; Academic Press: San Diego, 1995; pp 475-84. (C19) Heberle, J.; Zscherp, C. Appl. Spectrosc. 1996, 50, 588-96. (C20) Fu, F.-N.; DeOliveira D. B.; Trumble, W. R.; Sarkar, H. K.; Singh, B. R. Appl. Spectrosc. 1994, 48, 1432-41. (C21) Ball, A.; Jones, R. A. L. Langmuir 1995, 11, 3542-8. (C22) Buijs, J.; Norde, W.; Lichtenbelt, J. W. T. Langmuir 1996, 12, 1605-13. (C23) Baty, A. M.; Suci, P. A.; Tyler, B. J.; Geesey, G. G. J. Colloid Interface Sci. 1996, 177, 307-15. (C24) Boulkanz, L.; Balcar, N.; Baron, M.-H. Appl. Spectrosc. 1995, 49, 1737-46. (C25) Chan, H.-K.; Ongpipattanakul, B.; Au-Yeung, J. Pharm. Res. 1996, 13, 238-42. (C26) Tatulian, S. A.; Hinterdorfer, P.; Baber, G.; Tamm, L. K. Embo J. 1995, 14, 5514-23. (C27) Bramanti, E.; Benedetti, E. Biopolymers 1996, 38, 639-53. (C28) Kumosinski, T. F.; Unruh, J. J. Talanta 1996, 43, 199-219. (C29) Kumosinski, T. F.; Unruh, J. J. In Molecular Modeling; Kumosinski, T. F., Liebman, M. N., Eds.; ACS Symposium Series 576; American Chemical Society: Washington, DC, 1994; pp 71-99. (C30) Pribic, R. Anal. Biochem. 1994, 223, 26-34. (C31) Baumruk, V.; Pancoska, P.; Keiderling, T. A. J. Mol. Biol. 1996, 259, 774-91. (C32) Tonan, K.; Ikawa, S.-I. J. Am. Chem. Soc. 1996, 118, 6960-5. (C33) Liu, Y.; Cho, R.-K.; Sakurai, K.; Miura, T.; Ozaki, Y. Appl. Spectrosc. 1994, 48, 1249-54. NUCLEAR MAGNETIC RESONANCE (D1) Freund, S. M. V.; Bycroft, M. Molecular Biology: Current Innovations and Future Trends Part 2; Horizon Scientific Press: Wymondham, U.K., 1995; pp 155-69. (D2) Aswad, D. W.; Guzzetta, A. W. In Deamidation and Isoaspartate Formation in Peptides and Proteins; Aswad, D. W., Ed.; CRC: Boca Raton, FL, 1995; pp 7-29. (D3) Chazin, W. J.; Kossiakoff, A. A. In Deamidation Isoaspartate Formation in Peptides and Proteins; Aswad, D. W., Ed.; CRC: Boca Raton, FL, 1995; pp 193-206. (D4) Asakura, T.; Taoka, K.; Demura, M.; Williamson, M. P. J. Biomol. NMR 1995, 6, 227-36. (D5) Celda, B.; Biamonti, C.; Arnau, M. J.; Tjero, R.; Montelione, G. T. J. Biomol. NMR 1995, 5, 161-72. (D6) Lee, M. S.; Cao, B. Protein Eng. 1996, 9, 15-25. (D7) Danielson, M. A.; Falke, J. J. Annu. Rev. Biophy. Biomol. Struct. 1996, 25, 163-95. (D8) Heintz, D.; Kany, H.; Kalbitzer, H. R. Biochemistry 1996, 35, 12686-93.
(D9) Mumenthaler, Ch.; Braun, W. J. Mol. Biol. 1995, 254, 46580. (D10) Van Geerestein-Ujah, E. C.; Slijper, M.; Boelens, R.; Kaptein, R. J. Biomol. NMR 1995, 5, 67-78. (D11) Adzhubei, A. A.; Laughton, C. A.; Neidle, S. Protein Eng. 1995, 8, 615-25. (D12) Beckman, R. A.; Zuiderweg, E. R. P. J. Magn. Reson. Ser. A 1995, 113, 223-31. (D13) Hoogstraten, C. G.; Markley, J. L. J. Mol. Biol. 1996, 258, 334-48. (D14) Malliavin, T. E.; Delsuc, M. A.; Orekhov, V. Yu.; Arseniev, A. S. J. Biomol. NMR 1995, 5, 193-201. (D15) Schwager, M.; Bodenhausen, G. J. Magn. Reson. Ser. B 1996, 111, 40-9. (D16) Kar, L.; Sherman, S. A.; Johnson, M. E. In Molecular Modeling: From Virtual Tools to Real Problems; Kumosinski, T. F., Liebman, M. N., Eds.; ACS Symposium Series 576; American Chemical Society: Washington, DC, 1994; pp 446-65. (D17) Hoogstraten, C. G.; Westler, W. M.; Macura, S.; Markley, J. L. J. Am. Chem. Soc. 1995, 117, 5610-1. (D18) Canet, D.; Decorps, M. In Dynamics of Solutions and Fluid Mixtures by NMR; Delpuech, J., Ed.; Wiley: Chichester, U.K., 1995; pp 309-43. (D19) Hurd, R. E.; Deese, A.; Johnson, M. O.; Sukumar, K.; van Zijl, P. C. M. J. Magn. Reson. Ser. A 1996, 119, 285-8. (D20) Gozansky, E. K.; Gorenstein, D. G. J. Magn. Reson. Ser. B 1996, 111, 94-6. (D21) Van, Q. N.; Shaka, A. J. J. Magn. Reson. Ser. A 1996, 119, 295-301. (D22) Stilbs, P.; Paulsen, K.; Griffiths, P. C. J. Phys. Chem. 1996, 100, 8180-9. (D23) Wu, D.; Chen, A.; Johnson, C. S., Jr. J. Magn. Reson. Ser. A 1995, 115, 260-4. (D24) Wu, D.; Chen, A.; Johnson, C. S., Jr. J. Magn. Reson. Ser. A 1996, 121, 88-91. (D25) Lin, M.; Shapiro, M. J. J. Org. Chem. 1996, 61, 7617-9. (D26) Wu, D.; Chen, A.; Johnson, C. S., Jr. Bull. Magn. Reson. 1995, 17, 21-6. (D27) Lin, M.; Jayawickrama, D.; Rose, R. A.; DelViscio, J. A.; Larive, C. K. Anal. Chim. Acta 1995, 307, 449-57. (D28) Liu, M.; Nicholson, J. K.; Lindon, J. C. Anal. Chem. 1996, 68, 3370-6. (D29) Alteri, A. S.; Hinton, D. P.; Byrd, R. A. J. Am. Chem. Soc. 1995, 117, 7566-7. (D30) Lin, M.; Larive, C. K. Anal. Biochem. 1995, 229, 214-20. (D31) Stevens, A.; Wang, S. X.; Caines, G. H.; Schleich, T. Biochim. Biophys. Acta 1995, 1246, 82-90. (D32) Brown, R. A.; Grant, D. M. J. Magn. Reson. Ser. B 1995, 106, 253-60. (D33) Keifer, P. A. J. Org. Chem. 1996, 61, 1558-9. (D34) Sarkar, S. K.; Garigipati, R. S.; Adams, J. A.; Keifer, P. A. J. Am. Chem. Soc. 1996, 118, 2305-6. (D35) Anderson, R. C.; Jarema, M. A.; Shapiro, M. J.; Stokes, J. P.; Ziliox, M. J. Org. Chem. 1995, 60, 2650-1. (D36) Anderson, R. C.; Stokes, J. P.; Shapiro, M. J. Tetrahedron Lett. 1995, 36, 5311-4. (D37) Boutin, J. A.; Hennig, P.; Lambert, P.-H.; Bertin, S.; Petit, L.; Mahieu, J.-P.; Serkiz, B.; Volland, J.-P.; Fauchere, J.-L. Anal. Biochem. 1996, 234, 126-41. (D38) Lindon, J. C.; Nicholson, J. K.; and Wilson, I. D. Adv. Chromatogr. 1996, 36, 315-86. (D39) Wu, N.; Webb, A.; Peck, T. L.; Sweedler, J. V. Anal. Chem. 1995, 67, 3101-7. (D40) Behnke, B.; Schlotterbeck, G.; Tallarek, U.; Strohschein, S.; Tseng, L.-H.; Keller, T.; Albert, K.; Bayer, E. Anal. Chem. 1996, 68, 1110-5. (D41) Bariat, H.; Morris, G. A.; Newman, J. M.; Swanson, A. G. J. Magn. Reson. Ser. A 1996, 119, 115-9. (D42) Smallcombe, S. H.; Patt, S. L.; Keifer, P. A. J. Magn. Reson. Ser. A 1995, 117, 295-303. (D43) Chen, J. K.; Schreiber, S. L. Angew. Chem., Int. Ed. Engl. 1995, 34, 953-69. (D44) Siligard, G.; Drake, A. F. Biopolymers (Pept. Sci.) 1995, 37, 281-92. (D45) Wagner, G. Nature Struct. Biol. 1995, 2, 255-7. (D46) Shuker, S. B.; Hajduk, P. J.; Meadows, R. P.; Fesik, S. W. Science 1996, 274, 1531-4. (D47) Oldfield, E. J. Biomol. NMR 1995, 5, 217-25. (D48) Zheng, Z.; Czaplicki, J.; Jardetzky, O. Biochemistry 1995, 34, 5212-23. (D49) Epstein, D. M.; Benkovic, S. J.; Wright, P. E. Biochemistry 1995, 34, 11037-48. (D50) Kay, L. E.; Muhandiram, D. R.; Farrow, N. A.; Aubin, Y.; Forman-Kay, J. D. Biochemistry 1996, 35, 361-8. (D51) Shaw, G. L.; Davis, B.; Keeler, J.; Fersht, A. R. Biochemistry 1995, 34, 2225-33. (D52) Pervushin, K. V.; Orekhov, V. Yu.; Korzhnev, D. M.; Arseniev, A. S. J. Biomol. NMR 1995, 5, 383-96. (D53) Pascal, S. M.; Yamazaki, T.; Singer, A. U.; Kay, L. E.; FormanKay, J. D. Biochemistry 1995, 34, 11353-62. (D54) Henry, G. H.; Sykes, B. D. J. Biomol. NMR 1995, 6, 59-66. (D55) Huyghues-Despointes, B. M. P.; Klinger, T. M.; Baldwin, R. L. Biochemistry 1995, 34, 13267-71. (D56) Baranowska, H. M.; Olszewski, K. J. Biochim. Biophys. Acta 1996, 1289, 312-4.
(D57) Baguet, E.; Chapman, B. E.; Kuchel, P. W. Bull. Magn. Reson. 1995, 17, 180-1. (D58) Fukuzaki, M.; Miura, N.; Shinyashiki, N.; Kurita, D.; Shioya, S.; Haida, M.; Mashimo, S. J. Phys. Chem. 1995, 99, 431-5. (D59) Wang, Y.-X.; Freedberg, D. I.; Grzesiek, S.; Torchia, D. A.; Wingfield, P. T.; Kaufman, J. D.; Stahl, S. J.; Chang, C.-H.; Hodge, C. N. Biochemistry 1996, 35, 12694-704. (D60) Areˆas, E. P. G.; Areˆas, J. A. G.; Hamburger, J.; Peticolas, W. L.; Santos, P. S. J. Colloid Interface Sci. 1996, 180, 578-89. (D61) Andrew, E. R.; Szczesniak, E. Prog. Nucl. Magn. Reson. Spectrosc. 1995, 28, 11-36. (D62) Garbow, J. B.; Gullion, T. In Carbon-13 NMR Spectroscopy of Biological Systems; Beckmann, N., Ed.; Academic Press: San Diego, CA, 1995; pp 65-115. (D63) Vandenbussche, G.; Johansson, J.; Clercx, A.; Curstedt, T.; Ruysschaert, J.-M. EXS 1995, 73, 27-47. (D64) Wu, C. H.; Ramamoorthy, A.; Gierasch, L. M.; Opella, S. J. J. Am. Chem. Soc. 1995, 117, 6148-9. (D65) Gu, Z.; Ridenour, C. F.; Bronnimann, C. E.; Iwashita, T.; McDermott, A. J. Am. Chem. Soc. 1996, 118, 822-9. (D66) McDowell, L. M.; Lee, M.; McKay, R. A.; Anderson, K. S.; Schaefer, J. Biochemistry 1996, 35, 3328-34. (D67) Schmidt-Rohr, K. J. Am. Chem. Soc. 1996, 118, 7601-3. CIRCULAR DICHROISM (E1) Miller, I. R.; Brumfeld, V.; Korenstein, R. In Biomembrane Electrochemistry; Blank, M., Vodyanoy, I., Eds.; Advances in Chemistry Series 235; American Chemical Society: Washington, DC, 1994; pp 107-27. (E2) Szendrei, G. I.; Fabian, H.; Mantsch, H. H.; Lovas, S.; Nyeki, O.; Schoen, I.; Otvos, L., Jr. Eur. J. Biochem. 1994, 226, 91724. (E3) Higashi, N.; Yamamoto, T.; Yokoyama, K.; Niwa, M. Macromolecules 1995, 28, 2585-7. (E4) Li, S.-C.; Goto, N. K.; Williams, K. A.; Deber, C. M. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 6676-81. (E5) Yamashita, H.; Nakatsuka, T.; Hirose, M. J. Biol. Chem. 1995, 270, 29806-12. (E6) Moench, S. J.; Saterlee, J. D. Protein Chem. 1995, 14, 56782. (E7) Gortazar, P.; Ravina, M.; Vazquez, J. T. J. Pharm. Sci. 1995, 84, 1316-21. (E8) Bracete, A. M.; Kadkhodayan, S.; Sono, M.; Huff, A. M.; Zhuang, C.; Cooper, D. K.; Smith, K. M.; Chang, C. K.; Dawson, J. H. Inorg. Chem. 1994, 33, 5042-9. (E9) Fasman, G. D., Ed. Cicular Dichroism and the Conformation Analysis of Biomolecules; Plenum Press: New York, 1996. (E10) Galat, A. Eur. J. Biochem. 1996, 236, 428-35. (E11) Greenfield, N. J. Anal. Chem. 1996, 235, 1-10. (E12) Bohm, G.; Muhr, R.; Jaenicke, R. Protein Eng. 1992, 5, 1915. (E13) Dalmas, B.; Bannister, W. H. Anal. Biochem. 1995, 225, 3948. (E14) Cascio, M.; Wallace, B. A. Protein Pept. Lett. 1994, 1, 13640. (E15) Cascio, M.; Wallace, B. A. Anal. Biochem. 1995, 227, 90100. (E16) Manning, M.; Woody, R. W. Biopolymers 1991, 31, 569-86. (E17) Toniolo, C.; Polese, A.; Formaggio, F.; Crisma, M.; Kamphuis, J. J. Am. Chem. Soc. 1996, 118, 2744-5. (E18) Formaggio, F.; Crisma, M.; Toniolo, C.; Kamphuis, J. Biopolymers 1996, 38, 301-4. (E19) Keiderling, T. A.; Pancoska, P.; Baumruk, V.; Urbanova, M.; Gupta, V. J.; Dukor, R. K.; Huo, D. In Molecular Modeling; Kumosinski, T. F., Liebman, M. N., Eds.; ACS Symposium Series 576; American Chemical Society: Washington, DC, 1994; pp 61-70. (E20) Keiderling, T. A.; Wang, B.; Urbanova, M.; Pancoska, P.; Dukor, R. K. Faraday Discuss. 1994, 99, 263-85. (E21) Pancoska, P.; Bitto, E.; Janota, V.; Keiderling, T. A. Faraday Discuss. 1994, 99, 287-310. (E22) Baumruk, V.; Pancoska, P.; Keiderling, T. A. J. Mol. Biol. 1996, 259, 774-91. (E23) Bjo ¨rling, S. C.; Goldbeck, R. A.; Paquette, S. J.; Miller, S. J.; Kliger, D. S. Biochemistry 1996, 35, 8619-27. (E24) Su, Z.-D.; Arooz, M. T.; Chen, H. M.; Gross, C. J.; Tsong, T. Y. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 2539-44. (E25) Goldberg, M. E.; Chaffotte, A. F. In Statistical Mechanics, Protein Structure, and Protein Substrate Interactions; Doniach, S., Ed.; NATO ASI Series B; Plenum Press: New York, 1994; Vol. 325, pp 19-29. (E26) Kuwajima, K. Methods Mol. Biol. 1995, 40, 115-35. (E27) Zitzewitz, J. A.; Bilsel, O.; Luo, J.; Jones, B. E.; Matthews, C. R. Biochemistry 1995, 34, 12812-9. NONENZYMATIC POSTTRANSLATORIAL MODIFICATIONS (F1) Chang, B. S.; Reeder, G.; Carpenter, J. F. Pharm. Res. 1996, 13, 243-9. (F2) Chang, B. S.; Beauvais, R. M.; Dong, A.; Carpenter, J. F. Arch. Biochem. Biophys. 1996, 331, 249-58. (F3) Son, K.; Kwon, C. Pharm. Res. 1995, 12, 451-4. (F4) Deamidation and Isoaspartate Formation in Peptides and Proteins; Aswad, D. W., Ed.; CRC Press: Boca Raton, FL, 1995.
Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
55R
(F5) Brennan, T. V.; Clarke, S. Int. J. Pept. Protein Res. 1995, 45, 547-53. (F6) Cacia, J.; Keck, R.; Presta, L. G.; Frenz, J. Biochemistry 1996, 35, 1897-903. (F7) Derby, P. L.; Aoki, K. H.; Katta, V.; Rohde, M. F. In Techniques in Protein Chemistry VII; Marshak, D. R., Ed.; Academic Press: San Diego, CA, 1996; pp 109-19. (F8) Volkin, D. B.; Verticelli, A. M.; Bruner, M. W.; Marfia, K. E.; Tsai, P. K.; Sardana, M. K.; Middaugh, C. R. J. Pharm. Sci. 1995, 84, 7-11. (F9) Wood, T. D.; Chen, L. H.; Kelleher, N. L.; Little, D. P.; Kenyon, G. L.; MacLafferty, F. W. Biochemistry 1995, 34, 16251-4. (F10) Darrington, R. T.; Anderson, B. D. Pharm. Res. 1994, 11, 78493. (F11) Darrington, R. T.; Anderson, B. D. J. Pharm. Sci. 1995, 84, 275-82. (F12) Darrington, R. T.; Anderson, B. D. Pharm. Res. 1995, 12, 1077-84. (F13) Capasso, S.; Kirby, A. J.; Salvadori, S.; Sica, F.; Zagari, A. J. Chem. Soc., Perkin Trans. 2 1995, 437-42. (F14) Xie, M.; Vander Velde, D.; Morton, M.; Borchardt, R. T.; Schowen, R. L. J. Am. Chem. Soc. 1996, 118, 8955-6. (F15) Radkiewicz, J. L.; Zipse, H.; Clarke, S.; Houk, K. N. J. Am. Chem. Soc. 1996, 118, 9148-55. (F16) Orpiszewski, J.; Aswad, D. W. J. Biol. Chem. 1996, 271, 22965-8. (F17) Najbauer, J.; Orpiszewski, J.; Aswad, D. W. Biochemistry 1996, 35, 5183-90. (F18) Zhang, J.; Yueksel, K. U.; Gracy, R. W. Arch. Biochem. Biophys. 1995, 317, 112-20. (F19) Sun, A.-Q.; Yueksel, K. U.; Gracy, R. W. Arch. Biochem. Biophys. 1995, 322, 361-8. (F20) Delboni, L. F.; Mande, S. C.; Rentier-Delrue, F.; Mainfroid, V.; Turley, S.; Vellieux, F. M. D.; Martial, J. A.; Hol, V. G. J. Protein Sci. 1995, 4, 2594-604. (F21) Capasso, S.; Di Donato, A.; Esposito, L.; Sica, F.; Sorrentino, G.; Vitagliano, L.; Zagari, A.; Mazzarella, L. J. Mol. Biol. 1996, 257, 492-6. (F22) Miesbauer, R. L.; Zhou, X.; Yang, Z.; Yang, Z.; Sun, Y.; Smith, D. L.; Smith, J. B. J. Biol. Chem. 1994, 269, 12494-502. (F23) Xu, M.-Q.; Comb, D. G.; Paulus, H.; Noren, C. J.; Shao, Y.; Perler, F. EMBO J. 1994, 13, 5517-22. (F24) Shao, Y.; Xu, M.-Q.; Paulus, H. Biochemistry 1995, 34, 1084450. (F25) Levine, R. L.; Williams, J. A.; Stadtman, E. R.; Shacter, E. Methods Enzymol. 1994, 233, 346-57. (F26) Reznik, A. Z.; Packer, L. Methods Enzymol. 1994, 233, 35763. (F27) Cao, G.; Cutler, R. G. Arch. Biochem. Biophys. 1995, 320, 10614. (F28) Lyras, L.; Evans, P. J.; Shaw, P. J.; Ince, P. G.; Halliwell, B. Free Radical Res. 1996, 24, 397-406. (F29) Bowling, A. C.; Schulz, J. B.; Brown, R. H., Jr.; Beal, M. F. J. Neurochem. 1993, 61, 2322-5. (F30) Shaw, P. J.; Ince, P. G.; Falkous, G.; Mantle, D. Ann. Neurol. 1995, 38, 691-5. (F31) Shacter, E.; Williams, J. A.; Lim, M.; Levine, R. L. Free Radical Biol. Med. 1994, 17, 429-37. (F32) Sukalski, K. A.; LaBerge, T. P.; Johnson, W. T. Free Radical Biol. Med. 1997, 22, 835-42. (F33) Cabiscol, E.; Levine, R. L. J. Biol. Chem. 1995, 270, 14742-7. (F34) Oliver, C. N.; Ahn, B.; Moerman, E. J.; Goldstein, S.; Stadtman, E. R. J. Biol. Chem. 1987, 262, 5488-91. (F35) Stadtman, E. R.; Oliver, C. N. J. Biol. Chem. 1991, 266, 20058. (F36) Carney, J. M.; Starke-Reed, P. E.; Oliver, C. N.; Landum, R. W.; Cheng, M. S.; Wu, J. F.; Floyd, R. A. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 3633-6. (F37) Oliver, C. N.; Starke-Reed, P. E.; Stadtman, E. R.; Liu, G. J.; Carney, J. M.; Floyd, R. A. Proc. Natl. Acad. Sci. U.S.A. 1990, 87, 5144-7. (F38) Smith, C. D.; Carney, J. M.; Starke-Reed, P. E.; Oliver, C. N.; Stadtman, E. R.; Floyd, R. A.; Markesbery, W. R. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 10540-3. (F39) Smith, C. D.; Carney, J. M.; Tatsumo, T.; Stadtman, E. R.; Floyd, R. A.; Merkesbery, W. R. In Aging and Cellular Defense Mechanisms; Franceschi, C., Crepaldi, G., Cristofalo, V. J., Vijg, J., Eds.; Ann. N.Y. Acad. Sci. 1992; Vol. 663, pp 110-6. (F40) Sohal, R. S.; Sohal, B. H.; Orr, W. C. Free Radical Biol. Med. 1995, 19, 499-504. (F41) Tian, L.; Cai, Q.; Bown, R.; Wei, H. Free Radical Biol. Med. 1995, 19, 859-65. (F42) Cini, M.; Moretti, A. Neurobiol. Aging 1995, 16, 53-7. (F43) Ischiropoulos, H.; Al-Mehdi, A. B.; Fisher, A. B. Am. J. Physiol. 1995, 269, L158-64. (F44) Hensley, K.; Hall, N.; Subramaniam, R.; Cole, P.; Harris, M.; Aksenov, M.; Aksenova, M.; Gabbita, S. P.; Wu, J. F.; Carney, J. M.; Markesbery, W. R.; Butterfield, D. A. J. Neurochem. 1995, 65, 2146-56. (F45) Sohal, R. S.; Agarwal, S.; Sohal, B. H. Mech. Age Develop. 1995, 81, 15-25. (F46) Dubey, A.; Forster, M. J.; Sohal, R. S. Arch. Biochem. Biophys. 1995, 324, 249-54. (F47) Sato, E.; Oda, N.; Ozaki, N.; Hashimoto, S. I.; Kurokawa, T.; Ishibashi, S. Mech. Age. Develop. 1996, 86, 105-14. 56R
Analytical Chemistry, Vol. 69, No. 12, June 15, 1997
(F48) Yelinova, V.; Glazachev, Y.; Khramtsov, V.; Kudryashova, L.; Rykova, V.; Salganik, R. Biochem. Biophys. Res. Commun. 1996, 221, 300-3. (F49) Forster, M. J.; Dubey, A.; Dawson, K. M.; Stutts, W. A.; Lal, H.; Sohal, R. S. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 47659. (F50) Martinez, M.; Hernandez, A. I.; Martinez, N.; Ferrandiz, M. L. Brain Res. 1996, 731, 246-8. (F51) Dubey, A.; Forster, M. J.; Lal, H.; Sohal, R. S. Arch. Biochem. Biophys. 1996, 333, 189-97. (F52) Howard, B. J.; Yatin, S.; Hensley, K.; Allen, K. L.; Carney, J. M.; Butterfield, D. A. J. Neurochem. 1996, 67, 2045-50. (F53) Sohal, R. S.; Weindruch, R. Science 1996, 273, 59-63. (F54) Stadtman, E. R. Annu. Rev. Biochem. 1993, 62, 797-821. (F55) Davies, M. J.; Fu, S.; Dean, R. T. Biochem. J. 1995, 305, 6439. (F56) Simic, M. G. Methods Enzymol. 1990, 186, 89-100. (F57) Gebicki, S.; Gebicki, J. M. Biochem. J. 1993, 289, 743-9. (F58) Fu, S.; Hick, L. A.; Sheil, M. M.; Dean, R. T. Free Radical Biol. Med. 1995, 19, 281-92. (F59) Fu, S.; Gebicki, S.; Jessup, W.; Gebicki, J. M.; Dean, R. T. Biochem. J. 1995, 311, 821-7. (F60) Stadtman, E. R. Biochemistry 1990, 29, 6323-31. (F61) Uchida, K.; Szweda, L. I.; Chae, H. Z.; Stadtman, E. R. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 8742-6. (F62) Cohn, J. A.; Tsai, L.; Friguet, B.; Szweda, L. I. Arch. Biochem. Biophys. 1996, 328, 158-164. (F63) Fukuda, A.; Osawa, T.; Hitomi, K.; Uchida, K. Arch. Biochem. Biophys. 1996, 333, 419-26. (F64) Berliner, J. A.; Heinecke, J. W. Free Radical Biol. Med. 1996, 20, 707-27. (F65) Itabe, H.; Takeshima, E.; Iwasaki, H.; Kimura, J.; Yoshida, Y.; Imanaka, T.; Takano, T. J. Biol. Chem. 1994, 269, 15274-9. (F66) Itabe, H.; Yamamoto, H.; Suzuki, M.; Kawai, Y.; Nakagawa, Y.; Suzuki, A.; Imanaka, T.; Takano, T. J. Biol. Chem. 1996, 271, 33208-17. (F67) Itabe, H.; Yamamoto, H.; Imanaka, T.; Shimamura, K.; Uchyiama, H.; Kimura, J.; Sanaka, T.; Hata, Y.; Takano, T. J. Lipid Res. 1996, 37, 45-53. (F68) Uchida, K.; Toyokuni, S.; Nishikawa, K.; Kawakishi, S.; Oda, H.; Hiai, H.; Stadtman, E. R. Biochemistry 1994, 33, 1248794. (F69) Bolgar, M. S.; Yang, C.-H.; Gaskell, S. J. J. Biol. Chem. 1996, 271, 27999-8001. (F70) Bolgar, M. S.; Gaskell, S. J. Anal. Chem. 1996, 68, 2325-30. (F71) Al-Abed, Y.; Liebich, H.; Voelter, W.; Bucala, R. J. Biol. Chem. 1996, 271, 2892-6. (F72) Chen, P.; Wiesler, D.; Chmelı´k, J.; Novotny, M. Chem. Res. Toxicol. 1996, 9, 970-9. (F73) Fu, M.-X.; Requena, J. R.; Jenkins, A. J.; Lyons, T. J.; Baynes, J. W.; Thorpe, S. R. J. Biol. Chem. 1996, 271, 9982-6. (F74) Bucala, R.; Mitchell, R.; Arnold, K.; Innerarity, T.; Vlassara, H.; Cerami, A. J. Biol. Chem. 1995, 270, 10828-32. (F75) Hunt, J. V. Redox Rep. 1996, 2, 5-8. (F76) Nakamura, Y.; Horii, Y.; Nishino, T.; Shiiki, H.; Sakaguchi, Y.; Kagoshima, T.; Dohi, K.; Makita, Z,; Vlassara, H.; Bucala, R. Am. J. Pathol. 1993, 143, 1649-56. (F77) Kume, S.; Takeya, M.; Mori, T.; Araki, N.; Suzuki, H.; Horiuchi, S.; Kodama, T.; Miyauchi, Y.; Takahashi, K. Am. J. Pathol. 1995, 147, 654-67. (F78) Reddy, S.; Bichler, J.; Wells-Knecht, K. J.; Thorpe, S. R.; Baynes, J. W. Biochemistry 1995, 34, 10872-8. (F79) Ikeda, K.; Higashi, T.; Sano, H.; Jinnouchi, Y.; Yoshida, M.; Araki, T.; Ueda, S.; Horiuchi, S. Biochemistry 1996, 35, 807583. (F80) Burlet, O.; Yang, C.-Y.; Guyton, J. R.; Gaskell, S. J. J. Am. Soc. Mass Spectrom. 1995, 6, 242-7. (F81) Hazell, L.; Arnold, L.; Flowers, D.; Waeg, G.; Malle, E.; Stocker, R. J. Clin. Invest. 1996, 97, 1535-44. (F82) Leeuwenburgh, C.; Hardy, M. M.; Hazem, S. L.; Wagner, P.; Oh-ishi, S.; Steinbrecher, U. P.; Heinecke, J. W. J. Biol. Chem. 1997, 272, 1433-6. (F83) Giessauf, A.; Steiner, E.; Esterbauer, H. Biochim. Biophys. Acta 1995, 1256, 221-32. (F84) Meyer, D. F.; Mayans, M. O.; Groot, P. H. E.; Suckling, K. E.; Bruckdorfer, K. R.; Perkins, S. J. Biochem. J. 1995, 310, 41726. (F85) Meyer, D. F.; Nealis, A. S.; MacPhee, C. H.; Groot, P. H. E.; Suckling, K. E.; Bruckdorfer, K. R.; Perkins, S. J. Biochem. J. 1996, 319, 217-27. (F86) Lapolla, A.; Fedele, D.; Catinella, S.; Traldi, P. NATO ASI Ser., Ser. C 1996, 475, 317-49. (F87) Wells-Knecht, K. J.; Zyzak, D. V.; Litchfield, J. E.; Thorpe, S. R.; Baynes, J. W. Biochemistry 1995, 34, 3702-9. (F88) Wells-Knecht, K. J.; Brinkmann, E.; Baynes, J. W. J. Org. Chem. 1995, 60, 6246-7. (F89) Brinkmann, E.; Wells-Knecht, K. J.; Thorpe, S. R.; Baynes, J. W. J. Chem. Soc., Perkin Trans. 1 1995, 2817-8. (F90) Nagaraj, R. H.; Shipanova, I. N.; Faust, F. M. J. Biol. Chem. 1996, 271, 19338-45. (F91) Nagaraj, R. H.; Monnier, V. M. Biochim. Biophys. Acta 1995, 1253, 75-84. (F92) DiFranco, E.; Subbanagounder, G.; Kim, S.; Murthi, K.; Taneda, S.; Monnier, V. M. Chem. Res. Toxicol. 1995, 8, 61-7. (F93) Portero-Otin, M.; Nagaraj, R. H.; Monnier, V. M. Biochim. Biophys. Acta 1995, 1247, 74-80.
(F94) Wells-Knecht, M. C.; Thorpe, S. R.; Baynes, J. W. Biochemistry 1995, 34, 15134-41. (F95) Glomb, M. A.; Monnier, V. M. J. Biol. Chem. 1995, 270, 10017-26. (F96) Elgawish, A.; Glomb, M.; Friedlander, M.; Monnier, V. M. J. Biol. Chem. 1996, 271, 12964-71. (F97) Khalifah, R. G.; Todd, P.; Booth, A. A.; Yang, S. X.; Mott, J. D.; Hudson, B. G. Biochemistry 1996, 35, 4645-54. (F98) Zyzak, D. V.; Richardson, J. M.; Thorpe, S. R.; Baynes, J. W. Arch. Biochem. Biophys. 1995, 316, 547-54. (F99) Horiuchi, T.; Kurokawa, T.; Saito, N. Agric. Biol. Chem. 1989, 53, 103-10. (F100) Horiuchi, T.; Kurokawa, T. Agric. Biol. Chem. 1991, 55, 3338. (F101) Gerhardinger, C.; Marion, M. S.; Rovner, A.; Glomb, M.; Monnier, V. M. J. Biol. Chem. 1995, 270, 218-24. (F102) Yim, H.-S.; Kang, S.-O.; Hah, Y.-C.; Chock, P. B.; Yim. M. B. J. Biol. Chem. 1995, 270, 28228-33. (F103) Cervantes-Laurean, D.; Jacobson, E. L.; Jacobson, M. K. J. Biol. Chem. 1996, 271, 10461-9. (F104) Vasan, S.; Zhang, X.; Zhang, X.; Kapurniotu, A.; Bernhagen, J.; Teichberg, S.; Basgen, J.; Wagle, D.; Shih, D.; Terlecki, I.; Bucala, R.; Cerami, A.; Egan, J.; Ulrich, P. Nature 1996, 382, 275-8. (F105) Selkoe, D. J. J. Biol. Chem. 1996, 271, 18295. (F106) Butterfield, D. A.; Hensley, K.; Harris, M.; Mattson, M.; Carney, J. M. Biochem. Biophys. Res. Commun. 1994, 200, 710-5. (F107) Mattson, M. P. Neurobiol. Aging 1995, 16, 679-82. (F108) Aksenov, M. Y.; Aksenova, M. V.; Harris, M. E.; Hensley, K.; Butterfield, D. A.; Carney, J. M. J. Neurochem. 1995, 65, 1899902. (F109) Mark, R. J.; Hensley, K.; Butterfield, D. A.; Mattson, M. P. J. Neurosci. 1995, 15, 6239-49. (F110) Harris, M. E.; Hensley, K.; Butterfield, D. A.; Leedle, R. A.; Carney, J. M. Exp. Neurol. 1995, 131, 193-202. (F111) Barger, S. W.; Ho¨rster, D.; Furukawa, K.; Goodman, Y.; Krieglstein, J.; Mattson, M. P. Proc. Natl. Acad. Sci. U.S.A. 1995, 92, 9328-32. (F112) Butterfield, D. A.; Martin, L.; Carney, J. M.; Hensley, K. Life Sci. 1996, 58, 217-28. (F113) Yan, S. D.; Chen, X.; Fu, J.; Chen, M.; Zhu, H.; Roher, A.; Slattery, T.; Zhao, L.; Nagashima, M.; Morser, J.; Migheli, A.; Nawroth, P.; Stern, D.; Schmidt, A. M. Nature 1996, 382, 68591. (F114) Stamler, J. S.; Singel, D. J.; Loscalzo, J. Science 1992, 258, 1898-902. (F115) Rubbo, H.; Darley-Usmar, V.; Freeman, B. A. Chem. Res. Toxicol. 1996, 9, 809-20. (F116) Ischiropoulos, H.; Beckman, J. S.; Crow, J. P.; Ye, Y. Z.; Royall, J. A.; Kooy, N. W. Methods 1995, 7, 109-15. (F117) Crow, J. P.; Beckman, J. S. Methods 1995, 7, 116-20. (F118) Beckman, J. S. Chem. Res. Toxicol. 1996, 9, 836-44. (F119) Greis, K. D.; Zhu, S.; Matalon, S. Arch. Biochem. Biophys. 1996, 335, 396-402. (F120) Beckman, J. S.; Ye, Y. Z.; Anderson, P. G.; Chen, J.; Accavitti, M. A.; Tarpey, M. M.; White, C. R. Biol. Chem. Hoppe-Seyler 1994, 375, 81-8. (F121) Ischiropoulos, H.; Al-Mehdi, A. B.; Fisher, A. B. Am. J. Physiol. 1995, 269, L158-64.
(F122) Schulz, J. B.; Matthews, R. T.; Muqit, M. M. K.; Browne, S. E.; Beal, M. F. J. Neurochem. 1995, 64, 936-9. (F123) Schulz, J. B.; Matthews, R. T.; Jenkins, B. G.; Ferrante, R. J.; Siwek, D.; Henshaw, D. R.; Cipolloni, P. B.; Mecocci, P.; Kowall, N. W.; Rosen, B. R.; Beal, M. F. J. Neurosci. 1995, 15, 841929. (F124) Matthews, R. T.; Beal, M. F. Brain Res. 1996, 718, 181-4. (F125) Evans, T. J.; Buttery, L. D. K.; Carpenter, A.; Springall, D. R.; Polak, J. M.; Cohen, J. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 9553-8. (F126) MacMillan-Crow, L. A.; Crow, J. P.; Kerby, J. D.; Beckman, J. S.; Thompson, J. A. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 11853-8. (F127) Fukuyama, N.; Takebayashi, Y.; Hida, M.; Ishida, H.; Ichimori, K.; Nakazawa, H. Free Radical Biol. Med. 1997, 22, 771-4. (F128) Viner, R. I.; Ferringto, D. A.; Hu ¨ hmer, A. F. R.; Bigelow, D. J.; Scho¨neich, Ch. FEBS Lett. 1996, 379, 286-90. (F129) Eiserich, J. P.; Cross, C. E.; Jones, A. D.; Halliwell, B.; van der Vliet, A. J. Biol. Chem. 1996, 271, 19199-208. (F130) Lymar, S. V.; Jiang, Q.; Hurst, J. K. Biochemistry 1996, 35, 7855-61. (F131) Gow, A.; Duran, D.; Thom, S. R.; Ischiropoulos, H. Arch. Biochem. Biophys. 1996, 333, 42-8. (F132) Denicola, A.; Freeman, B. A.; Trujillo, M.; Radi, R. Arch. Biochem. Biophys. 1996, 333, 49-58. (F133) Lymar, S. V.; Hurst, J. K. Chem. Res. Toxicol. 1996, 9, 84550. (F134) Van der Vliet, A.; Eiserich, J. P.; O’Neill, C. A.; Halliwell, B.; Cross, C. E. Arch. Biochem. Biophys. 1995, 319, 341-9. (F135) Ramezanian, M. S.; Padmaja, S.; Koppenol, W. H. Chem. Res. Toxicol. 1996, 9, 232-40. (F136) Gow, A. J.; Duran, D.; Malcolm, S.; Ischiropoulos, H. FEBS Lett. 1996, 385, 63-6. (F137) Kong, S. K.; Yim, M. B.; Stadtman, E. R.; Chock, P. B. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 3377-82. (F138) Van der Vliet, A.; O’Neill, C. A.; Halliwell, B.; Cross, C. E.; Kaur, H. FEBS Lett. 1994, 339, 89-92. (F139) Berlett, B. S.; Friguet, B.; Yim, M. B.; Chock, P. B.; Stadtman, E. R. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 1776-80. (F140) Alvarez, B.; Rubbo, H.; Kirk, M.; Barnes, S.; Freeman, B. A.; Radi, R. Chem. Res. Toxicol. 1996, 9, 390-6. (F141) Gatti, R. M.; Radi, R.; Augusto, O. FEBS Lett. 1994, 348, 28790. (F142) Crow, J. P.; Beckman, J. S.; McCord, J. M. Biochemistry 1995, 34, 3544-52. (F143) Pryor, W. A.; Squadrito, G. L. Am. J. Physiol. 1995, 268, L699722. (F144) Padmaja, S.; Squadrito, G. L.; Lemercier, J.-N.; Cueto, R.; Pryor, W. A. Free Radical Biol. Med. 1996, 21, 317-22. (F145) Radi, R. Chem. Res. Toxicol. 1996, 9, 828-35. (F146) Jia, L.; Bonaventura, C.; Bonaventura, J.; Stamler, J. S. Nature 1996, 380, 221-6. (F147) Perutz, M. Nature 1996, 380, 205-6. (F148) Gow, A. J.; Buerk, D. G.; Ischiropoulos, H. J. Biol. Chem. 1997, 272, 2841-5.
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