Quantitative Assessment of In-solution Digestion Efficiency Identifies Optimal Protocols for Unbiased Protein Analysis

The majority of mass spectrometry-based protein quantification studies uses peptide-centric analytical methods and thus strongly relies on efficient and unbiased protein digestion protocols for sample preparation. We present a novel objective approach to assess protein digestion efficiency using a combination of qualitative and quantitative liquid chromatography-tandem MS methods and statistical data analysis. In contrast to previous studies we employed both standard qualitative as well as data-independent quantitative workflows to systematically assess trypsin digestion efficiency and bias using mitochondrial protein fractions. We evaluated nine trypsin-based digestion protocols, based on standard in-solution or on spin filter-aided digestion, including new optimized protocols. We investigated various reagents for protein solubilization and denaturation (dodecyl sulfate, deoxycholate, urea), several trypsin digestion conditions (buffer, RapiGest, deoxycholate, urea), and two methods for removal of detergents before analysis of peptides (acid precipitation or phase separation with ethyl acetate). Our data-independent quantitative liquid chromatography-tandem MS workflow quantified over 3700 distinct peptides with 96% completeness between all protocols and replicates, with an average 40% protein sequence coverage and an average of 11 peptides identified per protein. Systematic quantitative and statistical analysis of physicochemical parameters demonstrated that deoxycholate-assisted in-solution digestion combined with phase transfer allows for efficient, unbiased generation and recovery of peptides from all protein classes, including membrane proteins. This deoxycholate-assisted protocol was also optimal for spin filter-aided digestions as compared with existing methods.MS-based proteomics is an indispensable technology for the characterization of complex biological systems, including relative or absolute protein expression levels and protein post-translational modifications. The most popular method for analyzing medium to high complexity protein samples in large-scale proteomics relies on protein digestion by using the endoprotease trypsin. Analysis and sequencing of tryptic peptides by liquid chromatography-tandem MS (LC-MS/MS)¹ then enables identification and determination of protein expression levels based on the peptide ion abundance level or the (fragment) ion intensities of identified peptides. This peptide-centric approach thus strongly relies on efficient, unbiased and reproducible protein digestion protocols. Efficiency is required to maximize the number of detectable peptides per protein (coverage) to distinguish unique proteins within protein families with similar sequences and/or sequence variants, and to detect post-translational modifications. Unbiased generation of peptides is required for the resulting data set to most accurately reflect the relative (stoichiometry) and absolute protein abundance in a sample. A particular protocol should be unbiased with respect to abundance, molecular weight, hydrophobicity and protein class. Membrane proteins for example are often suspected to be underrepresented. For MS-based proteomics approaches several critical steps can be distinguished: (a) disruption and solubilization of cells and protein complexes, (b) protein denaturation and enzymatic proteolysis, (c) MS-compatible peptide recovery, which normally entails removal of reagent leftovers and desalting before MS analysis, (d) adequate peptide separation (achieved by liquid chromatography), and (e) MS peptide analysis and sequencing (MS/MS), including the chosen data acquisition strategy.Comparative evaluations of digestion protocols generally consist of qualitative studies using standard tandem mass spectrometry. These approaches may reveal efficiency (i.e. more identifications), but are unable to reveal digestion protocol induced bias with respect to peptide and protein abundance, including membrane proteins. In addition, most data-dependent acquisition workflows are intrinsically biased, which is detrimental for making comparisons. The aim of the present study was to systematically assess efficiency and bias of trypsin-based protocols applying both standard qualitative and label-free quantitative MS approaches.The in-gel digestion protocol for proteomics, established over 15 years ago (1), has been the cornerstone method affording robust protein identifications from many sample types. Although sodium dodecyl sulfate (SDS) interferes with trypsin digestion and hampers LC-MS analysis, this powerful detergent can still be used to achieve complete protein solubilization as gel-separation is an effective way to remove interfering substances. Gel-based approaches are however not optimal for protein samples of increasing complexity and dynamic range (2). Inherent and practical limitations include, for example, concentration-dependent, incomplete peptide recovery and error-prone handling procedures (3–6). This hampers throughput, reproducibility and unbiased protein analysis, which in recent years has prompted a shift toward the application and optimization of in-solution digestion procedures.Previous comparative studies revealed that for in-solution digestions, the acid labile and MS-compatible detergent RapiGest performed most favorably compared with buffer only, urea, other detergents and organic solvents (7–9). Sodium deoxycholate (SDC), naturally found in mammalian bile (10), has emerged as a cheaper MS-compatible detergent for in-solution digestion (11). Unlike other detergents, SDC was found to enhance trypsin activity almost fivefold at a concentration of 1% (12). Like RapiGest, SDC can also be removed by acidification, but potentially without detrimental peptide loss if a phase separation protocol involving organic solvent is applied (12).An alternative strategy is to perform protein digestion on spin filter devices, introduced a few years ago by Manza and co-workers (13), and further developed by Wisniewski et al. (14). This approach allows the use of SDS to first achieve complete protein solubilization followed by removal of the detergent through repeated washes with urea (14). This is an effective way to remove interfering chemicals and small molecules after protein solubilization, and before digestion, without substantial sample loss. Although this protocol is touted to be a highly effective and universal method for any type of sample, digestion is performed using urea or buffer only and has so far not been evaluated in combination with detergents such as SDC.For our comparative study we selected protocols and methods based on spin filter-aided and standard in-solution digestion that were previously reported optimal and we also report novel optimized protocols. We investigated several experimental parameters including reagents for protein solubilization and denaturation (SDS, SDC, urea), spin filter aided removal of SDS before digestion (urea, SDC, buffer), trypsin digestion conditions (buffer, RapiGest, SDC, urea), and methods for removal of detergents before analysis of peptides (acid precipitation or phase separation with ethyl acetate).Mitochondria are organelles carrying out key metabolic processes fundamental for cellular function (15). The mitochondrial proteome is predicted to contain up to a thousand proteins (16) and is very heterogeneous with a wide range of protein pI, molecular weight and hydrophobicity values (17). We selected mitochondrial preparations to serve as model sample of medium complexity, containing a favorable combination of peptide and protein classes, including soluble and insoluble membrane-anchored or integral proteins.Using standard qualitative as well as data-independent quantitative LC-MS/MS workflows we demonstrate that SDC-based protocols combined with phase separation are the most optimal for both in-solution and filter-aided tryptic digestion, yielding the highest efficiency and lowest bias. This workflow enabled quantitative and objective assessment of various protein digestion conditions, identifying optimal protocols for efficient and unbiased protein analysis.     

Phosphotyrosine profiling of human cerebrospinal fluid

Background

Cerebrospinal fluid (CSF) is an important source of potential biomarkers that affect the brain. Biomarkers for neurodegenerative disorders are needed to assist in diagnosis, monitoring disease progression and evaluating efficacy of therapies. Recent studies have demonstrated the involvement of tyrosine kinases in neuronal cell death. Thus, neurodegeneration in the brain is related to altered tyrosine phosphorylation of proteins in the brain and identification of abnormally phosphorylated tyrosine peptides in CSF has the potential to ascertain candidate biomarkers for neurodegenerative disorders.

Methods

In this study, we used an antibody-based tyrosine phosphopeptide enrichment method coupled with high resolution Orbitrap Fusion Tribrid Lumos Fourier transform mass spectrometer to catalog tyrosine phosphorylated peptides from cerebrospinal fluid. The subset of identified tyrosine phosphorylated peptides was also validated using parallel reaction monitoring (PRM)-based targeted approach.

Results

To date, there are no published studies on global profiling of phosphotyrosine modifications of CSF proteins. We carried out phosphotyrosine profiling of CSF using an anti-phosphotyrosine antibody-based enrichment and analysis using high resolution Orbitrap Fusion Lumos mass spectrometer. We identified 111 phosphotyrosine peptides mapping to 66 proteins, which included 24 proteins which have not been identified in CSF previously. We then validated a set of 5 tyrosine phosphorylated peptides in an independent set of CSF samples from cognitively normal subjects, using a PRM-based targeted approach.

Conclusions

The findings from this deep phosphotyrosine profiling of CSF samples have the potential to identify novel disease-related phosphotyrosine-containing peptides in CSF.

     

Rapid isolation of desired sequences from lone linker PCR amplified cDNA mixtures: Application to identification and recovery of expressed sequences in cloned genomic DNA

A simple and efficient method for the rapid isolation of specific sequences from PCR-amplified cDNA mixtures has been developed. cDNA mixtures obtained using lone linker PCR (Ko et al. 1990) appeared to be highly representative even though the starting material, 100 ng-2 g of total RNA, is much less than is required for making an ordinary cDNA library. With this method, cDNA mixtures were obtained from limited materials, including early mouse embryos and primordial germ cells. For selective enrichment of desired cDNAs, biotinylated probe was hybridized with the lone linker-linked cDNA in solution and the resulting probe-cDNA hybrid was captured by Streptavidin-coated magnetic beads. After appropriate washing, cDNA was released from the beads and subjected to amplification followed by cloning into a vector. Using genomic fragments isolated during chromosomal walking in the T/t complex of mouse Chromosome (Chr) 17, cDNAs encoding novel germ cell specific genes have been readily isolated by the above procedures. The method, termed random access retrieval of genetic information through PCR (RARGIP), will streamline the entire process from RNA to cDNA greatly. Its application potentials in various areas of molecular genetics will be discussed.     

Refining the Definition of Plant Mitochondrial Presequences through Analysis of Sorting Signals,N-Terminal Modifications,and Cleavage Motifs

Mitochondrial protein import is a complex multistep process from synthesis of proteins in the cytosol, recognition by receptors on the organelle surface, to translocation across one or both mitochondrial membranes and assembly after removal of the targeting signal, referred to as a presequence. In plants, import has to further discriminate between mitochondria and chloroplasts. In this study, we determined the precise cleavage sites in the presequences for Arabidopsis (Arabidopsis thaliana) and rice (Oryza sativa) mitochondrial proteins using mass spectrometry by comparing the precursor sequences with experimental evidence of the amino-terminal peptide from mature proteins. We validated this method by assessments of false-positive rates and comparisons with previous available data using Edman degradation. In total, the cleavable presequences of 62 proteins from Arabidopsis and 52 proteins from rice mitochondria were determined. None of these proteins contained amino-terminal acetylation, in contrast to recent findings for chloroplast stromal proteins. Furthermore, the classical matrix glutamate dehydrogenase was detected with intact and amino-terminal acetylated sequences, indicating that it is imported into mitochondria without a cleavable targeting signal. Arabidopsis and rice mitochondrial presequences had similar isoelectric points, hydrophobicity, and the predicted ability to form an amphiphilic α-helix at the amino-terminal region of the presequence, but variations in length, amino acid composition, and cleavage motifs for mitochondrial processing peptidase were observed. A combination of lower hydrophobicity and start point of the amino-terminal α-helix in mitochondrial presequences in both Arabidopsis and rice distinguished them (98%) from Arabidopsis chloroplast stroma transit peptides. Both Arabidopsis and rice mitochondrial cleavage sites could be grouped into three classes, with conserved −3R (class II) and −2R (class I) or without any conserved (class III) arginines. Class II was dominant in both Arabidopsis and rice (55%–58%), but in rice sequences there was much less frequently a phenylalanine (F) in the −1 position of the cleavage site than in Arabidopsis sequences. Our data also suggest a novel cleavage motif of (F/Y)↓(S/A) in plant class III sequences.Plant mitochondria play a key role in energy production and metabolism that requires the import and assembly of at least 1,000 proteins. Protein import into mitochondria begins with synthesis of the precursor protein in the cytosol, followed by binding to various proteins in the cytosol, binding to receptors on the outer mitochondrial membrane, translocation across one or both mitochondrial membranes, removal of the targeting signal, termed a presequence, and intraorganellar sorting and assembly. A variety of studies have shown that there is no primary amino acid sequence conservation among presequences, but they do have a high proportion of positively charged residues and the capacity to form an amphiphilic α-helix (Roise et al., 1986; von Heijne, 1986). Many mitochondrial presequences have a loosely conserved motif near the cleavage site comprising an Arg residue at the −2 and/or −3 position (von Heijne et al., 1989; Schneider et al., 1998). This Arg has been experimentally shown to be an important recognition site for the mitochondrial processing peptidase (MPP; Arretz et al., 1994; Ogishima et al., 1995; Tanudji et al., 1999). MPP is a heterodimeric enzyme that contains two similar subunits: α-MPP is involved in binding precursor proteins and β-MPP catalyzes the cleavage of the presequence (Kitada et al., 1995; Luciano et al., 1997). In yeast and mammals, MPP is a soluble protein located in the matrix, but in plants, MPP is integrated into the inner membrane-bound cytochrome b/c₁ complex (Braun et al., 1992; Eriksson et al., 1994; Glaser and Dessi, 1999).The mechanism through which the targeting signal binds to a receptor protein has been revealed by NMR studies and the crystal structure of rat Tom20 (for translocase of the outer membrane) with a bound presequence (Abe et al., 2000; Saitoh et al., 2007). A dynamic binding model in which different hydrophobic residues in the presequence interact with Tom20 has been proposed. Thus, the presequence has mobility in the binding site via hydrophobic interactions, with several different binding states being possible. This model accounts for the ability of a single Tom20 in yeast to bind to a diverse array of presequences. Although plants contain a protein that is called Tom20 and that has a receptor function in mitochondrial import, it is not orthologous to yeast or mammalian Tom20 (Perry et al., 2006; Lister et al., 2007). However, the NMR structure of plant Tom20 reveals a similar hydrophobic binding pocket. This has been highlighted as a case of convergent evolution of a receptor that uses a similar mechanism of binding to recognize presequences (Lister and Whelan, 2006). Although structural studies reveal the importance of hydrophobic residues for presequence binding, several studies on yeast, mammals, and plants reveal an important role for positively charged residues in presequences for import into mitochondria (Lister et al., 2005; Neupert and Herrmann, 2007). These positively charged residues may play a role in positioning the amphiphilic α-helix for binding to Tom20 and also in subsequent translocation into and across the pores forming proteins of the TOM and TIM (for translocase of the inner membrane) complexes. Movement of the presequence into and across a translocase is explained by the binding chain hypothesis (Pfanner and Geissler, 2001). According to this hypothesis, a presequence binds to higher affinity sites in the import apparatus until it is “trapped” on the inside of the inner membrane by a combination of electrostatic interactions, the net negative charge on the inside of the inner membrane, and binding to matrix-located HSP70 (Zhang and Glaser, 2002).In addition to the fact that plant Tom20s are not orthologous to other Tom20s, plant mitochondria also lack the other two receptor components that have been functionally characterized in yeast, namely Tom70 and Tom22 (Lister et al., 2007). Furthermore, mitochondrial and plastid targeting signals contain significant similarities in plants; thus, plant mitochondrial presequences have evolved to differentiate from the large number and abundant nature of plastid proteins requiring import from the cytosol (Macasev et al., 2000). This raises the question of how similar plant mitochondrial targeting signals are to those of yeast and how they are differentiated from plastid transit peptides. To adequately address these questions, a large number of presequences need to be assembled to define motifs that differentiate presequence classes. Traditionally, the N-terminal sequences of plant mitochondrial proteins have been obtained by Edman degradation either from purified mitochondrial protein complexes or in proteome studies (Braun and Schmitz, 1995; Jänsch et al., 1996; Millar et al., 1998, 1999; Kruft et al., 2001; Bardel et al., 2002). The presequences could only be obtained by comparison of these N-terminal sequences with the preprotein sequence deduced from full-length cDNA sequences, which were only available in a small number of cases. Glaser et al. (1998) presented a list of approximately 100 plant mitochondrial presequences; these were mainly derived from prediction and/or comparisons in homologous cDNA-derived protein sequences with a core set of 31 experimentally proven presequences for plant mitochondrial proteins. Later analysis of 58 experimentally proven plant mitochondrial presequences deposited in the Swiss-Prot database revealed two major classes containing an Arg residue at positions −2 and −3 and one class without any conserved Arg residues (Zhang et al., 2001; Zhang and Glaser, 2002). However, this data set relied on the sequences available at the time that were from a variety of plant species and contained redundant orthologs from similar proteins. This data set also clearly focused on dicot plants, as less than 20% of the sequences were from monocot species.In the chloroplast, N-terminal modification of chloroplast proteins has been shown to be important for protein viability (Pesaresi et al., 2003). N-terminal acetylation can be detected by high-resolution mass spectrometry (MS) through a change in mass of the N-terminal peptide. The recent systematic analysis of the Arabidopsis (Arabidopsis thaliana) chloroplast proteome revealed 47 stroma proteins with N-acetylated residues and 62 without N-acetylated residues (Zybailov et al., 2008). The detection of N-terminal and non-N-terminal acetylated proteins by identifications of semitryptic peptides also allowed analysis of the cleavage sites and potential motifs for cleavage by processing peptidases (Zybailov et al., 2008). However, no systematic experimental analysis of N-terminal modifications and potential cleavage sites of plant mitochondrial proteins has been carried out to date using such an MS approach.In this study, we have determined Arabidopsis and rice (Oryza sativa) mitochondrial protein-targeting presequences and cleavage sites using an MS approach after gel- or liquid chromatography (LC)-based separation and also identified a range of N-terminal modifications of mitochondrial proteins. Validation of this method was performed by false-positive analysis and comparison with previous results in Arabidopsis using an Edman degradation approach (Kruft et al., 2001). We compared the characteristics of the generated Arabidopsis and rice mitochondrial presequences and the cleavage site motifs. Comparison with experimentally proven yeast mitochondrial presequences and Arabidopsis plastid stroma transit peptides allowed consideration of some evolutionary questions and insights into the different signal-recognizing mechanism(s) used to distinguish between organelles.     

SISCAPA Peptide Enrichment on Magnetic Beads Using an In-line Bead Trap Device

A SISCAPA (stable isotope standards and capture by anti-peptide antibodies) method for specific antibody-based capture of individual tryptic peptides from a digest of whole human plasma was developed using a simplified magnetic bead protocol and a novel rotary magnetic bead trap device. Following off-line equilibrium binding of peptides by antibodies and subsequent capture of the antibodies on magnetic beads, the bead trap permitted washing of the beads and elution of bound peptides inside a 150-μm-inner diameter capillary that forms part of a nanoflow LC-MS/MS system. The bead trap sweeps beads against the direction of liquid flow using a continuous succession of moving high magnetic field-gradient trap regions while mixing the beads with the flowing liquid. This approach prevents loss of low abundance captured peptides and allows automated processing of a series of SISCAPA reactions. Selected tryptic peptides of α₁-antichymotrypsin and lipopolysaccharide-binding protein were enriched relative to a high abundance serum albumin peptide by 1,800 and 18,000-fold, respectively, as measured by multiple reaction monitoring. A large majority of the peptides that are bound nonspecifically in SISCAPA reactions were shown to bind to components other than the antibody (e.g. the magnetic beads), suggesting that substantial improvement in enrichment could be achieved by development of improved inert bead surfaces.MS is the method of choice for identification of peptides in digests of biological samples based on the power of MS to detect the chemically well defined masses of both peptides and their fragments produced by processes such as CID. This high level of structural specificity is also critical in improving peptide (and protein) quantitation because it overcomes the well known problems inherent in classical immunoassays related to limited antibody specificity, dynamic range, and multiplexability. In principle, a quantitative peptide assay using MRM¹ detection in a triple quadrupole mass spectrometer should have nearly absolute structural specificity, a dynamic range of ∼1e+4, and the ability to multiplex measurements of hundreds of peptides per sample (1). These properties suggest that MS-based methods could ultimately replace classical immunoassay technologies in many research and clinical applications.An important limitation of present peptide MRM measurements is sensitivity. The most sensitive widely used quantitative MS platforms use nanoflow chromatography and ESI to deliver trace amounts of peptides to the mass spectrometer. However, these processes are limited in the total amount of peptide that can be applied while retaining maximum sensitivity (typically limited to ∼1 μg of total peptide sample, i.e. the product obtained from digesting ∼14 nl of plasma). The lower cutoff for detecting proteins in a digest of unfractionated plasma by this approach appears to be in the neighborhood of 1–20 μg/ml plasma concentration, which would restrict analysis to the top 100 or so proteins in plasma (1).The sensitivity of MS assays can be substantially increased by fractionating the sample at the level of intact proteins, the tryptic peptides derived from them, or both. For example, immunodepletion of the six most abundant plasma proteins, removes ∼85% of the protein mass (2) and results in an increase of ∼7-fold in the signal-to-noise of MRM measurements of peptides from the remaining proteins after digestion (1). Similarly chromatographic fractionation by strong cation exchange provides another major improvement in sensitivity (3). However, increased sample fractionation brings with it the disadvantages of increased cost and time, the risk of losing specific components, and the continued requirement for very high resolution (lengthy, low throughput) reversed phase nanoflow chromatography en route to the ESI source.An alternative fractionation approach, used in the SISCAPA method, enriches specific target peptides through capture by anti-peptide antibodies, thus circumventing these disadvantages for preselected targets (4). In its initial implementation, SISCAPA used very small (∼10-nl) columns of POROS chromatography support carrying covalently bound rabbit antibodies and provided ∼100-fold enrichment of target peptides with respect to others (4). These columns were, like immunoaffinity depletion columns (2), recyclable many times. However, the potential for sample-to-sample carryover, limitations in the amount of sample digest that could be pumped over nanoaffinity columns at flow rates slow enough to permit peptide binding, and limited flexibility in changing and multiplexing antibodies were problematic. This led us to explore an alternative approach using magnetic beads as the antibody support (5). In this case, the binding reaction can be carried out off line, allowing equilibrium binding; the magnetic beads can be removed from the digest sample and washed; and the bound peptides can be eluted in 96-well plates either manually or using automated equipment such as a KingFisher Magnetic Particle Processor (ThermoFisher). One potential pitfall remains in the handling of eluted peptides. If the anti-peptide antibodies have very high selectivity, as desired in the SISCAPA approach, then in the case of low abundance peptides, only a very small amount of peptide will be eluted from the antibody. Such small amounts of peptide are easily lost through irreversible binding to the walls of vessels such as 96-well plate wells, and the smaller the amount of peptide (i.e. the more specific the capture), the worse the problem may be.To address this issue, we report here a hybrid approach in which peptide binding occurs off line (to equilibrium), whereas the subsequent washing and elution steps are carried out within a capillary that forms part of the nanoflow LC system, thus ensuring that peptide eluted from the antibodies on the beads will not be “lost” between elution and the ESI source. Although there is extensive literature on macroscopic and microfluidic devices for manipulating magnetic beads (6–8) we were unable to find components adaptable to the small scales and high pressures required for integration into nanoflow HPLC. We therefore developed a novel “bead trap” device that satisfies the following requirements: 1) the need to retain beads in a “trap” region against the flow of liquid (loading, wash, and elution buffers for example) in a vessel of capillary dimensions, 2) the need to ensure that beads do not escape from the trap region to contaminate downstream apparatus or columns, 3) the need to ensure that beads are effectively mixed with the flowing fluids (required for efficient washing and elution), and 4) the need to ensure that all beads can be efficiently ejected from the trap region in preparation for a subsequent cycle. The device provides multiple sequential magnetic trapping regions capable of sweeping commonly used 2.8- and 1-μm magnetic beads against liquid flow to prevent escape of beads through the trapping device (i.e. the second downstream trapping zone captures beads swept by the liquid stream past the first trap and so on). In addition, the bead trap device allows the movement of these trapping regions to agitate the trapped bead mass and mix it with fluids flowing past. Finally the device allows reversal of the sweeping action to effectively eject beads from the trap into the fluid stream. The bead trap capillary can be plumbed at various points in conventional nanoflow LC systems (e.g. in place of a sample loop or connecting tube), and the device can be controlled directly by the LC-MS/MS instrument software through contact closures. We show that the bead trap provides an effective method of implementing SISCAPA experiments.     

Flexibility contra Stiffness: The Phragmoplast as a Physical Barrier for Beads But Not for Vesicles

In plant cells, Golgi vesicles are transported to the division plane to fuse with each other, forming the cell plate, the initial membrane-bordered cell wall separating daughter cells. Vesicles, but not organelles, move through the phragmoplast, which consists of two opposing cylinders of microtubules and actin filaments, interlaced with endoplasmic reticulum membrane. To study physical aspects of this transport/inhibition process, we microinjected fluorescent synthetic 1,2-dioleoyl-sn-glycero-3-phospho-rac-1-glycerol (DOPG) vesicles and polystyrene beads into Tradescantia virginiana stamen hair cells. The phragmoplast was nonselective for DOPG vesicles of a size up to 150 nm in diameter but was a physical barrier for polystyrene beads having a diameter of 20 and 40 nm and also when beads were coated with the same DOPG membrane. We conclude that stiffness is a parameter for vesicle transit through the phragmoplast and discuss that cytoskeleton configurations can physically block such transit.Cells and their constituents are physical entities, and next to chemical interactions, cell structures are determinants of cell behavior. Therefore, apart from techniques to image living cells at the subcellular level, experiments are needed that probe physical parameters important in cell function in vivo. We took the plant phragmoplast structure to answer the question whether the physical aspect “stiffness” is a factor in the inhibition of transport through this structure by microinjecting synthetic vesicles and polystyrene beads in Tradescantia virginiana stamen hair cells during cytokinesis, when the phragmoplast is essential for partitioning the cytoplasm between two daughter cells. Plant cells partition by producing a cell plate made of fused 60- to 80-nm-diameter vesicles (Staehelin and Hepler, 1996; Jürgens, 2005) proven to be Golgi vesicles (Reichardt et al., 2007). Their content becomes the new cell wall and their membranes become the daughter cell plasma membranes. The phragmoplast consists of two opposing cylinders of microtubules and actin filaments, interlaced with similarly aligned endoplasmic reticulum (ER) membranes. This phragmoplast cytoskeleton is the transport vehicle for Golgi vesicles to the plane where the cell plate is being formed (Staehelin and Hepler, 1996; Valster et al., 1997), keeps them in this plane (Esseling-Ozdoba et al., 2008b), where they fuse with each other (Samuels et al., 1995; Otegui et al., 2001; Seguí-Simarro et al., 2004), and assists in the proper attachment of the cell plate to the parental cell wall (Valster et al., 1997; Molchan et al., 2002). Transit of organelles, including Golgi bodies, is inhibited (Staehelin and Hepler, 1996; Nebenführ et al., 2000; Seguí-Simarro et al., 2004). Most of these data are known from static electron microscopy images. Electron microscopy after high-pressure freezing and freeze substitution (Thijsen et al., 1998) and electron tomography studies (Otegui et al., 2001; Seguí-Simarro et al., 2004; Austin et al., 2005) show that, in the early stage of cell plate formation in the center and later at the phragmoplast border, microtubules are aligned parallel to each other at distances of 20 to 100 nm. Keeping in mind that also actin filaments and ER membranes, aligned in the same orientation, are present between the microtubules, this leaves little room for the cell plate-forming vesicles during their transport through this phragmoplast.Clearly, during the past decade, significant progress has been made in the elucidation of the structural organization of cell plate-forming phragmoplasts, which has set the stage for studies to elucidate physical properties of phragmoplasts. The experimental approach we use is injecting particulate and vesicular fluorescent probes into living and dividing cells and observing the extent to which such probes can enter the phragmoplast and can be transported to the cell plate region. We have shown before that synthetic lipid 1,2-dioleoyl-sn-glycero-3-phospho-rac-1-glycerol (DOPG) vesicles of 60 nm diameter are transported through the phragmoplast, accumulate, and are kept in the cell plate region but do not fuse (Esseling-Ozdoba et al., 2008b). Now, we asked whether similar, flexible, synthetic lipid (DOPG) vesicles of various sizes, smaller and larger than endogenous vesicles, as well as stiff polystyrene beads, and such beads coated with the DOPG membrane, are transported through the phragmoplast and enter the plane where the cell plate is being formed, a question pertaining to a physical property of the phragmoplast. Our principal finding is that injected synthetic vesicles up to 150 nm diameter can enter and be transported to the cell plate region, where they accumulate but do not become incorporated into the cell plate. In contrast, polystyrene beads, the noncoated ones and those coated with the same lipid as the vesicles with diameters of 20 and 40 nm, can enter phragmoplasts but cannot be transported to the cell plate region, and the 40-nm beads slow cell plate formation, possibly by interfering with the delivery of normal, cell plate-forming vesicles to the cell plate.     

Fabrication of Stable Packed Capillary Reversed-Phase Columns for Protein Structural Analysis

Capillary column (320-m ID) liquid chromatography is an essential tool for the separation and concentration of low-picomole amounts of proteins and peptides for mass-spectrometric based structural analysis. We describe a detailed procedure for the fabrication of stable and efficient 50- to 180-m ID polyimide fused-silica columns. Columns were packed by conventional slurry packing with reversed-phase silica-based supports followed by column bed consolidation with acetonitrile and sonication. PVDF membrane or internal fused-silica particles were employed for column end-frit construction. The ability of these columns to withstand high back pressures (300–400 bar) enabled their use for rapid chromatography (>3400 cm/hr; i.e., 40 l/min for 200-m ID columns) and the loading of large sample volumes (up to 500 l). The accurate low flow rates (0.4–4.0 l/min) and precise gradient formation necessary to operate these columns were achieved by a simple modification of conventional HPLC systems [Moritz et al. (1992), J. Chromatogr. 599, 119–130]. Column performance was evaluated for ability to resolve low-fmol amounts of all components of a mixture of PTH-amino acids and to separate peptides for on-line LC/MS analysis of peptide mixtures derived from in situ digestion of 2-DE resolved protein spots.     

Activation of PKC-ε counteracts maturation and apoptosis of HL-60 myeloid leukemic cells in response to TNF family members

Protein kinase C (PKC)-ε, a component of the serine/threo-nine PKC family, has been shown to influence the survival and differentiation pathways of normal hematopoietic cells. Here, we have modulated the activity of PKC-ε with specific small molecule activator or inhibitor peptides. PKC-ε inhibitor and activator peptides showed modest effects on HL-60 maturation when added alone, but PKC-ε activator peptide significantly counteracted the pro-maturative activity of tumor necrosis factor (TNF)-α towards the monocytic/macrophagic lineage, as evaluated in terms of CD14 surface expression and morphological analyses. Moreover, while PKC-ε inhibitor peptide showed a reproducible increase of TNF-related apoptosis inducing ligand (TRAIL)-induced apoptosis, PKC-ε activator peptide potently counteracted the pro-apoptotic activity of TRAIL. Taken together, the anti-maturative and anti-apoptotic activities of PKC-ε envision a potentially important proleukemic role of this PKC family member.Key words: acute myeloid leukemia, surface antigens, HL-60 cells, apoptosis, maturation.Activation of all protein kinase C (PKC) family of serine and threonine isoenzymes is associated with binding to the negatively charged phospholipids, phosphatidylserine, while different PKC isozymes have varying sensitivities to Ca²⁺ and lipid-derived second messengers such as diacylglycerol (Gonelli et al., 2009). Upon activation, PKC isozymes translocate from the soluble to the particulate cell fraction, including cell membrane, nucleus and mitochondria (Gonelli et al., 2009). PKC primary sequence can be broadly separated into two domains: the N-terminal regulatory domain and the conserved C-terminal catalytic domain.The regulatory domain of PKC is composed of the C1 and C2 domains that mediate PKC interactions with second messengers, phospholipids, as well as inter and intramolecular protein-protein interactions. Differences in the order and number of copies of signaling domains, as well as sequence differences that affect binding affinities, result in the distinct activity of each PKC isozyme (Gonelli et al., 2009).In recent years, a series of peptides derived from PKC have been shown to modulate its activity by interfering with critical protein-protein interactions within PKC and between PKC and PKC-binding proteins (Brandman et al., 2007, Souroujon and Mochly-Rosen, 1998). Focusing on PKC-ε isozyme and using a rational approach, one C2-derived peptide that acts as an isozyme-selective activator (Dorn et al., 1999) and another that acts as a selective inhibitor (Johnson et al., 1996) of PKC-ε, have been identified.These findings are particularly interesting since besides being involved in the physiology of normal cardiac (Braun and Mochly-Rosen, 2003, Johnson et al., 1996, Li et al., 2006), hematopoietic (Gobbi et al., 2009, Mirandola et al., 2006, Racke et al., 2001), and neuronal (Borgatti et al., 1996) cell models, mounting experimental evidences have linked altered PKC-ε functions to solid tumor development (Okhrimenko et al., 2005, Gillespie et al., 2005, Lu et al., 2006). Therefore, taking advantage of the recent availability of small molecule peptides able to activate or inhibit specifically PKC-ε by disrupting protein/protein interactions (Dorn et al., 1999, Johnson et al., 1996), which open important therapeutic perspectives, we have investigated the effects of both PKC-ε activator and PKC-ε inhibitor peptides on the maturation and survival of leukemic cells, using as a model system the HL-60 myeloblastic leukemia cell line, which can be induced to undergo terminal differentiation or apoptotic cell death by a variety of chemical and biological agents (Breitman et al., 1980, Zauli et al., 1996).     

Surrogate Antigen Processing Mediated by TAP-dependent Antigenic Peptide Secretion

MHC class I proteins assemble with peptides in the ER. The peptides are predominantly generated from cytoplasmic proteins, probably by the action of the proteasome, a multicatalytic proteinase complex. Peptides are translocated into the ER by the transporters associated with antigen processing (TAP), and bind to the MHC class I molecules before transport to the cell surface. Here, we use a new functional assay to demonstrate that peptides derived from vesicular stomatitis virus nucleoprotein (VSV-N) antigen are actively secreted from cells. This secretion pathway is dependent on the expression of TAP transporters, but is independent of the MHC genotype of the donor cells. Furthermore, the expression and transport of MHC class I molecules is not required. This novel pathway is sensitive to the protein secretion inhibitors brefeldin A (BFA) and a temperature block at 21°C, and is also inhibited by the metabolic poison, azide, and the protein synthesis inhibitor, emetine. These data support the existence of a novel form of peptide secretion that uses the TAP transporters, as opposed to the ER translocon, to gain access to the secretion pathway. Finally, we suggest that this release of peptides in the vicinity of uninfected cells, which we term surrogate antigen processing, could contribute to various immune and secretory phenomena.Protein secretion has traditionally been thought to involve the action of the translocon located in the membrane of the ER of eukaryotic cells. Proteins are recognized cotranslationally when a signal sequence or a signal–anchor sequence emerges from the ribosome (Walter and Johnson, 1994). These sequences are recognized and bound by the signal recognition particle, and the resulting ribosomal complex then interacts with the signal recognition particle receptor on the ER membrane at the translocon (Andrews and Johnson, 1996). This results in the inclusion of proteins within the secretory pathway. This pathway is by far the best described route of protein secretion in eukaryotic cells. Recently, it has been proposed that some proteins are recognized by a component of the translocon, sec 61, exit the ER, and are transported into the cytoplasm where they are degraded (Wiertz et al., 1996).The translocation into the ER of antigenic peptides for presentation by major histocompatibility complex (MHC)¹ class I molecules is largely independent of the translocon. This form of translocation involves the action of two gene products that are members of the ATP binding cassette family. These genes encode transporters associated with antigen processing 1 and 2 (TAP-1 and -2), and have been implicated in the translocation of peptides from the cytoplasm to the lumen of the ER (Deverson et al., 1990; Bahram et al., 1991; Spies and DeMars, 1991; Spies et al., 1992; Gabathuler et al., 1994). After translocation into the ER, antigenic peptides bind to MHC class I molecules composed of a heavy chain (46-kD) and a light chain (12-kD) called β₂m (Nuchtern et al., 1989; Yewdell and Bennink, 1989; van Bleek and Nathenson, 1990; Matsumura et al., 1992), before transport to the cell surface. The assembly and transport of MHC class I molecules appears to be regulated by a series of chaperones that includes calnexin (Degen and Williams, 1991), calreticulin, and tapasin (Sadasivan et al., 1996).High performance liquid chromatography analysis of peptides eluted from acid-treated whole cells or MHC class I molecules has allowed the identification and characterization of the peptides associated with MHC class I molecules (Falk et al., 1990; Rötzschke et al., 1990; van Bleek and Nathenson, 1990). It is proposed that MHC class I molecules determine the final identity of MHC- restricted peptides and have an instructive role, in addition to a selective role, in peptide selection (Wallny et al., 1992). MHC binding to larger peptides followed by protected proteolytic trimming is a possible mechanism that could account for the observed MHC dependency of cellular peptides (Falk et al., 1990). Peptides unable to bind MHC class I because they are in excess or lack the correct MHC binding motif for the MHC haplotype are thus far undetectable by the techniques commonly used in the field, and are presumed to be short lived and degraded (Falk et al., 1990; Rötzschke et al., 1990). Recent results, however, suggest that peptides not able to bind to a MHC class I molecule intracellularly may be found bound to heat shock proteins (HSPs) such as gp96 (grp94; Arnold et al., 1995). These authors show that cellular antigens are represented by peptides associated with gp96 molecules independently of the MHC class I expressed, confirming earlier results (Udono and Srivastava, 1993, 1994). Gp96 extracted from a specific cell is able to induce cross-priming (Udono and Srivastava, 1993, 1994). Finally, two studies have demonstrated that peptides transported into the lumen of the ER, and do not bind to MHC class I molecules, can be transported out of the ER into the cytoplasm again by a process called “efflux” (Momburg et al., 1994; Schumacher et al., 1994), which may involve the action of the TAP molecules or the sec 61 protein associated with the translocon (Wiertz et al., 1996).We have developed a new bioassay to test the hypothesis that peptides translocated into the ER by the action of the TAP molecules become secreted. Using this assay, we present evidence of an alternative secretion pathway that exists in various mammalian cell types. These observations revise the model of peptide catabolism, and may provide an explanation for various immune and secretion phenomena.     

In-depth Qualitative and Quantitative Profiling of Tyrosine Phosphorylation Using a Combination of Phosphopeptide Immunoaffinity Purification and Stable Isotope Dimethyl Labeling

Several mass spectrometry-based assays have emerged for the quantitative profiling of cellular tyrosine phosphorylation. Ideally, these methods should reveal the exact sites of tyrosine phosphorylation, be quantitative, and not be cost-prohibitive. The latter is often an issue as typically several milligrams of (stable isotope-labeled) starting protein material are required to enable the detection of low abundance phosphotyrosine peptides. Here, we adopted and refined a peptidecentric immunoaffinity purification approach for the quantitative analysis of tyrosine phosphorylation by combining it with a cost-effective stable isotope dimethyl labeling method. We were able to identify by mass spectrometry, using just two LC-MS/MS runs, more than 1100 unique non-redundant phosphopeptides in HeLa cells from about 4 mg of starting material without requiring any further affinity enrichment as close to 80% of the identified peptides were tyrosine phosphorylated peptides. Stable isotope dimethyl labeling could be incorporated prior to the immunoaffinity purification, even for the large quantities (mg) of peptide material used, enabling the quantification of differences in tyrosine phosphorylation upon pervanadate treatment or epidermal growth factor stimulation. Analysis of the epidermal growth factor-stimulated HeLa cells, a frequently used model system for tyrosine phosphorylation, resulted in the quantification of 73 regulated unique phosphotyrosine peptides. The quantitative data were found to be exceptionally consistent with the literature, evidencing that such a targeted quantitative phosphoproteomics approach can provide reproducible results. In general, the combination of immunoaffinity purification of tyrosine phosphorylated peptides with large scale stable isotope dimethyl labeling provides a cost-effective approach that can alleviate variation in sample preparation and analysis as samples can be combined early on. Using this approach, a rather complete qualitative and quantitative picture of tyrosine phosphorylation signaling events can be generated.Reversible tyrosine phosphorylation plays an important role in numerous cellular processes like growth, differentiation, and migration. Phosphotyrosine signaling is tightly controlled by the balanced action of protein-tyrosine kinases and protein-tyrosine phosphatases. Aberrant tyrosine phosphorylation has been suggested to be an underlying cause in multiple cancers (1). Therefore, the identification of tyrosine phosphorylated proteins and the investigation into their involvement in signaling pathways are important. Several groups have attempted to comprehensively study tyrosine phosphorylation by proteomics means (2–5). However, large scale identification of tyrosine phosphorylation sites by MS can be hindered by the low abundance of tyrosine phosphorylated proteins. Especially, signaling intermediates are usually low abundance proteins that show substoichiometric phosphorylation levels. In addition, the identification by mass spectrometry of phosphopeptides from a complex cellular lysate digest is often complicated by ion suppression effects due to a high background of non-phosphorylated peptides. Enrichment of tyrosine phosphorylated proteins or peptides prior to mass spectrometric detection is therefore essential. Traditionally, antibodies against phosphorylated tyrosine have been used to immunoprecipitate tyrosine phosphorylated proteins from cultured cells (2–4, 6–8). This phosphoprotein immunoaffinity purification method has for example been used to study the global dynamics of phosphotyrosine signaling events after EGF¹ stimulation using stable isotope labeling by amino acids in cell culture (SILAC) (2). This approach led to the identification of known and previously unidentified signaling proteins in the EGF receptor (EGFR) pathway, including their temporal activation profile after stimulation of the EGFR, providing crucial information for modeling signaling events in the cell. However, as the identification and quantification of these phosphorylated proteins in these studies were not necessarily based on tyrosine phosphorylated peptides but largely on non-phosphorylated peptides, little information is derived on the exact site(s) of tyrosine phosphorylation. Also, binding partners of tyrosine phosphorylated proteins, which themselves are not tyrosine phosphorylated, might be co-precipitated and impair the tyrosine phosphorylation quantification. Immunoaffinity purification of phosphotyrosine peptides, rather than proteins, using anti-phosphotyrosine antibodies (5, 9–16) significantly facilitates the identification of the site(s) of phosphorylation as it greatly alleviates most of the above mentioned problems because the tyrosine phosphorylated site can be directly identified and quantified.Accurate MS-based quantification is typically performed by stable isotope labeling. The isotopes can be incorporated metabolically during cell culture as in SILAC (17) or chemically as in an isobaric tag for relative and absolute quantitation (iTRAQ) (18) or stable isotope dimethyl labeling (19–21). Typically, the most precise quantification can be obtained by metabolic labeling as the different samples can be combined at the level of intact cells (22). However, metabolic labeling is somewhat limited to biological systems that can be grown in culture, and the medium may have an effect on the growth and development of the cells. iTRAQ has been used in conjunction with phosphotyrosine peptide immunoprecipitation (5). As the chemical labeling is performed before immunoprecipitation, the differentially labeled samples can be precipitated together, thereby neutralizing the potentially largest source of variation. However, as this phosphotyrosine peptide immunoprecipitation is typically performed on several hundreds of micrograms to milligrams of protein sample, iTRAQ provides in these cases a rather cost-prohibitive means.Here, we present an optimized immunoaffinity purification approach for the analysis of tyrosine phosphorylation combined with stable isotope dimethyl labeling (19–21, 23). We efficiently enriched and identified by MS 1112 unique phosphopeptides derived from 4 mg of starting protein material without any further affinity chromatographic enrichment whereby up to 80% of the peptides analyzed in the final LC run were phosphotyrosine peptides. We further advanced the method by introducing triplex stable isotope dimethyl labeling prior to immunoprecipitation. We quantified differences in tyrosine phosphorylation upon pervanadate treatment or EGF stimulation to detect site-specific changes in tyrosine phosphorylation. 128 unique phosphotyrosine peptides were identified and quantified upon pervanadate treatment. By using an internal standard comprising both mock and pervanadate-treated samples, we could more confidently identify and quantify phosphorylation sites that are strongly regulated and on-off situations. Analysis of EGF-stimulated HeLa cells resulted in the quantification of 73 unique phosphotyrosine peptides. Most of the up-regulated phosphotyrosine peptides that were identified have been reported previously to be involved in the EGFR signaling pathway, validating our approach. However, for the first time, we found TFG to also become highly tyrosine phosphorylated upon EGF stimulation together with some tyrosine phosphorylation sites on for example IRS2, SgK269, and DLG3 that have not been firmly established earlier to be involved in EGFR signaling.In general, we show that the combination of immunoaffinity purification of tyrosine phosphorylated peptides with large scale chemical stable isotope dimethyl labeling provides a cost-effective approach that can alleviate variation in immunoprecipitation and LC-MS as samples can be combined before immunoprecipitation and the necessity of performing additional enrichment is removed by an optimization of the protocol. With only a single LC-MS run, already a rather complete qualitative and quantitative picture of a signaling event can be generated.     

Rapid isolation of yeast genomic DNA: Bust n' Grab

Background

Mutagenesis of yeast artificial chromosomes (YACs) often requires analysis of large numbers of yeast clones to obtain correctly targeted mutants. Conventional ways to isolate yeast genomic DNA utilize either glass beads or enzymatic digestion to disrupt yeast cell wall. Using small glass beads is messy, whereas enzymatic digestion of the cells is expensive when many samples need to be analyzed. We sought to develop an easier and faster protocol than the existing methods for obtaining yeast genomic DNA from liquid cultures or colonies on plates.

Results

Repeated freeze-thawing of cells in a lysis buffer was used to disrupt the cells and release genomic DNA. Cell lysis was followed by extraction with chloroform and ethanol precipitation of DNA. Two hundred ng – 3 μg of genomic DNA could be isolated from a 1.5 ml overnight liquid culture or from a large colony. Samples were either resuspended directly in a restriction enzyme/RNase coctail mixture for Southern blot hybridization or used for several PCR reactions. We demonstrated the utility of this method by showing an analysis of yeast clones containing a mutagenized human β-globin locus YAC.

Conclusion

An efficient, inexpensive method for obtaining yeast genomic DNA from liquid cultures or directly from colonies was developed. This protocol circumvents the use of enzymes or glass beads, and therefore is cheaper and easier to perform when processing large numbers of samples.

     

Analysis of Monoclonal Antibody Sequence and Post-translational Modifications by Time-controlled Proteolysis and Tandem Mass Spectrometry

Methodology for sequence analysis of ∼150 kDa monoclonal antibodies (mAb), including location of post-translational modifications and disulfide bonds, is described. Limited digestion of fully denatured (reduced and alkylated) antibody was accomplished in seconds by flowing a sample in 8 m urea at a controlled flow rate through a micro column reactor containing immobilized aspergillopepsin I. The resulting product mixture containing 3–9 kDa peptides was then fractionated by capillary column liquid chromatography and analyzed on-line by both electron-transfer dissociation and collisionally activated dissociation mass spectrometry (MS). This approach enabled identification of peptides that cover the complete sequence of a murine mAb. With customized tandem MS and ProSightPC Biomarker search, we verified 95% amino acid residues of this mAb and identified numerous post-translational modifications (oxidized methionine, pyroglutamylation, deamidation of Asn, and several forms of N-linked glycosylation). For disulfide bond location, native mAb is subjected to the same procedure but with longer digestion times controlled by sample flow rate through the micro column reactor. Release of disulfide containing peptides from accessible regions of the folded antibody occurs with short digestion times. Release of those in the interior of the molecule requires longer digestion times. The identity of two peptides connected by a disulfide bond is determined using a combination of electron-transfer dissociation and ion–ion proton transfer chemistry to read the two N-terminal and two C-terminal sequences of the connected peptides.Monoclonal antibodies (mAbs)¹ and related biological molecules constitute one of the most rapidly growing classes of human therapeutics. These large proteins (Fig. 1) have molecular weights near 150 kDa and are composed of two identical ∼50 kDa heavy chains (HC) and two identical ∼25 kDa light chains (LC) (1). They also contain at least 16 disulfide bonds that maintain three-dimensional structure and biological activity (2). Although sharing similar secondary protein structures, different mAbs differ greatly in the sequence of variable regions, especially in the complementarity determining regions (CDRs) which are responsible for the diversity and specificity of antibody-antigen binding. Changes to the mAb structure introduced during the manufacturing process or storage may influence the therapeutic efficacy, bio-availability and -clearance, and immunogenic properties and thus alter drug safety (3–5). Comprehensive characterization of mAbs primary structure, post-translational modifications (PTMs), and disulfide linkages is critical to the evaluation of drug efficacy and safety, as well as understanding the structure/function relationships (4, 6). Presented in this work is a novel protein analytical platform that consists of innovative methods for mass spectrometry (MS) characterization of mAbs. The methodology reported here will have a dramatic impact on the whole field of antibody characterization.Open in a separate windowFig. 1.Diagram of a murine monoclonal antibody structure.Typical MS characterization of proteins uses a “Bottom-Up” approach. This method involves tryptic digestion of the protein(s) into small peptides (mostly below 2500 Da) followed by high-performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) analyses of the resulting peptides (7). Although sensitive for MS analysis, small tryptic peptides often have issues such as weak retention in liquid chromatography, difficulties in assigning peptides to specific gene products, and loss of combinatorial PTM information (8). Recent years have seen developments in direct MS analysis of intact proteins (often called “Top-Down” MS). Despite increasing success in characterization of small to medium-sized proteins, MS analysis of intact proteins larger than 50 kDa, including mAbs, is still unsatisfactory because of inefficient gas-phase protein fragmentation and complex fragment ions that restrict efficient data interpretation (9, 10).A “compromise” between Bottom-Up and Top-Down approaches is the “Middle-Down” (or “Middle-Up”) method. Middle-Down analysis typically involves proteolysis using proteases (e.g. Lys-C) or chemicals that hydrolyze proteins at a single type of amino acid residue. This approach aims to generate 3–15 kDa peptides which are compatible with high resolution MS/MS analysis on a chromatographic time scale. The Middle-Down approach inherits some of the advantages of Top-Down analysis, yet has less demanding instrumental requirements compared with intact protein MS in achieving sufficient signal-to-noise ratio (S/N) of fragment ions for sequence mapping (11–15).However, limitations of currently available tools for Middle-Down protein analysis are also obvious. First, none of the twenty amino acids is evenly distributed along a polypeptide. Protein digestion at single-type amino acid residues can still produce very small (<1000 Da) or ultra large (>15 kDa) peptides, which deviates from the original intention of the Middle-Down approach (16). Second, the enzymatic digestion efficiency is often low for proteins with highly folded structure or low solubility. Although high concentrations of chaotropic agents such as 8 m urea are often used for protein denaturation, this harsh condition quickly deactivates many commonly used proteases. Third, traditional data-dependent ETD or electron-capture dissociation MS/MS analyses adopt a single reaction parameter for gas-phase dissociation and select only several abundant ions regardless of their charge states. As these methods were previously optimized for tryptic peptide ions that typically carry +2 or +3 charges, they are incompatible with the analysis of large, highly charged peptides that require optimized ETD to achieve high sequence coverage and PTM mapping (12).Herein we report a “time-controlled” proteolysis method for tailored Middle-Down MS analysis of mAb. To hydrolyze the 150 kDa mAb into large peptides for HPLC-MS analysis, we fabricated a capillary enzyme reactor column that contains a specified length of immobilized protease (supplemental Fig. S1 and S2A). Precise control of the sample flow rate leads to defined digestion time of the substrate protein in the reactor. A short digestion time results in a small number of “cuts” along the protein chain and consequently the formation of large peptides (supplemental Fig. S2B). The Bruening group previously demonstrated a similar concept using a nylon membrane electrostatically adsorbed with pepsin or trypsin. Pushing a protein solution (protein dissolved in 5% formic acid solution) through the membrane-based enzyme reactor in less than 1 s breaks the protein into large peptides that facilitate sequence mapping of horse apomyoglobin (17 kDa) and bovine serum albumin (66 kDa) by infusion electrospray ionization MS/MS (17). The advantages of their enzyme disc include simple preparation procedures, as well as the low back pressure in the thin disc that allows for rapid sample flow rate. In our present work, we designed a more robust enzyme reactor that digests alkylated or native mAb into 3–12 kDa peptides in a buffer containing 8 m urea (a condition incompatible with most widely used proteases), and characterized their amino acid sequences, PTMs, as well as the disulfide linkages using HPLC-MS/MS.We chose a rarely used protease, aspergillopepsin I, for the enzyme reactor. Aspergillopepsin I, also known as Aspergillus saitoi acid proteinase, generally catalyzes the hydrolysis of substrate proteins at P1 and P1′ of hydrophobic residues, but also accepts Lys at P1 (18). There are several innovative aspects of employing this enzyme: (1) Aspergillopepsin I is active in 8 m urea at pH 3–4 for at least 1 h. This extreme chaotropic condition may disrupt the higher-order structure of proteins to a great extent and allows for easy access of the protease to most regions of the substrate protein once the disulfide bonds are reduced. (2) Compared with proteases with dual- or single-type amino acid specificity, aspergillopepsin I provides more cleavage sites along an unfolded substrate protein. Allowing limited time for the substrate protein to interact with immobilized aspergillopepsin I should generate large peptides with a relatively narrow size distribution because of similar numbers of missed cleavages on these peptides. (3) The enzyme reactor automatically “quenches” proteolysis as the sample flows out of the column. This is in great contrast to in-tube digestion using solubilized proteases that are active in acidic conditions. In the latter case, digestion is difficult to quench or control because of the sustained enzymatic activity in an acidic condition. (4) Compared with electrostatic or hydrophobic interactions for enzyme immobilization, covalent conjugation of the protease onto porous beads should prevent the replacement of enzymes by upcoming substrate proteins. (5) The enzyme beads can be stored at 4 °C for at least half a year once water is removed, allowing the production of hundreds of disposable enzyme reactors from one batch of beads. In addition, we introduced a new cysteine (Cys) alkylation reagent, N-(2-aminoethyl)maleimide (NAEM) for protein MS analysis. This reagent improves ETD (19) of peptides containing Cys residues by adding a basic, readily protonated side chain to thiol groups.The above features of our new strategy led to the generation of large, highly charged peptides that cover the entire murine mAb. Analyzing ETD and collisionally activated dissociation (CAD) fragments from the most abundant large peptides by ProSightPC revealed near complete sequence coverage of the mAb and multiple PTMs. Furthermore, we digested the native mAb into large fragments of disulfide-bonded peptides using time-controlled digestion. The ETD/ion-ion proton transfer (IIPT) technique (20) allowed facile identification of the N- and C-terminal sequences of two disulfide-bonded peptides and localization of the disulfide bond(s) within/connecting different mAb domains.     

Measuring Arabidopsis Chromatin Accessibility Using DNase I-Polymerase Chain Reaction and DNase I-Chip Assays

DNA accessibility is an important layer of regulation of DNA-dependent processes. Methods that measure DNA accessibility at local and genome-wide scales have facilitated a rapid increase in the knowledge of chromatin architecture in animal and yeast systems. In contrast, much less is known about chromatin organization in plants. We developed a robust DNase I-polymerase chain reaction (PCR) protocol for the model plant Arabidopsis (Arabidopsis thaliana). DNA accessibility is probed by digesting nuclei with a gradient of DNase I followed by locus-specific PCR. The reduction in PCR product formation along the gradient of increasing DNase I concentrations is used to determine the accessibility of the chromatin DNA. We explain a strategy to calculate the decay constant of such signal reduction as a function of increasing DNase I concentration. This allows describing DNA accessibility using a single variable: the decay constant. We also used the protocol together with AGRONOMICS1 DNA tiling microarrays to establish genome-wide DNase I sensitivity landscapes.Chromatin has a major impact on genome organization and gene activity. Differential accessibility of DNA is thought to be a major consequence of locally different chromatin composition and structure (Li et al., 2007). Chromatin sensitivity to nucleases has proven to be a powerful tool to probe DNA accessibility in chromatin. Frequently used nucleases include DNase, micrococcal nuclease, and restriction enzymes. The resolution of restriction enzymes is limited by their sequence specificity, and micrococcal nuclease is more often used to determine nucleosome occupancy (Schones and Zhao, 2008). Chromatin sensitivity to DNase I has often been used to define the “openness” of chromatin relative to its higher order structures. Its applicability has been manifested by detecting regulatory elements, such as promoters, enhancers, and insulators, as DNase I-hypersensitive sites (Wang and Simpson, 2001; Crawford et al., 2004, 2006; Dorschner et al., 2004; Sabo et al., 2006; Boyle et al., 2008; Naughton et al., 2010; Pique-Regi et al., 2011). DNase I sensitivity can also be used as a measure for the general accessibility of chromatin (Weil et al., 2004).Initially, the chromatin accessibility of local genomic regions to DNase I was probed by Southern blotting (Mather and Perry, 1983; Bender et al., 2000; Wang and Simpson, 2001; Bulger et al., 2003). However, Southern blotting is tedious and lacks sensitivity, and the interpretation of results can be challenging. Therefore, analysis methods based on PCR have been developed (Pfeifer and Riggs, 1991; Feng and Villeponteau, 1992; McArthur et al., 2001; Dorschner et al., 2004; Martins et al., 2007). In recent years, DNase I assays were coupled to high-throughput genome-wide profiling strategies such as genome tiling arrays and next-generation sequencing (Crawford et al., 2004, 2006; Sabo et al., 2004, 2006; Weil et al., 2004). While much has been learned about the accessibility of chromatin in animal and yeast systems, our knowledge of chromatin accessibility in plants is limited. Most studies have focused on selected genomic regions such as the general regulatory factor1 (GRF1) gene and the alcohol dehydrogenase1 (Adh1) and Adh2 genes in maize (Zea mays; Paul and Ferl, 1998a, 1998b) or the GRF gene, the Adh gene, and an 80-kb genomic region harboring 30 protein-coding genes in Arabidopsis (Arabidopsis thaliana; Vega-Palas and Ferl, 1995; Paul and Ferl, 1998a, 1998b; Kodama et al., 2007). The technique used in these reports was exclusively DNase I treatment and analysis of accessibility using Southern blotting. More recently, we have combined the DNase I sensitivity assay with whole-genome tiling arrays in Arabidopsis to generate a genome-wide chromatin accessibility profile (Shu et al., 2012).Here, we present a robust, optimized DNase I sensitivity assay protocol for Arabidopsis tissues based on PCR. This protocol can be adapted to different samples or experimental objectives; the strategies for optimizing each step are also discussed. Analysis of relatively large fragments by PCR has proven to be highly robust as a first step in probing DNase I sensitivity in any region of the genome. We also introduce a new strategy for presenting the DNase I sensitivity of the tested regions using a decay constant calculated by fitting PCR product intensity values from a gradient digestion. In this way, the sensitivity of each region is characterized by a single value, facilitating comparisons between different regions or samples. Finally, we describe how our protocol can be combined with genomic techniques for genome-wide profiles of chromatin accessibility.     

Septins recognize micron-scale membrane curvature

How cells recognize membrane curvature is not fully understood. In this issue, Bridges et al. (2016. J. Cell Biol. http://dx.doi.org/10.1083/jcb.201512029) discover that septins, a component of the cytoskeleton, recognize membrane curvature at the micron scale, a common morphological hallmark of eukaryotic cellular processes.Eukaryotic cells have dedicated proteins that sense membranes, depending on their curvature (Antonny, 2011). Sensors of membrane curvature are important because they organize a wide variety of cellular functions, including vesicle trafficking and organelle shaping (McMahon and Gallop, 2005). Curvature-sensing proteins, for example, the Bin-Amphiphysin-Rvs (BAR) domain–containing proteins, have been mostly described to work at the nanometer scale (Zimmerberg and Kozlov, 2006). In contrast, a clear mechanism of sensing membrane curvature at the micron scale in eukaryotic cells has not been described. In this issue, Bridges et al. discover that septins, a poorly understood component of the cytoskeleton, recognize plasma membrane curvature at the micron scale and serve as landmarks for eukaryotic cells to know their local shape.Septins are an evolutionarily conserved family of GTP-binding proteins that assemble into nonpolar filaments and rings (John et al., 2007; Sirajuddin et al., 2007; Bertin et al., 2008). Septins have been implicated in diverse membrane organization events where micron-scale curvature takes place (Saarikangas and Barral, 2011; Mostowy and Cossart, 2012), including the cytokinetic furrow, the annulus of spermatozoa, the base of cellular protrusions (e.g., cilium and dendritic spines), and the phagocytic cup surrounding invasive bacterial pathogens (Fig. 1). However, the precise role of septin–membrane interactions remains elusive. It was first suggested in 1999 that the interaction of human septins with phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is important for septin localization (Zhang et al., 1999). More recently, work using recombinant septins from budding yeast Saccharomyces cerevisiae assembled on PI(4,5)P2 lipid monolayers showed that septins interact with membrane to facilitate filament assembly (Bridges et al., 2014). Membrane-facilitated septin assembly has also been observed using phospholipid liposomes, and in this case septins were also shown to induce membrane tubulation (Tanaka-Takiguchi et al., 2009). Given that (a) septins can interact with membrane, (b) septin assembly is membrane facilitated, and (c) septin assemblies are associated with a variety of membrane organization events from yeast to mammals, Bridges et al. (2016) hypothesized that septins serve as a mechanism to recognize membrane curvature.Open in a separate windowFigure 1.Morphological hallmarks of eukaryotic cells characterized by micron-scale membrane curvature and septin assembly. Septins have been implicated in membrane organization events where micron-scale curvature takes place. (A) A septin ring acts as a scaffold for cytokinesis proteins and forms a diffusion barrier at the cytokinetic furrow of a dividing cell. (B) A septin ring forms a diffusion barrier at the annulus of a mammalian spermatozoon, which separates the anterior and posterior tail. (C) A septin ring forms a diffusion barrier at the base of a cilium to separate the ciliary membrane from the plasma membrane. (D) In neurons, a septin-dependent diffusion barrier can localize at the base of dendritic spine necks. (E) During phagocytosis, a cup is formed at the plama membrane; septin rings assemble at the base of the phagocytic cup to regulate entry.In their new work, Bridges et al. (2016) provide several lines of evidence to support the hypothesis that septins recognize micron-scale curvature. First, using the filamentous fungus Ashbya gossypii, they performed in vivo localization studies and showed that the fungal septin Cdc11a concentrates in regions of positive micron-scale curvature and that the degree of concentration is proportional to the degree of curvature. Moreover, septins localize to curved membranes that also recruit septin-interacting proteins (e.g., the signaling protein Hsl7). These findings indicate that, by acting as curvature-sensing proteins, septins can localize signaling platforms in the cell. To test if septins can differentiate among micron-scale curvatures, Bridges et al. (2016) developed an elegant model system for septin assembly in vitro. They decorated silica beads with anionic phospholipid bilayers and measured the interaction affinity between purified fungal septin complexes and beads of different curvatures. Interestingly, septins were maximally recruited to “intermediate” sized beads (1.0–3.0 µm in diameter), with little to no recruitment to either very large (5.0–6.5 µm in diameter) or very small (0.3 µm in diameter) beads. These results indicate that septin filaments preferentially localize to a curvature (κ) of 0.7–2.0 µm⁻¹ in the absence of other cellular factors. To provide additional information on the mechanism of sensing, the authors purified mutant septin complexes that fail to polymerize into filaments and showed that the affinity of septins for micron-scale membrane curvature does not require filament formation per se. However, septins must polymerize into filaments for stable membrane association. Collectively, in vivo experiments using A. gossypii and in vitro experiments using silica beads highlight that septins have the intrinsic ability to recognize membrane curvature at the micron scale.Finally, to study the recognition of micron-scale membrane curvature beyond fungi, Bridges et al. (2016) turn their attention to human septins. Using tissue culture cells, they observe that the abundance of septins is associated with the degree of membrane curvature. To confirm these observations in vitro, they purified human septins and analyzed their binding affinity to silica beads with phospholipid bilayers. As seen with A. gossypii septins, human septins also showed a preference for beads ∼1.0 µm in diameter, strongly suggesting an evolutionarily conserved property of septins for sensing membrane curvature at the micron scale.Based on their findings, Bridges et al. (2016) propose that septins provide eukaryotic cells with a mechanism to recognize curvature at the micron scale. This feature differentiates septins from other sensor proteins that strictly detect curvature at the nanometer scale (e.g., BAR domain–containing proteins). However, it is likely that septins do more than recognize membrane, and the precise role of septins in membrane recognition remains unknown. The highly conserved structural and biochemical properties of septins suggest they organize, stabilize, and functionalize membrane domains (Caudron and Barral, 2009; Kusumi et al., 2012; Bridges and Gladfelter, 2015). Although we are far from knowing the full repertoire of septin function, this new work by Bridges et al. (2016) reminds us that understanding how membranes can specify septin assembly is essential to understand the role of septins in eukaryotic cells.     

Large-Scale Phosphoprotein Analysis in Medicago truncatula Roots Provides Insight into in Vivo Kinase Activity in Legumes

Nitrogen fixation in legumes requires the development of root organs called nodules and their infection by symbiotic rhizobia. Over the last decade, Medicago truncatula has emerged as a major model plant for the analysis of plant-microbe symbioses and for addressing questions pertaining to legume biology. While the initiation of symbiosis and the development of nitrogen-fixing root nodules depend on the activation of a protein phosphorylation-mediated signal transduction cascade in response to symbiotic signals produced by the rhizobia, few sites of in vivo phosphorylation have previously been identified in M. truncatula. We have characterized sites of phosphorylation on proteins from M. truncatula roots, from both whole cell lysates and membrane-enriched fractions, using immobilized metal affinity chromatography and tandem mass spectrometry. Here, we report 3,457 unique phosphopeptides spanning 3,404 nonredundant sites of in vivo phosphorylation on 829 proteins in M. truncatula Jemalong A17 roots, identified using the complementary tandem mass spectrometry fragmentation methods electron transfer dissociation and collision-activated dissociation. With this being, to our knowledge, the first large-scale plant phosphoproteomic study to utilize electron transfer dissociation, analysis of the identified phosphorylation sites revealed phosphorylation motifs not previously observed in plants. Furthermore, several of the phosphorylation motifs, including LxKxxs and RxxSxxxs, have yet to be reported as kinase specificities for in vivo substrates in any species, to our knowledge. Multiple sites of phosphorylation were identified on several key proteins involved in initiating rhizobial symbiosis, including SICKLE, NUCLEOPORIN133, and INTERACTING PROTEIN OF DMI3. Finally, we used these data to create an open-access online database for M. truncatula phosphoproteomic data.Medicago truncatula has become a model for studying the biology of leguminous plants such as soybean (Glycine max), alfalfa (Medicago sativa), and clover (Trifolium spp.; Singh et al., 2007). Most members of this vast family have the ability to fix atmospheric nitrogen by virtue of an endosymbiotic association with rhizobial bacteria, through which legumes undergo nodulation, the process of forming root nodules (Jones et al., 2007). Legumes are central to modern agriculture and civilization because of their ability to grow in nitrogen-depleted soils and replenish nitrogen through crop rotation. Consequently, there is great interest in understanding the molecular events that allow legumes to recognize their symbionts, develop root nodules, and fix nitrogen. Nod factors are lipochitooligosaccharidic signals secreted by the rhizobia and are required, in most legumes, for intracellular infection and nodule development. In recent decades, an elegant combination of genetics, biochemistry, and cell biology has shown that Nod factors activate intricate signaling events within cells of legume roots, including protein phosphorylation cascades and intracellular ion fluxes (Oldroyd and Downie, 2008).Protein phosphorylation is a central mechanism of signal transfer in cells (Laugesen et al., 2006; Peck, 2006; Huber, 2007). Several characterized protein kinases are required for symbiosis signal transduction in M. truncatula roots (Lévy et al., 2004; Yoshida and Parniske, 2005; Smit et al., 2007). A recent antibody-based study of cultured M. truncatula cells observed protein phosphorylation changes at the proteomic level in response to fungal infection (Trapphoff et al., 2009); however, the target residues of the phosphorylation events were not determined. A variety of studies have determined in vitro phosphorylation sites on legume proteins and demonstrated the biological importance of the target residues by mutagenesis (Yoshida and Parniske, 2005; Arrighi et al., 2006; Lima et al., 2006; Miyahara et al., 2008; Yano et al., 2008). To our knowledge, only six sites of in vivo protein phosphorylation have been detected for M. truncatula (Laugesen et al., 2006; Lima et al., 2006; Wienkoop et al., 2008), demonstrating the need for the identification of endogenous protein phosphorylation sites in legume model organisms on a proteome-wide scale.While considerable advancements have been made in the global analysis of protein phosphorylation (Nita-Lazar et al., 2008; Macek et al., 2009; Piggee, 2009; Thingholm et al., 2009), phosphoproteomics in plants has lagged years behind that of the mammalian systems (Kersten et al., 2006, 2009; Peck, 2006), which have more fully sequenced genomes and better annotated protein predictions. Arabidopsis (Arabidopsis thaliana), the first plant genome sequenced (Arabidopsis Genome Initiative, 2000), is now predicted to have over 1,000 protein kinases (Finn et al., 2008), approximately twice as many as in human (Manning et al., 2002). Because many of the kinases in the commonly studied mammalian systems are not conserved in the plant kingdom, there is significant need for large-scale phosphoproteomic technologies to discern the intricacies of phosphorylation-mediated cell signaling in plants. With the high mass accuracy afforded by the linear ion trap-orbitrap hybrid mass spectrometer (Makarov et al., 2006; Yates et al., 2006), recent studies in Arabidopsis have reported 2,597 phosphopeptides from suspension cell culture (Sugiyama et al., 2008) and 3,029 phosphopeptides from seedlings (Reiland et al., 2009).All previous large-scale plant phosphoproteomic studies have relied solely on collision-activated dissociation (CAD) during tandem mass spectrometry (MS/MS) and have not taken advantage of the more recently developed methods (Kersten et al., 2009) electron capture dissociation (Kelleher et al., 1999) or electron transfer dissociation (ETD; Coon et al., 2004; Syka et al., 2004). Mapping sites of posttranslational modifications, such as phosphorylation, is often more straightforward using electron-based fragmentation methods, as they frequently produce a full spectrum of sequence-informative ions without causing neutral loss of the modifying functional groups (Meng et al., 2005; Chi et al., 2007; Khidekel et al., 2007; Molina et al., 2007; Wiesner et al., 2008; Chalkley et al., 2009; Swaney et al., 2009). With an ETD-enabled hybrid orbitrap mass spectrometer (McAlister et al., 2007, 2008), we previously compared the performance of CAD and ETD tandem MS for large-scale identification of phosphopeptides (Swaney et al., 2009). ETD identified a greater percentage of unique phosphopeptides and more frequently localized phosphorylation sites. Still, the low overlap of identified phosphopeptides indicates that the two methods are highly complementary. With this in mind, we recently developed a decision tree-driven tandem MS algorithm to select the optimal fragmentation method for each precursor (Swaney et al., 2008).Here, we utilize this technology to map sites of in vivo protein phosphorylation in roots of M. truncatula Jemalong A17 plants. Phosphoproteins, from both whole-cell lysate and membrane-enriched fractions, were analyzed after digestion with a variety of different enzymes individually. Utilizing the complementary fragmentation methods of ETD and CAD, we report 3,404 nonredundant phosphorylation sites at an estimated false discovery rate (FDR) of 1%. Analysis of these data revealed several phosphorylation motifs not previously observed in plants. The phosphorylation sites identified provide insight into the potential regulation of key proteins involved in rhizobial symbiosis, potential consensus sequences by which kinases recognize their substrates, and critical phosphorylation events that are conserved between plant species.